The Synthesis and in Vitro Toxicological Evaluation of the Mitoxantrone Naphthoquinoxaline Metabolite: A Comparative Study with the Parent Drug

ANA FILIPA DOS REIS MENDES The synthesis and in vitro toxicological evaluation of the mitoxantrone naphthoquinoxaline metabolite: a comparative study with the parent drug Dissertação do 2º Ciclo de Estudos Conducente ao Grau de Mestre em Toxicologia Analítica, Clínica e Forense, Faculdade de Farmácia, Universidade do Porto Trabalho realizado sob a orientação de: Professora Doutora Vera Marisa Costa Professora Doutora Emília Sousa Setembro de 2014 ii FULL REPRODUCTION OF THIS THESIS IS AUTHORIZED ONLY FOR THE PURPOSE OF RESEARCH, BY MEANS OF A WRITTEN STATEMENT OF THE INTERESTED PART, TO SUCH A COMMITMENT. iii Aos meus pais, Isaura e Paulino, Às minhas irmãs, Susana e Fátima, E ao Miguel. iv v ACKNOWLEDGMENTS Esta dissertação não poderia ter sido escrita sem a ajuda e a supervisão científica da Professora Doutora Vera Costa e Professora Doutora Emília Sousa, que não só me incentivaram como orientadoras como também me desafiaram para aumentar o meu conhecimento nestas duas grandes áreas. Ajudaram-me com o material de pesquisa, guiaram-me no laboratório e na dissertação, nunca aceitando menos do que os meus melhores esforços. Agradeço a ambas. À professora Doutora Vera Costa, minha orientadora neste trabalho, pelo tempo despendido, pela disponibilidade e paciência revelada, pelo espírito crítico, pela supervisão e orientação desta dissertação. Muito obrigada pelo profissionalismo, pela sincera amizade e pela total disponibilidade que sempre revelou para comigo. O seu apoio foi determinante na elaboração desta dissertação. À professora Doutora Emília Sousa, minha co-orientadora, pelo acompanhamento de todo o processo de síntese dos compostos e pelo apoio prestado. Agradeço também pela boa disposição em todos os momentos. A sua sabedoria foi essencial para que chegasse ao fim deste trabalho com um enorme sentimento de satisfação. À Mestre Ana Sara Gomes, pela sua colaboração ao longo de todo o trabalho realizado no laboratório de Química. Obrigada pela amizade, companheirismo e ajuda, factores muito importantes na realização desta dissertação, que permitiu que cada dia fosse encarado com motivação. À Doutora Sara, pelos conhecimentos transmitidos de cromatografia e pela prontidão com que esclareceu todas as minhas dúvidas. À Faculdade de Farmácia da Universidade do Porto, ao Laboratório de Química Orgânica e Farmacêutica (departamento de Ciências Químicas), e ao Laboratório de Toxicologia (departamento de Ciências Biológicas) por me terem proporcionado as condições necessárias à realização desta dissertação. À professora Doutora Maria Lourdes Bastos, coordenadora do mestrado em Toxicologia Analítica Clínica e Forense, e aos restantes professores que lecionaram neste mestrado, e que me incutiram um interesse cada vez maior na área da toxicologia. Agradeço a oportunidade e o privilégio que tive em frequentar este Mestrado que muito contribuiu para o enriquecimento da minha formação académica e científica. À professora Doutora Madalena Pinto, responsável do Laboratório de Química Orgânica e Farmacêutica por toda a simpatia e disponibilidade. vi Aos meus colegas e a todos os colaboradores destes laboratórios, por toda a ajuda prestada. Aos meus familiares e aos meus amigos, pelo incentivo prestado ao longo desta dissertação. Um agradecimento especial, ao Miguel por todo apoio e ajuda demonstrada, pela transmissão de confiança e de força, em todos os momentos. A todos, o meu sincero obrigado. This work was developed in REQUIMTE, Departamento de Toxicologia (Rede de Química e Tecnologia) and in Centro de Química Medicinal of University of Porto (CEQUIMED-UP) and CIIMAR – Interdisciplinary Centre of Marine and Environmental Research, Departamento de Ciências Químicas (Laboratório de Química Orgânica e Farmacêutica) both laboratories of the Faculty of Pharmacy, University of Porto. The work was funded through national funds from FCT – Fundação para a Ciência e a Tecnologia and also funded by EU/FEDER funds through the COMPETE program under the project FCT [EXPL/DTP-FTO/0290/2012]. The author of this thesis acknowledges all the participating laboratories and funding’s. 1 The results presented in this thesis led to the following works: Panel communications  A. Reis-Mendes, E. Sousa, F. Remião, F. Carvalho, M.L. Bastos, and V.M. Costa, “Mitoxantrone causes timeand concentration-dependent toxicity in H9c2 differentiated cells at pharmacological relevant concentrations”, 7th Meeting of young researches of U. Porto (IJUP ’14), Porto, Portugal, February 2014.  A. Reis-Mendes, E. Sousa, F. Remião, F. Carvalho, M.L. Bastos, and V.M. Costa, “At pharmacology relevant concentrations, mitoxantrone causes timeand concentration-dependent toxicity in H9c2 differentiated cells with autophagy involvement” 50th Congress of the European Societies of Toxicology, Edinburgh (Eurotox 2014), September 2014. Abstract in international journals: A. Reis-Mendes, E. Sousa, F. Remião, F. Carvalho, M.L. Bastos, and V.M. Costa, “At pharmacology relevant concentrations, mitoxantrone causes timeand concentrationdependent toxicity in H9c2 differentiated cells with autophagy involvement” Toxicol Lett 229, Supplement: S246. 2 3 ABSTRACT As anticancer treatment becomes more effective, a larger number of patients survive to cancer and the long-term side effects of anticancer therapy become an increasingly important issue. The ideal anticancer drug with maximal activity and neglectful toxicity to non-cancer cells is not yet found. Thus, anticancer treatment side effects still affect healthy organs. Mitoxantrone (MTX) has been used in patients with advanced breast cancer, prostate cancer, acute leukaemia, lymphoma and in multiple sclerosis. Cardiotoxicity leading to heart failure is a serious side effect that can affect up to 18% of MTX-treated patients, although its’ cardiotoxicity mechanisms are poorly known. The toxicity of the metabolites of anticancer drugs has been proved to be, at least, partially responsible for their reported cardiotoxicity. Regarding MTX, data regarding the synthesis and toxicological evaluation of its metabolites are scarcely available. Therefore, this work aims to synthetize MTX naphthoquinoxaline metabolite and to assess the toxicity of MTX and of the synthetized metabolite of MTX in an in vitro model. The synthesis of MTX naphthoquinoxaline metabolite was accomplished by the horseradish peroxidase (HRP) catalyzed H2O2 oxidation. With the enzymatic oxidation of MTX, several derivatives could be observed by chromatographic analysis, and two purified derivatives were isolated and identified as naphthoquinoxaline (NAPHT), a well-known metabolite of MTX, and naphthoquinoxaline dicarboxylic acid (NAPHTdi). In this study, several spectroscopic methods for the identification of the metabolites obtained in the synthesis were performed, namely infrared, high performance liquid chromatographywith diode array detector, proton nuclear magnetic resonance, and high resolution mass spectrometry. The purity obtained was 97% for NAPHT and 75% for NAPHTdi. Moreover, the toxicological evaluation of MTX and of the synthetized metabolite, NAPHT, was performed in differentiated H9c2 myocytes. The effects toward energetics, mitochondria, nuclei and lysosomes, and cell death mechanisms were primarily focused. H9c2 cells were differentiated using Dulbecco's Modified Eagle Medium supplemented with 10 nM retinoic acid (RA) and 1% foetal bovine serum (FBS) for 7 days (medium changed every two-days). After the differentiation protocol, cells were incubated with MTX (0.01 µM to 5 µM) or NAPHT (1 to 5 µM) for two time-points (24 and 48h), and cytotoxicity tests [lactate dehydrogenase leakage (LDH) assay, the 3-(4, 5-dimethylthiazol-2-yl)-2, 5-diphenyl tetrazolium bromide (MTT) assay and the neutral red (NR) uptake assay] were performed. Phase contrast microscopy and several staining’s were also used to evaluate cytotoxicity. Moreover, adenosine triphosphate (ATP) levels were assessed after MTX or NAPHT 10 The role of metabolism in the mitoxantrone’s toxicity ................................... 62 2. AIMS ....................................................................................... 67 3. MATERIALS AND METHODS ................................................... 71 3.1. Materials and chemicals ............................................................................. 71 3.2. Methods used in the synthesis of mitoxantrone metabolites ................... 72 3.3. Synthesis and purification of mitoxantrone metabolite naphthoquinoxaline .................................................................................................................... 72 3.4. High performance liquid chromatography analysis of mitoxantrone derivatives .................................................................................................................... 73 3.5. Cell culture model used in the toxicological evaluation ............................ 73 Subculturing H9c2 ........................................................................................... 74 H9c2 cells differentiation ................................................................................ 74 3.6. Cytotoxicity assays ..................................................................................... 74 Morphological evaluation ............................................................................... 74 3.7. Cytotoxicity tests ........................................................................................ 75 Lactate dehydrogenase kinetic leakage assay................................................ 75 MTT reduction assay ....................................................................................... 76 Neutral Red lysossomal uptake assay ............................................................ 77 3.8. Evaluation of mitochondrial potential ....................................................... 77 3.9. Determination of cellular ATP levels .......................................................... 78 3.10. Protein determination ............................................................................ 78 3.11. Statistical analysis ................................................................................... 79 4. RESULTS .................................................................................. 83 11 4.1. Synthesis of the mitoxantrone metabolite naphthoquinoxaline............... 83 4.2. Structure determination of naphthoquinoxaline and naphthoquinoxaline dicarboxylic acid ............................................................................................................... 85 4.3. Differentiation alters cell division and microscopic characteristics of H9c2 . .................................................................................................................... 91 4.4. Toxicological evaluation of mitoxantrone ................................................. 92 Microscopic evaluation of mitoxantrone-incubated H9c2 cells ..................... 92 Mitoxantrone led to a timeand concentration-dependent mitochondrial dysfunction ........................................................................................................................ 94 Mitoxantrone was only able to cause significant loss of membrane integrity at the highest concentrations tested ............................................................................................ 95 Mitoxantrone caused a significant lysosome uptake dysfunction in H9c2 differentiated cells .................................................................................................................... 96 Buthionine sulphoximine, an inhibitor of gamma-glutamylcysteine synthetase, had no effect on cellular death caused by mitoxantrone incubation in H9c2 cells .................. 96 N-acetyl cysteine, a glutathione precursor and reactive species scavenger, was not able to revert the cell death caused by mitoxantrone incubation ..................................... 97 L-carnitine, a mitochondrial enhancer was not able to revert cell death caused by mitoxantrone incubation ..................................................................................................... 98 3-Methyladenine, an autophagy inhibitor, did not inhibit cell death caused by incubation with mitoxantrone .................................................................................................. 99 The 3-methyladenine, an autophagy inhibitor, led to a partial protection to the impairment caused in lysossomal uptake by mitoxantrone..................................................... 99 The caspase inhibitor, Ac-LETD-CHO, did not inhibit the cell death caused by mitoxantrone in H9c2 cells ..................................................................................................... 100 Cycloheximide, a protein synthesis inhibitor, did not inhibit the cell death caused by incubation with mitoxantrone ............................................................................... 101 1-Aminobenzotriazole, a suicide CYP450 inhibitor, did not revert the cytotoxicity caused by mitoxantrone ..................................................................................... 102 12 Metyrapone, a competitive CYP450 inhibitor, did not inhibit the cytotoxicity caused by mitoxantrone in H9c2 cells .................................................................................... 102 Mitoxantrone was able to alter the ATP levels in H9c2 cells in a concentrationindependent manner .............................................................................................................. 103 4.5. Toxicological evaluation of naphthoquinoxaline ..................................... 104 Microscopic evaluation of naphthoquinoxaline incubated H9c2 cells .......... 104 Naphthoquinoxaline caused a timeand concentration-dependent mitochondrial dysfunction ...................................................................................................... 106 Naphthoquinoxaline was able to cause significant loss of membrane integrity ...................................................................................................................... 107 Naphthoquinoxaline caused a significant lysosome uptake dysfunction in H9c2 differentiated cells .................................................................................................................. 108 N-acetyl cysteine incubation increased cell death caused by naphthoquinoxaline ................................................................................................................ 108 3-Methyladenine, an autophagy inhibitor, caused a partial protection to the damage caused by naphthoquinoxaline ................................................................................. 109 Naphthoquinoxaline did not cause any significant alteration on ATP levels at an early time-point ................................................................................................................. 110 5. DISCUSSION AND CONCLUSIONS ......................................... 113 5.1. Synthesis and purification of the mitoxantrone metabolite naphthoquinoxaline ....................................................................................................... 113 5.2. Decreased foetal bovine serum and the addition of retinoic acid to H9c2 cells caused cell differentiation ................................................................................... 115 5.3. Cytotoxicity effects of mitoxantrone in differentiated H9c2 cells ........... 116 5.4. Evaluation of several pharmacological active molecules towards the cytotoxic effects of mitoxantrone .................................................................................. 118 5.5. Mechanisms of cell death induced by mitoxantrone ............................... 119 13 5.6. Evaluation of ATP levels and mitochondrial transmembrane potential of mitoxantrone.................................................................................................................. 122 5.7. The role of the cytochrome metabolism on mitoxantrone ..................... 122 5.8. Incubation naphthoquinoxaline in differentiated H9c2 cells .................. 124 5.9. Toxicological comparison between mitoxantrone and naphthoquinoxaline .................................................................................................................. 125 5.10. Final conclusions ................................................................................... 126 6. REFERENCES ......................................................................... 129 14 15 INDEX OF FIGURES Figure 1. Chemical structure of DOX and DNR. .............................................................. 33 Figure 2. The general metabolism of anthracyclines. ....................................................... 38 Figure 3. Oxidative stress related mechanisms for DOX-induced cardiotoxicity. Adapted from Costa et al., 2013. (SOD: superoxide dismutase) ............................................. 41 Figure 4. Iron involvement on DOX cardiotoxicity. Adapted from Torres and Simic, 2012. ................................................................................................................................. 43 Figure 5. Doxorubicin formation of aglycones. ................................................................. 47 Figure 6. Main metabolites of DNR. ................................................................................. 