Antioxidant and cytotoxic activity of portuguese propolis
Abstract
O própolis é uma mistura resinosa e balsâmica natural produzida por abelhas rica em flavonóides e compostos fenólicos, que são geralmente associados às atividades antioxidante, citotóxica e antimicrobiana. Mesmo sendo o própolis um produto reconhecido e bem estudado mundialmente, ainda há poucos estudos sobre o própolis português, sendo este um produto subvalorizado. O presente trabalho é uma extensão de trabalhos realizados anteriormente sobre o própolis português do apiário do Pereiro, Beira Alta. Nesses trabalhos foi observada uma diferença nos efeitos de extratos etanólicos de amostras colhidas em 2010 (P10.EE) e em 2013 (P13.EE) no modelo biológico Saccharomyces cerevisiae, sendo que P13.EE apresentava um efeito pouco citotóxico e um efeito antioxidante, enquanto P10.EE se apresentou bastante citotóxico e pouco antioxidante. Estas diferenças entre amostras do mesmo apiário levaram à questão de quais seriam os seus mecanismos de ação e o que os causaria. Os resultados anteriores sugeriram que o P10.EE induz um tipo de morte celular regulada, que poderia ser apoptótica ou necrótica, e, também, que este extrato causa danos no DNA. No presente estudo, no caso do P13.EE, os resultados de viabilidade e de análise da oxidação intracelular por citometria de fluxo com a estirpe parental BY4741 e mutantes deficientes na resposta ao stresse oxidativo demonstraram um efeito anti e pró oxidante dependente da concentração, possivelmente devido à presença de ester fenetil do ácido cafeico, pinobanksina e pinocembrina no extrato, apresentando atividade antioxidante em concentrações menores, que, por sua vez parece estar envolvida com a eliminação direta de radicais livres e a ativação das defesas antioxidantes celulares. Já no estudo de P10.EE, os resultados de ensaios distintos de microscopia de fluorescência com os marcadores 4’,6-diamino-2-fenil-indol e iodeto de propídio sugeriram que o extrato não causa morte celular por apoptose, mas sim por uma via necrótica. A análise do ciclo celular por citometria de fluxo indicou que o extrato causa uma paragem reversível nas fases G1/S do ciclo celular. Em suma, P13.EE e P10.EE exibem atividades antioxidante e citotóxica, respetivamente; porém, tais diferenças não estarão relacionadas com o seu teor fenólico, uma vez que os cromatogramas das análises cromatográficas de ultra eficiência de P10.EE e P13.EE não revelaram diferenças entre os dois extratos. Este estudo contribuiu para um maior conhecimento dos seus mecanismos de ação, para o crescimento do valor nacional deste produto português, e para a sua possível aplicação em diversas indústrias.
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Mariana Dias Pereira Antioxidant and cytotoxic activity of Portuguese propolis outubro de 2022 UMinho | 2022 Mariana Dias Pereira Antioxidant and cytotoxic activity of Portuguese propolis Universidade do Minho Escola de Ciências
Mariana Dias Pereira Antioxidant and cytotoxic activity of Portuguese propolis Dissertação de Mestrado Mestrado em Biologia Molecular, Biotecnologia e Bioempreendedorismo em Plantas Trabalho efetuado sob a orientação de Professora Doutora Cristina Alexandra Almeida Aguiar Professor Doutor Rui Pedro Soares de Oliveira Universidade do Minho Escola de Ciências outubro de 2022
ii DIREITOS DE AUTOR E CONDIÇÕES DE UTILIZAÇÃO DO TRABALHO POR TERCEIROS Este é um trabalho académico que pode ser utilizado por terceiros desde que respeitadas as regras e boas práticas internacionalmente aceites, no que concerne aos direitos de autor e direitos conexos. Assim, o presente trabalho pode ser utilizado nos termos previstos na licença abaixo indicada. Caso o utilizador necessite de permissão para poder fazer um uso do trabalho em condições não previstas no licenciamento indicado, deverá contactar o autor, através do RepositóriUM da Universidade do Minho. Licença concedida aos utilizadores deste trabalho Atribuição-NãoComercial-SemDerivações CC BY-NC-ND https://creativecommons.org/licenses/by-nc-nd/4.0/
iii ACKNOWLEDGEMENTS Firstly, I would like to thank professor Rui Oliveira and professor Cristina Aguiar, for guiding me through this work in the most supporting, encouraging and friendly way that I could have ever wished for. It was a remarkable experience to work with the both of you and I cannot thank you enough. I would also like to thank my family, who made it possible for me to pursue my goals and always wanted what was best for me, but a special “thank you” to my mom, Agnès Dias, who always supported me lovingly and unconditionally throughout my life and who never failed to encourage me, especially during the writing of this work, words cannot thank you enough. I am also immensely grateful for my other half, Jójó, who really had to put up with my temper most of the times and still was able to always make me laugh and give me all of the love, encouragement and help I needed, for this and for still being here with the same positivity as from the beginning, thank you. For my friends, Inês F. Ferreira, Filipa Silva, Cristiana Oliveira, and especially Inês M. Ferreira, my best friend and house mate for pretty much our whole academic years, and who I know nothing could separate from me, you really showed me what true friendship is about and I know this would have been a far bigger challenge if it weren’t for your love and support, thank you. I must also thank my lab partners, Ana Teixeira, Luara Simões, Rita Caetano, Juliana Silva, Catarina Baltazar, João Lopes, José Sousa, João Barbosa, João Gonçalves, João Noversa, and Ricardo Alves, for all the help, counselling, and company you gave me. I also thank all of the Biology Department technical and auxiliary staff, especially Cristina Ribeiro, Luís Correia, Manuela Rodrigues, Maria Isabel Gomes, and Inês Pinheiro, for all the help and availability, and Márcia Morais, for the support in the flow cytometry analysis. A final thanks goes to Doctor Susana Cardoso, from Aveiro University, for the support provided in the chemical analysis of the propolis extract.
iv STATEMENT OF INTEGRITY I hereby declare having conducted this academic work with integrity. I confirm that I have not used plagiarism or any form of undue use of information or falsification of results along the process leading to its elaboration. I further declare that I have fully acknowledged the Code of Ethical Conduct of the University of Minho.
v Atividade antioxidante e citotóxica de própolis português RESUMO O própolis é uma mistura resinosa e balsâmica natural produzida por abelhas rica em flavonóides e compostos fenólicos, que são geralmente associados às atividades antioxidante, citotóxica e antimicrobiana. Mesmo sendo o própolis um produto reconhecido e bem estudado mundialmente, ainda há poucos estudos sobre o própolis português, sendo este um produto subvalorizado. O presente trabalho é uma extensão de trabalhos realizados anteriormente sobre o própolis português do apiário do Pereiro, Beira Alta. Nesses trabalhos foi observada uma diferença nos efeitos de extratos etanólicos de amostras colhidas em 2010 (P10.EE) e em 2013 (P13.EE) no modelo biológico Saccharomyces cerevisiae , sendo que P13.EE apresentava um efeito pouco citotóxico e um efeito antioxidante, enquanto P10.EE se apresentou bastante citotóxico e pouco antioxidante. Estas diferenças entre amostras do mesmo apiário levaram à questão de quais seriam os seus mecanismos de ação e o que os causaria. Os resultados anteriores sugeriram que o P10.EE induz um tipo de morte celular regulada, que poderia ser apoptótica ou necrótica, e, também, que este extrato causa danos no DNA. No presente estudo, no caso do P13.EE, os resultados de viabilidade e de análise da oxidação intracelular por citometria de fluxo com a estirpe parental BY4741 e mutantes deficientes na resposta ao stresse oxidativo demonstraram um efeito anti e pró oxidante dependente da concentração, possivelmente devido à presença de ester fenetil do ácido cafeico, pinobanksina e pinocembrina no extrato, apresentando atividade antioxidante em concentrações menores, que, por sua vez parece estar envolvida com a eliminação direta de radicais livres e a ativação das defesas antioxidantes celulares. Já no estudo de P10.EE, os resultados de ensaios distintos de microscopia de fluorescência com os marcadores 4’,6-diamino-2-fenil-indol e iodeto de propídio sugeriram que o extrato não causa morte celular por apoptose, mas sim por uma via necrótica. A análise do ciclo celular por citometria de fluxo indicou que o extrato causa uma paragem reversível nas fases G1/S do ciclo celular. Em suma, P13.EE e P10.EE exibem atividades antioxidante e citotóxica, respetivamente; porém, tais diferenças não estarão relacionadas com o seu teor fenólico, uma vez que os cromatogramas das análises cromatográficas de ultra eficiência de P10.EE e P13.EE não revelaram diferenças entre os dois extratos. Este estudo contribuiu para um maior conhecimento dos seus mecanismos de ação, para o crescimento do valor nacional deste produto português, e para a sua possível aplicação em diversas indústrias. Palavras-chave: antioxidante, citotoxicidade, mecanismo de ação, própolis, Saccharomyces cerevisiae
vi Antioxidant and cytotoxic activity of Portuguese propolis ABSTRACT Propolis is a natural resinous and balsamic mixture made by honeybees, rich in flavonoids and phenolics, which are generally associated with propolis’ antioxidant, cytotoxic and antimicrobial activities. Although propolis is generally recognized and well-studied worldwide, Portuguese propolis research is still scarce and this product is nationally undervalued. The present work is an extension to previous works on Portuguese propolis from the Pereiro apiary, located in Beira Alta. Ethanol extracts of Pereiro propolis samples harvested in 2010 (P10.EE) and 2013 (P13.EE) were found to have different effects on the biological model Saccharomyces cerevisiae , with P13.EE presenting a low cytotoxic effect and a promising antioxidant effect, while P10.EE exhibited high cytotoxicity and very low antioxidant effects. These intriguing differences in samples from the same apiary led to the question of what their mechanisms of action could be and what could be causing these differences. The previous studies were able to hint on the P10.EE mechanism of action, inferring that the extract induces a type of regulated cell death, that could either be apoptotic or necrotic, and that it also causes DNA damage. In the present work, regarding the study of the P13.EE, results from viability assays and flow cytometry intracellular oxidation analysis with the parent strain BY4741 and derived mutants with a deficient oxidative stress response suggested a dose-dependent anti and pro-oxidant effect, that could be related to the presence of compounds like caffeic acid phenethyl ester, pinobanksin and pinocembrin in the extract, presenting antioxidant activity at lower concentrations, which in turn appears to result from both direct radical scavenging and activation of the cellular antioxidant defenses. Regarding the study of P10.EE, results from separate fluorescent microscopy assays with 4’,6-diamidino-2-phenylindole and propidium iodide staining suggested that the extract is not inducing cell death through apoptosis but instead through a necrotic pathway. Cell cycle analysis through flow cytometry indicated that P10.EE causes a reversible arrest in the G1/S cell cycle phases. In conclusion, P13.EE and P10.EE exhibit promising antioxidant and cytotoxic activities, respectively, but these differences are probably not related to their phenolic content, since the chromatograms from the ultra-performance chromatographic analysis of both extracts did not reveal differences between the two. This study was able to successfully uncover parts of their intriguing mechanisms of action, contributing to the growth of Portuguese propolis national value and their possible applications in various industries. Keywords: antioxidant, cytotoxic, mechanism of action, propolis, Saccharomyces cerevisiae
vii INDEX DIREITOS DE AUTOR E CONDIÇÕES DE UTILIZAÇÃO DO TRABALHO POR TERCEIROS ...... ii ACKNOWLEDGEMENTS ................................................................................................... iii STATEMENT OF INTEGRITY .............................................................................................. iv RESUMO ............................................................................................................................ v ABSTRACT ........................................................................................................................ vi LIST OF ABBREVIATIONS AND ACRONYMS .......................................................................x LIST OF FIGURES ............................................................................................................ xiii LIST OF TABLES ............................................................................................................. xix CHAPTER 1. General Introduction .................................................................................... 1 1. Propolis .................................................................................................................................. 2 2. Composition of propolis ........................................................................................................... 4 2.1. Phenolic compounds ....................................................................................................... 4 2.1.1. Flavonoids ........................................................................................................... 5 2.1.1.1. Flavones/flavonols ........................................................................................... 5 2.1.1.2. Flavanones/dihydroflavonols ............................................................................. 6 2.1.2. Phenolic acids ..................................................................................................... 7 3. Biological activities of propolis ................................................................................................. 8 3.1. Antitumoral, immunomodulatory, and cytotoxic activities .................................................. 8 3.2. Antioxidant activity ........................................................................................................... 9 3.3. Anti-inflammatory activity ............................................................................................... 10 3.4. Antimicrobial activity ...................................................................................................... 11 3.4.1. Antibacterial activity ........................................................................................... 11 3.4.2. Antifungal activity ............................................................................................... 12 3.4.3. Antiprotozoal activity .......................................................................................... 12 3.4.4. Antiviral activity .................................................................................................. 13 4. Cellular effects of propolis and its mechanism of action - Saccharomyces cerevisiae as a biological model ……………………………………………………………………………………………………………………………..15 5. Portuguese propolis ............................................................................................................... 16 6. Aim of the dissertation .......................................................................................................... 19 7. References ............................................................................................................................ 20
