In vitro evaluation of antidotes for Amanita Phalloides intoxications

(1)

FAC U LD A D E D E FA R M ÁC IA

D

anie

la F

er

re

ira R

od

rigu

es

. In v

itr

o

ev

alua

tio

n o

f a

nti

do

tes f

or

A

m

an

ita

p

ha

llo

id

es

in

to

xic

atio

ns

In v

itro

e

va

lu

at

io

n o

f a

nt

id

ot

es f

or

A

m

ani

ta

ph

allo

id

es

in

to

xic

at

ion

s

D

an

ie

la F

er

re

ira R

od

rig

ues

In vitro

evaluation of antidotes for

Amanita phalloides

intoxications

Daniela Ferreira Rodrigues

M

2018

M

.FFUP

2018

MESTRADO EM CONTROLO DE QUALIDADE

(2)

(3)

Daniela Ferreira Rodrigues

In vitro evaluation of antidotes for

Amanita phalloides intoxications

Dissertação do 2º Ciclo de Estudos Conducente ao Grau de Mestre em Controlo de Qualidade, na área de especialização em Fármacos e Plantas Medicinais

Trabalho realizado sob a orientação do Professor Doutor Félix Carvalho e co-orientação da Professora Doutora Vera Marisa Costa

(4)

DE ACORDO COM A LEGISLAÇÃO EM VIGOR, NÃO É PERMITIDA A REPRODUÇÃO DE QUALQUER PARTE DESTA DISSERTAÇÃO.

(5)

iii

Agradecimentos

Agradeço a todos que contribuíram para a elaboração desta dissertação e com ela para o meu crescimento tanto a nível científico como a nível pessoal.

Ao Professor Doutor Félix Carvalho agradeço a oportunidade de integrar o seu grupo de trabalho, o rigor científico e a constante motivação.

À Professora Doutora Vera Marisa Costa agradeço o acompanhamento, a paciência, a disponibilidade e a exigência. O seu apoio foi fundamental para a realização desta dissertação.

À Professora Doutora Beatriz Oliveira, coordenadora do Mestrado em Controlo de Qualidade, agradeço a disponibilidade e apoio no decurso deste mestrado.

Agradeço a todos membros do Laboratório de Toxicologia da FFUP em especial à Cátia e à Ana Margarida por toda a disponibilidade e apoio logístico, e aos meus caríssimos colegas de gabinete à Ana Rita, Bárbara, Brandon, Eva, Filipa, Flávio, João, Jorge, Margarida, Maria, Rita e Vera pela partilha de dúvidas e por todos os momentos de boa disposição.

Ao Diogo agradeço o suporte, a paciência, a jovialidade e a ajuda em todos os momentos que precisei.

O meu mais especial agradecimento vai para a minha família, para os meus pais Laurentina e José, os meus irmãos, Jorge, Ângela e António e para os meus queridos sobrinhos, Sofia, Francisco e Afonso, por sempre me terem apoiado.

(6)

iv

This work was supported by FEDER funds through the Operational Programme for Competitiveness Factors – COMPETE and by national funds by the Fundação para a Ciência e Tecnologia (FCT) within the project “PTDC/DTP-FTO/4973/2014– POCI-01-0145-FEDER-016545”.

(7)

v

Abstract

Mushroom poisoning has emerged as a serious health problem. Among poisonous mushrooms, species containing cyclopeptides, such as Amanita phalloides, are the most toxic. Amatoxins and specially α-amanitin are responsible for the major deleterious effects. The liver and the kidney are the main targets for α-amanitin cytotoxicity, but hepatocellular effects represent the most lethal and the least treatable manifestation of amatoxin toxicity. Furthermore, there are no worldwide accepted guidelines for the treatment of amatoxins-intoxicated patients and none of the procedures or antidotes used so far has been clearly proven to have great clinical efficiency. Consequently, lethality is still high among intoxications with mushroom containing amatoxins.

The main aims of this dissertation were to characterize α-amanitin cytotoxicity in two human hepatic cell lines (HepG2 and HepaRG) and to evaluate the effect of several putative pharmacological active molecules towards α-amanitin toxicity, since the establishment of an accurate cell model for the evaluation of new antidotes for α-amanitin poisoning is critical in this field. Cytotoxicity of α-amanitin (0.01 – 20µM) in high and low density HepG2 and confluent and differentiated HepaRG cells was evaluated by the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) reduction and neutral red (NR) uptake assays, following a 24 or 48-h incubation period. Additionally, t48-he morp48-hological c48-hanges induced by α-amanitin in 48-hig48-h and low density HepG2 and in confluent and differentiated HepaRG cells were also assessed by phase contrast microscopy following a 48-h incubation period. The characterization of α-amanitin cytotoxicity in high density HepG2 cells was complemented by assessing: 1) the production of reactive species with the 2′,7′-dichlorofluorescin diacetate (DCFH-DA) probe; 2) total glutathione levels; 3) mitochondrial membrane potential; 4) ATP levels and 5) the effect cycloheximide (CHX), a protein synthesis inhibitor, and the effect of buthionine sulfoximine (BSO), an inhibitor of gamma-glutamylcysteine synthetase, on α-amanitin cytotoxicity following a 48-h incubation. Thereafter, the effect of previously described antidotes used in α-amanitin intoxications, namely N-acetylcysteine (NAC), silibinin (SIL) and benzylpenicillin (BPNC) and new putative antidotes polymyxin B (Pol) and cyclosporine A (CsA) was assessed towards the toxic effects of α-amanitin in both high and low density HepG2 and confluent and differentiated HepaRG cells.