50 Figure 7. The main metabolites of cyclophosphamide (ALDH1: aldehyde dehydrogenase type 1). ..................................................................................................................... 54 Figure 8. The chemical structure of MTX. ........................................................................ 56 Figure 9. The major human metabolites of MTX. ............................................................. 60 Figure 10. The isolated products of the incubation of MTX with HRP: NAPHT and NAPHTdi. ................................................................................................................................. 83 Figure 11. Representative HPLC chromatograms (left) [= 254 nm, C18, linear gradient from 10 to 80% of eluent B within 30 min (eluent A: 0.1% aqueous solution of CF3COOH; eluent B: 100% MeOH)] and UV spectra (right) of MTX and isolated products NAPHT and NAPHTdi. ............................................................................... 84 Figure 12. Representative HPLC chromatogram [= 244 nm, C18, linear gradient from 10 to 80% of eluent B within 30 min (eluent A: 0.1% aqueous solution of CF3COOH; eluent B: 100% MeOH)] of 6.63 mg/mL solution of the crude product. MTX (k = 5.1), NAPHT (k = 6.3), NAPHTdi (k = 7.5). .................................................................................... 85 Figure 13. IR spectra of MTX. ......................................................................................... 87 Figure 14. IR spectra of NAPHT. ..................................................................................... 87 Figure 15. IR spectra of NAPHTdi. .................................................................................. 88 Figure 16. 1H NMR spectra of MTX. ................................................................................ 90 Figure 17. 1H NMR spectra of NAPHT. ............................................................................ 90 Figure 18. 1H NMR spectra of NAPHTdi. ......................................................................... 91 16 Figure 19. Fluorescence microscopy (Hoechst 33258 staining, A and C) and phase contrast microscopy images (B and D) of undifferentiated (A and B) and differentiated (C and D) H9c2 cells for 7 days. Images are representative of three independent experiments (scale bar 100 μm). .................................................................................................. 92 Figure 20. Fluorescence microscopy (Hoechst 33258 staining) (A, B, C), phase contrast microscopy (D, E, F) images of differentiated H9c2 cells after incubation with 2 and 5 μM of MTX for 24h. Control (A and D), MTX 2 μM (B and E), MTX 5 μM (C and F). Images are representative of three independent experiments (scale bar 100 μm). .. 93 Figure 21. Ethidium bromide and acridine orange (A, B, C) images of differentiated H9c2 cells in control and in cells incubated with 2 and 5 μM of MTX for 24h. Control (A), MTX 2 μM (B), MTX 5 μM (C). Images are representative of three independent experiments (scale bar 100 μm). .................................................................................................. 94 Figure 22. Mitochondrial transmembrane potential (A, B, C) images of differentiated H9c2 cells after incubation with 2 and 5 μM of MTX for 12h. Control (A), MTX 2 μM (B), MTX 5 μM (C). Images are representative of three independent experiments (scale bar 100 μm). ......................................................................................................................... 94 Figure 23. Mitochondrial dysfunction evaluated by MTT reduction assay in differentiated H9c2 cells incubated with 0.01, 0.1, 1, 2 and 5 μM of MTX for 24 (A) and 48h (B). Results are presented as mean ± SD of 5 independent experiments (total of 30 wells). Statistical analysis were performed using Kruskal–Wallis test, followed by the Dunn’s post hoc test (*p < 0.05, ***p < 0.001, ****p < 0.0001 vs. control; ##p < 0.01, ####p < 0.0001 vs. 0.01 µM; $p < 0.05, $$$$p < 0.0001 vs. 0.1 µM; +p < 0.05 vs. 2 µM). ......... 95 Figure 24. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 0.01, 0.1, 1, 2 and 5 μM of MTX for 24 (A) and 48h (B). Results are presented as mean ± SD of 5 independent experiments (total of 30 wells). Statistical analysis was done using the Kruskal–Wallis test, followed by the Dunn’s post hoc test (****p < 0.0001 vs. control; ####p < 0.0001 vs. 0.01 µM; $$p < 0.01, $$$$p < 0.0001 vs. 0.1 µM; &&p < 0.01, &&&p < 0.001 vs. 1 µM). .. 96 Figure 25. NR uptake (% of control) in differentiated H9c2 cells incubated with 0.01, 0.1, 1, 2 and 5 μM of MTX for 24 (A) and 48h (B). Results are presented as mean ± SD of 5 independent experiments (total of 30 wells). Statistical analysis was done using the Kruskal–Wallis test, followed by the Dunn’s post hoc test (****p < 0.0001 vs. control; ##p < 0.01, ####p < 0.0001 vs. 0.01 µM; $p < 0.05, $$p < 0.01, $$$$p < 0.0001 vs. 0.1 µM; &&&p < 0.001, &&&&p < 0.0001 vs. 1 µM). .................................................................... 96 17 Figure 26. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 2 μM of MTX and buthionine sulphoximine (BSO), an inhibitor of gamma-glutamylcysteine synthetase at 50 µM and BSO + MTX, for 48h. Results are presented as mean ± SD of 4 independent experiments (total of 20 wells). Statistical analysis was done using the Kruskal–Wallis test, followed by the Dunn’s post hoc test (****p < 0.0001 vs. control; $$$$p < 0.0001 vs. BSO 50 µM). ............................................................................................................ 97 Figure 27. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 2 μM of MTX and N-acetyl cysteine (NAC), an antioxidant and reactive species scavenger at 1 mM and NAC + MTX for 48h. Results are presented as mean ± SD of 4 independent experiments (total of 24 wells). Statistical analysis was done using the Kruskal–Wallis test, followed by the Dunn’s post hoc test (****p < 0.0001 vs. control; $$$$p < 0.0001 vs. NAC 1 mM). 98 Figure 28. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 2 μM of MTX, and LCarnitine (L-Carn), a mitochondrial enhancer at 2 mg/mL and L-Carn + MTX for 48h. Results are presented as mean ± SD of 4 independent experiments (total of 24 wells). Statistical analysis was done by using the Kruskal–Wallis test, followed by the Dunn’s post hoc test (****p < 0.0001 vs. control; $$$$p < 0.0001 vs. L-Carnitine 2 mg/mL. .... 99 Figure 29. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 2 μM of MTX, 3methyladenine (3-MA) 2.5 mM and 3-MA + MTX for 48h. DMSO (final concentration of 0.1% v/v) was used as vehicle. Results are mean ± SD of 4 independent experiments (24 wells). Statistical analysis: Kruskal–Wallis test, followed by the Student–Newman– Keuls post hoc test (****p < 0.0001 vs. vehicle; $$$$p < 0.0001 vs. 3-MA 2.5 mM). .... 99 Figure 30. NR uptake (% of vehicle) in differentiated H9c2 cells incubated with 2 μM of MTX, 3-methyladenine (3-MA) 2.5 mM and 3-MA + MTX for 24 (A) and 48h (B). DMSO (final concentration of 0.1% v/v) was used as vehicle. Results are mean ± SD of 7 independent experiments (42 wells). Statistical analysis: Kruskal–Wallis test, followed by the Student–Newman–Keuls post hoc test (****p < 0.0001 vs. vehicle; $$$$p < 0.0001 vs. 3-MA 2.5 mM; #p < 0.05 vs. 2 µM). ................................................................... 100 Figure 31. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 2 μM of MTX, Ac-LETD-CHO 100 µM (A) or 200 µM (B) and Ac-LETD-CHO + MTX for 48h. Results are mean ± SD of 4 independent experiments (24 wells). Statistical analysis: Kruskal–Wallis test, 18 followed by the Student–Newman–Keuls post hoc test (****p < 0.0001 vs. control; $$$$p < 0.0001 vs. Ac-LETD-CHO). ................................................................................. 101 Figure 32. Mitochondrial dysfunction evaluated by MTT reduction assay in differentiated H9c2 cells incubated with 2 μM of MTX, cycloheximide (Cyc.) 10 µg/mL and Cyc. + MTX for 48h. Results are mean ± SD of 3 independent experiments (18 wells). Statistical analysis: Kruskal–Wallis test, followed by the Student–Newman–Keuls post hoc test (***p < 0.001; ****p < 0.0001 vs. control; $$p < 0.01; $$$$p < 0.0001 vs. cycloheximide 10 µg/mL). ...................................................................................... 102 Figure 33. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) (A) in differentiated H9c2 cells incubated with 2 μM of MTX, 1aminobenzotriazole (1-abtz) 0.5 mM and 1-abtz + MTX for 48h. NR uptake (% control) (B) in differentiated H9c2 cells incubated with 2 μM of MTX, 1-abtz and 1-abtz + MTX for 48h. Results are mean ± SD of 6 independent experiments (36 wells). Statistical analysis were performed using Anova test, followed by the Tukey’s post hoc test (****p < 0.0001 vs. control; $$$$p < 0.0001 vs. 1-abtz 0.5 mM). ......................................... 102 Figure 34. Mitochondrial dysfunction evaluated by MTT reduction assay (A) in differentiated H9c2 cells incubated with 2 μM of MTX, metyrapone (MTP) 0.5 mM and MTP + MTX for 48h. Results are presented as mean ± SD of 5 independent experiments (30 wells). Statistical analysis was performed using Anova test, followed by the Tukey’s post hoc test (****p < 0.0001 vs. vehicle; $$$$p < 0.0001 vs. MTP; &&&&p < 0.0001 vs. 2 µM MTX. NR uptake (% of vehicle) (B) in differentiated H9c2 cells incubated with 2 μM of MTX, MTP and MTP + MTX for 48h. Results are mean ± SD of 4 independent experiments (24 wells). Statistical analysis: Kruskal–Wallis test, followed by the Student–Newman– Keuls post hoc test (****p < 0.0001 vs. vehicle; $$$$p < 0.0001 vs. MTP). DMSO (final concentration of 0.1% v/v) was used as vehicle. .................................................... 103 Figure 35. ATP levels in differentiated H9c2 cells incubated with 2 and 5 μM of MTX for 24h. Results, in nmol/mg protein, are presented as mean ± SD of 6 independent experiments. Statistical analysis was performed using Anova test, followed by the Tukey’s post hoc test (****p < 0.0001 vs. control). .................................................. 104 Figure 36. Fluorescence microscopy (Hoechst 33258 staining) (A, B, C), and phase contrast microscopy (D, E, F) images of differentiated H9c2 cells after incubation with 2 and 5 μM of NAPHT for 24h. DMSO (final concentration of 0.1% v/v) was used as vehicle (A and D), NAPHT 2 μM (B and E), and NAPHT 5 μM (C and F). Images are representative of three independent experiments (scale bar 100 μm). .......................................... 105 19 Figure 37. Ethidium bromide and acridine orange (A, B, C) images of differentiated H9c2 cells after incubation with 2 and 5 μM of NAPHT for 24h. DMSO (final concentration of 0.1% v/v) was used as vehicle (A), NAPHT 2 μM (B), NAPHT 5 μM (C). Images are representative of three independent experiments (scale bar 100 μm). ................... 105 Figure 38. Images of mitochondrial transmembrane potential of differentiated H9c2 cells after incubation with 2 and 5 μM of NAPHT for 12h (A, B, C). DMSO (final concentration of 0.1% v/v) was used as vehicle (A), NAPHT 2 μM (B), NAPHT 5 μM (C). Images are representative of three independent experiments (scale bar 100 μm). ................... 106 Figure 39. Mitochondrial dysfunction evaluated by MTT reduction in differentiated H9c2 cells incubated with 1, 2 and 5 μM of NAPHT for 24 (A) and 48h (B). DMSO (final concentration of 0.1% v/v) was used as vehicle. Results are presented as mean ± SD of 6 independent experiments (total of 35 - 36 wells). Statistical analysis were performed using the Anova test, followed by the Tukey’s post hoc test (*p < 0.05, **p < 0.01, ***p < 0.001 vs. vehicle; ##p < 0.01, ####p < 0.0001 vs. 1 µM; $$$$p < 0.0001 vs. 2 µM). ........................................................................................................................ 107 Figure 40. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 1, 2 and 5 μM of NAPHT for 24 (A) and 48h (B). DMSO (final concentration of 0.1% v/v) was used as vehicle. Results are presented as mean ± SD of 6 independent experiments (total of 35 - 36 wells). Statistical analysis was performed at 24h using the Kruskal–Wallis test, followed by the Student–Newman–Keuls post hoc test. For 48h, the Anova test, followed by the Tukey’s post hoc test, was used (***p < 0.001, ****p < 0.0001 vs. vehicle; ###p < 0.001, ####p < 0.0001 vs. 1 µM; $$p < 0.01, $$$$p < 0.0001 vs. 2 µM). ................................. 107 Figure 41. NR uptake (% of vehicle) in differentiated H9c2 cells incubated with 1, 2 and 5 μM of NAPHT for 24 (A) and 48h (B). DMSO (final concentration of 0.1% v/v) was used as vehicle. Results are presented as mean ± SD of 5 independent experiments (total of 30 wells). Statistical analysis was performed using the Anova test, followed by the Tukey’s post hoc test for analysis at 24h and Kruskal–Wallis test, followed by the Student–Newman–Keuls post hoc test, for analysis at 48h (**p < 0.01, ****p < 0.0001 vs. vehicle; #p < 0.05, ####p < 0.0001 vs. 1 µM; $$p < 0.01, $$$$p < 0.0001 vs. 2 µM).108 Figure 42. Cellular viability at 48h evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 2 μM of NAPHT and N-acetyl cysteine (NAC), an antioxidant, at 1 mM. DMSO (final concentration of 0.1% v/v) was used as vehicle. Results are presented as mean ± SD of 4 independent experiments (total of 24 wells). Statistical analysis was performed using the Anova test, 26 27 1. INTRODUCTION 28 29 1.1. The problem of cardiotoxicity caused by anticancer drugs Presently, cancer is one of the most concerning health issues worldwide. Over 3.45 million new cases of cancer (excluding non-melanoma skin cancers) and 1.75 million deaths were registered in Europe in 2012, with breast and prostate cancers counting for more than 800.000 new cases (Ferlay et al., 2013). Cancer is currently the second leading cause of death in Portugal, after cardiovascular diseases. The colorectal cancer is the leading cause of cancer-related mortality in Portugal (Adão et al., 2013). Briefly, cancer is a disequilibrium between cell division and cell death, accompanied with dysregulation of cell differentiation, which results in a limitless replicative potential and uncontrolled multiplication (Almeida et al., 2005, Ruddon, 2007). Most drugs used in cancer chemotherapy affect neoplastic transformation, cell proliferation, invasion and metastasis, and/or tumour-related angiogenesis. Ideally, targeted therapy spares normal cells and, therefore, would be better tolerated. The higher selectivity should translate into a wider therapeutic window that better enables the use of sufficient drug dosages to accomplish the therapeutic goals of tumour eradication or control, while minimizing the risk of resistance or toxicity (Bruce and Lin, 1969, Faivre et al., 2006). From the pharmacological point of view, agents used in anticancer therapy can be divided in chemotherapeutic agents (with several subgroups), biological or immunological agents. Radiation can also be used against cancer (Ewer and Suter, 2010). Chemotherapy is still the most frequent currently used therapy and it has largely contributed to the significant decrease of morbidity and mortality in several types of cancer. Even so, chemotherapy has general disadvantages, namely the potential development of cancer resistance and adverse effects due to the lack of selectivity of the drugs used (Espinosa and Raposo, 2010). Presently, the increase on the number of cancer survivors has led to the increase in toxic effects in non-target organs, especially long-term toxic effects. Several antineoplastic agents, namely cytotoxic antimetabolites, topoisomerase II inhibitors, alkylating and agents that interfere with tubulin, tyrosine kinase inhibitors, hormonal agents, and biological agents, among others, are widely reported to cause cardiac toxicity (Sá et al., 2009, Costa et al., 2013). The heart is an organ with limited potential for regeneration and, therefore, its damage may have long-term importance (Ewer and Suter, 2010). The cardiotoxicity resulting from antineoplastic therapy is a growing concern for oncologists and cardiologists. The cardiotoxicity resulting from the administration of anticancer drugs is multifactorial and complex, as it is influenced by factors as dose, possible drug interactions, potential synergism by co-administration or sequential 1. Introduction 30 administration of chemotherapeutic cardiotoxic drugs or the application of other therapeutics with potential cardiac toxicity (for example: radiotherapy in the mediastinum or immunotherapy) (Table 1) (Ewer and Suter, 2010, Senkus and Jassem, 2011, Octavia et al., 2012). Moreover, the biological variability of the patient and its health in the moment of treatment and thereafter has also a significant impact on the cardiovascular effects of the chemotherapeutic agent/combination of agents to be used. Therefore, when an attempt is made to diagnose or predict future cardiovascular effects after chemotherapy or radiation therapy, a large number of variables must be taken into account (Colombo and Cardinale, 2013). Several cardiotoxic manifestations can arise from xenobiotic exposure, namely damage in electrophysiology mechanisms, in the cardiac muscle and/or changes in the lumen diameter of coronary arteries. From the clinical perspective, the cardiotoxicity caused by anticancer agents is associated with temporary functional impairment, permanent loss of contractile elements, abnormalities of rhythm formation and/or conduction, as well as vascular effects that include vasospasms, alterations of blood pressure and thromboembolic sequelea (Senkus and Jassem, 2011). One of the most common cardiotoxic manifestations associated with the antineoplastic agents is the development of left ventricular dysfunction, namely decreased left ventricular ejection fraction (LVEF), which can progress to congestive heart failure. Heart failure is a severely debilitating disease that results, in a few years, on the death of the majority of the patients diagnosed (Colombo and Cardinale, 2013). The cardiac adverse effects of anticancer drugs may or may not be permanent. According to damage caused chemotherapy agents are divided in two groups: type I and type II (Table 1). The type I agents show cardiotoxicity associated with the total cumulative dose of the drug administered and a high propensity for cardiac cell death occurs after exposure (Ewer and Suter, 2010). Type I chemotherapy-related cardiac dysfunction, typically anthracycline-induced, is irreversible and typically associated with significant ultrastructural changes at biopsy (Todaro et al., 2013). Type II agents can cause a reversible cardiac dysfunction (Ewer and Suter, 2010). In fact, type II chemotherapy-related cardiac dysfunction (often termed trastuzumab-induced) is associated with reversible myocardial dysfunction rather than structural damage, it is highly reversible (up to 79%) and generally is not dose-related. However, a synergistic cardiotoxic effect between anthracyclines and trastuzumab has been demonstrated, especially when the two compounds are administered over a short period of time. Type II agents are not described as causing cell death; however, when used in combination with anthracyclines or other type I agents or in high-risk cardiac patients, some degree of cell death can actually occur (Todaro et al., 2013). 