xiv exhibiting less fluorescence emission (cells to the left of the gating). These values represent the mean ± SD of n = 3 independent replicates. One-way ANOVA was performed, followed by Dunnett’s test for multiple comparisons. Asterisks represent significant differences between treatments and C+ (* p < 0.05, ** p < 0.01). .................................................................................................................................... 41 Figure 6. P13.EE decreases intracellular oxidation of cells from S. cerevisiae mutant strains Δ yap1 , Δ sod1 , and Δ sod2 . H2DCFDA-loaded cells from the S. cerevisiae mutant strains Δ yap1 (A-B), Δ sod1 (C-D), and Δ sod2 (E-F) , were treated with P13.EE at 50 (P13_50), 100 (P13_100), 200 (P13_200), or 500 μg/mL (P13_500) and with 5 mM H2O2, and incubated at 30 °C for 20 min. For the negative control (C-) cells were treated with absolute EtOH, at the same volume as for 500 μg/mL P13.EE, and for the positive control (C+) cells were treated with 5 mM H2O2. (A, C, E) Histograms of one representative experiment out of three independent replicates. (B, D, F) Percentage of cells with decreased intracellular oxidation. A gating was drawn at the peak of C+ to obtain the percentage of cells exhibiting less fluorescence emission (cells to the left of the gating). These values represent the mean ± SD of n = 3 independent replicates. One-way ANOVA was performed, followed by Dunnett’s test for multiple comparisons. Asterisks represent significant differences between treatments and C+ (* p < 0.05, ** p < 0.01). .................................................................................................................................... 43 Figure 7. Chromatographic profile of P13.EE at 280 nm obtained by UPLC-DAD-ESI/MS. Numbers in the figure represent the fractions that were further analyzed by ESI/MS. ......................... 45 CHAPTER 3 Figure 8. Summary of the morphological and biochemical characteristics of the yeast RCD types apoptosis, necrosis and autophagy. ................................................................................ 55 Figure 9. Effects of P10.EE on the viability of S. cerevisiae . Cells from the S. cerevisiae BY4741 strain were treated with P10.EE at the concentrations of 200, 500, or 750 μg/mL. For the negative control (C-) cells were treated with absolute EtOH, at the same volume as for the P10.EE. Aliquots were collected after 0, 30, 60, and 90 min, and serially diluted from 10-1 to 10-4, and 5 μL drops of each dilution were placed on YPDA plates and incubated at 30 °C for 48 h. Cytotoxicity is observed by a decrease in number of colonies when compared to C-. ........................................................................................ 61 Figure 10. P10.EE-treated S. cerevisiae cells do not present apoptotic features. Percentage of S. cerevisiae BY4741 cells presenting apoptotic features, such as chromatin condensation and/or
xv fragmentation, after 0, 30, 60, and 90 min of incubation with 500 μg/mL P10.EE. At each time-point, aliquots were collected and the cells were heat-fixed and stained with DAPI. For the negative control (C-) cells were treated with absolute EtOH, at the same volume as for the P10.EE, for the apoptosis control (C Apoptosis) cells were treated with H2O2 at 10 mM, and for the necrosis control (C Necrosis) cells were treated with H2O2 at 180 mM. These values represent the mean ± SD of n = 3 independent replicates. Two-way ANOVA was performed, followed by Tukey’s test for multiple comparisons. Asterisks represent significant differences between treatments and C- (**** p < 0.0001). ................................................ 63 Figure 11. Analysis of chromatin condensation and nuclei fragmentation through fluorescence microscopy. S. cerevisiae BY4741 cells treated with absolute EtOH (C-), 10 mM H2O2 (C Apoptosis), 180 mM H2O2 (C Necrosis) and 500 μg/mL P10.EE, after 90 min of incubation, were heatfixed and stained with DAPI. Images were obtained through BF and fluorescence microscopy (magnification of 100x) and are from one representative experiment, out of three independent replicates. ........................................................................................................................................................ 64 Figure 12. P10.EE-treated S. cerevisiae cells exhibit loss of membrane integrity. Percentage of S. cerevisiae BY4741 PI positive cells, after 0, 30, 60, and 90 min of incubation with 500 μg/mL P10.EE. At each time-point, aliquots were collected and the cells were stained with PI. For the negative control (C-) cells were treated with absolute EtOH, at the same volume as for the P10.EE, for the apoptosis control (C Apoptosis) cells were treated with 10 mM H2O2, and for the necrosis control (C Necrosis) cells were treated with 180 mM H2O2. These values represent the mean ± SD of n = 4 independent replicates. Two-way ANOVA was performed, followed by Tukey’s test for multiple comparisons. Asterisks represent significant differences between treatments and C- (**** p < 0.0001). ................................................ 65 Figure 13. Analysis of plasma membrane integrity through fluorescence microscopy. S. cerevisiae BY4741 cells treated with absolute EtOH (C-), 10 mM H2O2 (C Apoptosis), 180 mM H2O2 (C Necrosis) and 500 μg/mL P10.EE, after 90 min of incubation, were stained with PI. Images were obtained through BF and fluorescence microscopy (magnification of 100x) and are from one representative experiment, out of four independent replicates. ................................................................................. 67 Figure 14. P10.EE affects S. cerevisiae cell cycle progression. (A) Using the CytExpert 2.5 software, three gates were drawn to define the percentage of S. cerevisiae BY4741 cells, prior to HU synchronization, in the cell cycle phases G1, S and G2/M, based on the amount of fluorescence and number of cells – a first peak with less fluorescence represented G1 phase, a second peak with higher
xvi amount of fluorescence represents G2/M phases, and the cells in between the two peaks represent S phase. The histogram presented is of one representative experiment out of three independent replicates. (B) Percentage of S. cerevisiae BY4741 cells in the cell cycle phases G1, S, and G2/M, after 0, 30, 60, 90, 120, and 180 min of treatment with 200 μg/mL P10.EE. Aliquots were collected at each time-point and cells were prepared for flow cytometry analysis and stained with SYTOXTM Green Nucleic Acid Stain (Invitrogen). For the negative control (C-) cells were treated with absolute EtOH, at the same volume as for the P10.EE. These values represent the mean ± SD of n = 3 independent replicates. Two-way ANOVA was performed, followed by Sidak’s test for multiple comparisons. Asterisks represent significant differences between treatments and C- (* p < 0.05, *** p < 0.001, **** p < 0.0001). ....................... 69 APPENDIX 1 Figure A1. Chromatographic profile of P10.EE at 280 nm obtained by UPLC-DAD-ESI/MS. ........................................................................................................................................................ 80 APPENDIX 2 Figure A2. Analysis of chromatin condensation and nuclei fragmentation through fluorescence microscopy. S. cerevisiae BY4741 cells treated with absolute EtOH (C-), 10 mM H2O2 (C Apoptosis), 180 mM H2O2 (C Necrosis) and 500 μg/mL P10.EE, after 0 min of incubation, were heatfixed and stained with DAPI. Images were obtained through BF and fluorescence microscopy (magnification of 100x) and are from one representative experiment, out of three independent replicates. ........................................................................................................................................................ 82 Figure A3. Analysis of chromatin condensation and nuclei fragmentation through fluorescence microscopy. S. cerevisiae BY4741 cells treated with absolute EtOH (C-), 10 mM H2O2 (C Apoptosis), 180 mM H2O2 (C Necrosis) and 500 μg/mL P10.EE, after 30 min of incubation, were heatfixed and stained with DAPI. Images were obtained through BF and fluorescence microscopy (magnification of 100x) and are from one representative experiment, out of three independent replicates. ........................................................................................................................................................ 83 Figure A4. Analysis of chromatin condensation and nuclei fragmentation through fluorescence microscopy. S. cerevisiae BY4741 cells treated with absolute EtOH (C-), 10 mM H2O2 (C Apoptosis), 180 mM H2O2 (C Necrosis) and 500 μg/mL P10.EE, after 60 min of incubation, were heat-
xvii fixed and stained with DAPI. Images were obtained through BF and fluorescence microscopy (magnification of 100x) and are from one representative experiment, out of three independent replicates. ........................................................................................................................................................ 84 Figure A5. Analysis of plasma membrane integrity through fluorescence microscopy. S. cerevisiae BY4741 cells treated with absolute EtOH (C-), 10 mM H2O2 (C Apoptosis), 180 mM H2O2 (C Necrosis) and 500 μg/mL P10.EE, after 0 min of incubation, were stained with PI. Images were obtained through BF and fluorescence microscopy (magnification of 100x) and are from one representative experiment, out of four independent replicates. ................................................................................. 85 Figure A6. Analysis of plasma membrane integrity through fluorescence microscopy. S. cerevisiae BY4741 cells treated with absolute EtOH (C-), 10 mM H2O2 (C Apoptosis), 180 mM H2O2 (C Necrosis) and 500 μg/mL P10.EE, after 30 min of incubation, were stained with PI. Images were obtained through BF and fluorescence microscopy (magnification of 100x) and are from one representative experiment, out of four independent replicates. ................................................................................. 86 Figure A7. Analysis of plasma membrane integrity through fluorescence microscopy. S. cerevisiae BY4741 cells treated with absolute EtOH (C-), 10 mM H2O2 (C Apoptosis), 180 mM H2O2 (C Necrosis) and 500 μg/mL P10.EE, after 60 min of incubation, were stained with PI. Images were obtained through BF and fluorescence microscopy (magnification of 100x) and are from one representative experiment, out of four independent replicates. ................................................................................. 87 Figure A8. Analysis of P10.EE effect on the viability of S. cerevisiae cells. Percentage of S. cerevisiae BY4741 cell viability after 0, 30, 60, 90, 120, 180, and 240 min of treatment with 200 μg/mL P10.EE. Aliquots were collected at each timepoint, serially diluted from 10-1 to 10-4 and three 40 μL drops of the 10-4 dilution were placed on YPDA plates and incubated at 30 °C for 48 h. For the negative control (C-) cells were treated with absolute EtOH, at the same volume as for the P10.EE. Viability of each timepoint was calculated as the percentage of colony-forming units, taking the 0 min time-point as 100 %. These values represent the mean ± SD of n = 3 independent replicates............................................. 88 Figure A9. Analysis of P10.EE effect on S. cerevisiae cell cycle progression. HU synchronized S. cerevisiae BY4741 cells were treated with 200 μg/mL P10.EE for 180 min. After 0, 30, 60, 90, 120, and 180 min of incubation, aliquots were collected and cells were prepared for flow cytometry analysis and stained with SYTOXTM Green Nucleic Acid Stain (Invitrogen). For the negative control (C-), cells were
xviii treated with absolute EtOH, at the same volume as for the P10.EE. Histograms presented are of one representative experiment out of three independent replicates. .......................................................... 89 Figure A10. Synchronization of S. cerevisiae cell cycle. Percentage of S. cerevisiae BY4741 cells in the cell cycle phases G1, S, and G2/M, after synchronization of the cells with HU at 200 mM. Synchronized and unsynchronized cells were prepared for flow cytometry analysis and stained with SYTOXTM Green Nucleic Acid Stain (Invitrogen). These values represent the mean ± SD of n = 3 independent replicates. Two-way ANOVA was performed, followed by Sidak’s test for multiple comparisons. Asterisks represent significant differences between synchronized and unsynchronized cells (* p < 0.05, **** p < 0.0001). .......................................................................................................... 91
xix LIST OF TABLES CHAPTER 2 Table 1. Yeast strains used in this work and their respective genotypes and origin. ......... 35 Table 2. Chemical characterization of phenolic content of P13.EE obtained by UPLC-DADESI/MS. ......................................................................................................................................... 45
1 CHAPTER 1. General Introduction
2 1. Propolis Propolis is a natural product produced by honeybees (mainly Apis mellifera ), consisting of lipophilic resinous materials collected from branches, flowers, pollen, buds and exudates of various plant sources, that are partially digested with the bees’ salivary enzyme -glucosidase and then mixed with beeswax and other compounds of bees’ metabolism. With a gluey and resinous consistency, propolis displays a strong odor and usually a brown, green, or red tint, depending on the type of propolis (Burdock, 1998; Bankova et al. , 2016; Peixoto et al. , 2021). Because this natural product is mainly produced from plant material, its precise composition depends greatly on the local flora diversity, being recognized several propolis types (Silva-Carvalho et al. , 2015) with diverse chemical compositions that provide different biological activities, such as cytotoxic, antitumoral, antioxidant, and antimicrobial activities (Kujumgiev et al. , 1999; Bankova, 2005; Gómez-Caravaca et al. , 2006; Castro et al. , 2011; Valente et al. , 2011; Bankova et al. , 2014; Capoci et al. , 2015; Silva-Carvalho et al. , 2015). The word propolis comes from the Greek pro- (defense of) and polis- (city), which means defense of the city, or the hive (Ghisalberti, 1979). Due to its mechanical and biological properties, propolis is used by bees in many ways to defend the hive, for example to fix holes and cracks in the honeycombs, to smooth its internal walls, and to protect the hive from intruders – propolis is used to encapsulate the invaders, killing them by asphyxia, but also to conserve their bodies, containing putrefaction and its resultant plagues (Bankova et al. , 2002). Although propolis is a very popular natural remedy nowadays, its usage goes back to at least 300 BC, being recognized by many ancient civilizations (Ghisalberti, 1979): Egyptians used propolis to embalm cadavers due to its anti-putrefactive properties (Kuropatnicki et al. , 2013); the Greeks and Romans used it for wound treatment and as mouth disinfectant, due to its antiseptic and healing properties, and it was recognized for its medical properties by many physicians, such as Aristoteles, Hippocrates, Dioscorides, Pliny, and Galen (Castaldo & Capasso, 2002; Kuropatnicki et al. , 2013). Arab physicians recognized propolis properties as well, using it for the treatment of eczemas, myalgia and rheumatism (Kuropatnicki et al. , 2013). Propolis continued being used for its therapeutic properties during medieval times, being very present in traditional folk medicine in Eastern Europe (Kuropatnicki et al. , 2013). It became a very popular drug in Europe between the 17th and 19th centuries, due to its antibacterial and healing properties, and it was also used during the Second World War for tuberculosis treatment, as well as for the treatment of wounds, burns, sore throats and stomach ulcers (Wollenweber et al. , 1990; Fokt et al. , 2010). Although most propolis application records are due to its medicinal
3 properties, it can also be used for other purposes, for example, Antonio Stradivari used this bee product in the varnish for musical instruments (Burdock, 1998). Nowadays, we can easily find an array of products made with propolis in many health-food stores, being available in different formats, such as capsules (either in pure form or combined with other compounds), extracts (hydroalcoholic or glycolic), mouthwash solutions (combined with other substances), creams, powder, and as a purified product (Castaldo & Capasso, 2002; Fokt et al. , 2010). Due to its extensive biological and medicinal properties, propolis popularity has been growing, and so has the scientific interest in it, with aims to study it further and possibly apply to human and veterinary medicine, pharmacology, food and cosmetics.