Evaluation of the cytotoxic effects in HepG2 and HepaRG cells revealed that following a 48-h incubation, α-amanitin was more cytotoxic in differentiated HepaRG cells, which can be

(8)

vi

accounted to the presence and functional activity of the Na+_{-taurocholate cotransporting}

polypeptide (NTCP) uptake transporter. Concerning the characterization of α-amanitin cytotoxicity in high density HepG2 cells, this amatoxin (2 or 5µM) did not induce changes in mitochondrial membrane potential but α-amanitin 5µM caused a significant increase in intracellular ATP levels. CHX had no effect on the cytotoxicity caused by α-amanitin 5µM following a 48-h incubation. Regarding the cell redox status, α-amanitin (1, 2 or 5µM) did not induce changes in reactive species production but α-amanitin 2 and 5µM caused a tendency to increase total glutathione levels. Moreover, the inhibition of gamma-glutamylcysteine synthetase with BSO led to a partial protection to the impairment caused by α-amanitin 5µM in the MTT reduction assay, following a 48-h incubation period. Concerning the possible protective effect of previously described antidotes for α-amanitin poisoning, NAC had no significant effect towards the cytotoxic effects of α-amanitin in high and low density HepG2 cells but showed a tendency to decrease the cytotoxicity caused by α-amanitin 1µM in both confluent and differentiated HepaRG cells, regarding the cytotoxic effects observed in the MTT reduction assay. On the other hand, SIL, at a non-cytotoxic concentration, pronouncedly aggravated the cytotoxic effects of amanitin in high and low density HepG2 cells and slightly aggravated α-amanitin toxicity in differentiated HepaRG cells, while it had no effect on α-α-amanitin cytotoxic effects in confluent HepaRG cells. Pol had no protective effect on α-amanitin cytotoxicity in high and low density HepG2 cells, but the lowest Pol concentrations tested showed a tendency to decrease the cytotoxicity caused by α-amanitin 1µM in both confluent and differentiated HepaRG cells, when cytotoxicity was assessed by the MTT reduction assay. CsA had no significant protective effects on the cytotoxicity of α-amanitin in high and low density HepG2 and differentiated HepaRG cells, but CsA 2.5µM conferred partial protection against α-amanitin damage in confluent HepaRG cells in the NR uptake assay, which might be mediated by the inhibition of the NTCP uptake transporter.

HepG2 cells have proven to be an interesting model to evaluate the mechanisms of α-amanitin hepatotoxicity; however, studies with HepaRG cells gave powerful insights regarding α-amanitin uptake transport that should be further explored. Moreover, some putative antidotes work in dissimilar manner in these two cell lines and those mechanisms need to be further investigated.

Keywords: HepG2, HepaRG, α-amanitin, mushroom poisoning, Na+_{-taurocholate}

(9)

vii

Resumo

As intoxicações por ingestão de cogumelos venenosos têm aumentado significativamente nos últimos anos, surgindo como um problema de saúde pública. As espécies de cogumelos que contêm ciclopéptidos, tal como a Amanita phalloides, são as mais tóxicas, sendo que as amatoxinas, e particularmente a α-amanitina, são as principais responsáveis pelos devastadores efeitos tóxicos observados. Os principais órgãos alvos de toxicidade são o fígado e o rim. No entanto, a toxicidade hepática representa a consequência mais difícil de tratar e por isso a mais letal. Para além do referido, ainda não existe um consenso para o tratamento de intoxicações com amatoxinas e tanto as medidas terapêuticas como os antídotos disponíveis têm-se revelado pouco eficazes. Consequentemente, a mortalidade causada por intoxicações com cogumelos contendo amatoxinas é ainda consideravelmente alta.

Os principais objetivos desta dissertação foram caraterizar a citotoxicidade da α-amanitina em duas linhas celulares hepáticas (células HepG2 e HepaRG) e avaliar o efeito de vários compostos na citotoxicidade causada pela α-amanitina. A citotoxicidade da α-amanitina (0,01 – 20µM) foi avaliada nas células HepG2 (em duas densidades celulares diferentes) e nas células HepaRG (confluentes e diferenciadas) após incubação por 24 ou 48h, através da utilização de dois ensaios de citotoxicidade, o ensaio de redução do MTT (3-(4,5-dimetilltiazol-2-yl)-2,5-difenil brometo de tetrazólio) e o ensaio de captação do vermelho neutro. Além disso, as alterações morfológicas causadas pela α-amanitina foram avaliadas por microscopia de contraste de fase, após incubação por 48h. Foi ainda efetuada a caraterização da citotoxicidade da α-amanitina nas células HepG2 através da avaliação de: 1) a produção de espécies reativas com a sonda 2′,7′-diclorofluoresceína diacetato (DCFH-DA); 2) os níveis totais de glutationa; 3) potencial da membrana mitocondrial; 4) os níveis de ATP; 5) o efeito na citotoxicidade causada pela α-amanitina 5µM da cicloheximida (CHX), um inibidor da síntese proteica e da butionina sulfoximina (BSO), um inibidor da gama-glutamilcisteína sintetase, após um período de incubação de 48h. Subsequentemente foi avaliado, quer nas células HepG2 como nas HepaRG, o efeito de antídotos na citotoxicidade causada pela α-amanitina, nomeadamente os atualmente usados nas intoxicações por Amanita phalloides, como a N-acetilcisteina (NAC), silibinina (SIL) e benzilpenicilina (BPNC) e de novos possíveis antídotos, como a polimixina B (Pol) e ciclosporina A (CsA).

(10)

viii

A avaliação dos efeitos citotóxicos causados pela α-amanitina revelou que, após incubação por 48h, a α-amanitina foi mais citotóxica nas células HepaRG diferenciadas. A maior sensibilidade das células HepaRG diferenciadas pode dever-se à presença e atividade funcional do transportador, polipéptido cotransportador de Na+_{-taurocolato (NTCP).}

Relativamente à caraterização da citotoxicidade nas células HepG2, a incubação com α-amanitina (2 ou 5µM) não causou a alterações significativas no potencial da membrana mitocondrial, mas a incubação da α-amanitina 5µM causou um aumento significativo nos níveis intracelulares de ATP após incubação por 24h. A incubação da CHX não reverteu a citotoxicidade causada pela α-amanitina 5µM após uma incubação de 48h. Em termos do estado redox, a α-amanitina não causou alterações na produção de espécies reativas em qualquer das concentrações testadas (1, 2 ou 5µM), mas a incubação com α-amanitina 2 e 5µM apresentou uma tendência para aumentar os níveis totais de glutationa. A inibição da gama-glutamilcisteína sintetase conduziu a uma proteção parcial dos efeitos citotóxicos causados pela α-amanitina 5µM após incubação por 48h, no ensaio de redução do MTT. Relativamente a um possível efeito protetor de antídotos previamente descritos para intoxicações com amanitina, a NAC não protegeu contra a citotoxicidade causada pela α-amanitina nas células HepG2, mas causou uma ligeira proteção contra o dano causado pela α-amanitina 1µM nas células HepaRG confluentes e diferenciadas, relativamente à toxicidade observada no ensaio de redução do MTT, mas sem atingir significância estatística. Por outro lado, a SIL, numa concentração não citotóxica, agravou a citotoxicidade causada pela α-amanitina nas células HepG2 e HepaRG diferenciadas, mas não apresentou qualquer efeito sobre a citotoxicidade causada pela α-amanitina nas células HepaRG confluentes. A Pol não conferiu qualquer efeito protetor relativo à toxicidade causada pela α-amanitina nas células HepG2, mas as mais baixas concentrações testadas de Pol mostraram uma tendência para conferir proteção relativamente à toxicidade causada pela α-amanitina 1µM nas células HepaRG confluentes e diferenciadas, quando esta citotoxicidade foi avaliada pelo ensaio de redução de MTT após incubação por 48h. A CsA não conferiu um efeito protetor relativo à citotoxicidade causada pela α-amanitina nas células HepG2 e nas células HepaRG diferenciadas, mas nas células HepaRG confluentes a CsA na concentração de 2,5µM protegeu parcialmente contra o dano causado pela α-amanitina, quando este foi avaliado pelo ensaio de captação de vermelho neutro. Tal efeito pode dever-se à inibição do transportador NTCP.