31 Table 1. Comparison of cardiotoxicity caused by agents of type I and type II. Table adapted from Ewer and Suter, 2010, Senkus and Jassem, 2011, and Adão et al., 2013. Type I (myocardial damage) example: Doxorubicin Type II (myocardial dysfunction) example: Trastuzumab Predominantly cell death Cellular dysfunction Biopsy changes: vacuoles, sarcomere/ myofibrillar disruption, apoptosis and necrosis No typical anthracycline-like biopsy changes: benign ultrastructure appearance Cumulative dose-related Not cumulative dose-related Permanent (myocyte death, bad prognosis) and irreversible cell damage; may stabilize, but subclinical damage persists Predominantly reversible (myocyte dysfunction, good prognosis), high probability of recovery after therapy interruption Risk factors:  Chemotherapy of combination prior or concomitant  Age  Previous cardiac disease  Hypertension Risk factors:  Chemotherapy prior or concomitant with anthracyclines or taxanes  Age  Previous cardiac disease  Obesity Mechanisms of cardiotoxicity:  Reactive oxygen species formation, oxidative stress/damage, energy collapse, and calcium and iron dysregulation Mechanisms of cardiotoxicity:  Block the human epidermal growth factor receptor 2 (HER2) signalling pathway, elimination of HER2/HER4related survival factors (gene expression, growth, glucose uptake, sarcomere turnover) Cardiovascular effects:  May result in heart failure and death Cardiovascular effects:  Low probability of cardiovascular mortality The best well-characterized cardiotoxic agents are the anthracyclines, namely doxorubicin (DOX), but other oncology drugs can also cause serious cardiovascular complications (Colombo and Cardinale, 2013). The anticancer drugs that adversely affect the cardiovascular system are systematized in Table 2. 32 Table 2. Agents used in anticancer therapy associated with cardiovascular toxicity. Table adapted from Ewer and Suter, 2010. (I) Agents associated with impaired left ventricular contractility (a) Agents associated with type I related cardiac dysfunctions (i) Anthracyclines: Doxorubicin, daunorubicin, epirubicin, idarubicin (ii) Anthraquinones: Mitoxantrone (b) Agents associated with type II related cardiac dysfunction (agents associated with potentially reversible dysfunction of the myocyte contractile elements) (i) Trastuzumab, lapatinib, sunitinib and others monoclonal antibodies/ tyrosine kinase inhibitors (c) Other cardiodepressant agents (i) Cyclophosphamide (especially high dose) (ii) α-Interferon (II) Agents associated with ischemia (a) 5-Fluorouracil (b) Capecitabine (c) Vinblastine (d) Vincristine (e) Bleomycin (f) Cisplatin (g) Biological response modifiers (III) Agents associated with hypertension or hypotension (a) Bevacizumab (b) Sunitinib (c) Sorafenib (d) Homoharringtonine (e) Interleukin-2 (IV) Miscellaneous cardiotoxic agents (a) Paclitaxel (bradycardia) (b) Arsenic trioxide (QT prolongation and torsades de points) (c) Radiation (pericardial damage, but may damage all cardiac structures) Several drugs have shown to cause toxicity towards the heart. Anthracyclines and mitoxantrone (MTX) have been widely described as cardiotoxic (Seiter, 2005, Menna et al., 2008, Adão et al., 2013, Todaro et al., 2013). In the introduction of this thesis, anthracyclines [DOX and daunorubicin (DNR)], cyclophosphamide and MTX will be addressed for their cardiotoxicity in independent sections: anthracyclines for their frequent clinical use and the 33 wide literature available; cyclophosphamide whose cardiotoxicity and antineoplastic effects strongly relates to its metabolism; and MTX as it is the drug of study in the present dissertation. 1.2. Anticancer drugs that lead to cardiotoxicity Anthracyclines Anthracyclines and related compounds are intercalating agents that are effective in treating many malignant diseases. DOX and DNR belong to the anthracyclines group. This family of compounds share a similar chemical structure (Figure 1) (Monsuez et al., 2010, Menna et al., 2012). According to the World Health Organization, to the group of anthracyclines belong 17 essential drugs that are used to treat cancers (Sikora et al., 1999). The skeleton of anthracyclines is composed by a tetracyclic system with quinone and hydroquinone moieties (often named anthraquinone skeleton), an amino sugar (daunosamine) attached by a glycosidic bond to C-7, and a side chain with a carbonyl group at C-13 (Figure 1) (Menna et al., 2012). Compounds R1 R2 R3 R4 DOX OH OCH3 OH H DNR H OCH3 OH H Figure 1. Chemical structure of DOX and DNR. 34 The anthracyclines were first isolated in the early 1950s from Streptomyces peucetius (Minotti et al., 2004a). During the decade of 1960, DNR was found to be very effective in treating leukaemia’s and lymphomas (Tan et al., 1967). Also in the sixties, a resembled molecule of DNR, the 14-hydroxydaunomycin or adriamycin (later named as DOX) was isolated and shown to be a more effective anticancer agent than DNR in solid tumours (Arcamone et al., 1969, Di Marco et al., 1969). Anthracyclines are shown to have a high therapeutic efficacy towards several types of cancers, although cardiotoxicity is one of their most significant adverse effects (Bonadonna et al., 1970). Over the years, the search for new compounds produced by the same bacteria led to the discovery of five new anthracyclines, however all still cardiotoxic (Takemura and Fujiwara, 2007). Anthracyclines are often used in combination with other antineoplastic agents, in the treatment of Hodgkin's disease, non-Hodgkin's lymphomas, acute leukaemia’s, bone and soft-tissue sarcoma, neuroblastoma, Wilm's tumour, and malignant neoplasms of the bladder, breast, lung, ovary, and stomach (Gewirtz, 1999, Minotti et al., 2004a). 1.2.1.1. Pharmacological mechanisms and general adverse effects of anthracyclines Several mechanisms for the anticancer proprieties of anthracyclines have been suggested: they are deoxyribonucleic acid (DNA) intercalating agents and disruptors of topoisomerase II, they cause oxidative stress, show alkylating properties and/or cause direct effects on the cell membrane. Anthracyclines have also been reported as inhibitors of the helicase, DNA polymerase, besides topoisomerase II. They induce apoptosis as a result of topoisomerase II inhibition (Uchegbu et al., 1995, Gewirtz, 1999, Minotti et al., 2004a, Takemura and Fujiwara, 2007, Ewer and Suter, 2010). Topoisomerase II is generally recognized to be the cellular main target of DOX. Anthracyclines act by stabilizing a reaction intermediate in which DNA strands are cut and covalently linked to this enzyme, therefore disrupting the DNA from returning to its natural structure (Simunek et al., 2009). Anthracyclines administration is accompanied by adverse drug reactions arising from their limited selectivity towards cancer cells (Aronson, 2006). Particularly common are bone marrow depression, which is dose-limiting, and gastrointestinal problems. Alopecia occurs in the majority of patients. Occasional hypersensitivity reactions may also occur. However, a cumulative-dose dependent cardiac toxicity has been the major limitation of DOX usage (Aronson, 2006). 1.2.1.2. Cardiotoxicity of anthracyclines 35 Anthracyclines cardiotoxicity was not detected during the preclinical animal testing; however, it was described during the initial clinical trials with DNR and DOX (Bonadonna et al., 1970). Since then, the cardiotoxicity associated with anthracyclines has been intensively studied. Anthracyclines can cause acute, subacute and late cardiac damage. Acute morbidity occurs during or shortly after drug infusion and includes arrhythmias (supraventricular tachycardia, ventricular ectopy) accompanied in some patients by heart failure, pericarditis-myocarditis syndrome, as well as non-specific ST–T changes and QT prolongation (Senkus and Jassem, 2011). This acute toxicity is usually reversible and not dose-dependent. Acute cardiotoxicity occurs in 1% of patients and no development of chronic heart failure occurs later in life (Todaro et al., 2013). The subacute cardiac toxicity occurs within a few weeks, clinically resembles myocarditis (with oedema and thickening of the left ventricle walls), and is accompanied by diastolic dysfunction with a 60% mortality rate. These acute and subacute forms of cardiac morbidity are, however, rare (1-4%) (Senkus and Jassem, 2011). Anthracyclines can cause short-term adverse effects that may occur up to one year after the start of therapy, while the adverse long-term effects can be diagnosed years or decades after the end of anticancer therapy. In fact, clinically the most relevant side effect of anthracyclines is chronic cardiac toxicity leading to left ventricular dysfunction and most importantly congestive heart failure (Senkus and Jassem, 2011). Late-onset chronic left ventricular dysfunction occurs in 1.6–5% of patients at least 1 year to 10 to 30 years after completion of therapy. A study documented chronic left ventricular dysfunction later in life in 23% of children treated with anthracyclines (Todaro et al., 2013). A strong correlation between chronic left ventricular dysfunction and the cumulative dose of anthracyclines is widely demonstrated (Todaro et al., 2013). The prognosis in anthracyclinerelated heart failure is poor, with 50% 2-year mortality in untreated established left ventricular dysfunction (Senkus and Jassem, 2011, Colombo and Cardinale, 2013). However, early diagnosis and appropriate management can significantly improve the outcome and the quality of life of the patients (Colombo and Cardinale, 2013). The risk factors for cardiac toxicity of anthracyclines are: the cumulative dose of the anthracycline administered, age, prior radiation therapy, concomitant (or previous) administration of other cardiotoxic anticancer agents and heart disease or pre-existing factors of cardiac risk (de Graaf et al., 1997, Pai and Nahata, 2000, Octavia et al., 2012, Adão et al., 2013) (Table 3). Clinical cases and pharmacovigilance data already obtained suggest that the most important risk factor for the development of delayed cardiotoxicity by anthracyclines is the maximum cumulative dose taken during all lifespan. After the absolute maximum cumulative dose threshold, the cardiotoxicity increases exponentially. This absolute cumulative dose varies according to the different anthracyclines (de Graaf et al., 42 dismutase (SOD). Hydrogen peroxide has a low toxic potential, being eliminated from the body by enzymatic antioxidant defence system (catalase and glutathione peroxidase). However, in the presence of the transition metals, especially iron, H2O2 and O2●- can generate the highly toxic ●OH. This process occurs during the Haber-Weiss reaction, which is actually quite slow. The presence of iron strongly catalyses the formation of ●OH, in a two-step process. In the first step, ferric ion (Fe3+) is reduced in to ferrous ion (Fe2+) by O2●- . Ferrous ion then reacts with H2O2 (formed during DOX reduction) and results in in the formation of ●OH (Figure 4). Unlike H2O2 and superoxide anion radical, ●OH is extremely reactive and cannot be neutralized by antioxidant enzymes (Kotamraju et al., 2002, Minotti et al., 2004b, Halliwell and Gutteridge, 2007). Moreover, DOX can directly bind to iron and, in the presence of oxygen, it can cycle between the Fe2+ and Fe3+ states (Figure 4). The DOX-Fe3+ complex can be reduced to the DOX-Fe2+ complex in the presence of NADPH CYP450 reductase, glutathione, or cysteine. These reactions are accompanied by the formation of O2●- and the conversion of the anthracycline quinone moieties to the semiquinone free radical (Doroshow et al., 1980, Rajagopalan et al., 1988, Vasquez-Vivar et al., 1997, Sawyer et al., 1999, Xu et al., 2005, Takemura and Fujiwara, 2007). As stated, the oxidation of the semiquinone again to the quinone results in the formation of the superoxide radical anion. It is believed that this radical is the initiator of oxidative stress caused by DOX in the heart. The reactive oxygen species formed, in particular the ●OH, can begin the peroxidation process of unsaturated membrane lipids and cause cellular damage in the myocardium (Kotamraju et al., 2002, Minotti et al., 2004b). In addition, reactive oxygen species can seriously affect nucleic acids and proteins, particularly ion channels and ion transporters (Halliwell and Gutteridge, 2007). It has been reported that DOX decreases the levels of enzymatic antioxidant defences (vitamin C, SOD, glutathione, catalase) making the heart more susceptible to oxidative stress (Costa et al., 2013). 43 Figure 4. Iron involvement on DOX cardiotoxicity. Adapted from Torres and Simic, 2012. Importantly, it was also found that NOS3 also contributes to cardiac dysfunction and injury observed after DOX administration of a single dose of 20 mg/kg, in a study using mice (Neilan et al., 2007). The results strongly suggest that DOX induces cardiac reactive species production via a NOS3-dependent mechanism. When NOS3 is deprived of Larginine or cofactors, superoxide can be generated from the oxygenase domain instead of ●NO, and this uncoupling may be associated with dissociation of NOS3 dimers. An alternative mechanism by which DOX can induce NOS3 to generate superoxide radical anion involves the conversion of DOX by the NOS3 reductase domain to an unstable semiquinone intermediate, which in turn produces superoxide radical anion in the presence of oxygen (Neilan et al., 2007). 1.2.2.3. Hypothesis of impairment of calcium homeostasis and cell death pathways by doxorubicin 44 An alternative hypothesis for the cardiotoxicity caused by DOX is the hypothesis of mitochondrial damage and changes on mitochondrial calcium concentration. The change in mitochondrial calcium is associated with reactive oxygen species production and dissipation of the mitochondrial membrane potential, which in turn may result in depletion of cellular adenosine triphosphate (ATP) (Zhou et al., 2001, Geisberg et al., 2012). This change in the transport of calcium leads to an irreversible decrease in storage ability and cell death, tissue damage, and consequently compromised cardiac contraction (Zhou et al., 2001). It is important to notice that the changes caused by the accumulation of reactive oxygen species, the alteration in calcium homeostasis and the mitochondrial damage observed are not isolated events, but can occur sequentially or simultaneously after DOX exposure (Kalyanaraman et al., 2002). Intracellular calcium is involved in the mechanisms of cell death. Usually, it is known that if exposed to low concentrations of anthracyclines, cardiomyocytes undergo apoptosis, whereas if exposed to high concentrations cardiomyocytes undergo necrosis (Doroshow et al., 1980, Rajagopalan et al., 1988, Vasquez-Vivar et al., 1997, Sawyer et al., 1999, Takemura and Fujiwara, 2007). When cell exposure to anthracyclines leads to apoptosis, one of the mediators of this process is the transcription factor and DNA damage sensor p53, accompanied with the activation of caspases (proteases) (Geisberg et al., 2012). However, ceramide, a lipid degradation product and the Fas receptor-ligand system have also been implicated in the damage caused by anthracyclines (Friesen et al., 1996). 