4 2. Composition of propolis Propolis is a complex resinous and balsamic mixture, generally composed of resins and plant balsams (50 %), beeswax (30 %), essential and aromatic oils (10 %), pollen (5 %), and other substances and materials, including mineral and organic compounds (5 %) (Bankova 2005; Pasupuleti et al. 2017). Phenolic acids (cinnamic and caffeic acid), flavonoids, their esters (flavones, flavanones, flavonols, and dihydroflavonols chalcones), terpenes, aromatic aldehydes and alcohols, fatty acids, stilbenes, and 𝛽steroids are the main organic compounds present in propolis (Huang et al. , 2014). As mentioned above, propolis chemical composition varies among different types of propolis, and is difficult to standardize, since it depends on many different factors, like the geographic location, environmental conditions, vegetation available, and the season of collection (Bankova, 2005). In total, there have been identified over 800 compounds in propolis samples, for example: polyphenols, phenolic aldehydes, sesquiterpenes, quinines, coumarins, amino acids, steroids, and inorganic compounds, but many more are expected to be identified as new samples get chemically characterized (Castro et al. , 2011; Huang et al. , 2014; Kasote et al. , 2022). Due to chemical complexity and diversity, several types of propolis have been classified according to its plant source and chemical composition: Poplar propolis (Europe, North-America, non-tropic regions of Asia, China and New Zealand), Birch propolis (Russia), Green (Alecrim) propolis (Brazil), Red propolis (Brazil, Cuba and Mexico), “Clusia” propolis (Cuba and Venezuela), “Pacific” propolis (Okinawa, Taiwan, Hawaii, Indonesia and Myanmar), “Canarian” propolis (Canary Islands), Mediterranean propolis (Sicily and the Adriatic Coast), Australian propolis, Thailand propolis and African propolis (Bankova, 2005; Silva-Carvalho et al. , 2015; Miłek et al. , 2022). Propolis chemical profile can be characterized by three parameters: total flavone and flavonols content, total flavanone and dihydroflavonol content, and total phenolics content (Bankova, 2005). Bankova (2005) analyzed various samples of Poplar propolis, the most thoroughly studied type of propolis, and proposed the characteristics of typical poplar propolis as 8 ± 4 % flavones/flavonols, 6 ± 2 % flavanones/dihydroflavonols and 28 ± 9 % of total phenolics. 2.1. Phenolic compounds Phenolic compounds are greatly present in plants and account for one of the most notable groups of plant secondary metabolites. Their chemical structure is of one or more aromatic rings, with at least one hydroxyl group, being classified as simple phenols if containing a single phenol unit, or as polyphenols if the molecule contains more than one (Roleira et al. , 2018). The main groups of phenolic compounds
11 and the proliferation of T cells (Araujo et al. , 2011; Oryan et al. , 2018). Flavonoids isolated from an ethanol extract of propolis from Nepal have shown an inhibitory effect on the expression of many inflammatory genes in bone marrow-derived mast cells, such as IL-6 , TNF-𝛼 , and IL-13 (reviewed by SilvaCarvalho et al. , 2015). Brazilian green propolis has been demonstrated to decrease the number of macrophages and neutrophils in a lipopolysaccharide-induced pulmonary inflammation model, inducing a decrease in IL-6 and TNF-𝛼 secretion, and an increase in IL-10 and transforming growth factor-𝛽 (TGF𝛽) (Machado et al. , 2012). Additionally, an EE of Portuguese propolis has exhibited anti-inflammatory effects, both in vitro , inhibiting bovine serum albumin (BSA) denaturation, and in vivo , improving mechanical hyperalgesia and gait pattern parameters in an osteoarthritis-induced model (Araújo et al, 2022). 3.4. Antimicrobial activity Antimicrobial activity is one of the most studied properties in propolis and it has been well documented against bacteria, fungi, protozoa, and viruses (Sforcin et al. , 2000; Gekker et al. , 2005; Búfalo et al. , 2009; Sforcin & Bankova, 2011; Silva-Carvalho et al. , 2015; Asfaram et al. , 2021). 3.4.1. Antibacterial activity Bacterial resistance to antibiotics is a rising concern nowadays and investigating natural products with antibacterial properties is crucial to cope with such problem (Campos et al. , 2015). Propolis has long been a contender for this purpose, due to the vast reports on its antibacterial activity (reviewed by Silva-Carvalho et al. , 2015). This property of propolis depends greatly on its chemical composition and place of collection, but it is mostly due to its flavonoids, aromatic acids and esters (Marcucci, 1995; Hegazi et al. , 2000; Seidel et al. , 2008). Compounds such as galangin, pinocembrin and pinostrobin have been strongly associated with propolis antibacterial activity, but caffeic and ferulic acid play a role in it too (Dimov et al. , 1992; Marcucci, 1995). Propolis is much more active against Gram-positive than Gram-negative bacteria, which could be a consequence of the differences in cell wall structure of these two bacterial types, since Gram-negative bacteria have a negatively charged lipopolysaccharide (LPS) layer that acts as a barrier (Kujumgiev et al. , 1999; Cushnie & Lamb, 2005; Fokt et al. , 2010;). Mirzoeva et al. (1997) proposed that propolis can affect the ion permeability of the inner bacterial membrane, leading to the dissipation of the membrane potential and inhibiting bacterial motility. Propolis has been reported to be effective against many Gram-
12 positive bacteria, such as Bacillus subtilis , Staphylococcus aureus , Staphylococcus epidermidis , several species of the genera Streptococcus , Enterococcus spp., Micrococcus luteus , among others (Popova et al. , 2009; Righi et al. 2011; Schmidt et al. , 2014; Bittencourt et al. , 2015; Campos et al. , 2015; AL-Ani et al. , 2018). Although not common in many propolis extracts, Turkish and Brazilian propolis ethanol extracts have shown significant activity against Escherichia coli , a Gram-negative bacterium (Fernandes Jr. et al. , 2006; Katircioǧlu & Mercan, 2006). Moreover, synergetic effects between propolis and several antibiotics have been reported (Orsi et al. , 2006; Al-Waili et al. , 2012; Lavigne et al. , 2020; Freitas et al. , 2021; Ramata-Stunda et al. , 2022), suggesting that a combination of both could potentiate the antibiotics’ effect and allow for dose reduction. 3.4.2. Antifungal activity Much like the antibacterial, propolis antifungal activity is affected by its chemical composition and site of collection (Kujumgiev et al. , 1999). This property of propolis has been studied among several fungi, and has been mostly ascribed to propolis volatile compounds and flavonoids, such as pinocembrin and galangin (Quiroga et al. , 2006; Fokt et al. , 2010; Bankova et al. , 2014; Silva-Carvalho et al. , 2015). Propolis has been reported to be effective against several Candida strains, including Candida albicans , but also Pichia ohmeri , Rhodotorula spp., Trichosporon spp., Aspergillus fumigatus , several species of the genera Trichophyton and Microsporum , Epidermophyton floccosum , Malassezia spp. among others (Fernandes Jr. et al. , 1995; Oliveira et al. , 2006; Wagh, 2013; Falcão et al. , 2014; Cerqueira et al. , 2022). The effectiveness against C. albicans is of great interest, and Brazilian propolis EE has been reported to inhibit the growth and biofilm formation of this strain in vulvovaginal candidiasis (Dota et al. , 2011; Capoci et al. , 2015). It has been proposed that propolis affects C. albicans via metacaspase and by the Ras pathway (Castro et al. , 2013). Moreover, Portuguese propolis EEs have been reported to act as biocontrol agents in fruit fungal infections, such as brown spot disease in pear, caused by the fungus Stemphylium vesicarium (Loebler et al. , 2020), and apple blue mold, caused by the fungus Penicillium expansum (Pereira et al. , 2022), being a great candidate for agri-food applications. 3.4.3. Antiprotozoal activity Propolis antiprotozoal activity, although not as deeply investigated as its other antimicrobial activities, is of great importance, as protozoan parasitic infections represent a worldwide public health issue (Sundar & Chakravarty, 2013). Propolis has been reported to be effective against a few protozoan
13 parasites: Leishmania spp., the causer of leishmaniasis – propolis inhibits the proliferation of the parasite and has an anti-inflammatory action through the inhibition of NO• production (Asfaram et al. , 2021); Plasmodium spp., the causer of malaria – propolis inhibits the parasitemia (quantitative measure of parasites in the blood), which causes an improvement in the anemia caused by the infection, but also controls the infection through immunomodulation (Olayemi, 2014; AlGabbani et al. , 2017); Trypanosoma spp., the causer of trypanosomiasis – propolis has been reported to act in a similar way as for Plasmodium spp. (Nweze et al. , 2017, Asfaram et al. , 2019); Blastocystis spp., a common lower intestinal opportunistic pathogen – propolis could be activating apoptotic mechanisms (Asfaram et al. , 2021). A phenolic extract of propolis from Central Portugal was reported to be active against Plasmodium falciparum , presenting a half maximal inhibitory concentration (IC50) of 8.83 μg/mL (Falcão et al. , 2014), while Brazilian green propolis presented an IC50 of 20 μg/mL (Filho et al. , 2009). Samples of propolis from North-eastern and central Portugal both presented an IC50 of 8.11 μg/mL against Leishmania infantum (Falcão et al. , 2014), while two Turkish samples presented an IC50 of 125 and 325 μg/mL (Duran et al. , 2011) and Brazilian green propolis an IC50 of 49 μg/mL (Filho et al. , 2009). Propolis samples from North-eastern and central Portugal presented an IC50 of 6.16 and 8.59 μg/mL against Trypanosoma cruzi , respectively (Falcão et al. , 2014), while in Bulgarian and Brazilian propolis samples IC50 ranged from 108.8 to 1065 μg/mL (Prytzyk et al. , 2003) and from 421 to 1437 μg/mL (Cunha et al. , 2004a), respectively. Finally, propolis from North-eastern Portugal was reported to be very active against Trypanosoma brucei , presenting an IC50 of 1.70 μg/mL (Falcão et al. , 2014), which fits in the value range of the antitrypanosomal reference drug suramin (Otoguro et al. , 2012). These disparities in activities of propolis samples from different countries are caused by their distinct chemical compositions, although it is not yet clear what propolis compounds are behind it (Paula et al. , 2021). Otogura et al. (2012) observed that a ß-phenylethyl caffeate isolate from propolis presented a considerable activity against T. brucei , with an IC50 of 0.013 μg/mL, but it is suggested that propolis antiprotozoal activity originates from a combination of its antibacterial, anti-inflammatory, and immunomodulatory activities (Paula et al. , 2021). 3.4.4. Antiviral activity Propolis from different geographic locations has been reported to be active against several viruses, such as influenza, herpes simplex (HSV) types 1 and 2, hepatitis B, and even immunodeficiency
14 virus (HIV) (Amoros, et al. , 1992a,b; Serkedjieva et al. , 1992; Marcucci, 1995; Gekker et al. , 2005; Shimizu et al. , 2008; Schnitzler et al. , 2010; Sartori et al. , 2012). Many compounds present in propolis have been associated with its antiviral action: quercetin, kaempferol, CAPE, and isoprenyl caffeate have been shown to act against HIV by inhibiting the enzyme HIV-1 integrase, which is essential for viral DNA integration into the host genome (Fesen et al. , 1993; Harish et al. , 1997; Xu et al. , 2000; Chiu & Davies, 2004). Flavonoids have also been associated with propolis action against picornavirus, preventing the decapsidation of viral particles and the release of RNA in the cells (Tait et al. , 2006). Propolis from Czech Republic was tested against HSV-1, and galangin and chrysin proved to be the main bioactive compounds (Schnitzler et al. , 2010). Isopentyl ferulate, an ester of a substituted cinnamic acid, inhibited the activity of influenza virus A (Serkedjieva et al. , 1992). Recently, propolis has been proposed to be effective against SARS-CoV-2, although this has only been tested in silico (molecular docking), and further studies would be necessary to confirm it (Berretta et al. , 2020). Many propolis components, such as CAPE, galangin, chrysin, and caffeic acid could serve as potential drugs against this virus, as they are predicted to interact with MPRO, an enzyme essential for coronavirus’ life cycle and processing of proteins (Hashem, 2020). SARS-CoV-2 also binds firmly to the enzyme ACE2, using it as receptor and for the replication in host cells (Hoffmann et al. , 2020; Zhou et al. , 2020); Guler et al. (2021) used molecular docking to test if some propolis flavonoids and phenolic compounds would bind to it: although rutin had the most favourable binding energy to ACE2, myricetin, CAPE, hesperetin and pinocembrin were great candidates too.