(11)

ix

As células HepG2 demonstraram ser um modelo interessante para a avaliação dos mecanismos de hepatotoxicidade da α-amanitina, no entanto os estudos com as células HepaRG forneceram dados relevantes sobre a entrada da α-amanitina mediada por transportadores, o que deve ser futuramente aprofundado. De facto, alguns dos possíveis antídotos apresentaram efeitos diferentes sobre a citotoxicidade observada após incubação com a α-amanitina nas duas linhas celulares e esses mecanismos necessitam de ser futuramente investigados.

Palavras-chave: HepG2, HepaRG, α-amanitina, intoxicação por cogumelos, polipéptido cotransportador de Na+_{-taurocolato.}

(12)

x

4.1.2.1 Only the highest concentration of α-amanitin tested caused a significant cytotoxic effect in low density HepG2 cells ... 48 4.1.2.2 α-Amanitin caused mitochondrial and lysosomal uptake dysfunction in low density HepG2 cells ... 49 4.1.2.3 N-Acetylcysteine, a glutathione precursor and reactive species scavenger, had no effect on the cytotoxicity caused by α-amanitin in low density HepG2 cells .... 50 4.1.2.4 Silibinin, an antioxidant and inhibitor of the OATP1B3 uptake transporter, aggravated the cytotoxicity caused by α-amanitin in low density HepG2 cells ... 51 4.1.2.5 Benzylpenicillin, an antibiotic and inhibitor of the OATP1B3 uptake transporter, slightly aggravated lysosomal disfunction caused by α-amanitin in low density HepG2 cells ... 52 4.1.2.6 Polymyxin B, an antibiotic and in silico inhibitor of α-amanitin binding to RNAP II, had no effect on the cytotoxicity caused by α-amanitin in low density HepG2 cells………... 53 4.1.2.7 Cyclosporine A, an inhibitor of the OATP1B3 and NTCP uptake transporters, slightly aggravated the mitochondrial dysfunction caused by α-amanitin in low density HepG2 cells ... 55

Toxicological evaluation of α-amanitin in HepaRG cells ... 57

Toxicological evaluation of α-amanitin in confluent HepaRG cells ... 58 4.2.1.1 α-Amanitin caused concentration dependent cytotoxicity in confluent HepaRG cells ... 58 4.2.1.2 α-Amanitin caused a time dependent mitochondrial and lysosomal dysfunction in confluent HepaRG cells... 60 4.2.1.1 N-Acetylcysteine, a glutathione precursor and reactive species scavenger, had no significant effect on the cytotoxicity caused by α-amanitin in confluent HepaRG cells………... 61 4.2.1.2 Silibinin, an antioxidant and inhibitor of the OATP1B3 uptake transporter, had no effect on α-amanitin cytotoxicity in confluent HepaRG cells ... 62 4.2.1.3 Benzylpenicillin, an antibiotic and inhibitor of the OATP1B3 uptake transporter, had no significant effect on α-amanitin cytotoxicity in confluent HepaRG cells………... 63 4.2.1.4 Polymyxin B, an antibiotic and in silico inhibitor of α-amanitin binding to RNAP II, had no significant effect on the cytotoxicity caused by α-amanitin in confluent HepaRG cells ... 64 4.2.1.5 Cyclosporine A, an inhibitor of the OATP1B3 and NTCP uptake transporters, partially reverted the lysosomal dysfunction induced by α-amanitin in confluent HepaRG cells………... 65 Toxicological evaluation of α-amanitin in differentiated HepaRG cells ... 66 4.2.2.1 α-Amanitin caused concentration dependent cytotoxicity in differentiated HepaRG cells ... 66

(15)

xiii

4.2.2.2 α-Amanitin caused a time dependent mitochondrial and lysosomal

dysfunction in differentiated HepaRG cells ... 69

4.2.2.3 N-Acetylcysteine, a glutathione precursor and reactive species scavenger, had no effect on the cytotoxicity caused by α-amanitin in differentiated HepaRG cells………... 70

4.2.2.4 Silibinin, an antioxidant and inhibitor of the OATP1B3 uptake transporter, slightly aggravated the lysosomal dysfunction caused by α-amanitin in differentiated HepaRG cells ... 71

4.2.2.5 Benzylpenicillin, an antibiotic and inhibitor of the OATP1B3 uptake transporter, had no significant effect on α-amanitin cytotoxicity in differentiated HepaRG cells……….. ... 72

4.2.2.6 Polymyxin B, an antibiotic and in silico inhibitor of α-amanitin binding to RNAP II, had no significant effect on the cytotoxicity caused by α-amanitin in differentiated HepaRG cells ... 73

4.2.2.7 Cyclosporine A, an inhibitor of the OATP1B3 and NTCP uptake transporters, had no significant effect on the cytotoxicity caused by α-amanitin in differentiated HepaRG cells ... 74

α-Amanitin was more cytotoxic in differentiated HepaRG cells at the latter time point…. ... 76

5. Discussion and conclusions ... 77

Cytotoxicity effects of α-amanitin in HepG2 and HepaRG cells ... 78

Evaluation of the effect of α-amanitin on cellular energetics of high density HepG2 ... 83