1.2.2.4. Damage caused by metabolites of doxorubicin There are reports that attribute anthracyclines cardiotoxicity not only to oxidative stress, but also to the damage caused by their metabolites, namely their secondary alcohols derivatives and aglycones. DOX is metabolized by aldo/ketoses reductases that convert DOX to a secondary alcohol, the doxorubicinol (Figure 3). Those carbonyl reductases are present in the cytosol of cardiac cells (Boucek et al., 1987, Minotti et al., 1998). Cytosolic fractions from human myocardium have NADPH-dependent aldo/ketoses reductases capable of adding two electrons to the C-13 carbonyl group of DOX (Minotti et al., 1995). The doxorubicinol formed after enzymatic catalysis accumulates in the cardiac tissue and it is eliminated more slowly from the heart than the original drug, DOX (Gambliel et al., 2002). Stewart et al. demonstrated that doxorubicinol levels are higher (60%) than the levels of DOX in samples of heart tissue obtained from autopsies of patients treated with the parent compound (Stewart et al., 1993). Moreover, doxorubicinol levels are positively related to the cumulative administered dose of DOX (Stewart et al., 1993), which was identified as the main risk factor for the development of chronic cardiotoxicity since the early studies (Minow and Gottlieb, 45 1975). Autopsy tissues were collected from 35 patients who had received DOX at any time ante-mortem (Stewart et al., 1993). The major species found in human autopsy cardiac tissues were doxorubicinol (median concentration 92 ng/g, range 0 to 484 ng/g), and DOX (median 58 ng/g, range 0-1665 ng/g). Of the ten organs studied, heart ranked fifth regarding the median DOX concentration and ranked fourth regarding median doxorubicinol concentration. These results showed that heart accumulates large amounts of DOX and of its alcohol metabolite (Stewart et al., 1993). Behnia and Boroujerdi used phenobarbital as an inhibitor of aldo/ketoses reductases after which they used DOX at 10 mg/kg, intravenously. They demonstrated a decrease of doxorubicinol levels in vivo, as well as a reduction in cardiac damage as assessed by serum levels of creatine kinase in the rat, when phenobarbital was administered (Behnia and Boroujerdi, 1999). The correlation between secondary alcohols and cardiotoxicity has also been clearly shown by the use of transgenic animals. Mice with only one functional copy of the gene carbonyl reductase are healthy and grossly normal despite having decreased levels of carbonyl reductase transcript and protein (Olson et al., 2003). To control wild-type animals and to carbonyl reductase transgenic mice were administered DOX at 20 mg/kg intraperitoneally. Within 2 weeks, 91% of wild-type mice were severely affected by DOX when compared to 18% of mice with only one functional copy of the gene carbonyl reductase. Transgenic mice showed decreased circulating levels of the metabolite, doxorubicinol, after administration. Echocardiography and histological analysis showed that those transgenic mice were protected from gross and cellular level pathologies associated with DOX treatment (Olson et al., 2003). Another study with transgenic mice that overexpressed a heart-specific human carbonyl reductase was performed (Forrest et al., 2000); it was found that the administration of a single injection of DOX (15 mg/kg) in these animals resulted in a higher cardiotoxicity than in control wild-type mice (Forrest et al., 2000). Moreover, levels of doxorubicinol were found to be four times higher in the human carbonyl reductase expresser hearts than in the nonexpressers. Acute cardiotoxicity was evident by a 60% increase in serum creatine kinase activity and a 5-fold increase in cardiac damage seen in the transgenic mice measured by electron microscopy. The human carbonyl reductase expressers survived for 5 weeks when compared with 12 weeks for the controls DOX-treated animals. Electrocardiograph profiles and necropsies showed the cause of death to be the development of cardiomyopathies leading to congestive heart failure. Electron microscopy data showed swelling and major structural damage of the mitochondria in the human carbonyl reductase expresser’s mice that had DOX treatment (Forrest et al., 2000). Doxorubicinol plays an important role in DOX cardiotoxicity, possibly by amplifying the damage induced by other species (e.g., reactive oxygen species). Thus, abnormal 46 increases of doxorubicinol formation in the heart can lead to an acceleration in the progression of cardiotoxicity and cause the development of heart failure at lower cumulative doses of DOX than predicted (Minotti et al., 2004a). Doxorubicinol is an inhibitor of cardiac contractility, acts on the calcium transport ATPase of the sarcoplasmic reticulum and in the calcium release promoted by the calcium channel of the sarcoplasmic reticulum (Olson et al., 1988). In preparations obtained from cardiac tissue, doxorubicinol is a negative inotropic agent even more potent than DOX (Olson et al., 1988). It should be added that doxorubicinol interferes with ferritin (Minotti et al., 1995). Doxorubicinol is able to release Fe2+ from ferritin, which is the major site of iron storage in cytosolic medium of human cardiomyocytes (Minotti et al., 1995, Minotti et al., 1996). Another work suggested that this secondary alcohol may also contribute to the cardiotoxicity through its ability to impair the iron regulatory protein/ aconitase system involved in the homeostasis of the intracellular iron (Minotti et al., 1998). Changes in iron homeostasis have been implicated in the cardiotoxicity induced by the DOX. Moreover, doxorubicinol or reactive oxygen species may contribute to cardiotoxicity by inactivating iron regulatory proteins that modulate the fate of the messenger ribonucleic acid (RNA) for the transferrin receptor and ferritin (Minotti et al., 2001). Doxorubicinol is also able to remove iron from the catalytic Fe-S cluster of cytoplasmic aconitase (Minotti et al., 2001). The comparison between the properties of DOX and doxorubicinol has shown how a simple hydroxylation can strongly affect the toxic properties of those compounds (Heibein et al., 2012). Generally, doxorubicinol is less prone than DOX to cause oxidative stress. However, since it has a high cardiac accumulation (Stewart et al., 1993), doxorubicinol can lead to a large accumulation in the cardiac tissue when compared to the parent drug (Platel et al., 2000). DOX and doxorubicinol may still undergo reduction via the reaction of glucosidase generating metabolites called aglycones (Boucek et al., 1987) (Figure 5). The deglycosylation of the daunosamine sugar at C-7 aglycone of DOX may lead to 7hydroxydoxorubicin aglycone and 7-hydroxydoxorubicinol aglycone (Figure 5). The resulting hydroxyl group at C-7 can be metabolized to obtain 7-deoxydoxorubicin aglycone or 7-deoxydoxorubicinol aglycone (Takanashi and Bachur, 1976). 47 Figure 5. Doxorubicin formation of aglycones. Aglycones are powerful mitochondrial toxins and mitochondrial dysfunction further amplifies oxidative stress (Costa et al., 2013). The aglycones formed lead to increased permeability of the inner membrane in the mitochondria of the heart, being this phenomenon accompanied by the release of mitochondrial calcium, by mitochondrial swelling, disfunction of the membrane potential, and mitochondrial oxidation of pyridine nucleotides (Sokolove, 1988). Aglycones can also modify mitochondrial sulfhydryl groups and induce a calcium independent oxidation of mitochondrial NADPH, which appears to redirect electron transport from NADH to oxygen and result in the production of superoxide radical anion (Sokolove, 1994). Calcium release and sulfhydryl modification are shown to depend similarly on aglycone concentration and on the C-7 substituent of the anthracycline ring (Sokolove, 1988). In particular, the 7-hydroxydoxorubicin aglycone decreased the amount of calcium required to trigger the mitochondrial permeability increase. 7-Deoxydoxorubicin aglycone, the most prominent aglycone metabolite of DOX, was similarly effective in inducing calcium release, and both aglycones were substantially more effective than the parent drug (Sokolove and Shinaberry, 1988). Sulfhydryl modification induced in mitochondria by 7-deoxydoxorubicin aglycone is more efficient than by 7hydroxydoxorubicin aglycone. High potency of aglycones towards thiols may reflect increased access to key membrane thiols, due to their enhanced hydrophobicity (Sokolove, 1988). Moreover, Sokolove et al. examined the effects of DOX and its aglycone derivatives 48 on the pyridine nucleotide redox status of isolated, intact heart mitochondria. DOX aglycones induced the slow, calcium-independent oxidation of mitochondrial NADPH with 7-deoxydoxorubicin aglycone showing greater or equal potency than 7-hydroxydoxorubicin aglycone. That ability was much greater than DOX (Sokolove, 1991). Studies on laboratory animals have limited value in the prediction of the structure and reactivity of toxic metabolites in humans. Therefore, some authors use ethically cytosolic fractions of myocardial samples obtained during surgery for coronary bypass grafting (Licata et al., 2000). After reconstitution with NADPH and incubation with DOX, these fractions generate the alcohol metabolite doxorubicinol as well as 7-deoxydoxorubicin aglycone and 7-hydroxydoxorubicinol aglycone. These data show that the reduction of the side chain carbonyl group, reductase-type deglycosylation of the anthracycline, and the hydrolase-type deglycosylation followed by carbonyl reduction, respectively, occurs in human samples. The authors have suggested through structure-activity considerations that aglycones and doxorubicinol may inflict cardiac damage by inducing oxidative stress and by perturbing iron homeostasis, respectively (Figure 5) (Licata et al., 2000). 1.2.2.5. Hypothesis associated with immune and genetic mechanisms It was suggested by some authors that the cardiac damage caused by anthracyclines would increase due to their ability to activate the immune response. The damage caused by DOX in the cytoplasmic membrane of cardiomyocytes alone would compromise cardiac function, but the involvement of the immune system, with the consequent inflammatory response, exacerbates the cardiac dysfunction (Zhang et al., 1993). Zhang et al. showed that there was evidence of inflammatory responses in mice treated with DOX, since they exhibited increased dendritic antigen presenting cells. These results indicated that DOX treatment triggers immune reactions in which interstitial dendritic cells function as the antigen-presenting cells of the heart (Zhang et al., 1993). In addition to this inflammatory response, the release of vasoactive amines (histamine, catecholamines, prostaglandins, etc), after DOX administration, causes chemotaxis, further aggravating cardiac dysfunction (Huber, 1990, Zhang et al., 1993). Another accepted mechanism for anthracyclines toxicity results from altered expression of cardiac proteins or the activation of some genetic programs. DOX negatively influences the expression of genes involved in the production of constitutive proteins of the cardiac muscle (cardiac troponin, myosin, etc) and mitochondrial proteins and other proteins (Takemura and Fujiwara, 2007), thus affecting the quality of cardiac work. Several transcription factors, including factor nuclear kappa B, are sensitive to the redox state of the myocyte (Valen et al., 2001) and redox mechanisms may explain, at least in part, the effect on the expression of this gene induced by DOX. 49 Daunorubicin and its’ general pharmacokinetic data DNR was the first discovered active substance in the anthracyclines group (Iwamoto et al., 1968). This drug is for injectable use and has significant activity on acute myelogenous leukaemia, being rarely used in other types of tumours. DNR is also used in the treatment of neuroblastomas (Infarmed, 2011). DNR and DOX differ only in a hydroxyl group (Figure 1); however, their activity spectrum greatly differs (Infarmed, 2011). The recommended administration schedule for DNR is 25 to 45 mg/m2 for three days. Total doses higher than 550 mg/m2 are associated with a high cardiotoxicity risk (Table 3) (Chabner et al., 2012). DNR is widely distributed among tissues, mainly spleen, kidneys, liver, lungs, and heart (Trillet et al., 1985). DNR undergoes rapid and extensive metabolism in the liver and other tissues, mainly through cytoplasmic aldo/ketoses reductases. One hour after its administration, the predominant metabolite in the plasma is the metabolite daunorubicinol (Bachur and Gee, 1971) that has a plasma half-life of 26.7h and has antineoplasic activity (Takanashi and Bachur, 1975, Roche, 2002). Other metabolic routes for DNR include the glycosidic bond breakdown resulting in the production of aglycones, which have little or no antiproliferative activity and are further demethylated and conjugated via sulfate or glucuronide (Figure 6) (Di Marco et al., 1967, Alberts et al., 1971). In mouse and rabbit hepatocytes in suspension, DNR was shown to be metabolized mainly into daunorubicinol and deoxydaunorubicinol aglycones. The deoxydaunorubicin aglycone was rarely observed in both species. The anthracyclines conjugates were not observed in hepatocytes of any species (Gewirtz and Yanovich, 1986). The overall metabolites of DNR were identified as: daunorubicinol, daunorubicinol aglycone, deoxydaunorubicin aglycone, deoxydaunorubicinol aglycone, demethyl deoxydaunorubicinol aglycone, deoxydaunorubicinol aglycone 13-O-β-glucuronide, demethyl deoxydaunorubicinol aglycone 4-O-sulfate and demethyl deoxydaunorubicinol aglycone 4-O-β-glucuronide. The human DNR metabolism involves mainly reduction of the carbonyl, reductive glycosidic cleavage, O-demethylation, O-sulfation and Oglucuronidation (Figure 6) (Takanashi and Bachur, 1975). 50 Figure 6. Main metabolites of DNR. DNR and its metabolites are excreted in urine and bile (approximately 40% of the administered dose) (Rusconi et al., 1968, Alberts et al., 1971). 1.2.3.1. Cardiotoxicity of daunorubicin and daunorubicinol The risk of developing congestive heart failure increases even in the absence of other cardiac risk factors when the total cumulative dose of DNR exceeds 550-800 mg/m2 in adults (Adão et al., 2013), 300 mg/m2 in children over two years of age, or 10 mg/kg in children under two years of age (Mortensen et al., 1992). These cardiotoxicity effects are long known (Bonadonna and Monfardini, 1969, Marmont et al., 1969) and the risk factors for cardiotoxicity resemble other anthracyclines (Table 3). Several studies were performed in animal models to better highlight the cardiotoxicity of DNR and its main metabolite, daunorubicinol. In a rat model, and after 51 treatment with 6 doses 3 mg/kg intraperitoneally, DNR showed a strong ability of decreasing functional cardiac parameters, while daunorubicinol administered in the same dose and administration route did not cause any observed cardiotoxicity (Platel et al., 2001). Both treatments led to a similar accumulation of daunorubicinol in the myocardium while DNR was found only in the hearts of rats treated with this drug and not with daunorubicinol (Platel et al., 2001). Direct infusion to isolated rat hearts of the same dose (10 µM) of DNR or daunorubicinol induced a depression of heart function in both groups; however, only DNR induced a progressive increase in diastolic pressure. On the other hand, in cardiac sarcoplasmic reticulum vesicles, daunorubicinol at a concentration of 5.5 µ/g tissue or 10 µM inhibited calcium uptake by 39 ± 3%, whereas 10 µM of DNR did not cause any detectable inhibition (Cusack et al., 1993). Another study, however, demonstrated that both DNR and daunorubicinol were several fold more potent at inhibiting than they were at stimulating sarcoplasmic reticulum calcium release in canine heart vesicles (Olson et al., 2000). Concentrations that cause 50% inhibitory effect (IC50) on caffeine-induced calcium release of DNR and daunorubicinol were 1.2 and 0.6 µM, respectively, and for spontaneous calcium release were 3 and 1 µM, respectively (Olson et al., 2000). The model used and the amounts of calcium used for loading may explain these dissimilar results. After DNR or daunorubicinol administration with 6 doses of 3 mg/kg intraperitoneally on rat, DNR strongly decreased the cardiac functional parameters, while daunorubicinol did not induce cardiotoxicity (Platel et al., 2001). In summary, DNR follows the same metabolic route of DOX, namely regarding the reduction to daunorubicinol (Loveless et al., 1978). However, in contrast to DOX, DNR administration leads to much higher concentrations of this metabolite in the plasma and tissues than the parent compound (Robert and Gianni, 1993, Platel et al., 2001), whereas the formed alcohol metabolite is less cardiotoxic than the initial drug (Platel et al., 2001). 