15 4. Cellular effects of propolis and its mechanism of action - Saccharomyces cerevisiae as a biological model The biological model chosen in this work to study propolis effects is the unicellular yeast Saccharomyces cerevisiae . It is among the most commonly used eukaryotic model organisms, mainly because of its cellular structure and simplicity to work with, its fully sequenced genome that allows the researcher to easily delete or replace specific genes, and the fact that many genes present in this yeast’s genome are orthologues to genes associated with human diseases and, for most of them, the mammalian homologue is functional (Goffeau et al. , 1996; Heinicke et al. , 2007; Karathia et al. , 2011; Wloch-Salamon & Bem, 2013). Saccharomyces cerevisiae has been used as a model to study many biological processes, such as cell cycle, metabolism, cell death, regulation of gene expression, and neurodegenerative disorders, among others (Nasheuer et al. , 2002; Brocard-Masson & Dumas, 2006; López-Mirabal & Winther, 2008; Owsianowski et al. , 2008; Karathia et al. , 2011; Wloch-Salamon & Bem, 2013; Scariot et al. , 2018). It can be grown on a small volume of defined media and its cell cycle lasts between 90 and 120 min, allowing for a rapid growth of vast cell populations (Goffeau et al. , 1996; Hanson, 2018). Saccharomyces cerevisiae has been used as a model on several studies with propolis regarding its biological properties (Castro et al. , 2011; Marques, 2015; Cruz et al. , 2016; Alves, 2018; Freitas et al. , 2019; Peixoto et al. , 2021). As mentioned above, propolis biological properties depend greatly on its composition. Castro et al. (2011) reported a cytotoxic effect from Green propolis (from Brazil) in S. cerevisiae , inducing cell death by apoptosis and even necrosis, when the exposure to propolis is longer. Green propolis was also reported to induce vacuolar acidification and the translocation of Atg8p (a hallmark of autophagy) to the vacuole – this pH imbalance could lead to necrosis, since pro-necrotic proteases are active in acidified cytosol (Eisenberg et al. , 2010). Saccharomyces cerevisiae was later reported to have many cell cycle related genes with a decreased messenger RNA (mRNA) accumulation, when exposed to this propolis, suggesting an effect on transcriptional checkpoint controls (Castro et al. , 2012).
16 5. Portuguese propolis Presently, as reported on the Portugal National Beekeeping Program 2020-2022 (Gabinete de Planeamento, Políticas e Administração Geral, 2019), there are 11,883 registered beekeepers in Portugal, corresponding to around 42,000 apiaries and 768,000 hives. Although beekeeping activity is spread along the whole national territory, most beekeepers are centered in Northern and Central Portugal, and their main focus is on honey production and trade, and not so much on other bee products, such as propolis. Despite propolis' broad list of valuable properties related to human health and well-being, Portuguese propolis has not deserved much attention and it still is an undervalued product, most of the times being discarded or stored without being used (Peixoto et al. , 2021). Propolis production is limited, reaching from 10 to 300 g per hive per year. Yet, propolis economic value in Portugal is still very low, compared to other countries, since studies regarding its chemical composition and biological properties are scarce and still at an early stage (Dias et al. , 2012; Tsagkarakis et al. , 2017). As mentioned above, propolis chemical composition can vary depending on the local flora, climate conditions and season of harvest, even in the same country (Falcão et al. , 2012). Differences between Portuguese samples harvested from the same apiary but from different years or seasons have been reported (Miguel et al. , 2010; Marques, 2015; Peixoto et al. , 2022) – these differences have also been reported for other propolis samples worldwide (Simões-Ambrosio et al. , 2010; Isla et al. , 2012; Neto et al. , 2017). Falcão et al. (2013) characterized the phenolic compounds present in 40 propolis samples from different regions of Portugal – north, central interior, central coast, south, Azores archipelago, and Madeira island – and separated the samples in two groups: the ones that resemble common temperate propolis, containing several compounds typically found in poplar propolis, such as flavonoids and their methylated forms, and phenylpropanoid acids and their esters (Falcão et al. , 2010) (Type I); and the ones with an uncommon composition (Type II), that besides common flavonoids, also contained quercetin and kaempferol glycosides, with many of them not previously described in propolis. These differences in composition are likely due to a different plant origin other than poplar buds and could be related to diversified biological activities (Dias et al. , 2012; Falcão et al. , 2013; Silva-Carvalho et al. , 2015). Portuguese propolis has been associated with many biological properties, such as antimicrobial, antioxidant, and antitumoral activities (Moreira et al. , 2008; Fokt et al. , 2010; Miguel et al. , 2010; Valente et al. , 2011; Dias et al. , 2012; Valença et al. , 2013; Calhelha et al. , 2014; Silva-Carvalho et al. , 2014,
17 2015; Falcão et al. , 2014; Cruz et al. , 2016; Freitas et al. , 2019; Peixoto et al. , 2021, 2022; Oliveira et al, 2022). Regarding antimicrobial activity, Portuguese propolis has been mainly reported to be active against bacteria and fungi (Silva et al. , 2012; Freitas et al. , 2019). Ethanol extracts of propolis from Gerês harvested from 2011 to 2014 have been reported to exhibit antibacterial activity against Gram-positive bacteria, with a strong effect against the spore forming bacteria of Bacillus genus, presenting minimum inhibitory concentration (MIC) values of 50 μg/mL (Freitas et al. , 2019). Falcão et al. (2014) reported that although propolis EEs from Bragança and Leiria were not very effective against the yeast Candida albicans , they exhibited an effect against the Gram-positive bacterium Staphylococcus aureus and the fungus Trichophyton rubrum . The most studied biological property of this natural product in Portugal is definitely its antioxidant activity. Cruz et al. (2016) reported that an EE of propolis from Côa has a protective effect in yeast cells against H2O2-induced oxidative stress at a concentration of 100 μg/mL and has also caused a decrease in H2O2-induced intracellular oxidation, in a dose dependent manner (100 and 300 μg/mL). Propolis EEs from Bornes and Fundão were able to reduce lipid peroxidation and hemolysis caused by peroxyl radicals in human erythrocytes in a dose dependent manner, starting at 5 μg/mL all the way to 40 μg/mL (Valente et al. , 2011), which are very low concentrations, when compared to the already tested concentrations for other types of propolis (Campos et al. , 2014; Bonamigo et al. , 2017). Moreover, many extracts of propolis from all over the country, such as Algarve, Gerês, Bragança and Leiria have shown high antioxidant activity in in vitro assays like DPPH scavenging activity (Falcão et al. 2014; Peixoto et al. , 2021, 2022), O2scavenging activity and iron chelating activity (Miguel et al. , 2010; Freitas et al. , 2019). A property that has recently been receiving more and more attention in Portuguese propolis studies because of its importance in health sciences is its antitumoral and cytotoxic activity. Valente et al. (2011) tested the ability of Bornes and Fundão propolis EE to suppress the proliferation of human primary renal cancerous cells, with an MTT assay, reporting 85 % (Bornes) and 91 % (Fundão) cell proliferation inhibition at an extract concentration of 100 μg/mL. Chloroform and residual ethanol fractions from Azores propolis EE were shown to decrease the proliferation of the human colon carcinoma cell line HCT15 in a dose-dependent manner, starting at the concentrations of 10 μg/mL for the chloroform fraction and 15 μg/mL for the residual ethanol fraction, but also caused a disturbance in HCT-15 glycolytic metabolism (Valença et al. , 2013). Phenolic extracts of propolis from Bragança and São Miguel, Azores, were too shown to exhibit high cytotoxicity against the HCT-15 cell line, with the concentrations necessary to inhibit 50 % of the net cell growth of 10 ± 1 (Bragança) and 9 ± 1 μg/mL (São Miguel; Calhelha et al. ,
18 2014). A Beira Alta propolis EE from 2010 was reported to decrease proliferation and migration of the cancer cell lines MDA-MB-231 (breast cancer) and DU145 (prostate cancer), in a dose-dependent manner and starting at the concentrations of 15 μg/mL for MDA-MB-231 cell line and 7 μg/mL for the DU145 cell lines (Silva-Carvalho et al. , 2014). Recently, an n-butanol (n-BuOH) fraction of Gerês propolis EE from 2018 was found to show high cytotoxicity on BRAF-mutated cell lines, one of the most aggressive melanoma types, with an IC50 of 8.14 ± 0.03 μg/mL for the A375 cell line and of 11.22 ± 1.66 μg/mL for the WM9 cell line – this fraction has high quantities of CAPE, a compound known for its anticancer properties, suggesting its involvement in Gerês propolis cytotoxicity against melanoma (Oliveira et al. , 2022). An ethyl acetate fraction of the same Gerês propolis EE was also shown to be highly cytotoxic against the A498 and 786-O renal cell carcinoma cell lines, with an IC50 of 0.162 μg/mL for the A498 cell line and of 0.271 μg/mL for the 786-O cell line (Freitas et al. , 2022). It is crucial to keep studying Portuguese propolis, diving deeper into its properties and exact composition, in order to promote its use and increase its national value.