Evaluation of the effect of α-amanitin on high density HepG2 cells redox status ………84

Evaluation of putative antidotes towards the cytotoxic effects of α-amanitin in HepG2 and HepaRG cells ... 85

N-Acetylcysteine ... 85 Silibinin ... 86 Benzylpenicillin ... 88 Polymyxin B ... 90 Cyclosporine A ... 91 Final conclusions ... 92 References ... 94 Appendix ... 104

(16)

xiv

List of figures

Figure 1 – Top view of RNAP II structure. ... 8 Figure 2 – Interaction of α-amanitin with RNAP II. ... 9 Figure 3 – Mechanisms of α-amanitin toxicity. ... 16 Figure 4 – Phase contrast microscopy of high density HepG2 cells after incubation with 5, 10 or 20µM of α-amanitin for 48h. ... 36 Figure 5 – α-Amanitin cytotoxicity in high density HepG2 cells following a 24-h incubation period by assessing alterations in MTT reduction (A) and NR uptake (B). ... 37 Figure 6 – α-Amanitin cytotoxicity in high density HepG2 cells following a 48-h period incubation by assessing alterations in MTT reduction (A) and NR uptake (B). ... 38 Figure 7 – Mitochondrial membrane potential evaluated by JC-1 staining in high density HepG2 cells incubated with 2 or 5µM of α-amanitin after a 24-h exposure. ... 38 Figure 8 – ATP levels in high density HepG2 cells treated with 2 or 5µM of α-amanitin for 24h. ... 39 Figure 9 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of high density HepG2 cells pre-incubated with 10nM of cycloheximide (CHX) and then incubated with 5µM of α-amanitin (AMA) for 48h. ... 40 Figure 10 – Cellular redox status evaluated with the fluorescent DCFH-DA probe in high density HepG2 cells treated with 1, 2 or 5µM of α-amanitin. ... 41 Figure 11 – Total glutathione (tGSH) levels in high density HepG2 cells treated with 2 or 5µM of α-amanitin (AMA), or pre-incubated with buthionine sulfoximine (BSO) 25µM and incubated with AMA 2 or 5µM for 24h. ... 42 Figure 12 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of high density HepG2 cells pre-incubated with 25µM of buthionine sulfoximine (BSO) and then incubated with 5µM of α-amanitin (AMA) for 48h. ... 43 Figure 13 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of high density HepG2 cells pre-incubated with 1mM of N-acetyl cysteine (NAC) and then incubated with 5µM of α-amanitin (AMA) for 48h. ... 44 Figure 14 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of high density HepG2 cells pre-incubated with 10µM of silibinin (SIL) and then incubated with α-amanitin (AMA) 5µM for 48h. ... 45

(17)

xv

Figure 15 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of high density HepG2 cells pre-incubated with 0.5mM of benzylpenicillin (BPNC) and then incubated with α-amanitin (AMA) 5µM for 48h. ... 46 Figure 16 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of high density HepG2 cells pre-incubated with polymyxin B (Pol) and then incubated with α-amanitin (AMA) 5µM for 48h. ... 47 Figure 17 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of high density HepG2 cells pre-incubated with cyclosporine A (CsA) and then incubated with 5µM of α-amanitin (AMA) for 48h. ... 48 Figure 18 – Phase contrast microscopy of low density HepG2 cells after incubation with 2 or 5µM of α-amanitin for 48h. ... 49 Figure 19 – α-Amanitin cytotoxicity in low density HepG2 cells following a 48-h incubation period by assessing alterations in MTT reduction (A) and NR uptake (B). ... 50 Figure 20 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of low density HepG2 cells pre-incubated with 1mM of N-acetyl cysteine (NAC) and then incubated with 2 or 5µM of α-amanitin (AMA) for 48h. ... 51 Figure 21 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of low density HepG2 cells pre-incubated with 10µM of silibinin (SIL) and then incubated with 2 or 5µM of α-amanitin (AMA) for 48h. ... 52 Figure 22 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of low density HepG2 cells pre-incubated with 0.5mM of benzylpenicillin (BPNC) and then incubated with 2 or 5µM of α-amanitin (AMA) for 48h. ... 53 Figure 23 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of low density HepG2 cells pre-incubated with polymyxin B (Pol) and then incubated with 2µM of α-amanitin (AMA) for 48h. ... 54 Figure 24 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of low density HepG2 cells pre-incubated with polymyxin B (Pol) and then incubated with 5µM of α-amanitin (AMA) for 48h. ... 54 Figure 25 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of low density HepG2 cells pre-incubated with cyclosporine A (CsA) and then incubated with 2µM of α-amanitin (AMA) for 48h. ... 55 Figure 26 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of low density HepG2 cells pre-incubated with cyclosporine A (CsA) and then incubated with 5µM of α-amanitin (AMA) for 48h. ... 56

(18)

xvi

Figure 27 – Phase contrast microscopy of HepaRG cells 24h after seeding (A, B), and in a confluent (C, D) or differentiated state (E, F). In a differentiated state two types of cells can be identified: hepatocyte (arrow head) and biliary (arrow) like cells. ... 57 Figure 28 – Phase contrast microscopy of confluent HepaRG cells after incubation with 1, 2, 5 or 10µM of α-amanitin for 48h. ... 59 Figure 29 – α-Amanitin cytotoxicity in confluent HepaRG cells following a 24-h incubation period by assessing alterations in MTT reduction (A) and NR uptake (B). ... 60 Figure 30 – α-Amanitin cytotoxicity in confluent HepaRG cells following a 48-h incubation period by assessing alterations in MTT reduction (A) and NR uptake (B). ... 61 Figure 31 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of confluent HepaRG cells pre-incubated with 1mM of N-acetyl cysteine (NAC) and then incubated with 1µM of α-amanitin (AMA) for 48h. ... 62 Figure 32 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of confluent HepaRG cells pre-incubated with 10µM of silibinin (SIL) and then incubated with α-amanitin (AMA) 1µM for 48h. ... 63 Figure 33 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of confluent HepaRG cells pre-incubated with 0.5mM of benzylpenicillin (BPNC) and then incubated with α-amanitin (AMA) 1µM for 48h. ... 64 Figure 34 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of confluent HepaRG cells pre-incubated with polymyxin B (Pol) and then incubated with 1µM of α-amanitin (AMA) for 48h. ... 65 Figure 35 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of confluent HepaRG cells pre-incubated with cyclosporine A (CsA) and then incubated with 1µM of α-amanitin (AMA) for 48h. ... 66 Figure 36 – Phase contrast microscopy of differentiated HepaRG cells after incubation with 1, 2, 5 and 10µM of α-amanitin for 48h. ... 68 Figure 37 – α-Amanitin cytotoxicity in differentiated HepaRG cells following a 24-h period incubation by assessing alterations in MTT reduction (A) and NR uptake (B). ... 69 Figure 38 – α-Amanitin cytotoxicity in differentiated HepaRG cells following a 48-h incubation period by assessing alterations in MTT reduction (A) and NR uptake (B). ... 70 Figure 39 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of differentiated HepaRG cells pre-incubated with 1mM of N-acetyl cysteine (NAC) and then incubated with 1µM of α-amanitin (AMA) for 48h. ... 71