1.3. Alkylating agents (i.e. cyclophosphamide) and their cardiotoxicity Alkylating agents are highly reactive compounds that readily bind to phosphates, amines, imidazole and hydroxyl groups found in nucleic acids. The cytotoxic and mutagenic effects of alkylating agents are directly related to DNA alkylation, which can lead to the replacement of base pairs, breaks or the formation of covalent bonds between DNA strands, stopping the DNA replication and causing cell death (Chabner et al., 2012). The most important pharmacological actions of the alkylating agents affect DNA synthesis and cell division. Therefore, they are more cytotoxic in rapidly proliferating tissues. However, these agents affect both healthy and cancer cells and notwithstanding act on the cells at any stage of the cycle (Chabner et al., 2012). Cyclophosphamide is an alkylating agent used orally or 58 clinical studies using a single intravenous infusion of MTX, the administered dose ranges between 5-14 mg/m2, while the administration schedule can vary according to the disease and response to therapy, ranging from 4-6 weeks for the treatment of acute myeloid leukaemia, 3 weeks for the treatment of prostate cancer (Fox, 2004), and 3 months for multiple sclerosis (Scott and Figgitt, 2004). The plasma concentration-time curve of MTX can be best fitted with three exponentials: a rapid half-life of 4.1 to 10.7 min, corresponding to the moment when MTX rapidly leaves the plasma and binds to the endothelial surface; an intermediate half-life of 0.3 to 3.1h, corresponding to the distribution phase; and a long terminal half-life of 8.9h to 9 days (Batra et al., 1986, Ehninger et al., 1990, Fox, 2004). Therefore, MTX exhibits a rapid initial distribution phase followed by a relatively slow elimination phase from the deep tissues (Fox, 2004). In summary, the pharmacokinetic characteristics of MTX are: rapid plasma clearance, long half-life elimination without the need for significant change in the case of kidney or liver dysfunctions and persistent tissue concentrations (Stewart et al., 1986). MTX has a very large volume of distribution, suggesting that most of the drug is accumulated in the tissues (Stewart et al., 1986). MTX is mostly bound to human plasma proteins (approximated 78%) (Batra et al., 1986). The elimination of the drug is slow, with a half-life of about 12 days (range 5-18 days) with persistent tissue concentrations in mice with tumour xenographs (Patel et al., 2013). One study examined the concentrations of MTX in autopsy tissue samples from 11 patients who had received the drug 10 to 272 days before death (Stewart et al., 1986). The total cumulative lifetime dose of MTX for these patients ranged from 6 to 100 mg/m2. MTX was detected in tissues from all patients, even 272 days after administration. The highest concentrations were found in the thyroid, liver, and heart. The lowest concentration was reported in the brain (Stewart et al., 1986). Also in humans, MTX was detected 35 days after a single dose of 12 mg/m2 in the liver (1140 ng/g) and heart (716 ng/g) (Batra et al., 1986). These studies were conducted in cancer patients, and the results may therefore be different in patients with multiple sclerosis who are dosed differently than cancer patients (Fox, 2004). The major route for the elimination of MTX from the body is biliary excretion, as renal clearance can account for only up to 10% of the total clearance of the drug (Fox, 2004). 14C-labeled MTX was administered to eight patients who had advanced soft tissue cancers (Alberts et al., 1985) and only 6.5% of the total MTX dose administered was excreted in the urine as unchanged drug over the next 5 days. The mean recovery of 14C-labeled material in faeces over 5 days was 18.3% of the administered dose. Thirty-five days after MTX administration, one of the patients died of progressive kidney cancer, and approximately 15% of the 14C dose was found in seven major organs (Alberts et al., 1985). 59 Two of the MTX metabolites were isolated from the urine of patients and characterized as of monoand dicarboxylic acids resulting from oxidation of the terminal methyl groups in the side chains (Figure 9) (Chiccarelli et al., 1986). Regarding the interspecies variability, the major metabolic difference between rat and humans relates to monoand dicarboxylic acids that are the main products of human metabolism of MTX, whereas in rat, they are residual (Blanz et al., 1991b, Richard et al., 1991). Therefore, the metabolic differences between the species can determine that electrophilic metabolites can be formed in higher amounts in rats. Controversial data still exists whether the MTX undergoes one electron reduction or reduction by two electrons (Wolf et al., 1986, Duthie and Grant, 1989, Fisher and Patterson, 1992). In fact, the reduction of MTX by flavin reductase is not facilitated, due to its low reduction potential (Fisher and Patterson, 1992). Thus, the preferred metabolism route of MTX seems to occur through two electron reduction as evidenced in studies where inhibition of CYP450-mediated metabolism is done (Duthie and Grant, 1989, Mewes et al., 1993, Li et al., 1995). It has been shown that a cyclic metabolite of MTX, the naphthoquinoxaline (NAPHT) (Figure 9) is a product of biotransformation of MTX in vivo in humans, pig, and rat (Blanz et al., 1991b, Bruck and Bruck, 2011). This metabolite has been described as the product of metabolism through systems containing CYP450 enzymes and peroxidases and it is found in urine (Blanz et al., 1991b, Bruck and Bruck, 2011). Other studies suggest that this metabolite has a significant role in the pharmacological anticancer activity of MTX (Mewes et al., 1993, Shipp et al., 1993, Feofanov et al., 1997, Panousis et al., 1997). After incubation of urine from MTX-treated patients with β-glucuronidase or sulfatase, no differences between control and test samples were found (Smyth et al., 1986). Therefore, glucuronidation and conjugation with sulfate did not seem to be part of the biotransformation of MTX in humans in that study (Smyth et al., 1986). However, other works confirm that the glucuronides are also urinary metabolites of MTX in humans (Batra et al., 1986, Blanz et al., 1991a). The human urinary glucuronide metabolite was detected in 1991 (Blanz et al., 1991a), whereas the MTX glutathione metabolites were only detected in animals (Rossato et al., 2013a) or in in vitro models (Mewes et al., 1993). In rat liver hepatic microsomes or S9 fraction, MTX formed glucuronic and glutathione conjugates (Wolf et al., 1986, Rossato et al., 2013a). Incubation of MTX with peroxidase/H2O2, in the presence of glutathione, led to the formation of two MTX glutathione conjugates (Blanz et al., 1991b). An in vivo study was performed to investigate whether the metabolites of MTX previously found in vitro were also present in liver and heart of rats, after one intraperitoneal administration of 7.5 mg/kg MTX 24h before (Rossato et al., 2013a). MTX and NAPHT metabolite were present in the heart and liver of male rats (Rossato et al., 2013a). Moreover, 60 in that study, an in vitro study was also performed using rat liver S9 fractions. In that system, MTX metabolism led to the formation of five metabolites all of them maintaining the chromophore group. Of those five metabolites, NAPHT and a new metabolite, an acetoxy ester derivative, were identified as resulting of MTX metabolism (Rossato et al., 2013a). Compounds R1 R2 R3 R4 Monocarboxylic acid of MTX COOHCH2OHHHDicarboxylic acid of MTX COOHCOOHHHGlucuronide metabolites of MTX CH2OHCH2OHHGlucuronideFigure 9. The major human metabolites of MTX. The cardiotoxicity of mitoxantrone Despite its initial objective, MTX is potentially cardiotoxic (Avasarala et al., 2003, Seiter, 2005). In 1993, Estorch et al. suggested that MTX was less cardiotoxic than anthracyclines (Estorch et al., 1993), but this conclusion has not been corroborated by other studies (Thomas et al., 2002, Avasarala et al., 2003). In fact, the cardiotoxicity of MTX remains a major concern for patients receiving intensive and/or prolonged therapy with MTX (Ehninger et al., 1990). Presently, the recommended maximum lifetime cumulative dose of MTX is 140 mg/m2, with 2.6 to 13% of patients developing cardiac toxicity (Seiter, 2005). As early as 1984, the cardiotoxicity of MTX was detected in 34 patients with advanced breast cancer that had never received previous chemotherapy. They were treated with MTX 61 (14 mg/m2 intravenous every 21 days) (Coleman et al., 1984). Before starting the MTX therapy, and every 3 months thereafter, radionuclide assessment of ventricular performance was obtained at rest and in response to stress. Ten patients showed a deterioration in ejection fraction and two of them developed congestive cardiac failure (Coleman et al., 1984). In another study, biopsies were examined in a small number of patients treated with MTX by Unverferth and colleagues. This study found cardiac histopathological changes similar to those found in treatment with anthracyclines. The nuclear chromatin changes included shrinkage, nucleolar agglutination and degeneration. They also observed mitochondrial swelling and tubular swelling in cardiomyocytes (Unverferth et al., 1983). In a study by Benjamin et al., biopsies from patients treated with MTX revealed dilatation of sarcoplasmic reticulum with vacuoles and myofibrillar changes (Benjamin et al., 1985). Those histological changes were also found to be initial cardiac findings in DOX therapy (Benjamin et al., 1985). In addition, an increase in cell damage with a higher dose of MTX was observed. Thus, the cardiac histological data of MTX indicates that it produces adverse changes in the human heart, which are similar to those produced by DOX (Benjamin et al., 1985). The clinical cardiotoxicity signs of MTX include decreased LVEF, congestive heart failure, ischemic chest pain, arrhythmias and conduction abnormalities on the ECG (Henderson et al., 1989, Estorch et al., 1993). Some predisposing factors for MTX cardiotoxicity include prior exposure to anthracyclines, thoracic radiation or pre-existing heart disease, although cardiotoxicity can occur in the absence of these risk factors (Benjamin et al., 1985) (Table 3). Cardiotoxicity of mitoxantrone in experimental models The chronic cardiotoxicity has been studied in animal models in an attempt to discover and counteract the cardiotoxic mechanisms associated (Zbinden and Beilstein, 1982, Perkins et al., 1984, Rossato et al., 2013a, Rossato et al., 2014). In a study by Zbinden and Beilstein with a twice a week administration of MTX, and then interrupted for 18 days and then resumed with four other injections, several evaluations were made (Zbinden and Beilstein, 1982). After about six weeks, the authors observed significant changes in the ECG, elevated serum marker enzymes of cardiac damage and marked changes in mitochondrial structure in heart tissue (Zbinden and Beilstein, 1982). In the same study the authors compared the mitochondrial alterations induced by DOX and MTX in vivo (Zbinden and Beilstein, 1982). In these experiments, MTX injections (1 mg/kg) and DOX (2 mg/kg) were administered twice per week for four weeks. Each week, a group of animals were sacrificed and mitochondria were isolated from the heart, being the oxygen uptake and oxidative phosphorylation activity measured. Under these conditions, the 62 mitochondrial function was inhibited in a time dependent manner, in both MTX and DOX. MTX caused a marked inhibition of the activity of the pump Na+/K+ ATPase (from 41% to 68% of the control) in the first week and that measurement remained unchanged thereafter (Zbinden and Beilstein, 1982). These results were in agreement with the findings of Neri and Cini-Neri (Neri and Cini-Neri, 1986); the later observed a progressive reduction in oxygen consumption and loss of ATP and phosphocreatine from rat heart samples incubated with DOX or MTX in vitro (Neri and Cini-Neri, 1986). Chronic MTX cardiotoxicity was also studied in mice with a twice per week of MTX administration (2 mg/kg) at weeks 1, 2, 5, 6 and 7 (Perkins et al., 1984). The rats were sacrificed during week 11 and the hearts were evaluated by histology. Of the 14 mice treated with MTX, 12 exhibited myocardial injury. The Bertazzoli scale was used to quantify the degree of cardiac damage and the rate 0 degree indicating that there is no difference between the control animals, while grade 4 indicates the most marked damage of myocardial fibres (Bertazzoli et al., 1979). In the Perkins et al. study, twelve animals showed myocardial changes: five with grade 1, four with grade 2 and three with grade 3. Infiltration focal mononuclear cells also occurred in three hearts (Perkins et al., 1984). In a more recent study, male Wistar rats were treated with 3 cycles of 2.5 mg/kg of MTX at day 0, 10, and 20 and then one treated group was euthanized on day 22 to evaluate the early MTX cardiac toxic effects, while the other group was euthanized on day 48, to allow the evaluation of MTX late cardiac effects (Rossato et al., 2014). Decreased levels of plasma total creatine kinase and creatine kinase-MB were detected in the early sacrificed animals while increased plasma levels of lactate were seen in the animals sacrificed later on. Increased cardiac relative mass and microscopic changes were evident in both treated groups, while activity of mitochondrial complexes was changed in both groups when compared to control. MTX induced an increase in the complex IV and complex V activities in early sacrificed treated group, while a decrease in the complex V activity was accompanied by the reduction in ATP content in the late-sacrificed rats (Rossato et al., 2014). The role of metabolism in the mitoxantrone’s toxicity Most studies on MTX cardiotoxicity in vitro were performed in order to evaluate the MTX cardiotoxicity compared to DOX, instead of trying to explain the biochemical basis of cardiotoxicity that is observed in MTX. However, these studies have contributed immensely to the overall understanding of the biochemical activity and the toxicity of the drug. Metabolism is involved in MTX anticancer proprieties, however, little is known about the influence of metabolism on MTX cardiotoxicity. 63 The oxidoreductive metabolism of MTX has a significant role on its antitumour effects. MTX has particular effectiveness in tumours with high contents of peroxidases (Blanz et al., 1991b, Bruck and Bruck, 2011), and it was verified that the inhibitory effect of MTX on cell growth was prevented by inhibiting the activity of CYP450 mixed oxidase function in a human hepatoma-derived cell line (Duthie and Grant, 1989). Similar results were obtained with rat hepatocytes (Mewes et al., 1993) and with human breast cancer cells (Li et al., 1995) incubated with MTX. Moreover, phorbol ester-stimulated human neutrophils can bioactive MTX through myeloperoxidase metabolism and the generated metabolites can form adducts with DNA, although the chemical nature the metabolites is unknown (Panousis et al., 1997). As stated, the role of MTX metabolism in its induced cardiotoxicity is poorly studied; however, two studies have been made that can give some hints on that subject (Shipp et al., 1993, Rossato et al., 2013a). In the work by Rossato et al., H9c2 cells were incubated with non-metabolized MTX or MTX after metabolization (Rossato et al., 2013a). It demonstrated that the cytotoxicity caused by MTX after metabolization was significantly higher than that observed in the H9c2 cells incubated with non-metabolized MTX. The coincubation of MTX with CYP450 and CYP2E1 inhibitors partially prevented the cytotoxicity caused by the drug, thus highlighting that the metabolism of MTX is relevant for its undesirable effects (Rossato et al., 2013a). The work by Shipp et al. demonstrates that the metabolite NAPHT has a higher potential on ATP depletion than MTX and, importantly, that the bioenergetics impairment hugely depends on iron availability (Shipp et al., 1993). Furthermore, the catechol ring of MTX and of the metabolite NAPHT seems to be relevant to the observed ATP decrease since ametantrone has no significant effect on ATP levels (Shipp et al., 1993). One can speculate by putting together all the scarce data available that MTX is not able in vivo to cause an initial redox cycle that is sufficient to cause an early and abrupt oxidative stress (as do the aglycones and semiquinones of DOX); however, MTX and its NAPHT metabolite rapidly accumulate in the heart (Ehninger et al., 1985, Rossato et al., 2013a) easily impairing ATP homeostasis in the heart, factors that seem to depend on iron (Shipp et al., 1993). 64 65 2. AIMS 66 67 The toxicity of anticancer drugs and particularly of MTX towards the cardiovascular system is well known. However, the mechanisms involved are largely unknown. Pharmacokinetic metabolism is involved in the cardiotoxicity of several anticancer drugs, whereas the data regarding to MTX metabolism and its relation to MTX-induced cardiotoxicity are scarce. The full understanding of the cardiotoxic effects induced by MTX and/or metabolites is important to reveal the underlying cardiac mechanisms, but also to pursue cardioprotective strategies when MTX is to be used. The main aim of this dissertation was to clarify the underlying mechanisms by which MTX and, its metabolite NAPHT, elicit cardiotoxicity. The following specific objectives were pursued:  Synthesis of MTX-metabolite, NAPHT,  Structure characterization of the obtained derivatives of MTX,  Evaluation of the cytotoxicity caused by MTX in different time-points and concentrations,  Evaluation of several pharmacological-active drugs towards MTX-elicited cytotoxicity in H9c2 differentiated cells,  Evaluation of cytotoxicity caused by NAPHT in different time-points and concentrations,  Evaluation of pharmacological-active drugs towards NAPHT-elicited cytotoxicity in H9c2 differentiated cells. 2. AIMS 74 model to assess in vitro cardiotoxicity (Kimes and Brandt, 1976), especially due to its metabolic features, which are comparable to those found in rat heart (Zordoky and El-Kadi, 2007). Subculturing H9c2 H9c2 cells were maintained in the proliferative state in the presence of 10% FBS. H9c2 cell were grown in complete medium: DMEM with high glucose supplemented with 10% FBS and antibiotics (10 000 units/mL penicillin and 10 000 µg/mL streptomycin) at 37°C with 5% CO2. Cell passaging was done by trypsinization. All experiments were carried out with cells before reaching 70–80 % confluence (Ruiz et al., 2012). The cell line was used between passage 15 and 30. H9c2 cells differentiation Differentiation into a “cardiac like” phenotype was induced with 1% FBS in culture medium supplemented with RA 10 nM (medium changed every two-days) (Pereira et al., 2011, Ruiz et al., 2012). The differentiation and decreased proliferation was assessed by contrast phase microscopy and the fluorescent nuclear dye Hoechst 33258 (Soares et al., 2013). 