19 6. Aim of the dissertation Because of their many biological properties, natural products have been on the forefront of pharmaceutical, cosmetic, agricultural and food research (Che & Zhang, 2019; Atanasov et al. , 2021; González-Manzano & Dueñas, 2021; Goyal & Jerold, 2021). Propolis is a natural product that presents many biological properties, such as antioxidant, antimicrobial and cytotoxic activities. Although it is gaining attention worldwide for its multiple purposes, its chemical composition varies greatly from region to region, presenting different biological activities, which is a setback for its application on health, cosmetic and food industries, since it needs to be chemically standardized. Portuguese propolis, besides the mentioned standardization problem, is yet to be recognized for its value by beekeepers and consumers. Ethanol extracts of Portuguese propolis samples from the Pereiro apiary harvested in 2010 (P10.EE) and 2013 (P13.EE) were reported to exhibit very different activities, as P13.EE exhibits antioxidant activity and P10.EE exhibits cytotoxic activity. The present work aims to evaluate these P13.EE and P10.EE bioactivities, in order to better understand their different mechanisms of action, and therefore contributing to the valorization of Portuguese propolis, that has so much potential and is many times overlooked. For this purpose, the biological model S. cerevisiae was used, and the work was divided in two parts, the study of the antioxidant properties of P13.EE, and the study of the cytotoxic properties of P10.EE. For the study of P13.EE, viability assays with S. cerevisiae BY4741 and mutant strains were done, to assess cytotoxicity. Intracellular oxidation was also analyzed, by flow cytometry, for the same yeast strains, to assess the extract’s antioxidant activity. For the study of P10.EE, viability assays with S. cerevisiae BY4741 strain were done, to confirm cytotoxicity, and, to assess its effect on regulated cell death, two fluorescence microscopy assays were done: chromatin condensation and fragmentation (apoptotic morphological features) was evaluated with 4’,6-diamidino-2-phenylindole (DAPI) staining; loss of plasma membrane integrity (common necrosis marker) was evaluated with propidium iodide (PI) staining. P10.EE effect on the yeast cell cycle was also studied, by flow cytometry, to assess if the extract’s cytotoxicity was causing an arrest or delay in the cell cycle.
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31 CHAPTER 2. Antioxidant activity of Pereiro propolis
32 1. Introduction 1.1. Reactive oxygen species Although aerobic eukaryotes depend on oxygen to survive, this molecule can also present a danger to their survival, when in supraphysiological concentrations. In fact, atomic oxygen has two unpaired electrons, making it vulnerable to the formation of radicals (Bardaweel et al. , 2018). Reactive oxygen species (ROS) is an umbrella term used to describe the reactive molecules and free radicals that originate from molecular oxygen, such as superoxide (•O2-), hydrogen peroxide (H2O2), hydroxyl radical (•OH) and hydroxyl ion (OH-) (Sies & Jones, 2020). ROS can be formed naturally in many ways and from various sources, and even though the mitochondrial electron transport chain contributes greatly to their formation, it has been evidenced that ROS are also produced by peroxisomes and some cellular enzymes, such as NADPH oxidases, cytochrome P450 oxidases and xanthine oxidases (Izyumov et al. , 2010; Lassègue et al. , 2012; Lismont et al. , 2015; Bezawork-Geleta et al. , 2017; Fransen et al. , 2017; Zhao et al. , 2019; Al-Shehri et al. , 2020; Wolf et al. , 2020; Zhang et al. , 2020). Although not a strong oxidizing agent, •O2is the precursor of all cellular ROS and is easily dismutated to H2O2 and O2 by superoxide dismutases (SOD) (Sies & Jones, 2020; Herb et al. , 2021). Hydrogen peroxide is considered to be the main ROS in redox regulation of biological activities, contributing to many cell processes, namely cell proliferation, differentiation, migration and angiogenesis (Sauer et al. , 2001; Jones & Sies, 2015; Sies et al. , 2017; Winterbourn, 2018). It is also involved in the formation of •OH, the most reactive ROS and an initiator of lipid peroxidation, by reacting with iron-sulfur (Fe-S) clusters through a metal-catalyzed Fenton reaction (Liochev & Fridovich, 1999; Imlay, 2003; Koppenol & Hider, 2019): Fe2+ + H2O2 → Fe3+ + •OH + OHWhen in physiological concentrations, ROS play many important cellular roles, such as intracellular redox signaling and contribution to blood pressure regulation, cognitive functions and immune responses (Yang et al. , 2013; Li et al. , 2017; Togliatto et al. , 2017; Zarse & Ristow, 2021). However, these are very reactive molecules, and a detoxification of ROS is crucial for cell survival, as they could cause harm when in uncontrolled concentrations – an imbalance between ROS cellular levels and the cell’s antioxidant response is referred to as oxidative stress (Bardaweel et al. , 2018). Antioxidant defense mechanisms have evolved in cells, allowing them to combat oxidative stress. As mentioned above, SOD converts •O2 into H2O2 and O2, but the enzyme catalase converts H2O2 into H2O and O2 in the peroxisomes (Alfonso-Prieto et al. , 2009). Glutathione peroxidase is another enzyme that also reduces
33 H2O2 into H2O (Chu et al. , 2020). Besides enzymatic defenses, there are many small molecule antioxidants that play a part in combating oxidative stress, such as glutathione, vitamin C (ascorbic acid), vitamin E (-tocopherol), carotenoids and flavonoids (Valko et al. , 2007). Regardless of the many defense systems, living aerobic organisms still suffer from oxidative damage, and oxidative stress is linked to a large variety of degenerative processes, diseases and syndromes (Pizzino et al. , 2017; Bardaweel et al. , 2018). Oxidative stress affects numerous biological mechanisms and structures, causing protein modification, inflammation, cell death, mitochondrial malfunction, among others, which can lead to and/or worsen serious pathologies, like Alzheimer’s disease, type 2 diabetes mellitus, chronic obstructive pulmonary disease, hypertension and cancer (Zinellu et al. , 2016; Butterfield & Halliwell, 2019; Oguntibeju, 2019; Barnes, 2020; Hayes et al. , 2020; Touyz et al. , 2020 Arfin et al. , 2021; Forman & Zhang, 2021; Misrani et al. , 2021; Bhatti et al. , 2022; Tain & Hsu, 2022). 1.2. Antioxidant activity of Pereiro propolis In this chapter, the focus will be on Pereiro propolis harvested in 2013 (P13). An ethanol extract of this sample (P13.EE) has been studied in previous works and has exhibited antioxidant properties (Marques et al. , 2015; Barroso et al. , 2017). P13.EE showed little effect on the viability of the yeast S. cerevisiae at the concentrations of 100, 200, and 500 μg/mL, but presented cytotoxicity at 750 μg/mL. These results were reported by Marques (2015) and their constancy was confirmed by Barroso (2017). Barroso (2017) also tested this extract mutagenicity with an Ames test and since P13.EE showed no significant differences when compared to the negative control, the author concluded that this extract is not mutagenic. Marques (2015) studied the effects of P13.EE on yeast viability under H2O2-induced oxidative stress, showing that this extract (at the concentrations of 50, 100 and 200 μg/mL) caused an increase in S. cerevisiae viability when compared to the H2O2 control, suggesting an antioxidant activity. Barroso (2017) assessed the antioxidant activity of P13.EE through in vitro assays, namely the DPPH radical scavenging activity at the concentrations of 100, 1000 and 10000 μg/mL, showing similar activity to the positive control ascorbic acid at the same concentrations of the extract, and the reducing power of Fe3+/ferricyanide complex to ferrous form, at the concentrations of 100, 500 and 1000 μg/mL, and although the results were significantly lower when compared to the positive control gallic acid (at the same concentrations of the extract), the extract still presented reducing power at all the tested concentrations. Both activities tested were shown to be dose-dependent and several studies have linked them to the presence of phenolic compounds and flavonoids (Moreira et al. , 2008; Petelinc et al. , 2013;
34 De Marco et al. , 2017). Barroso (2017) also tested the P13.EE Fe2+ chelating activity and plasmid DNA (pDNA) protection against Fe2+ damage. Remarkably, P13.EE was not able to chelate Fe2+ but presented a protective effect against the damage it caused on pDNA even at the lowest concentration of 10 μg/mL. It was suggested by the author that the DNA protection mechanism could be related to the extract ability to scavenge free radicals, since it is not able to directly chelate Fe2+. The combination of these results suggest that P13.EE has promising antioxidant properties, but it would be interesting to dive deeper into the study of such bioactivity. The aim of this chapter is to continue and complement this study, using the biological model S. cerevisiae to better understand the mechanism of action behind this antioxidant activity.
35 2. Materials and methods 2.1. Propolis samples and ethanol extract preparation The propolis sample used in this work was obtained from the Pereiro (P) apiary located at Beira Alta, Pinhel city, Guarda district (N 40° 44’57.135“, W 7° 0’59.403”, elevation: 672.6 m) – and was collected in the year 2013 (P13) from A. mellifera beehives by conventional scraping. An ethanol extract was then prepared from this sample, obtaining P13.EE, according to Alves (not published). Briefly, 100 mL of absolute ethanol (EtOH) (CARLO ERBA Reagents) were added to 15 g of raw propolis sample P13 in an Erlenmeyer and kept in an orbital shaker at 25 °C and 130 revolutions per minute (rpm), in the dark, for 24 h. This suspension was then filtered with Macherey-Nagel filter papers, utilizing a Buchner funnel and a Kitasato combined with a vacuum pump. The remaining residue went through a new extraction with 35 mL of absolute EtOH. The resulting filtrates were mixed and the EtOH was evaporated in a Büchi Rotavapor RE 121 with a water bath (Büchi 461), at 40 °C and 46 rpm. The dried ethanol extract was then kept in the dark, at -20 °C, until usage (Marques, 2015). Stock solutions were prepared with EtOH at the necessary concentrations for the different assays whenever necessary and kept at -20 °C. 2.2. Saccharomyces cerevisiae strains and conditions of media and culture growth The effects of P13.EE were evaluated using Saccharomyces cerevisiae BY4741 (Table 1). Mutants derived from this parent strain (Δ yap1, Δ sod1, Δ sod2 ) were also used in some assays (Table 1). Table 1. Yeast strains used in this work and their respective genotypes and origin. Cultures of the selected S. cerevisiae strains were prepared in YPD medium (1 % (w/v) yeast extract (DB BactoTM), 2 % (w/v) peptone (DB BactoTM), 2 % (w/v) glucose (Scharlau)) with incubation at 30 °C, 200 rpm and growth was monitored by optical density at 600 nm (OD600). Inocula for the liquid Strain Genotype Origin BY4741 MATa his3 Δ 1 leu2 Δ 0 met15 Δ 0 ura3 Δ 0 Euroscarf Δ yap1 BY4741; MATa his3 Δ 1 leu2 Δ 0 met15 Δ 0 ura3 Δ 0 YML007w::kanMX4 Euroscarf Δ sod1 BY4741; MATa his3 Δ 1 leu2 Δ 0 met15 Δ 0 ura3 Δ 0 YJR104c::kanMX4 Euroscarf Δ sod2 BY4741; MATa his3 Δ 1 leu2 Δ 0 met15 Δ 0 ura3 Δ 0 YHR008c::kanMX4 Euroscarf
36 cultures were obtained from single colonies cultured on solid YPDA medium (YPD medium complemented with 2 % (w/v) agar (LabChem Inc.)) at 30 °C for 48 h. Cultures in solid media were kept at 4 °C. To prepare cells for the different assays, liquid cultures were grown overnight at 30 °C and 200 rpm and then diluted in fresh medium to an OD600 of 0.1. The culture was then incubated under the same conditions for 4 h, guaranteeing two generations at exponential phase (OD600 between 0.4 and 0.8). 2.3. Evaluation of cellular viability A S. cerevisiae BY4741 YPD-culture was prepared as described above and after OD600 confirmation, aliquots of the cell suspension were placed in microtubes and different volumes of P13.EE were added, at the final concentrations of 200, 500 or 750 μg/mL. A negative control was prepared using the same volume of EtOH as for the 750 μg/mL P.EE. The cells were then incubated at 30 °C and 200 rpm for 90 min and at time-points 0, 30, 60 and 90 min, aliquots of 100 μL were collected and serially diluted from 10-1 to 10-4 in sterilized distilled H2O (dH2O) and 5 μL drops of each dilution were placed on YPDA medium plates, which were inverted and incubated at 30 °C for 48 h, in the dark. Photographs were made using ChemiDocTMXRS. Viability assays were also performed with S. cerevisiae BY4741 mutants Δ yap1 , Δ sod1 and Δ sod2 , to analyze the effect of P13.EE at the same concentrations tested for the parent strain. 2.4. Evaluation of intracellular oxidation A S. cerevisiae BY4741 YPD-culture was prepared as described above and, after OD600 confirmation, it was centrifuged at 4 °C and 4,400 g , for 3 min (Eppendorf Centrifuge 5804 R), and the pellet washed two times with and resuspended in PBS (137 mM NaCl (Honeywell FlukaTM), 2.7 mM KCl (Honeywell FlukaTM), 4.3 mM Na2HPO4 (PanReac AppliChem), 1.47 mM KH2PO4 (Merck KGaA); pH 7.4). The cell suspension was diluted to an OD600 of 0.04 in a volume of 10 mL, and 2 mL were collected for autofluorescence controls. Fifty micromolar 2’,7’-dichlorodihydrofluorescein diacetate (H2DCFDA; Molecular Probes) was added to the remaining suspension and it was incubated at 30 °C and 200 rpm, in the dark, for 60 min. The cell suspension was then distributed into microtubes, P13.EE was added to the final concentrations of 50, 100, 200 or 500 μg/mL, and H2O2 (CARLO ERBA Reagents) was added to a final concentration of 5 mM. Negative and positive controls were prepared, respectively, with absolute EtOH (same volume as for the 500 μg/mL P.EE) and 5 mM H2O2. The cells were then incubated at 30 °C and 200 rpm, in the dark, for 20 min. The samples were placed on a flat bottom 96-well plate and
43 (A) (B) (C) (D) (E) (F) Figure 6. P13.EE decreases intracellular oxidation of cells from S. cerevisiae mutant strains Δ yap1 , Δ sod1 , and Δ sod2 . H2DCFDA-loaded cells from the S. cerevisiae mutant strains Δ yap1 (A-B), Δ sod1 (C-D), and Δ sod2 (E-F) , were treated with P13.EE at 50 (P13_50), 100 (P13_100), 200 (P13_200), or 500 μ g/mL (P13_500) and with 5 mM H2O2, and incubated at 30 °C for 20 min. For the negative control (C-) cells were treated with absolute EtOH, at the same volume as for 500 μ g/mL P13.EE, and for the positive control (C+) cells were treated with 5 mM H2O2. (A, C, E) Histograms of one representative experiment out of three independent replicates. (B, D, F) Percentage of cells with decreased intracellular oxidation. A gating was drawn at the peak of C+ to obtain the percentage of cells exhibiting less fluorescence emission (cells to the left of the gating). These values represent the mean ± SD of n = 3 independent replicates. One-way ANOVA was performed, followed by Dunnett’s test for multiple comparisons. Asterisks represent significant differences between treatments and C+ (* p < 0.05, ** p < 0.01).