(19)

xvii

Figure 40 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of differentiated HepaRG cells pre-incubated with 10µM of silibinin (SIL) and then incubated with α-amanitin (AMA) 1µM for 48h. ... 72 Figure 41 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of differentiated HepaRG cells pre-incubated with 0.5mM of benzylpenicillin (BPNC) and then incubated with α-amanitin (AMA) 1µM for 48h. ... 73 Figure 42 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of differentiated HepaRG cells pre-incubated with polymyxin B (Pol) and then incubated with α-amanitin (AMA) 1µM for 48h. ... 74 Figure 43 – Cellular cytotoxicity evaluated by the MTT reduction (A) and NR uptake (B) assays of differentiated HepaRG cells pre-incubated with cyclosporine A (CsA) and then incubated with α-amanitin (AMA) 1µM for 48h. ... 75 Figure 44 – MTT reduction assay (A) and NR uptake assay (B) in high density (HD) and low density (LD) HepG2 cells, confluent (Conf) and differentiated (Dif) HepaRG cells exposed to 2 and 5µM of α-amanitin for 48h. ... 76

(20)

xviii

List of tables

Table 1 – Summary of putative in vitro antidotes for α-amanitin poisoning in hepatic cell models. ... 19

(21)

xix

Abbreviations

ATP – Adenosine 5'-triphosphate BPNC – Benzylpenicillin

BSA – Bovine serum albumin BSO – Buthionine sulfoximine CEFT – Ceftazidime

CHX – Cycloheximide CsA – Cyclosporine A DMSO – Dimethyl sulfoxide DOX – Doxorubicin

FBS – Fetal bovine serum GSH – Glutathione

GSSG – Glutathione disulfide

IL-Ra – Interleukin-receptor antagonist Ki – Inhibitory constant

LDH - Lactate dehydrogenase

MDCKII – Madin Darby canine kidney strain II MLD – Minimum lethal dose

MMP – Mitochondrial membrane potential

MTT – 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide NAC – N-Acetylcysteine

NADPH – Nicotinamide adenine dinucleotide 2′-phosphate NR – Neutral Red

NTCP/Ntcp – Na+_{-taurocholate cotransporting polypeptide}

OATP/Oatp – Organic anion-transporting polypeptide Pol – Polymyxin B

RNAP – RNA polymerase RS – Reactive species SIL – Silibinin

SRB – Sulforhodamine B tGSH – Total glutathione TNF – Tumor necrosis factor

(22)

1

1. Mushroom poisoning

Due to a higher interest in mushroom nutritional properties, mushroom poisoning has been increasing for the past few years, owed by the lack of efficient identification of toxic and edible mushroom by its collectors (Garcia et al., 2015b). While several toxic components have been identified in poisonous mushrooms, species containing cyclopeptides are the most toxic. Regarding cyclopeptides, amatoxins are important toxin agents since they are bicyclic heat-stable octapeptides which are also not degraded under acidic conditions in the stomach, being highly toxic in the liver and kidneys (Garcia et al., 2015b).

Owing to the lack of case reporting or misidentification, accurate estimates of worldwide poison by amatoxins-containing mushrooms are difficult to establish. Nevertheless, several studies show that amatoxin poisoning has emerged as a serious health problem. Data from the 2016 Annual Report of the American Association of Poison Control Centers' National Poison Data System (NPDS) reported 6421 mushroom intoxications of which 72 cases were attributed to mushroom containing cyclopeptides (Gummin et al., 2017). In Northern Italy, a 21-year retrospective analysis (from January 1996 to December 2016) identified 443 mushroom poisoning cases, of which 20 cases were attributed to amatoxin poisoning (Cervellin et al., 2018). A retrospective case study with data from human exposures to mushrooms notified to the Swiss Toxicological Information Centre between January 1995 and December 2009 reported a total of 32 amatoxin poisoning confirmed cases, of which five had a fatal outcome (Schenk-Jaeger et al., 2012). In Portugal, 93 cases of mushroom poisoning were reported between 1990 and 2008. Of these, 63.4% presented a hepatotoxic profile (a profile that has been associated with the consumption of amatoxin containing mushrooms), from which 11.8% had a fatal outcome (Brandao et al., 2011).

Amatoxins are present in three genera: Amanita (mainly Amanita phalloides, A. virosa and A. verna); Lepiota and Galerina. From these, A. phalloides is responsible for the majority of mushroom poisoning fatal cases (Garcia et al., 2015b). Amatoxins, and especially α-amanitin, are the main responsible for the toxic effects. The liver is the main target organ of toxicity, but the kidneys are also affected by amatoxin deleterious effects (Garcia et al., 2015b). In fact, amatoxin poisoning usually has a bad prognosis due to the high risk of liver failure. Still, there are no worldwide accepted guidelines regarding the treatment of amatoxins-intoxicated patients and none of the procedures or antidotes used so far has been clearly proven to have

(23)

2

great clinical efficacy (Enjalbert et al., 2002; Garcia et al., 2015b). Therefore, the discovery of a fully effective antidote is still a major unsolved issue.