3.6. Cytotoxicity assays After trypsinization, cells were placed in 48 well-plates, 12 well-plates, or 6 wellplates with the density of 24 000 cells/mL. After the 7-day differentiation protocol described, H9c2 cells were incubated with different chemicals. The first tests performed were the cytotoxicity curves of MTX and metabolite, combined with the morphological evaluation, using by phase contrast microscopy. After cell differentiation protocol, cells were incubated with MTX in concentrations that ranged from 0.01 to 5 µM and NAPHT in concentrations that ranged from 1 to 5 µM, and two time-points were selected (24 and 48h) for the cytotoxicity tests. The cytotoxicity tests used were: LDH leakage assay, the MTT assay and the NR uptake assay (Soares et al., 2013). Morphological evaluation 3.6.1.1. Contrast phase microscopy Cell cultures were assessed morphologically by phase contrast microscopy at the selected time-points in 6 or 12 well-plates. Cells were examined in a Nikon Eclipse TS100 equipped with a Nikon DS-Fi1 camera (Japan). 75 3.6.1.2. Hoescht nuclear staining For Hoechst staining, H9c2 cells were seeded in 48 well-plates with 24 000 cells/mL and differentiated or not for 7 days. After reaching the end-point, cells were fixed in 4% paraformaldehyde (10 min, 4ºC) and washed with PBS (three times). Cells were then stained with the nuclear dye Hoechst 33258 for 10 min at 37°C (protected from light), after which they were washed with PBS (three times) at room temperature. Cells were examined in Nikon Eclipse TS100 equipped with a Nikon DS-Fi1 camera using a fluorescent filter (λexcitation maximum = 346 nm and λemission maximum = 460 nm) (Soares et al., 2013). 3.6.1.3. Ethidium bromide and acridine orange staining The fluorescent DNA-intercalating dyes ethidium bromide and acridine orange are suitable to distinguish live from dead cells. Also, the ethidium bromide and acridine orange staining allows the microscopic morphological discrimination between necrotic and apoptotic cell death and acridine largely accumulates in acidic vesicles. Ethidium homodimers do not penetrate intact cellular membranes. Therefore, ethidium bromide only intercalates with nucleic acids if the outer cellular membrane has disintegrated. In contrast, the fluorescent cationic dye acridine orange diffuses through intact membranes of live cells. When bound to DNA, it reaches an emission maximum at 525 nm (Capela et al., 2013). The staining of H9c2 was done after removing all the growth medium of the 48-wells after the incubation period with MTX or NAPHT. To the cells were added 200 µL of heated PBS with Ca2+e Mg2+, and then 8 µL of the stock solution ethidium bromide and acridine orange solution (0.5 mg/mL). A 5 min incubation in the dark was followed. After removing the ethidium bromide and acridine orange, cells were washed with warm PBS with Ca2+e Mg2+ (200 µL/ well) and then new medium was added. Cells were examined in Nikon Eclipse TS100 equipped with a Nikon DS-Fi1 camera, using a standard fluorescein filter (λexcitation = 485 nm and λemission = 525 nm). 3.7. Cytotoxicity tests Lactate dehydrogenase kinetic leakage assay H9c2 cells were seeded in 48-well plates. For MTX and NAPHT evaluation of induced cell death, cells were incubated for 24 or 48h with different drug concentrations. After cell treatment, cell death was evaluated as the measurement of membrane integrity seen by the percentage of LDH release over the total LDH. For experiments with caspase 8 and 9 inhibitor, Ac-LETD-CHO (200 µM and 100 µM) (Shi and Shen, 2008), the antioxidant N-acetyl cysteine, a glutathione precursor and reactive species scavenger (1 mM) (Martins 76 et al., 2013, Rossato et al., 2013b), the buthionine sulphoximine, an inhibitor of gammaglutamylcysteine synthetase (50 µM) (Ferreira et al., 2013), L-carnitine, a mitochondrial enhancer (2mg/mL) (Rossato et al., 2013b), 3-methyladenine, an autophagy inhibitor (2.5 mM) (Soares et al., 2014), 1-aminobenzotriazole, a suicide substrate for CYP450 (0.5 mM) (Shi et al., 2011a), the addition of these compounds was done 1h prior to the addition of MTX 2 µM. In the case of NAPHT 2 µM incubation with N-acetyl cysteine (1 mM), this later molecule was added 1h prior to the addition of NAPHT. The quantification of LDH activity was made using a colorimetric method, based on the reversible reduction of pyruvate to lactate (last step of anaerobic glycolysis) in the presence of β-NADH, as described below (Capela et al., 2006). 3.7.1.1. Extracellular lactate dehydrogenase activity LDH can be found outside the cell only when there is cell death (apoptotic or necrotic type) or lysis of cytoplasmatic membrane. The extracellular activity of LDH was determined after the suitable amount of medium was added to β-NADH 0.15 mg/mL solution at room temperature, in 96-microplate wells. Finally, pyruvate 22.7 mM was added to start the reaction. NADH oxidation to NAD+ was measured at 340 nm for 4 and a half min, using a 96-well plate reader (BioTek Instruments, Powerwave X, USA). The slope of the curve obtained was used for the calculation of the % LDH leakage (Capela et al., 2006). 3.7.1.2. Total lactate dehydrogenase activity Total LDH activity was determined after adding 20 µL of Triton X-100 5% to each well and incubated at 37°C for 30 min. The cells were totally lysed by this procedure and then the activity of the LDH obtained was evaluated as described previously. Medium was added to 0.15mg/mL β-NADH solution at room temperature, in a 96-well microplate. Finally, pyruvate 22.7mM was added to start the reaction as described in the previous section (Capela et al., 2006). Cell death was quantified as a measurement of membrane integrity and evaluated by the percentage of LDH released over total LDH. MTT reduction assay The MTT assay was first done to evaluate the cytotoxicity of MTX and NAPHT in differentiated H9c2 cells. This colorimetric assay relies mostly on the ability of mitochondrial complexes (or other dehydrogenases) to convert the soluble yellow tetrazolium dye, MTT, into an insoluble blue formazan product that can be measured at 550 nm (Costa et al., 2009, Soares et al., 2013). After the removal of culture medium, to each well was added 200 µL of fresh medium (DMEM 1% FBS and 10 nM RA) and 20 µL of the MTT solution (final 77 concentration of MTT 500 µg/mL). Subsequently, cells were incubated at 37ºC for 4h. The reaction was stopped by adding 200 µL of 10% SDS in 0.01 M hydrochloric acid solution followed by an overnight incubation at 37ºC, to allow solubilization of the formed formazans. Finally, the formed formazans were measured at 550 nm using a 96-well plate reader (Capela et al., 2006). The percentage of MTT reduction of control/vehicle cells was set to 100%, and the effects resulting from the incubation with MTX or metabolite were expressed as the percentage to the respective controls or vehicle. The MTT test was also done to evaluate the effects of cycloheximide, a glutarimide antibiotic (10 µg/mL) (Gupta et al., 2006), and metyrapone, CYP450 inhibitor (0.5 mM) (Rossato et al., 2013b), towards MTX cytotoxicity. Cycloheximide and metyrapone were added 1h before MTX. Neutral Red lysossomal uptake assay For this assay, H9c2 cells were seeded in 48-well plates. The amount of NR dye incorporated in the cells represents lysossomal functionality, as this dye easily penetrates viable cell membranes and accumulates intracellularly in lysosomes (Soares et al., 2013). At the end of the incubation time, the cellular medium was removed and 250 µL/well of NR solution (33 mg/mL of NR in DMEM 1% FBS) was added. Then, plates were incubated for 3h at 37ºC, protected from light. After this time, NR solution was removed and 200 µL/well of the solution formed by 50% ethanol/1% acetic acid was added to extract the NR dye meanwhile captured within the cells. The plates were then placed in a microplate shaker for 15 min, at room temperature and protected from light. Absorbance was measured at 540 nm, in a 48-well plate reader [Biotech Synergy HT (VT, USA)] and results were compared to control/ vehicle wells whose media of values was set to 100% (Soares et al., 2013). The NR test was done to access the effects of 3-methyladenine (2.5 mM), 1aminobenzotriazole (0.5 mM) and metyrapone (0.5 mM) following the exposure to MTX (and also 3-methyladenine in the case of NAPHT) in H9c2 differentiated cells. Those compounds were added 1h before MTX (or NAPHT). 3.8. Evaluation of mitochondrial potential The evaluation of the mitochondrial potential was done according to Freitas et al., with some modifications (Freitas et al., 2013). Briefly, 7-days differentiated cells were incubated for 12h with MTX or NAPHT and then cells were incubated for 30 min at 37ºC with DiO6 at the final concentration of 50 µM per well. In each condition and experiment, a well was placed with DiO6 and the mitochondrial uncoupling agent, carbonyl cyanide 3chloro-phenylhydrazone, at the final concentration 20 µM. Also, a well in each condition was 78 tested without any DiO6 as to evaluate if any component of the medium or if MTX or NAPHT conditions had any residual fluorescence that interfered with the fluorescence readings. After the 30 min incubation time, the cells were then washed 2 times with warmed PBS with Ca2+e Mg2+. Photographs were taken in a fluorescent microscope (Nikon Eclipse TS100 equipped with a Nikon DS-Fi1 camera) with the standard fluorescein filter (λexcitation = 485 nm and λemission = 520 nm). 3.9. Determination of cellular ATP levels For this assay, H9c2 cells were seeded in 6-well plates. The ATP was evaluated in differentiated H9c2 cells incubated for 24h with 2 and 5 μM of MTX, and 2 and 5 μM NAPHT. After the incubation period, medium was discarded and cells were washed with cold PBS with Ca2+e Mg2+ (750 μL/well). The PBS used for washing was discarded. Another 750 μL/well cold PBS with Ca2+e Mg2+ was added and cells were scrapped. Two wells per each condition were collected and centrifuged at 5 000 rpm, 4°C, for 10 min. The supernatant was rejected and to the pellet was added 200 μL of cold 5% perchloric acid. Then, the obtained samples were vortexed and centrifuged at 13 000 rpm, for 10 min, at 4°C. The acidic supernatant was collected for ATP determination and placed at -80°C. The pellet was used for protein determination and stored at -20°C. The ATP levels were determined by bioluminescence through the reaction with the firefly luciferin-luciferase system (Costa et al., 2007). D-Luciferin 90.9 mg/L stock reagent and luciferase from Photinus pyralis (firefly) (3 000 000 U/mL final concentration) were prepared in luciferin-luciferase buffer (50 mM glycine, 10 mM MgSO4, 1 mM Tris, 0.55 mM EDTA, 0.1% bovine serum albumin, pH = 7.6) and light protected aliquots were stored at - 20ºC until use. Before the assay, luciferin-luciferase reaction system was constituted by adding the two solutions and then kept at room temperature. Briefly, the assay consisted of neutralizing 150 μL of the acidic samples, standards or blank with 150 μL of 0.76 M of KHCO3, followed by vortex, and centrifugation for 10 min at 13 000 rpm (4ºC). One hundred μL of neutralized supernatants was added to a 96-well white microtiter plate, followed by addition of 100 μL/well of the luciferin-luciferase reagent. The reading was immediately done in the plate reader [Biotech Synergy HT (VT, USA)]. Light output was given as the integral relative light units. The assay was made sequentially with few measurements in each reading in order to avoid loss of the bioluminescence signal (Costa et al., 2007). The levels of ATP were expressed as ATP levels per amount of protein. 3.10. Protein determination 79 The protein content assay was determined using the protein assay kit Bio-Rad RC DC, according to the manufacturer’s instructions. Bovine serum albumin was used as protein standard and the calibration curve varied from 200 μg/mL to 1 200 μg/mL in 0.3 M NaOH. Five μL of samples, standards or blank was added in duplicated to a 96-well microtiter plate, followed by addition of 25 μL of Reagent A and 200 μL of Reagent B, light protected. The microtiter plate was kept protected from the light for 15 min, after which the absorbance was measured at 750 nm [BioTek Instruments, Powerwave X, (VT, USA)]. 3.11. Statistical analysis The data is presented as means ± standard deviation (SD) of different independent experiments. Statistical analysis was carried out by the non-parametric Anova (KruskalWallis test) followed by a Dunn’s post hoc test once a significant p was achieved. When the distribution was normal, one-way Anova was performed followed by Tukey’s post hoc test. Statistical significance was considered with a p values < 0.05. All statistical analysis were assessed using the GraphPad Prism 6 software program (San Diego, CA, USA). 80 81 4. RESULTS 82 83 4.1. Synthesis of the mitoxantrone metabolite naphthoquinoxaline The synthesis of the MTX metabolites was accomplished through the horseradish peroxidase (HRP)-catalysed H2O2 oxidation of MTX and the products isolated from this reaction are depicted in Figure 10. Figure 10. The isolated products of the incubation of MTX with HRP: NAPHT and NAPHTdi. Although with the enzymatic oxidation of MTX, several derivatives could be observed by chromatographic analysis, only two purified derivatives were isolated and identified as NAPHT, corresponding to 8, 11-dihydroxy-4-(2-hydroxyethyl)-6-[[2-[(2hydroxyethyl)-amino]ethyl]-amino]-1,2,3,4,7,12-hexahydronaphtho-[2,3-f]-quinoxaline-7,12 -dione, and NAPHTdi, corresponding to 3-((2-((4-(2-carboxyethyl)-8,11-dihydroxy-7,12dioxo-1,2,3,4,7,12-hexahydronaphtho-[2,3-f]-quinoxalin-6-yl)amino)ethyl)amino) propanoic acid, already proposed as a product of the peroxidase-catalyzed H2O2 oxidation of MTX by Bruck and colleagues (Bruck and Bruck, 2011). First attempts to purify the crude product using normal-phase chromatography were unsuccessful due to the retention of MTX derivatives in the stationary phase. Thus, procedures involving reverse-phase (for both) and/or ion exchange chromatography (for NAPHT) that have shown great versatility and capability for retaining and separating a variety of charged polar compounds, were used in the isolation of derivatives NAPHT and NAPHTdi (see Methods section). 4. Results 90 Figure 16. 1H NMR spectra of MTX. Figure 17. 1H NMR spectra of NAPHT. 91 Figure 18. 1H NMR spectra of NAPHTdi. The multiplicity and coupling constants of the protons observed in the 1H NMR spectrum of MTX showed the existence of two symmetrical 1, 4-disubstituted benzene rings, the signals of the protons of the hydroxyl and amine groups and of the aliphatic chain. In turn, the 1H NMR spectrum of NAPHT showed a different pattern of substitution in the aromatic ring while the proton signals of methylene groups, characteristic of the aliphatic chains, were maintained. The fusion of the piperazine ring with C-5/C-6 of the aromatic ring of the anthraquinone is evidenced by the lack of H-6 signal and the presence of a singlet corresponding to H-7 and of two aromatic protons, corresponding to H-2 and H3. The proposed structure was in agreement with the result obtained from the HRMS. The amount of NAPHTdi isolated allowed obtaining the 1H spectrum, which revealed a similar pattern of substitution in the aromatic ring when compared to NAPHT while signals of deshielded protons were observed at 4.1 and 3.1 ppm for the methylene groups characteristic of the aliphatic chains for the proposed structure of NAPHTdi. The proposed structure was in agreement with the result obtained from the HRMS. 4.3. Differentiation alters cell division and microscopic characteristics of H9c2 92 The differentiation protocol was conducted with 1% FBS and RA for 7 days. The differentiation by 10 nM RA and 1% FBS by 7 days led to morphological changes making cells more resembled to the cardiac phenotype. There were substantial differences between undifferentiated cells and differentiated cells, which are illustrated in Figure 19. At panel 19B and D, two detailed Figures of both differentiated and undifferentiated H9c2 cells cultivated for 7 days in multi-well plates are presented. The differentiation protocol dramatically reduced cell division. In fact, cellular density was substantially higher in undifferentiated cells cultivated for 7 days with 10% FBS medium without RA. Also, while inhibition of cell division occurred, the cell bodies of differentiated cells became smaller and fusiform when compared to undifferentiated cells, which exhibit a large and flat cell body. Overall, differentiated cells showed to established a more organized network bearing a more cardiac-like morphology, as shown by phase-contrast morphology. The differentiation also resulted in a lower proliferation rate as verified by Hoechst 33258 staining. Figure 19. Fluorescence microscopy (Hoechst 33258 staining, A and C) and phase contrast microscopy images (B and D) of undifferentiated (A and B) and differentiated (C and D) H9c2 cells for 7 days. Images are representative of three independent experiments (scale bar 100 μm). 