44 Similarly to what was observed in the parental strain (Figure 5), cells of Cdisplayed very low levels of fluorescence, while cells exposed to H2O2 (C+) exhibited higher fluorescence, suggesting the presence of oxidative stress. Again, all of the mutant cells treated with P13.EE at the concentrations of 50 and 100 μg/mL did not exhibit significantly different fluorescence emission when compared to the C+ cells (Figures 6B, 6D and 6F), except for the Δ yap1 mutant, where the percentage of cells to the left of the C+ peak was significantly lower than the percentage of C+ cells itself, suggesting that, in this strain, the lower concentrations of P13.EE not only did not decrease intracellular oxidation, but they increased it. Reports of this effect were not found in the literature, but there is a possibility of a synergism occurring between the extract and H2O2, which appears to cease when the extract is at a concentration of 200 and 500 μg/mL. Further studies are required in order to confirm and interpret these results, possibly with isolated compounds of P13.EE. Contrastingly to what was observed in the parent strain, cells treated with the 200 μg/mL P13.EE did not exhibit a significant decrease in fluorescence emission when compared to the C+, however, P13.EE at 500 μg/mL still caused a significantly lower fluorescence emission on all of the tested strains, with a percentage of cells to the left of the C+ peak of 89.26 ± 5.41 % for Δ yap1 , 92.72 ± 6.06 % for Δ sod1 , and 97.28 ± 4.22 % for Δ sod2 , suggesting that it is causing a decrease in the intracellular oxidation of all of the mutants tested. The difference in the results from the parent strain and the mutants suggest that the antioxidant activity of P13.EE may be involved in the direct scavenging of free radicals, since it still is able to decrease intracellular oxidation when genes involved in the oxidative stress response are deleted, but they also suggest that the extract may additionally interact and activate the yeast antioxidant mechanisms that these genes are involved in, since without them, the extract is not able to have the same antioxidant effect. Additionally, since the yeast’s oxidative stress response involves several proteins, like the mentioned Sod1, Sod2 and Yap1, but also Ctt1, a catalase (Petrova et al. , 2004), the studied mutants are not entirely incapable of responding to oxidative stress, as they only lack a single specific protein and the rest remain active. 3.3. Chemical characterization of phenolic content from P13.EE Since P13.EE had not yet been chemically characterized, an UPLC-DAD-ESI/MS analysis was carried out in order to assess its chemical profile of phenolic compounds. This type of analysis allows for the identification of these compounds, by the determination of their chromatographic behavior, UV spectra, and mass spectrometry. Figure 7 represents the chromatographic profile of P13.EE at 280 nm, where 21 fractions were identified. These fractions were further analyzed by electrospray ionization/mass
45 spectrometry (ESI/MS), allowing for the identification of the probable compounds each fraction represents (Table 2), by comparison with data published in the literature. From the ESI/MS analysis, it was possible to suggest several phenolic compounds present in the P13.EE extract (Table 2). Although the most abundant compound in the extract is caffeic acid isoprenyl ester (peaks 9 and 10), several other compounds were identified, such as caffeic acid, p -coumaric acid and derivatives, ferulic acid, apigenin, pinocembrin, pinobanksin and derivatives, galangin, and derivatives of caffeic acid, like CAPE, along with some compounds which had not been referred in the literature but were also present in the ethanol extract of a propolis sample harvested in 2010 from the same apiary (peaks 18 and 19; Silva-Carvalho et al. , 2014). These compounds are commonly present in propolis samples and are associated with antioxidant activity (Russo et al. , 2002; Kowalczyk et al. , 2017; Okińczyc et al. , 2021; Kurek-Górecka et al. , 2022; Xu et al. , 2022). Furthermore, these results support the observed anti and pro-oxidant dose-dependent antagonistic effects of P13.EE, as the presence of CAPE along with antioxidants like pinobanksin and pinocembrin have been linked to this effect by Cruz et al. (2016). Table 2. Chemical characterization of phenolic content of P13.EE obtained by UPLC-DAD-ESI/MS. Peak tR (min) λmax (nm) [M - H] - Probable compound 1 2.2 216, 231sh, 296sh, 322 179 Caffeic acid 2 3.2 208, 224, 296sh, 309 163 p -Coumaric acid 3 4.0 216, 239sh, 297sh, 323 193 Ferulic acid 4 7.4 216, 236, 295sh, 321 207 3,4-Dimethyl-caffeic acid (DMCA) 5 11.1 226, 308 n.d. p -Coumaric acid derivative Figure 7. Chromatographic profile of P13.EE at 280 nm obtained by UPLC-DAD-ESI/MS. Numbers in the figure represent the fractions that were further analyzed by ESI/MS. 0 25 50 75 100 0 5 10 15 20 25 30 35 40 45 50 1 3 4 5 6 7 8 9 12 13 14 15 17 18 20 21
46 The chromatographic profile of the extract from the 2010 sample (P10.EE) was also analyzed in the same conditions as for the P13.EE (Figure A1, Appendix 1), for the purpose of comparing the two. Both extracts display very similar chromatographic profiles, suggesting that their differences in biological activities, which will be exhibited in the next chapter, are probably not related to their phenolic content, at least not in a qualitative manner. Additionally, total flavonoid and phenolics contents have been estimated for both extracts, with P13.EE having a total phenolic content (TPC) of 217.6 ± 6.6 mg gallic acid equivalent/g extract and a total flavonoid content (TFC) of 38 ± 2.0 mg quercetin equivalent/g extract (Peixoto et al. , 2022), and P10.EE with a TPC of 252.4 ± 13.4 mg gallic acid equivalent/g extract and TFC of 51.3 ± 2.4 mg quercetin/g extract (Silva-Carvalho et al. , 2014). Although comparison of TPC and TFC of both extracts should be cautious, since different compounds contribute to these chemical classes, both extracts present values in the same range, further supporting the conclusion that the biological activity differences might be due to other chemical families. 6 12.2 196, 209, 267, 294sh, 336 269 Apigenin 7 12.4 212, 226sh, 291 271 Pinobanksin 8 17.4 236, 311 313 Unknown 9 21.6 211, 245, 270, 297sh, 325 247 Caffeic acid isoprenyl ester 10 22.4 215, 236, 297sh, 325 247 Caffeic acid isoprenyl ester 11 23.0 210, 226sh, 289, 326 255 Pinocembrin 12 23.6 231, 266, 290sh, 355 269 Galangin 13 25.1 211, 227sh, 293 313 Pinobanksin-3O -acetate 14 25.6 243, 297sh, 325 283 Caffeic acid phenylethyl ester 15 28.7 234, 311 231 p -Coumaric acid isoprenyl ester 16 29.5 235, 311 231 p -Coumaric acid isoprenyl ester 17 30.0 238, 291sh, 325 231 p -Coumaric acid isoprenyl ester 18 35.3 211, 268, 309 387 621 Unknown Unknown 19 35.7 211, 268, 292, 357 279 387 Unknown Unknown 20 36.3 239, 268, 292, 357 341 621 Pinobanksin-3O -butyrate or isobutyrate Unknown 21 38.5 236, 269, 396sh, 344 565 p -Coumaric acid-4-hydroxyphenylethyl
47 4. Conclusions Antioxidant activity in natural products is usually related to the presence of polyphenolic compounds (Balasundam at al., 2006). Polyphenols have exhibited many beneficial effects on human health due to their antioxidant activity (Pandey & Rizvi, 2009), therefore natural products with this property are of great interest to the pharmaceutical and cosmetic industries. Propolis is well known for its strong antioxidant activity, due to its high polyphenolic content, making it a great contender for these industries. From the work presented in this chapter, it can be concluded, through viability assays, that P13.EE was able to maintain its low cytotoxic effect on S. cerevisiae over the years of storage (Figure 4A), and that effect appears dose-dependent, exhibiting a slight cytotoxic effect at higher concentrations. This observation was corroborated with the results from viability assays with the Δ yap1 , Δ sod1, and Δ sod2 mutants (Figures 4B, 4C and 4D), which suggested that P13.EE could be having a pro-oxidant effect at the concentration of 750 μg/mL. These effects could be related to the presence of CAPE, and antioxidants like pinobanksin and pinocembrin in the extract (Table 2), as reported by Cruz et al. (2016) Results from intracellular oxidation assays with S. cerevisiae BY4741 strain (Figure 5) are in accordance with the antioxidant activity reports previously made by Marques (2015) and Barroso (2017), as P13.EE was shown to reduce H2O2-induced intracellular oxidation at the concentrations of 200 and 500 μg/mL. However, the extract was less effective in the Δ yap1 , Δ sod1, and Δ sod2 mutants (Figure 6), suggesting that although the extract is able to directly scavenge free radicals, it may also interact with the genes YAP1 , SOD1 and SOD2 and/or with the proteins encoded by these genes for its antioxidant activity, possibly aiding in the activation of the yeast’s antioxidant mechanisms, as these genes are involved in the yeast’s oxidative stress response and each of the studied strains only lacked one specific gene, meaning that the rest are active and the cells are not entirely incapable of responding to oxidative stress.