Toxicokinetics of amatoxins

Signs and symptoms of amatoxin poisoning can extend from a simple gastroenterological disorder to death and are mainly attributable to the accumulation of α-amanitin in the liver and kidneys (Garcia et al., 2015b). Amatoxin poisoning is characterized by a long asymptomatic period (from 6 to 12h) and three clinical phases: 1) gastrointestinal phase, 2) latent period and 3) the hepatorenal phase (Enjalbert et al., 2002; Garcia et al., 2015b). The gastrointestinal phase (12-24h) is characterized by nausea, vomiting, diarrhea (occasionally bloody), abdominal pain, and hematuria. During the second phase (24-48h), clinical signs and biochemical makers of hepatic damage appear. In the hepatorenal phase (4-7 days), hemorrhages, convulsions, and fulminant hepatic failure occur resulting in coma and death (Enjalbert et al., 2002; Garcia et al., 2015b).

Amatoxins are rapidly absorbed from the intestinal tract, as they were detected in the gastric fluid and urine within 90-120 minutes after ingestion of wild mushrooms by a 15-year old boy who tried to commit suicide (Homann et al., 1986). Jaeger et al., quantified amatoxins (α-amanitin and β-amanitin) in plasma, urine, gastroduodenal fluid, feces, liver and kidney of 45 patients intoxicated with A. phalloides (Jaeger et al., 1993). Overall, amatoxins were often detectable in plasma (α-amanitin levels ranged from 88 to 190ng/mL) for up to 36h following ingestion but were present in the urine until 4 days after the ingestion has happened (Jaeger et al., 1993). α-Amanitin was present in the urine at high concentrations (the highest concentration observed was 48020ng/mL) and presented a mean excretion per hour of 32.18µg. Additionally, amatoxins were found in high concentrations in gastroduodenal aspiration fluid (α-amanitin concentration ranged from 492 to 4950µg/L) and α-amanitin was also present in feces, where the values ranged from 8.4 to 152µg. α-Amanitin was found in the liver and kidney, but kidney concentrations exceeded those of the liver. Liver concentrations ranged between 0 and 19ng/g, while kidney concentrations ranged from 122 to 1719ng/g [Method detection limit: 5 ng/mL] (Jaeger et al., 1993).

Experimental studies on amatoxin kinetics show that after dog intravenous administration, the plasma half-life of amatoxins was shown to be short, ranging from 26.7 to

(24)

3

49.6 minutes. Moreover, four to six hours after the administration, amatoxins were not detectable in the plasma. More than 80% of the dose injected was eliminated in the urine and less than 10% in the bile (Faulstich et al., 1985). In order to evaluate the toxicokinetic process of α-amanitin in rats, Li et al., quantified plasma α-amanitin concentrations after intraperitoneal administration of a single dose of 0.5 mg/kg to Sprague–Dawley rats, since rodents do not have oral bioavailability for amatoxins (Li et al., 2017). The α-amanitin plasma quantification showed that α-amanitin is quickly eliminated from plasma with an elimination half-life of 0.49 ± 0.18h. Moreover, α-amanitin was undetected 200 minutes after administration [Method detection limit 3.0ng/mL] (Li et al., 2017). Additionally, Garcia et al., quantified α-amanitin in kidney, liver and plasma samples 2 or 4h after the administration of a single intraperitoneal dose of α-amanitin (10 or 21.4mg/kg) to Wistar rats (Garcia et al., 2015a). Results from the UV-diode array detection at 305nm show that the highest concentration of α-amanitin was found in the kidney, whereas no detectable values were found in the plasma [Based on the calibration curve, the detection limit for plasma samples was 0.450µg/mL] (Garcia et al., 2015a). Interestingly, the accumulation of α-amanitin in the kidneys was dependent on the dose administered (Garcia et al., 2015a). In fact, while quantification 4h after α-amanitin administration in the liver was in the same range independently of the dose of α-amanitin administrated (10mg/kg – α-amanitin: 0.97µg/g; 21.4mg/kg – α-amanitin: 0.71µg/g), in the kidney α-amanitin levels dramatically increased for the highest dose. Quantification of α-amanitin in kidney 4h following administration of a 10mg/kg dose was of 1.40µg/g, while a dose of 21.4mg/kg resulted in a quantification of 26.58µg/g (Garcia et al., 2015a). In addition, 2h following the administration of 21.4mg/kg of α-amanitin, its levels remained undetected in the liver and where of 3.16µg/g in the kidney [Based on the calibration curve, the detection limit for liver samples was 0.330µg/g] (Garcia et al., 2015a).

Hepatocellular effects represent the most lethal and the least treatable manifestation of amatoxin toxicity. In fact, a retrospective analysis of 81 patients intoxicated by A. phalloides with simultaneous damage of liver and kidneys showed that even though renal damage was present with acute tubular necrosis in 16 patients, none of the patients died because of renal failure (Mydlik et al., 2013). Nevertheless, acute toxic hepatitis rapidly developed to liver insufficiency and coma, and 24 patients died (Mydlik et al., 2013). Damage to the liver as a result of α-amanitin poisoning is characterized by massive centrilobular necrosis, vacuolar degeneration and a positive acid-phosphatase reaction (Fineschi et al., 1996). Additionally, some authors have suggested that the clinical course of amanitin poisoning could be

(25)

4

significantly influenced by the enterohepatic recirculation of α-amanitin (Faulstich et al., 1980; Faulstich et al., 1985). To evaluate the clinical significance of the enterohepatic circulation in the course of high-dose and medium lethal dose (LD50) α-amanitin intoxication, Thiel et al.,

calculated the difference between portal and systemic plasma α-amanitin concentration (Δ-Amanitin: portal - systemic plasma concentration) following an intraportal administration of 0.35mg/kg or 0.15mg/kg of α-amanitin to female German landrace pigs. In the high dose group (0.35 mg/kg), 5h after intoxication Δ-Amanitin presented negative values representing the systemic plasma excess. Afterwards, values became slightly positive 16h after intoxication and trend to baseline levels within 24h. In the low dose group (0.15 mg/kg), 5h after intoxication Δ-amanitin presented positive value, being verified a rapid enterohepatic reuptake, suggesting that within the early phase of poisoning a larger amount of α-amanitin is excreted to the bile. Values decreased to baseline levels within 16h. Until 16-24h after of administration of α-amanitin, enterohepatic circulating amanitin might produce a delayed biliary excretion from hepatocytes into the intestine, but amanitin reuptake from the small intestine did not produce prolonged appearance of the toxin in portal plasma samples. After 16–24h, relevant toxin concentration and/or intestinal reuptake do not longer exist and thus, at this time point, interruption of the enterohepatic circulation might be poorly effective to intoxication treatment (Thiel et al., 2011).