4.4. Toxicological evaluation of mitoxantrone Microscopic evaluation of mitoxantrone-incubated H9c2 cells Cells were morphologically assessed after 24h exposure to MTX. H9c2 cells were incubated with MTX (2 and 5 μM), as described in the Methods section. Phase-contrast microscopy and Hoescht staining were used. Phase-contrast microscopy, after 24h 93 incubation, revealed cell death, with intensifying features of cell death when increasing concentrations (Figure 20). In particular, H9c2 cell exposed with 5 μM of MTX showed signs of loss of membrane integrity and decrease in cell number (Figure 20C and 20F). No signs of nuclear alteration were found by Hoescht staining. At 48h, the cell injury was more increased in MTX-incubated cells (data not shown). Figure 20. Fluorescence microscopy (Hoechst 33258 staining) (A, B, C), phase contrast microscopy (D, E, F) images of differentiated H9c2 cells after incubation with 2 and 5 μM of MTX for 24h. Control (A and D), MTX 2 μM (B and E), MTX 5 μM (C and F). Images are representative of three independent experiments (scale bar 100 μm). Ethidium bromide/ acridine orange staining was also used for evaluate MTX effects on H9c2 cells. Through this method, living cells appear with a regular-sized green fluorescent nucleus, as shown in control H9c2 in Figure 21A. The number of cells largely decreased with increased MTX concentration, but cells present in the field remained with green nucleus, although with cytoplasmatic injury (Figure 21C). 94 Figure 21. Ethidium bromide and acridine orange (A, B, C) images of differentiated H9c2 cells in control and in cells incubated with 2 and 5 μM of MTX for 24h. Control (A), MTX 2 μM (B), MTX 5 μM (C). Images are representative of three independent experiments (scale bar 100 μm). Assessment of mitochondrial membrane potential in differentiated H9c2 cells incubated with 2 μM of MTX (B) and 5 μM of MTX (C) for 12h (Figure 22) was done. Lower levels of fluorescence resulting from MTX treatment were observed, suggesting the depolarization of mitochondrial membrane potential at 12h. This depolarization effect seems dose dependent (Figure 22C). Figure 22. Mitochondrial transmembrane potential (A, B, C) images of differentiated H9c2 cells after incubation with 2 and 5 μM of MTX for 12h. Control (A), MTX 2 μM (B), MTX 5 μM (C). Images are representative of three independent experiments (scale bar 100 μm). Mitoxantrone led to a timeand concentration-dependent mitochondrial dysfunction The ability of MTX to interfere with MTT reduction was evaluated at 2 time-points (24 and 48h) and with several concentrations (0.01 to 5 µM). At 24h (Figure 23A), MTX at concentration 0.1 µM already caused significant cytotoxicity when compared to control cells, being the highest dose tested (5 µM) able to decrease the levels of MTT reduction to 85.99 ± 8.52%. At 48h, as can be seen in Figure 23B, the cytotoxicity elicited was even higher, when compared to control and the same concentrations in the earlier time point. 95 Figure 23. Mitochondrial dysfunction evaluated by MTT reduction assay in differentiated H9c2 cells incubated with 0.01, 0.1, 1, 2 and 5 μM of MTX for 24 (A) and 48h (B). Results are presented as mean ± SD of 5 independent experiments (total of 30 wells). Statistical analysis were performed using Kruskal–Wallis test, followed by the Dunn’s post hoc test (*p < 0.05, ***p < 0.001, ****p < 0.0001 vs. control; ##p < 0.01, ####p < 0.0001 vs. 0.01 µM; $p < 0.05, $$$$p < 0.0001 vs. 0.1 µM; +p < 0.05 vs. 2 µM). Mitoxantrone was only able to cause significant loss of membrane integrity at the highest concentrations tested The ability of MTX to cause LDH leakage to the extracellular medium was tested at 2 time-points (24 and 48h) and with several concentrations (0.01 to 5 µM). At 24h (Figure 24A), only the higher concentrations of MTX (1, 2 and 5 µM) caused significant cellular membrane integrity loss when compared to control cells. In Figure 24B, the data obtained after the 48h incubation of H9c2 differentiated cells is observed. MTX, at 48h, caused a significant decrease in cellular toxicity, being observed that at 1 µM MTX the viability was 64.08 ± 8.26%, 2 µM MTX the viability was 43.47 ± 7.49% and at 5 µM the viability was 27.98 ± 9.92%. 96 Figure 24. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 0.01, 0.1, 1, 2 and 5 μM of MTX for 24 (A) and 48h (B). Results are presented as mean ± SD of 5 independent experiments (total of 30 wells). Statistical analysis was done using the Kruskal–Wallis test, followed by the Dunn’s post hoc test (****p < 0.0001 vs. control; ####p < 0.0001 vs. 0.01 µM; $$p < 0.01, $$$$p < 0.0001 vs. 0.1 µM; &&p < 0.01, &&&p < 0.001 vs. 1 µM). Mitoxantrone caused a significant lysosome uptake dysfunction in H9c2 differentiated cells After 24 and 48h incubation with several concentrations of MTX (0.01 to 5 µM), the lysossomal uptake of NR was evaluated. At 24h (Figure 25A), the highest concentrations of MTX (1, 2 and 5 µM) caused a substantial impairment of lysossomal uptake of NR when compared to control cells. At 48h (Figure 25B), lysossomal functionality was even more impaired in MTX highest concentrations (1, 2 and 5 µM). At 48h, the H9c2 cells incubated MTX 1 µM had 58.98 ± 15.41%, 2 µM had 37.81 ± 7.34 %, and 5 µM 24.64 ± 4.07% NR uptake ability when compared to control cells (Figure 25B). Figure 25. NR uptake (% of control) in differentiated H9c2 cells incubated with 0.01, 0.1, 1, 2 and 5 μM of MTX for 24 (A) and 48h (B). Results are presented as mean ± SD of 5 independent experiments (total of 30 wells). Statistical analysis was done using the Kruskal–Wallis test, followed by the Dunn’s post hoc test (****p < 0.0001 vs. control; ##p < 0.01, ####p < 0.0001 vs. 0.01 µM; $p < 0.05, $$p < 0.01, $$$$p < 0.0001 vs. 0.1 µM; &&&p < 0.001, &&&&p < 0.0001 vs. 1 µM). Buthionine sulphoximine, an inhibitor of gammaglutamylcysteine synthetase, had no effect on cellular death caused by mitoxantrone incubation in H9c2 cells Buthionine sulphoximine, an inhibitor of gamma-glutamylcysteine synthetase, at 50 µM concentration, was pre-incubated in H9c2 cells, to access if the decrease of glutathione 97 synthesis caused any effect on the cytotoxicity caused by MTX. Buthionine sulphoximine did not cause any significant alteration on the cell death caused by the 48h incubation of 2 µM MTX (Figure 26). At 48h, 2 µM MTX had 53.95 ± 5.26% viability, whereas the condition 2 µM MTX plus buthionine sulphoximine had a viability of 50.96 ± 8.30% (Figure 26). The use of buthionine sulphoximine at 50 µM did not cause any significant change in cellular viability when compared to control cells. Figure 26. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 2 μM of MTX and buthionine sulphoximine (BSO), an inhibitor of gamma-glutamylcysteine synthetase at 50 µM and BSO + MTX, for 48h. Results are presented as mean ± SD of 4 independent experiments (total of 20 wells). Statistical analysis was done using the Kruskal–Wallis test, followed by the Dunn’s post hoc test (****p < 0.0001 vs. control; $$$$p < 0.0001 vs. BSO 50 µM). N-acetyl cysteine, a glutathione precursor and reactive species scavenger, was not able to revert the cell death caused by mitoxantrone incubation N-acetyl cysteine, a glutathione precursor and reactive species scavenger (Avantaggiato et al., 2014), at 1 mM concentration, was used in H9c2 cells. The antioxidant, N-acetyl cysteine, did not cause any significant alteration on the cell death caused by the 48h incubation of 2 µM MTX (Figure 27). At 48h, 2 µM MTX had 52.20 ± 4.38% viability, whereas the condition 2 µM MTX plus N-acetyl cysteine had a viability of 47.47 ± 6.82% (Figure 27). The use of N-acetyl cysteine at 1 mM did not cause any significant change in cellular viability when compared to control cells. 98 Figure 27. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 2 μM of MTX and N-acetyl cysteine (NAC), an antioxidant and reactive species scavenger at 1 mM and NAC + MTX for 48h. Results are presented as mean ± SD of 4 independent experiments (total of 24 wells). Statistical analysis was done using the Kruskal–Wallis test, followed by the Dunn’s post hoc test (****p < 0.0001 vs. control; $$$$p < 0.0001 vs. NAC 1 mM). L-carnitine, a mitochondrial enhancer was not able to revert cell death caused by mitoxantrone incubation L-carnitine acts as a carrier for fatty acids across the inner mitochondrial membrane for subsequent β-oxidation (Zammit et al., 2009). L-carnitine did not cause any significant alteration on the cell death caused by the 48h incubation with 2 µM MTX (Figure 28). At 48h, 2 µM MTX had 53.29 ± 5.67% viability, whereas the condition 2 µM MTX plus Lcarnitine had a viability of 52.83 ± 5.56% (Figure 28). The use of L-carnitine at 2 mg/mL did not cause any significant change in cellular viability when compared to control cells. 99 Figure 28. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 2 μM of MTX, and L-Carnitine (L-Carn), a mitochondrial enhancer at 2 mg/mL and L-Carn + MTX for 48h. Results are presented as mean ± SD of 4 independent experiments (total of 24 wells). Statistical analysis was done by using the Kruskal–Wallis test, followed by the Dunn’s post hoc test (****p < 0.0001 vs. control; $$$$p < 0.0001 vs. L-Carnitine 2 mg/mL. 3-Methyladenine, an autophagy inhibitor, did not inhibit cell death caused by incubation with mitoxantrone 3-Methyladenine was used to assess if autophagy was involved on MTX cytotoxicity. 3-Methyladenine did not inhibit the cell death caused by the 48h incubation of 2 µM MTX (Figure 29). At 48h, 2 µM MTX had 49.40 ± 4.00% viability, whereas the condition 2 µM MTX plus 3-methyladenine had a viability of 48.30 ± 2.92% (Figure 29). The use of 3methyladenine at 2.5 mM did not cause any significant change in cellular viability when compared to vehicle cells. Figure 29. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 2 μM of MTX, 3-methyladenine (3-MA) 2.5 mM and 3-MA + MTX for 48h. DMSO (final concentration of 0.1% v/v) was used as vehicle. Results are mean ± SD of 4 independent experiments (24 wells). Statistical analysis: Kruskal–Wallis test, followed by the Student–Newman–Keuls post hoc test (****p < 0.0001 vs. vehicle; $$$$p < 0.0001 vs. 3-MA 2.5 mM). The 3-methyladenine, an autophagy inhibitor, led to a partial protection to the impairment caused in lysossomal uptake by mitoxantrone 106 Assessment of mitochondrial membrane potential in differentiated H9c2 cells incubated with 2 μM of NAPHT (B) and 5 μM of NAPHT (C) for 12h (Figure 38) was also performed. Lower levels of fluorescence resulting from NAPHT treatment were observed, suggesting that the depolarization of mitochondrial membrane potential at 12h occurred after incubation with NAPHT. This depolarization effect seems to be dose dependent (Figure 38). Figure 38. Images of mitochondrial transmembrane potential of differentiated H9c2 cells after incubation with 2 and 5 μM of NAPHT for 12h (A, B, C). DMSO (final concentration of 0.1% v/v) was used as vehicle (A), NAPHT 2 μM (B), NAPHT 5 μM (C). Images are representative of three independent experiments (scale bar 100 μm). Naphthoquinoxaline caused a timeand concentrationdependent mitochondrial dysfunction The ability of NAPHT to interfere with MTT reduction was evaluated at 2 time-points (24 and 48h) and with several concentrations (1 to 5 µM). At 24h (Figure 39A), NAPHT at concentration 2 and 5 µM caused cytotoxicity when compared to control cells. At 48h, (Figure 39A) NAPHT at all the concentrations tested caused significant cytotoxicity when compared to vehicle cells, being the highest concentration tested (5 µM) able to decrease the levels of MTT reduction to 82.49 ± 4.05% when compared to vehicle cells (100 ± 3.85%). 107 Figure 39. Mitochondrial dysfunction evaluated by MTT reduction in differentiated H9c2 cells incubated with 1, 2 and 5 μM of NAPHT for 24 (A) and 48h (B). DMSO (final concentration of 0.1% v/v) was used as vehicle. Results are presented as mean ± SD of 6 independent experiments (total of 35 - 36 wells). Statistical analysis were performed using the Anova test, followed by the Tukey’s post hoc test (*p < 0.05, **p < 0.01, ***p < 0.001 vs. vehicle; ##p < 0.01, ####p < 0.0001 vs. 1 µM; $$$$p < 0.0001 vs. 2 µM). Naphthoquinoxaline was able to cause significant loss of membrane integrity The ability of NAPHT to cause cell death was tested at 2 time-points (24 and 48h) and with several concentrations (1 to 5 µM). At 24h (Figure 40A), all the concentrations of NAPHT (1, 2 and 5 µM) tested caused significant cellular membrane integrity loss when compared to control cells. In Figure 40B, the data obtained after the 48h incubation of NAPHT in H9c2 differentiated cells is observed. NAPHT at 48h caused a significant decrease in cellular integrity, being observed that at 2 µM NAPHT the viability was 73.02 ± 2.86% and at 5 µM the viability was 52.07 ± 6.83%, when compared with vehicle cells (81.91 ± 1.89%). Figure 40. Cellular viability evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 1, 2 and 5 μM of NAPHT for 24 (A) and 48h (B). DMSO (final concentration of 0.1% v/v) was used as vehicle. Results are presented as mean ± SD of 6 independent experiments (total of 35 - 36 wells). Statistical analysis was performed at 24h using the Kruskal–Wallis test, followed by the Student–Newman–Keuls post hoc test. For 48h, the Anova test, followed by the Tukey’s post hoc test, was used (***p < 0.001, ****p < 0.0001 vs. vehicle; ###p < 0.001, ####p < 0.0001 vs. 1 µM; $$p < 0.01, $$$$p < 0.0001 vs. 2 µM). 108 Naphthoquinoxaline caused a significant lysosome uptake dysfunction in H9c2 differentiated cells After 24 and 48h incubation with several concentrations of NAPHT (1 to 5 µM), the lysossomal uptake of NR was evaluated. At 24h (Figure 41A), only the highest concentrations of NAPHT (2 and 5 µM) caused a substantial impairment of lysossomal uptake of NR when compared to control cells. At 48h (Figure 41B) the H9c2 cells incubated with NAPHT 2 µM had 88.82 ± 5.53% of NR uptake, while 5 µM showed values of 68.83 ± 6.28% NR incorporation when compared to vehicle cells (100.00 ± 6.18%). Figure 41. NR uptake (% of vehicle) in differentiated H9c2 cells incubated with 1, 2 and 5 μM of NAPHT for 24 (A) and 48h (B). DMSO (final concentration of 0.1% v/v) was used as vehicle. Results are presented as mean ± SD of 5 independent experiments (total of 30 wells). Statistical analysis was performed using the Anova test, followed by the Tukey’s post hoc test for analysis at 24h and Kruskal–Wallis test, followed by the Student–Newman–Keuls post hoc test, for analysis at 48h (**p < 0.01, ****p < 0.0001 vs. vehicle; #p < 0.05, ####p < 0.0001 vs. 1 µM; $$p < 0.01, $$$$p < 0.0001 vs. 2 µM). N-acetyl cysteine incubation increased cell death caused by naphthoquinoxaline N-acetyl cysteine, a glutathione precursor and reactive species scavenger (Avantaggiato et al., 2014), at 1 mM concentration was used in H9c2 cells. N-acetyl cysteine increased cell death caused by 2 µM NAPHT (Figure 42). At 48h, 2 µM NAPHT had 74.09 ± 2.50% viability, whereas in the condition 2 µM NAPHT plus N-acetyl cysteine the viability observed was 68.16 ± 3.26% (Figure 42). The use of N-acetyl cysteine at 1 mM did not cause significant change in cellular viability when compared to vehicle cells. 109 Figure 42. Cellular viability at 48h evaluated using the LDH leakage assay (% of extracellular LDH/total LDH) in differentiated H9c2 cells incubated with 2 μM of NAPHT and N-acetyl cysteine (NAC), an antioxidant, at 1 mM. DMSO (final concentration of 0.1% v/v) was used as vehicle. Results are presented as mean ± SD of 4 independent experiments (total of 24 wells). Statistical analysis was performed using the Anova test, followed by the Tukey’s post hoc test (****p < 0.0001 vs. vehicle; $$$$p < 0.0001 vs. NAC; &&&&p < 0.0001 vs. 2 µM NAPHT). 3-Methyladenine, an autophagy inhibitor, caused a partial protection to the damage caused by naphthoquinoxaline After 48h incubation with 2 µM NAPHT, the lysossomal uptake of NR was evaluated. At 48h, 2 µM NAPHT showed values of 88.26 ± 6.93%, whereas the condition 2 µM NAPHT plus 3-methyladenine had the values of 94.24 ± 7.91% (Figure 43). Thus, 3-methyladenine caused a partial protection to the effects caused by NAPHT. Figure 43. NR uptake (% of vehicle) in differentiated H9c2 cells incubated with 2 μM of NAPHT, 3-methyladenine (3-MA) 2.5 mM and 3-MA + NAPHT for 48h. DMSO (final concentration of 0.1% v/v) was used as vehicle. Results are mean ± SD of 5 independent experiments (30 wells). Statistical analysis was performed using the Anova test, followed by the Tukey’s post hoc test (**p < 110 0.01, ****p < 0.0001 vs. vehicle; $$$p < 0.001, $$$$p < 0.0001 vs. 3-MA 2.5 mM; &&p < 0.01 vs. 2 µM NAPHT). Naphthoquinoxaline did not cause any significant alteration on ATP levels at an early time-point To understand if NAPHT has effects in cellular energetics, intracellular ATP levels were measured in H9c2 cell exposed to NAPHT for 24h. In the NAPHT tested concentration, and time point (24h), no significance changes were detected, as shown in the Figure 44. After the 24h incubation with NAPHT (2 and 5 μM), the ATP intracellular levels were about 14.94 ± 4.38 nmol/mg protein and 16.74 ± 4.21 nmol/mg protein, respectively, compared to 14.24 ± 3.48 nmol/mg protein for the vehicle group. Figure 44. ATP levels in differentiated H9c2 cells incubated with 2 and 5 μM NAPHT for 24h. DMSO (final concentration of 0.1% v/v) was used as vehicle. Results, in nmol/mg protein are presented as mean ± SD of 6 independent experiments. 111 5. DISCUSSION AND CONCLUSIONS 112 113 Xenobiotics that cause heart damage may have a part of their toxicity resulting from metabolization. This chemical change is mainly catalysed by enzymes, and usually favours the increase of hydrophilicity of parental xenobiotics. The majority of the metabolism in the body occurs in the liver, and the metabolites formed can reach the heart due to the distribution via blood stream. Moreover, the heart also has metabolizing enzymes, therefore in loco it generates metabolites that may cause cardiac toxicity (Wormhoudt et al., 1999). Several anticancer drugs can lead to cardiotoxicity. Some drugs, like DOX and cyclophosphamide, are shown to be cardiotoxic through their metabolites (Costa et al., 2013). Other anticancer drugs are scarcely studied regarding the cardiotoxicity of their metabolites. 