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52 CHAPTER 3. Cytotoxic activity of Pereiro propolis
59 2.5. Evaluation of plasma membrane integrity with propidium iodide staining Aliquots of a S. cerevisiae BY4741 YPD-culture prepared as described above and with OD600 = 0.4 were placed in microtubes and P10.EE was added to one of the microtubes, at the final concentration of 500 μg/mL. A negative control as well as apoptosis and necrosis controls were prepared as described (section 2.4. of the present chapter). The cells were then incubated at 30 °C and 200 rpm for 90 min and at time points 0, 30, 60, and 90 min, aliquots of 100 μL of each sample were stained with propidium iodide (PI; Sigma-Aldrich) at a final concentration of 2 μg/mL and incubated at room temperature for 10 min in the dark. The cells were observed using a Leica DM-5000B fluorescence microscope (Leica Microsystems), with the red channel filter and a 100x oil-immersion objective. Images were acquired with a Leica DCF350FX digital camera and processed with LAS AF Leica Microsystems software. 2.6. Analysis of cell cycle progression A S. cerevisiae BY4741 YPD-culture was prepared as described and, after OD600 confirmation, the yeast cell cycle was synchronized with the addition of hydroxyurea (HU; Sigma-Aldrich) at a final concentration of 200 mM, causing an arrest in early S phase (Rosebrock, 2017b). The cells were then incubated at 30 °C and 200 rpm for 2.5 h. To release the cells from the arrest, the culture was centrifuged at 4 °C and 4,400 g , for 3 min (Eppendorf Centrifuge 5804 R), the pellet washed two times with ultrapure H2O (upH2O) and resuspended in fresh YPD medium. Aliquots of 500 μL were collected before and after synchronization, for controls. The synchronized cell suspension was divided into Falcon® tubes and P10.EE was added to one of the tubes, at the final concentration of 200 μg/mL. A negative control was prepared with EtOH (same volume as for the P.EE). The cells were then incubated at 30 °C and 200 rpm for 180 min and at time points 0, 30, 60, 90, 120, and 180 min, aliquots of 500 μL were collected and centrifuged at 2,500 g for 4 min (Eppendorf 5453 MiniSpin Plus Centifuge). The supernatant was discarded and the pelleted cells were fixed with 70 % EtOH (added drop by drop, with frequent vortexing) and incubated at -20 °C overnight (Rosebrock, 2017a). After fixation, the samples were centrifuged at 10,000 g for 20 min (Eppendorf 5453 MiniSpin Plus Centifuge), and the pellets were resuspended in 500 μL of 50 mM sodium citrate buffer (50 mM Trisodium citrate dihydrate (PanReac AppliChem; pH 7.2) and incubated for 10 min at room temperature. The cells were centrifuged at 5,000 g for 5 min (Eppendorf 5453 MiniSpin Plus Centifuge), the supernatant discarded, and the washing was repeated. The resulting pellet was then resuspended in in
60 500 μL of 50 mM sodium citrate buffer (pH 7.2) with 20 μg/mL RNase A (PanReac AppliChem) and 2.5 μM SYTOXTM Green Nucleic Acid Stain (Invitrogen) and the cells were incubated at 37 °C for 60 min. Afterwards, 10 μL of 20 mg/mL proteinase K (Alfa Aesar) was added to the samples and incubated at 55 °C for 60 min followed by incubation at 4 °C overnight (Rosebrock, 2017a). Finally, the samples were sonicated in an ultrasonic bath, for 30 s at the highest intensity, and then placed on a flat bottom 96-well plate and analyzed in a flow cytometer (Beckman Coulter CytoFLEX V0-B4-R2), using a blue laser for excitation at 488 nm and the FITC filter. Data analysis was done with the program CytExpert 2.5. To confirm that the cells used for the cell cycle progression analysis were alive, a viability assay was conducted. At each time point, aliquots of 100 μL were collected and serially diluted from 10-1 to 10-4 in sterilized distilled H2O (dH2O) and 3 drops of 40 μL from the 10-4 dilution were placed on YPDA medium plates. After incubation at 30 °C for 48 h, in the dark, the colonies were counted to assay the viability of the culture over time. Viability of each time-point was calculated as the percentage of colonyforming units (CFU), taking time 0 min as 100 %. 2.7. Statistical analysis Unless otherwise stated, the experiments were done in at least three independent replicas, and the results are either presented as one representative experiment or as the mean ± SD. For all of the statistical tests, a 95 % confidence interval was assumed. Data was analyzed with Two-way ANOVA, followed by post hoc tests for multiple comparisons (Tukey or Sidak). The significance of differences between the mean values is represented by asterisks (* p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001). Data was analyzed and graphs were created using GraphPad Prism 8.0.
61 3. Results and discussion 3.1. Evaluation of cellular viability – P10.EE maintains its cytotoxicity against Saccharomyces cerevisiae As mentioned in this chapter, besides exhibiting low antioxidant activity, P10.EE stands out for its cytotoxic activity, when compared to EEs of propolis samples from the same apiary but from different years. Marques (2015) assessed the cytotoxic effect of this extract through viability assays, treating a YPD-culture of S. cerevisiae BY4741 cells with P.EE, collecting aliquots after 0, 30, 60 and 90 min of treatment and plating 10-1 to 10-4 dilutions of each aliquot on YPDA plates. This sample was collected in 2010, and its EE has been stored in a Falcon® tube at -20 °C since it was prepared, in 2011. In order to confirm if P10.EE maintained its effect on the viability of S. cerevisiae , after a prolonged storage, this assay was replicated in the present study (Figure 9). A cytotoxic effect of P10.EE can be observed by a decrease in number of colonies when compared to the C- (cells treated with absolute EtOH at the same volume as for the P10.EE), which is very clear at 60 and 90 min incubation with 500 μg/mL and at 30, 60 and 90 min with 750 μg/mL. As observed in Figure 9, P10.EE shows a strong cytotoxic effect at the concentrations of 500 and 750 μg/mL, visibly decreasing the viability of S. cerevisiae after 60 min of incubation for the concentration of 500 μg/mL and after 30 min for 750 μg/mL. These results are similar to the ones observed by Marques (2015), confirming that P10.EE was able to maintain its activity over the many years of storage at -20 °C. 10-1 10-2 10-3 10-4 10-1 10-2 10-3 10-4 10-1 10-2 10-3 10-4 10-1 10-2 10-3 10-4 0 30 60 90 Dilutions Time (min) Figure 9. Effects of P10.EE on the viability of S. cerevisiae . Cells from the S. cerevisiae BY4741 strain were treated with P10.EE at the concentrations of 200, 500, or 750 μ g/mL. For the negative control (C-) cells were treated with absolute EtOH, at the same volume as for the P10.EE. Aliquots were collected after 0, 30, 60, and 90 min, and serially diluted from 10-1 to 10-4, and 5 μ L drops of each dilution were placed on YPDA plates and incubated at 30 °C for 48 h. Cytotoxicity is observed by a decrease in number of colonies when compared to C-. C200 μg/mL 500 μg/mL 750 μg/mL
62 3.2. Evaluation of apoptotic characteristics with 4’,6-diamidino-2-phenylindole staining – P10.EE does not seem to induce apoptosis in Saccharomyces cerevisiae Previous results from Marques (2015) and Alves (2018) suggested that the cytotoxic activity exhibited by P10.EE is involved in a cell death inducing mechanism, but they were not conclusive on the type of cell death, as viability and DNA damage assays with mutants treated with the extract suggested apoptotic cell death (Marques, 2015) and the nucleo-cytosolic release of the Nhp6A protein on cells treated with P10.EE, observed by Alves (2018), suggested necrotic cell death. Yeast apoptotic characteristics, such as chromatin condensation and fragmentation, can be observed through fluorescence microscopy with the use of DNA fluorescent dyes, like DAPI (CarmonaGutierrez at al., 2018). DAPI is a blue fluorescent dye that binds to A-T rich regions in DNA, forming a fluorescent complex. However, DAPI does not pass efficiently through the cell membrane, therefore cells need to be fixed before staining (Wloch-Salamon & Bem, 2013). In order to analyze the effect of P10.EE on yeast cell nuclear morphology, S. cerevisiae BY4741 cells were treated with P10.EE at the concentration of 500 μg/mL and incubated for 90 min. Aliquots were collected at 0, 30, 60 and 90 min, and the cells were heat-fixed and stained with DAPI. For this assay, three controls were prepared: the negative control, with cells treated with absolute EtOH at the same volume as for the P10.EE; the apoptosis control, with cells treated with H2O2 at 10 mM (Ribeiro et al. , 2006); and the necrosis control, with cells treated with H2O2 at 180 mM (Madeo et al. , 1999). For each time-point and treatment, a total of 100 cells were randomly counted, and out of those, cells presenting chromatin condensation and/or fragmentation were counted. Graphical representation of these cells is shown in Figure 10.
63 As expected, the necrosis control showed no significant differences to the Cfor the percentage of cells presenting apoptotic features, at all the tested time-points, contrarily to what was observed for the apoptosis control, that presented 43.00 ± 2.00 % of cells with apoptotic features at 90 min of treatment, a value significantly higher when compared to the Cat the same time-point. Similarly to the Cand the necrosis control, cells treated with P10.EE did not exhibit apoptotic features at any of the tested timepoints, showing no significant differences to the Cat all of the time-points. . Images of a representative experiment obtained from bright filed (BF) and fluorescence microscopy of the yeast cells at 90 min of treatment can be observed in Figure 11, where chromatin condensation and fragmentation are clearly visible. Images of all the treatments from the time-points 0, 30, and 60 min of this representative experiment can be observed in Figures A2-4 (Appendix 2). These results suggest that P10.EE is not inducing cell death in S. cerevisiae through an apoptotic mechanism. Figure 10. P10.EE-treated S. cerevisiae cells do not present apoptotic features. Percentage of S. cerevisiae BY4741 cells presenting apoptotic features, such as chromatin condensation and/or fragmentation, after 0, 30, 60, and 90 min of incubation with 500 μ g/mL P10.EE. At each time-point, aliquots were collected and the cells were heat-fixed and stained with DAPI. For the negative control (C-) cells were treated with absolute EtOH, at the same volume as for the P10.EE, for the apoptosis control (C Apoptosis) cells were treated with H2O2 at 10 mM, and for the necrosis control (C Necrosis) cells were treated with H2O2 at 180 mM. These values represent the mean ± SD of n = 3 independent replicates. Two-way ANOVA was performed, followed by Tukey’s test for multiple comparisons. Asterisks represent significant differences between treatments and C- (**** p < 0.0001).
64 BF DAPI BF + DAPI C90 min C Apoptosis C Necrosis P10.EE Figure 11. Analysis of chromatin condensation and nuclei fragmentation through fluorescence microscopy. S. cerevisiae BY4741 cells treated with absolute EtOH (C-), 10 mM H2O2 (C Apoptosis), 180 mM H2O2 (C Necrosis) and 500 μ g/mL P10.EE, after 90 min of incubation, were heat-fixed and stained with DAPI. Images were obtained through BF and fluorescence microscopy (magnification of 100x) and are from one representative experiment, out of three independent replicates.
65 3.3. Evaluation of plasma membrane integrity with propidium iodide staining – P10.EE leads to loss of plasma membrane integrity in Saccharomyces cerevisiae Since P10.EE does not appear to be causing apoptotic cell death in S. cerevisiae , its effect in the cell membrane integrity was assessed, as loss in membrane integrity is a marker associated with necrotic cell death. For this assay, cells were stained with PI and analyzed through fluorescence microscopy, as PI is a red fluorescent dye that binds to DNA and it is commonly used to detect dead cells, because it is cell-impermeant and can only enter the cell when the plasma membrane has lost its integrity (Büttner et al. , 2007). Saccharomyces cerevisiae BY4741 cells were treated with P10.EE at the concentration of 500 μg/mL and incubated for 90 min. Aliquots were collected at 0, 30, 60 and 90 min, and the cells were stained with PI. Negative, apoptosis and necrosis control treatments were as described for the previous assay. For each time-point and treatment, a total of 100 cells were randomly counted, and out of those, cells emitting red fluorescence (PI positive cells) were counted. Graphical representation of these cells is shown in Figure 12. As expected, the apoptosis control showed no significant differences to the Cfor the percentage of PI positive cells, at all the tested time-points, contrarily to what was observed for the necrosis control, that presented almost 100 % of PI positive cells at 30, 60, and 90 min of treatment, which are values significantly higher when compared to the Cat the same time-point. Cells treated with P10.EE exhibited Figure 12. P10.EE-treated S. cerevisiae cells exhibit loss of membrane integrity. Percentage of S. cerevisiae BY4741 PI positive cells, after 0, 30, 60, and 90 min of incubation with 500 μ g/mL P10.EE. At each time-point, aliquots were collected and the cells were stained with PI. For the negative control (C-) cells were treated with absolute EtOH, at the same volume as for the P10.EE, for the apoptosis control (C Apoptosis) cells were treated with 10 mM H2O2, and for the necrosis control (C Necrosis) cells were treated with 180 mM H2O2. These values represent the mean ± SD of n = 4 independent replicates. Two-way ANOVA was performed, followed by Tukey’s test for multiple comparisons. Asterisks represent significant differences between treatments and C- (**** p < 0.0001).