Notwithstanding the major deleterious effects of α-amanitin in the liver, the main route of α-amanitin elimination is the kidney and α-amanitin kidney concentrations have been reported to be much higher than those of the liver (Garcia et al., 2015a; Jaeger et al., 1993). Lesions in the kidney caused by α-amanitin poisoning are characterized by acute tubular necrosis and hyaline casts in the tubules (Fineschi et al., 1996). Since in mouse kidney necrosis is only observed in the proximal tubules, Fiume et al., evaluated if necrosis could be caused by the reabsorption of amatoxin filtered in the glomeruli. For that, a conjugate of β-amanitin with albumin was produced and administrated by intraperitoneal injection to mice. As a result, all mice died from necrosis of the liver but without any lesions in the kidney, three days after administration, since albumin is not reabsorbed in the glomeruli. Thus, amanitin poisoning in mouse kidney depends on the reabsorption in the tubules (Fiume et al., 1969). Even though the liver is described as the major target for α-amanitin toxic effects, Fiume et al., reported that effects of α-amanitin in either the liver or mouse kidney are dependent on the dose administered (Fiume et al., 1969). In fact, the authors reported that administration of a minimum lethal dose (MLD - 3.5µg α-amanitin/10g body weight) of α-amanitin always causes necrosis in the kidneys

(26)

5

but never in the liver. Necrosis in the liver appeared with the administration of doses above the MLD, generally within two days. For a dose up to 3 MLD, necrosis of the kidneys never appeared less than 3 days after administration, thus when necrosis of the liver occurs, it can result in death before necrosis of the kidney has had time to develop (Fiume et al., 1969).

The differential toxic effects of α-amanitin in the liver and kidneys can be explained by transporter expression profiles as they are key determinants for drug disposition (Giacomini et al., 2010). In fact, the existence of uptake transporters can be particularly important for α-amanitin due to its chemical characteristics, namely size and hydrophilicity. Earlier studies on α-amanitin uptake suggested that sodium-dependent bile acid transporters may play a role in α-amanitin uptake (Kroncke et al., 1986). Afterwards, Gundala et al. showed that rat hepatocellular bile acid transporter, Na+_{-taurocholate cotransporter polypeptide (Ntcp), may be}

an important mediator of α-amanitin uptake in the liver (Gundala et al., 2004). Evaluation of the importance of the Ntcp transporter for α-amanitin uptake was performed by measuring the ability of α-amanitin to block cytokine-induced synthesis of interleukin-receptor antagonist (IL-Ra) mRNA in HepG2 transfected and non-transfected with rat Ntcp (Gundala et al., 2004). In non-transfected HepG2 cells, IL-1Ra mRNA expression was not affected by pre-treatment with α-amanitin, since IL-1Ra mRNA levels presented a 21.6-fold increase, whereas cells without α-amanitin presented an 8.6-fold increase. On the other hand, Ntcp-transfected HepG2 cells treated with α-amanitin presented a significant reduction of induced IL-1Ra mRNA expression. In fact, pre-treatment of HepG2 transfected cells with α-amanitin caused a reduction in IL-1RA expression to 1.9-fold, while transfected cells without α-amanitin pre-treatment presented a 15.6-fold increase (Gundala et al., 2004). Additionally, the authors showed that the presence of the Ntcp transporter had a significant effect on the cytotoxic effects of α-amanitin. Indeed, incubation of α-amanitin 0.75µM for seven days caused 90% cell death in Ntcp-HepG2 transfected cells, while non-transfected cells were more resistant to α-amanitin cytotoxic effects (Gundala et al., 2004). Additionally, Letschert et al., evaluated if members of the organic anion– transporting polypeptide (OATP) family localized in the sinusoidal membranes of human hepatocytes are also involved in amatoxin uptake (Letschert et al., 2006). For that, α-amanitin uptake was measured in Madin Darby canine kidney strain II (MDCKII) cells stably expressing human OATP1B3, OATP2B1 or OATP1B1 transporters. MDCKII cells expressing OATP2B1 and OATP1B1 showed little, if any, α-amanitin uptake, while OATP1B3-expressing MDCKII exhibited a significant higher uptake ratio (Letschert et al., 2006). In addition, the authors evaluated α-amanitin 0.1µM cytotoxicity in MDCKII transfected with the three uptake

(27)

6

transporters under study, using the Alamar Blue assay, following incubation for 24 to 72h. The results showed that viability of control MDCKII and cells transfected with OATP2B1 and -OATP1B1 transporters remained unaffected, while OATP1B3 transfected cells exhibited a time dependent decrease in cell viability. Moreover, the concentration dependent cytotoxicity caused by amanitin was also evaluated following incubation with α-amanitin (0.1 – 10µM) for a 24-h incubation period. MDCKII-OATP1B3 cells showed an α-amanitin concentration dependent viability decrease, with a LD50 of approximately 0.3µM, while control MDCKII and

-OATP2B1 and -OATP1B1 transfected cells, only exhibited a slight cytotoxic effect for α-amanitin 10µM (Letschert et al., 2006). To further support the role of OATP1B3 uptake transporter in α-amanitin cytotoxicity, inhibitors of OATP1B3, such as cyclosporine A (CsA), rifampicin and cholecystokinin octapeptide (CCK-8), conferred protection against the cellular damage induced by α-amanitin in MDCKII-OATP1B3 transfected cells (Letschert et al., 2006).

Mechanisms of α-amanitin toxicity

The high lethality of A. phalloides poisoning relies on the presence of powerful toxins such as cyclic octapeptides as previously mentioned. From these, amatoxins, and specially α-amanitin are responsible for the severe liver and kidney injury observed after A. phalloides poisoning (Garcia et al., 2015b). Several mechanisms have been proposed to describe α-amanitin toxicity. The main mechanism of toxicity is the inhibition of RNA polymerase (RNAP) II transcription process (Lindell et al., 1970). Nevertheless, other mechanisms have been suggested to contribute to α-amanitin potent toxic effects, such as inflammation and oxidative stress (Leist et al., 1994; Leist et al., 1997; Zheleva et al., 2007).

RNAP II transcriptional blockage

1.2.1.1 Briefing on RNAP II structure and function

RNAP II is responsible for the transcription of all protein-coding genes through mRNA production. Therefore, transcription by RNAP II is a key step in gene expression and a fundamental process in a living cell (Nygaard and Hovig, 2009).