5.1. Synthesis and purification of the mitoxantrone metabolite naphthoquinoxaline The full characterization of the metabolic profile and the elucidation of the metabolites structure in new chemical entities are often not completed during clinical development or even after the drugs’ introduction into the market (Fura et al., 2004). It is also often difficult to predict a priori whether after metabolization a certain compound could form a toxic or pharmacologically active metabolite. The degree of contribution of active metabolites to the overall observed therapeutic or toxic effects needs further study, but the rapid chemical synthesis of metabolites by traditional methods is often challenging (Fura et al., 2004). In this thesis, the synthesis of MTX metabolites was accomplished by the HRP catalysed H2O2 oxidation (Figure 10). Several derivatives were observed by chromatographic analysis of the crude product and two purified derivatives were isolated and identified as NAPHT, an already reported metabolite of MTX, and a newly isolated derivative, NAPHTdi. The enzymatically catalysed oxidation of MTX by HRP/H2O2 has already been demonstrated in other works (Reszka et al., 1986, Kolodziejczyk et al., 1988, Bruck and Bruck, 2011). In those works, the isolation of the metabolite NAPTH was achieved and it was partially characterized by 13C NMR and liquid chromatography-mass spectrometry. Herein, the isolation and structure characterization of NAPHT was accomplished and additionally the isolation of a possible oxidation product of MTX, NAPHTdi, was achieved for the first time. 5. Discussion and conclusions 114 In the study by Blanz et al., NAPHT metabolite was isolated by preparative HPLC from the urine of a patient and it was characterized by tandem mass spectrometry and UVvisible spectroscopy (Blanz et al., 1991b). Moreover, two studies in cell systems suggest that NAPHT may contribute to the cytotoxic activity of MTX (Duthie and Grant, 1989, Mewes et al., 1993). The proposed mechanism for the formation of NAPHT and NAPHTdi (seen in Figure 45) (Blanz et al., 1991b) established the reaction to proceed via a two-electron oxidation of the phenylenediamine substructure I to the formation of the highly reactive quinone II. The nonprotonated form IV of the equilibrium reacts by an intramolecular attack of the basic amino group of the side chain with the electrophilic centre at C-6 of MTX radical cation. Subsequent oxidation of the cyclized radical V led to the formation of NAPHT. As the reaction progressed, further oxidation of the NAPHT side-chains occurred, yielding the dicarboxylic acid derivative NAPHTdi. Figure 45. Proposed mechanism for the formation of NAPHT and NAPHTdi after the oxidation of MTX. Adapted from Blanz et al., 1991b. 115 The NAPHT and other metabolites resulting from oxidation of MTX are not normally purified after synthesis. For example, in the study of Bruck et al., the crude product of the HRP/H2O2 catalysed oxidation was used for the steady-state spectrokinetic and HPLC-MS analysis without further purification (Bruck and Bruck, 2011). Herein, with the purification methods used allowed to obtain a high purity NAPHT metabolite that allowed further toxicological evaluation. Nevertheless, the amount of NAPHTdi obtained did not allow further in vitro studies and the unequivocal structure characterization. 13C NMR analysis should be considered in a future work for its full chemical characterization. This compound has already been previously proposed by UV analysis to be formed with the HRP catalysed H2O2 oxidation (Bruck and Harvey, 2003). In this thesis, NAPHTdi was first time isolated and characterized. 5.2. Decreased foetal bovine serum and the addition of retinoic acid to H9c2 cells caused cell differentiation The H9c2 myoblast cell line, originated from rat ventricular tissue, has been used in vitro as a model for skeletal and cardiac muscle due to the morphological features and electrical/hormonal signalling properties of these cells (Kimes and Brandt, 1976, Hescheler et al., 1991). H9c2 cells show electrophysiological and biochemical properties of both skeletal and cardiac tissues, including depolarization in response to acetylcholine (Kimes and Brandt, 1976), and rapid activation of calcium currents through L-type channels (Hescheler et al., 1991, Mejia-Alvarez et al., 1994, Wang et al., 1999). An interesting feature of this cell line is its ability to differentiate from mono-nucleated myoblasts to myotubes upon reduction of serum concentration (Hescheler et al., 1991). With myotube formation the expression of myogenic transcription factors (Chun et al., 2000) and calcium channel proteins also occurs (Menard et al., 1999). Despite the wide use of the H9c2 cell line in the undifferentiated state, a cardiac-like phenotype can be obtained after differentiation in a low FBS (1%) and 10 nM RA - supplemented media. Because the adult heart tissue is composed mainly by differentiated cardiomyocytes, toxicological studies can have different outcomes depending on the developmental state of the biological model used (Branco et al., 2011). In the study by Menard et al., the cells were cultured in DMEM supplemented with 1% foetal calf serum (replaced every 2 days) and RA (Menard et al., 1999). RA-treated cells exhibited morphological changes with rather large and rounded cells. The treatment of H9c2 cells with RA maintained their cardiac phenotype and inhibited myogenic differentiation of those 122 time (Figure 30). The 3-methyladenine led to a partial protection in the NR test (Figure 30), which was not observed in the LDH assay (Figure 29), thus demonstrating that other mechanisms may be involved in MTX-membrane damage. 5.6. Evaluation of ATP levels and mitochondrial transmembrane potential of mitoxantrone To understand if MTX has any effects on cellular energetics, intracellular ATP levels were assessed through the bioluminescence test. In our study, levels of ATP increased with incubation of MTX at 24h in differentiated H9c2 cells (Figure 35). In other models results are different. Recently, Rossato et al. showed that MTX (100 nM and 1 μM) caused an important energetic imbalance evidenced by decreased ATP levels and hyperpolarization of the mitochondrial membrane potential (as referred in 5.5) after 24, 48, and 96h of MTX incubation in undifferentiated H9c2 cells (Rossato et al., 2013b). In the study of Shipp et al., they determined the sensitivity of cultured rat heart cells to MTX treatment. Those cells were treated with several drug concentrations (0.05 to 5 μg/mL) for 3h and then medium was replaced. ATP levels were measured 72h after initial exposure and a concentrationdependent decline in ATP levels was observed. ATP suppression was maximal at 72h for all MTX concentrations tested; at that time point, there was also a substantial loss of spontaneous synchronous contraction in the cultures (Shipp et al., 1993). In vivo, male Wistar rats treated with 3 cycles of 2.5 mg/kg MTX at day 0, 10, and 20 and euthanized on day 48, showed ATP decreases. That ATP decrease was shown to be related to mitochondrial disruption (Rossato et al., 2014). Therefore, ATP levels seem to be affected by MTX. In our study with MTX an increase in the levels of ATP was reported, which seem to be an adaptive response to MTX-incubation. Longer incubation time should be tested in our model to assess if it is an adaptive response. 5.7. The role of the cytochrome metabolism on mitoxantrone So far the following enzymes were detected in the heart: CYP450, NADPH cytochrome reductase, dihydropyrimidine dehydrogenase, epoxide hydrolase, DTdiaphorase, glutathione S-transferase and UDP-glucuronosyltransferases, among others. CYP450 is a superfamily of enzymes that are found in all living organisms. They are mixed function mono-oxygenases, which are involved in the oxidative metabolism of a wide range of xenobiotics and endogenous compounds (Elbekai and El-Kadi, 2006). Several CYP450 have been identified in human (Delozier et al., 2007) and animal heart tissues 123 (Imaoka et al., 2005) as well as different in vitro models such as primary rat cardiomyocytes (Thum and Borlak, 2000), immortalized H9c2 cells (Zordoky and El-Kadi, 2007), and recently in the atrial HL-1 cell line (Elshenawy et al., 2013). Cardiac expression of CYP450 subfamilies identified in mammals includes: CYP1A, CYP1B, CYP2A, CYP2B, CYP2D, CYP2E, CYP2J, CYP2R, CYP2S, CYP2U, CYP4A, CYP4B, CYP4F and CYP11B (Chaudhary et al., 2009). Herein, CYP450 inhibitors, namely metyrapone and 1-aminobenzotriazole, were also used to evaluate the role of the CYP450 metabolism on MTX cytotoxicity, since the oxidoreductive metabolism of MTX has a significant role on its antitumor effects and recent data suggest that MTX metabolism can be related to its cardiotoxicity (Blanz et al., 1991b, Bruck and Bruck, 2011, Rossato et al., 2013a). MTX has particular effectiveness in tumours with high contents of peroxidases (Blanz et al., 1991b, Bruck and Bruck, 2011). Duthie et al. showed that metyrapone is able to prevent the cytotoxic effect of MTX in human liver derived HepG2 hepatoma cells, thereby giving further evidence for the participation of the CYP450-dependent mixed function oxidase systems on MTX toxicity as referred above (Duthie and Grant, 1989). Similar results (metyrapone prevent the cytotoxic effect of MTX) were obtained with a rat hepatocytes model (Mewes et al., 1993) and with human breast cancer cells (Li et al., 1995) incubated with MTX and CYP450 inhibitors. In the study by Rossato et al. the influence of CYP450and CYP2E1-mediated metabolism for the cytotoxicity of MTX was assessed after 96h incubation with MTX (100 nM and 1 μM) in H9c2 undifferentiated cells. The co-incubation of MTX with CYP450 and CYP2E1 inhibitors partially prevented the cytotoxicity caused by the drug, thus highlighting that the metabolism of MTX is relevant for its undesirable effects in that model. They proved that the bioactivation of MTX mediated through CYP450 and namely through CYP2E1 metabolism occurs in loco in the cardiomyoblasts and exerts a significant role in the cellular damage promoted by MTX (Rossato et al., 2013a). Moreover, they demonstrated that the cytotoxicity caused by MTX after S9 hepatic fractions metabolization was significantly higher than that observed in the H9c2 cells incubated with MTX without S9 metabolization (Rossato et al., 2013a). In this thesis, metyrapone and 1-aminobenzotriazole did not prevent of the MTX cytotoxicity observed in differentiated H9c2 cells at 48h. The two inhibitors used have different mechanisms. 1-Aminobenzotriazole is suicide nonspecific inhibitor. It is a metabolism-based inactivator of CYP450 by the mechanism of N-alkylation of heme moiety. 1-Aminobenzotriazole has been implicated in the inhibition of CYP1A, 2A, 2B, 2C, 2E, 3A, 4A, and 4B in various organs of different species (Linder et al., 2009). Metyrapone is a competitive inhibitor of CYP450-mediated ω/ω-1 hydroxylase activity (Asakura and Shichi, 124 1992), and it is mainly an inhibitor of CYP11B1 (11-β-hydroxylase), but it also inhibits CYP11B2 (Sampath-Kumar et al., 1997) and CYP3A4 (Park et al., 2005). Rossato et al. described protection with CYP450 inhibitors at 96h in MTT assay (Rossato et al., 2013a), whereas in our work the time point was 48h. Thus, longer time point may be required for MTX metabolization to form significant amounts of toxic metabolites. On the other hand, to the best our knowledge, there are no studies regarding enzyme expression in differentiated H9c2 cells and the CYP450 expression can be altered by cellular differentiation. In undifferentiated H9c2 cells, there is only few knowledge about CYP isoenzymes (Zordoky and El-Kadi, 2007). In the study by Zordoky and El-Kadi, they showed that CYP1A1 and 1B1, CYP2B1, CYP2B2, CYP2E1, CYP2J3, CYP2C11, 2C13 and 2C23 were expressed in H9c2 cells. They concluded that multiple CYP genes are expressed in H9c2 cells at comparable level to those expressed in the heart or cardiomyocytes (Zordoky and El-Kadi, 2007). However, at this point no study has been made to clarify if differentiation alters CYP450 expression, and therefore different metabolites can be formed. 5.8. Incubation naphthoquinoxaline in differentiated H9c2 cells In this thesis, MTT (Figure 39), LDH (Figure 40), and NR (Figure 41) assays were done at 24h and 48h and with different NAPHT concentrations. For NAPHT incubation (1, 2, and 5 µM) cytotoxicity was timeand concentration-dependent in all assays, being LDH the most sensitive test. At 48h, all the concentrations tested caused significant cytotoxicity when compared to vehicle cells, being the highest concentration tested (5 µM) able to decrease the levels of MTT reduction to values that reached 82.49 ± 4.05%. All the concentrations of NAPHT of 1, 2, and 5 µM tested for 24h, caused significant cellular membrane integrity loss when compared to vehicle cells. NAPHT at 48h caused a significant decrease in cellular integrity, being observed that at 2 µM NAPHT the viability was 73.02 ± 2.86% and at 5 µM the viability was 52.07 ± 6.83%, when compared with vehicle cells (81.91 ± 1.89%). NAPHT at 2 and 5 µM caused a substantial impairment of lysossomal uptake of NR when compared to vehicle cells. At 48h, in the NR assay, the H9c2 cells incubated with NAPHT 2 µM (88.82 ± 5.53%) and 5 µM (68.83 ± 6.28%) showed a significant lysosome uptake dysfunction when compared to vehicle cells (100.00 ± 6.18%). Two pharmacological active molecules were tested to elucidate the mechanisms involved in NAPHT-induced cardiotoxicity, namely 3-methyladenine (an autophagy inhibitor) and N-acetyl cysteine (a glutathione precursor and reactive species scavenger). 3-Methyladenine caused a partial protection to the damage caused in lysossomal uptake 125 by NAPHT (Figure 43). NAPHT caused cardiotoxicity in micromolar concentration being autophagy involved. On the other hand, N-acetyl cysteine increased cell death caused by 2 µM NAPHT (Figure 42). This result may suggest a preconditioning stage that early reactive species formation can elicit and that N-acetyl cysteine may avoid (Ferreira et al., 2013). This mechanism needs further investigation. Phase contrast microscopy and several staining’s (Figure 36-38) were used to evaluate cytotoxicity of NAPHT. Phase microscopy allowed to observe that NAPHT caused cell damage at 24 or 48h incubation. Assessment of mitochondrial membrane potential in differentiated H9c2 cells incubated with 2 and 5 μM NAPHT for 12h showed significant effects: there was an evident decrease in mitochondrial potential seen in fluorescent microscopy (Figure 38). To understand if NAPHT had effects in cellular energetics, intracellular ATP levels were measured in H9c2 cell after a 24h incubation. In the NAPHT tested concentrations, no significance changes were detected (Figure 44). The work by Shipp et al. demonstrated that the metabolite NAPHT had a higher potential on ATP depletion than MTX in neonatal rat heart cells and, importantly, that the bioenergetics impairment hugely depends on iron availability (Shipp et al., 1993). The work by Shipp a co-workers is the only work available so far regarding to NAPHT cytotoxicity and used a different model and methodology than the work herein performed. 5.9. Toxicological comparison between mitoxantrone and naphthoquinoxaline For the highest concentrations of MTX (1, 2, and 5 µM) cytotoxicity was timeand concentration-dependent in all assays, being the NR assay more sensitive for the MTX toxic effects in H9c2 differentiated cells. On the other hand, for NAPHT (1, 2, and 5 µM) cytotoxicity was timeand concentration-dependent in all assays, being LDH the most sensitive test. MTX and NAPHT caused a timeand concentration-dependent cytotoxicity, although with different sensitivity towards the assays used. 3-Methyladenine caused a partial protection to the damage caused in lysossomal uptake by both NAPHT and MTX, suggesting that for both molecules autophagy has an important role. N-acetyl cysteine increased cell death caused by 2 µM NAPHT, whereas N-acetyl cysteine did not cause any alteration in the cytotoxicity observed after MTX incubation. Both molecules caused a decreased of mitochondrial membrane potential in differentiated H9c2 cells incubated in similar conditions (time and concentration). 126 Regarding ATP levels, drugs had different effects, since NAPHT had no effects and MTX caused a significant increase in MTX incubated cells, suggesting an adaptive response towards MTX in H9c2 cells. MTX and NAPHT share several mechanisms, but differ in other aspects that should be further investigated, mainly because the data on NAPHT are very scarce. 5.10. Final conclusions Based on the results presented in this study, we propose the following main conclusions:  With the enzymatic oxidation of MTX, several derivatives could be observed by HPLC and two purified derivatives were isolated and identified as NAPHT and NAPHTdi.  MTX and NAPHT caused a timeand concentration-dependent cytotoxicity, although with different sensitivity towards the assays used.  MTX and NAPHT caused cardiotoxicity at micromolar concentration being autophagy involved.  MTX and NAPHT caused a decreased of mitochondrial membrane potential in differentiated H9c2 cells incubated in similar conditions.  ATP levels were affected in a different manner in MTXand NAPHT-incubated H9c2 cells.  Thus, both drugs seem to impair cellular pathways in a dissimilar manner, although both cause early mitochondrial depolarization. 127 6. 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