66 a gradual increase in percentage of PI positive cells over the tested time-points, reaching a percentage of 37.33 ± 4.61 % at 90 min of treatment, which is a value significantly higher when compared to the Cat that time-point. Images of a representative experiment obtained from bright filed (BF) and fluorescence microscopy of the yeast cells at 90 min of treatment can be observed in Figure 13, where PI positive cells are clearly visible in the necrosis control and P10.EE treated cells. Images from the time-points 0, 30, and 60 min of this representative experiment can be observed in Figures A5-7 (Appendix 2). These results, together with the results from the previous assays and the ones reported by Alves (2018), suggest that P10.EE could be inducing cell death in S. cerevisiae through a necrotic mechanism. Yeast necrotic cell death induced by propolis has already been observed by Castro et al. (2011) in a Brazilian green propolis ethanol extract, that although initially induced apoptosis, an increased exposure to propolis caused an increase in necrotic cells. The author suggested that these effects were partially mediated by YCA1 metacaspase, as a mutant with a deletion of this gene proved to be more resistant to propolis, which goes accordingly to the results reported by Marques (2015).
67 3.4. Analysis of cell cycle progression – P10.EE provokes an arrest in the cell cycle of Saccharomyces cerevisiae The yeast cell cycle, like in other eukaryotes, is a tightly controlled process divided into four different phases: G1, where cells prepare for duplication; S, where cells begin to duplicate genetic information; G2, where cells get ready to be divided; and M, where cells are divided into two “daughter” BF PI BF + PI 90 min CC Apoptosis C Necrosis P10.EE Figure 13. Analysis of plasma membrane integrity through fluorescence microscopy. S. cerevisiae BY4741 cells treated with absolute EtOH (C-), 10 mM H2O2 (C Apoptosis), 180 mM H2O2 (C Necrosis) and 500 μ g/mL P10.EE, after 90 min of incubation, were stained with PI. Images were obtained through BF and fluorescence microscopy (magnification of 100x) and are from one representative experiment, out of four independent replicates.
68 cells (Jiménez et al. , 2015). During the normal progression of the cell cycle, there are various checkpoints that can cause an arrest or a delay in the cell cycle if DNA damage is detected (Demeter et al. , 2000). Marques (2015) has previously reported that P10.EE causes significant DNA damage, and to test if this damage affects the progression of the yeast cell cycle, S. cerevisiae BY4741 cells treated with P10.EE were analyzed through flow cytometry. In this assay, the tested concentration of extract was 200 μg/mL, as it does not affect the yeast viability and cells should be viable for a reliable cell cycle analysis. A viability test was conducted alongside every independent replicate of this assay, to confirm the viability of the treated cells (Figure A8, Appendix 2), and showed that cells were alive during the whole assay meaning that an arrest in the cell cycle would not be due to cell death, but instead to repair damage allowing for the cell survival. Firstly, the yeast cell cycle was synchronized with HU at a concentration of 200 mM – HU inhibits ribonucleotide reductase (RNR), causing an arrest in early S phase/G1 phase (Juanes, 2017). After the synchronization, cells were released from the arrest and treated with P10.EE. At different time-points, aliquots were collected and prepared as described for staining with SYTOXTM Green Nucleic Acid Stain (Invitrogen) and flow cytometry analysis. SYTOXTM Green Nucleic Acid Stain (Invitrogen) is a green fluorescent dye that binds to nucleic acids and allows for the identification of cell cycle phases through flow cytometry, as the amount of fluorescence is directly proportional to the amount of DNA and number of cells. For the C-, cells were treated with absolute EtOH at the same volume as for the P10.EE. From the histograms obtained (Figure A9, Appendix 2), and with the CytExpert 2.5 software, three gates were drawn to define the percentage of cells in the phases G1, S and G2/M, based on the amount of fluorescence and number of cells on the unsynchronized cells – a first peak with less fluorescence represents G1 phase, a second peak with higher amount of fluorescence represents G2/M phases, and the cells in between the two peaks represent S phase (Figure 14A). Graphical representation of this distribution for all of the tested time-points is shown in Figure 14B. Percentage of P10.EE cells in a certain cell cycle phase was compared to the percentage of Ccells in the same phase.
75 CHAPTER 4. Final remarks and future perspectives
76 1. Final remarks and future perspectives Propolis is a natural product produced by honeybees that has been recognized worldwide for its many biological properties, such as antioxidant, cytotoxic, and antimicrobial activities, yet depending on a very diverse chemical composition. Because of this plethora of biological activities, the demand for propolis research for potential pharmaceutical, medical, cosmetic, and food applications has been growing. Despite this increasing demand, Portuguese propolis research has been stagnant for many years, only starting to slowly gain ground recently. However, it is still an undervalued product in Portugal, creating a necessity to further investigate this Portuguese beehive product and its mechanism of action, in order to contribute to its national and international recognition and economic valorization. The aims of this work were to further study the intriguing differences previously observed in the ethanol extracts of two Portuguese propolis samples from the Pereiro apiary, P10.EE and P13.EE, by Marques (2015), Barroso (2017) and Alves (2018), diving deeper into P13.EE antioxidant activity and P10.EE cytotoxic activity, in an attempt to better understand their distinct mechanisms of action and what are the causes for these differences in samples from the same apiary. The present work was able to successfully confirm these activities and infer on their modes of action, although it is still not clear what the differences in the extracts’ chemical compositions are. As it is proposed in this work that such difference is not related to their phenolic content, a gas chromatography-mass spectrometry (GC-MS) analysis could be performed on both P13.EE and P10.EE, allowing for a more complete chemical characterization of the extracts, namely of different types of compounds like terpenes, which have already been associated with propolis antimicrobial (e.g. isocupressic acid, diphenylheptanoids) and anticancer (e.g. clerodane diterpenes) activities (reviewed by Aminimoghadamfarouj & Nematollahi, 2017). Although the conjunction of the results from this and the previous works allow for a much clearer view on the different extracts’ mechanisms of action, other approaches can be followed to further study and clarify these mechanisms. For the P13.EE, an intracellular oxidation assay could be done on an S. cerevisiae mutant with a combined deletion of the YAP1 , SOD1 and SOD2 genes, to observe how the P13.EE ability to decrease the yeast intracellular oxidation would be affected. For the P10.EE, an Annexin V/PI flow cytometry assay could be done, in order to confidently confirm the occurrence of necrotic cell death. Nonetheless, taken all together, the results observed for the assays with P13.EE suggest promising antioxidant properties, that could be further explored for possible pharmaceutical, cosmetic, and food applications. P10.EE cytotoxic activity, possibly through a necrotic pathway, could be of great interest for antitumoral applications or the development of anticancer drugs, as it has already been
77 reported by Silva-Carvalho et al. (2014) to decrease cell viability of the tumor cell lines MDA-MB-231 and DU145 while being less cytotoxic to non-tumoral cells, and especially since apoptosis resistance is becoming a problem during cancer treatment, as the conventional therapeutic pathway is pro-apoptotic (Galluzzi et al. , 2011; Gong et al. , 2019). Indeed, the development of a non-apoptotic treatment would be of an enormous scientific and medical significance. However, P10.EE effect needs to be further investigated in different cellular models, such as mammalian cell lines and cancer cell lines, to really comprehend how the extract works in more advanced organisms. Furthermore, since P10.EE exhibits cytotoxicity against S. cerevisiae , an application in biopesticides could be considered, after further studies on its antifungal activity.
78 2. References Alves, H. (2018). Genotoxicity of propolis extracts from Beira Alta: insights on the mode of action, using different biological models. (Master thesis). Department of Biology, School of Sciences, University of Minho, Braga, Portugal. Aminimoghadamfarouj, N., & Nematollahi, A. (2017). Propolis diterpenes as a remarkable biosource for drug discovery development: a review. International Journal of Molecular Sciences , 18 (6), 1290. Barroso, L. (2017). Analysis of the effects of propolis extracts on DNA damage and mutagenicity. (Master thesis). Department of Biology, School of Sciences, University of Minho, Braga, Portugal. Galluzzi, L., Vitale, I., Vacchelli, E., & Kroemer, G. (2011). Cell death signaling and and anticancer therapy. Frontiers in Oncology, 1 , 5. Gong, Y., Fan, Z., Luo, G., Yang, C., Huang, Q., Fan, K, Cheng, H, Jin, K., Ni, Q., Yu, X., & Liu, C. (2019). The role of necroptosis in cancer biology and therapy. Molecular Cancer, 18 , 100. Marques, R. (2015). Contributos para a elucidação do modo de ação de própolis português: o caso do própolis do Pereiro. (Master thesis). Department of Biology, School of Sciences, University of Minho, Braga, Portugal. Silva-Carvalho, R., Miranda-Gonçalves, V., Ferreira, A. M., Cardoso, S. M., Sobral, A. J. F. N., Almeida-Aguiar, C., & Baltazar, F. (2014). Antitumoural and antiangiogenic activity of Portuguese propolis in in vitro and in vivo models. Journal of Functional Foods , 11 , 160–171.
79 APPENDIX 1
80 Figure A1. Chromatographic profile of P10.EE at 280 nm obtained by UPLC-DAD-ESI/MS.
81 APPENDIX 2
82 BF DAPI BF + DAPI C0 min C Apoptosis C Necrosis P10.EE Figure A2. Analysis of chromatin condensation and nuclei fragmentation through fluorescence microscopy . S. cerevisiae BY4741 cells treated with absolute EtOH (C-), 10 mM H2O2 (C Apoptosis), 180 mM H2O2 (C Necrosis) and 500 μ g/mL P10.EE, after 0 min of incubation, were heat-fixed and stained with DAPI. Images were obtained through BF and fluorescence microscopy (magnification of 100x) and are from one representative experiment, out of three independent replicates.
83 BF DAPI BF + DAPI C30 min C Apoptosis C Necrosis P10.EE Figure A3. Analysis of chromatin condensation and nuclei fragmentation through fluorescence microscopy . S. cerevisiae BY4741 cells treated with absolute EtOH (C-), 10 mM H2O2 (C Apoptosis), 180 mM H2O2 (C Necrosis) and 500 μ g/mL P10.EE, after 30 min of incubation, were heat-fixed and stained with DAPI. Images were obtained through BF and fluorescence microscopy (magnification of 100x) and are from one representative experiment, out of three independent replicates.
84 BF DAPI BF + DAPI C60 min C Apoptosis C Necrosis P10.EE Figure A4. Analysis of chromatin condensation and nuclei fragmentation through fluorescence microscopy . S. cerevisiae BY4741 cells treated with absolute EtOH (C-), 10 mM H2O2 (C Apoptosis), 180 mM H2O2 (C Necrosis) and 500 μ g/mL P10.EE, after 60 min of incubation, were heat-fixed and stained with DAPI. Images were obtained through BF and fluorescence microscopy (magnification of 100x) and are from one representative experiment, out of three independent replicates.
91 Figure A10. Synchronization of S. cerevisiae cell cycle. Percentage of S. cerevisiae BY4741 cells in the cell cycle phases G1, S, and G2/M, after synchronization of the cells with HU at 200 mM. Synchronized and unsynchronized cells were prepared for flow cytometry analysis and stained with SYTOXTM Green Nucleic Acid Stain (Invitrogen). These values represent the mean ± SD of n = 3 independent replicates. Two-way ANOVA was performed, followed by Sidak’s test for multiple comparisons. Asterisks represent significant differences between synchronized and unsynchronized cells (* p < 0.05, **** p < 0.0001).