(28)

7

RNAP II is composed by 12 subunits (Rpb 1-12). Ten of the subunits form a catalytic core and the remaining two subunits Rpb4 and Rpb7 (Rpb4/7) form a subcomplex located on the periphery of the enzyme, the Rpb4/7 heterodimer (Figure 1) (Armache et al., 2005; Bushnell and Kornberg, 2003; Cramer et al., 2008). The ten-subunit catalytic core alone is capable of elongating the RNA transcript, but it requires the Rpb4/7 complex for transcript initiation (Cramer, 2004; Edwards et al., 1991; Orlicky et al., 2001).

Rpb1 is the largest subunit of RNAP II and it has a regulatory role on RNAP II activity through the transcription cycle, due to the presence of a C-terminal domain (CTD) (Corden et al., 1985). The CTD extends from the core enzyme to form a tail-like structure that provides a binding site for various factors involved in RNA transcription and processing (Hsin and Manley, 2012; Meinhart et al., 2005). The CTD is an unique feature of RNAP II and is composed of tandem heptad repeats, with the consensus sequence Tyr1-Ser2-Pro3-Thr4-Ser5-Pro6-Ser7 (Corden, 2016). The CTD suffers extensive modification through phosphorylation, glycosylation, ubiquitination, and methylation (Hsin and Manley, 2012). CTD modifications and structural plasticity enable CTD to serve as a binding platform for a variety of factors that regulate the transcription and maturation of nascent transcripts (Egloff et al., 2012; Hintermair et al., 2016). The ability of the CTD to be modified at each residue can generate a wide range of distinct combinations, which contain information that is crucial at different steps of RNA transcription and processing (Egloff et al., 2012; Heidemann et al., 2013). These different combinations have been liked to a readable code, the CTD code (Buratowski, 2003; Corden, 2007; Komarnitsky et al., 2000) that orchestrates the recruitment and interaction of factors with RNAP II (Heidemann et al., 2013). Of the possible modifications, CTD phosphorylation stands out, since the phosphorylation status is critical for RNAP II activity (Hsin and Manley, 2012; Malumbres, 2014): CTD in a hypophosphorylated form allows RNAP II to enter the preinitiation complex, whereas a hyperphosphorylated form is necessary for progressive elongation of the transcript, while dephosphorylation of the CTD leads to transcription termination (Egloff et al., 2012). Specifically, some modifications have been attributed to different stages of the transcription process. After transcription initiation, Ser5 and Ser7 are phosphorylated. Transition from initiation to elongation is marked by Ser2 and Thr4 phosphorylation, whereas the phosphate groups on Ser5 and Ser7 are gradually removed by phosphatases. As RNAP II proceeds for termination, Ser2 is dephosphorylated, regenerating unphosphorylated RNAP II that can be recycled for another round of transcription (Hsin and Manley, 2012).

(29)

8

Figure 1 – Top view of RNAP II structure.

The polymerase catalytic core is composed by ten subunits, and the remaining two subunits Rpb4 and Rpb7 (Rpb4/7) form a subcomplex located at the periphery of the enzyme, the Rpb4/7 heterodimer. Adapted from Armache et al., 2005.

1.2.1.2 α-Amanitin mediated RNAP II transcription inhibition

One of the mechanisms of α-amanitin toxicity is the potent and specific inhibition of transcription by RNAP II (Lindell et al., 1970). While RNAP II is extremely sensitive to α-amanitin, RNAP I is insensitive and RNAP III is hundred-fold less sensitive than RNAP II (Lindell et al., 1970; Weinmann and Roeder, 1974). RNAP III sensitivity to α-amanitin has been associated with the expression regulation of RNAP III class III genes by RNAP II (Raha et al., 2010). The affinity of α-amanitin for RNAP II is very high, with an equilibrium association constant in the order of 108_-1010_M-1 _{(Cochet-Meilhac and Chambon, 1974).}

The first structural studies of α-amanitin with RNAP II indicated that α-amanitin binds RNAP II in the interface between to the two largest subunits, Rpb1 and Rpb2, through an interaction with the bridge helix residues (Bushnell et al., 2002). This mode of interaction suggested that α-amanitin interfered with the bridge helix movement. The bridge helix movement promotes RNAP II activity by participating in the translocation of the DNA-RNA hybrid, an essential step needed to clear the space to load the next substrate nucleoside triphosphate into the active site, necessary for the next round of synthesis (Bushnell et al., 2002; Gong et al., 2004; Kaplan and Kornberg, 2008). However, the fact that α-amanitin binds

(30)

9

only to one residue in the bridge helix and since the residue is separated by one turn from the residues that apparently change conformation, raised the possibility that this mode of interaction could be insufficient to explain RNAP II translocation inhibition by α-amanitin (Brueckner and Cramer, 2008). An alternative model for transcription inhibition was proposed, suggesting that α-amanitin interferes with the movement of the trigger loop, which is involved in nucleotide selection and addition (Figure 2) (Brueckner and Cramer, 2008; Kaplan et al., 2008; Wang et al., 2006; Zhang et al., 2010). Even though prior studies suggested that α-amanitin did not influence nucleotide incorporation (Chafin et al., 1995; Cochet-Meilhac and Chambon, 1974), the interference with the trigger loop movement, shows that α-amanitin impairs nucleotide incorporation (Kaplan et al., 2008; Wang et al., 2006). Furthermore, in silico identification of the critical residues for α-amanitin binding to RNAP II support that α-amanitin interferes with the bridge helix residues and compromises the movement of the trigger loop (Garcia et al., 2014). Therefore, this amatoxin interferes in both nucleotide incorporation and RNAP II translocation, thereby stabilizing RNAP II in the elongation stage of transcription (Brueckner and Cramer, 2008; Kaplan et al., 2008; Wang et al., 2006).

Figure 2 – Interaction of α-amanitin with RNAP II.

α-Amanitin interacts with the bridge helix and trigger loop residues of RNAP II, leading to a disturbance in the transcription process due to the interference with nucleotide incorporation and RNAP II translocation. Source: Garcia et al., 2014 (license details and the terms and conditions provided by Elsevier and Copyright Clearance Center are presented in the appendix).

(31)

10

The interaction of α-amanitin with RNAP II causes a strong reduction of the elongation rate. Nevertheless, α-amanitin allows synthesis of multiple phosphodiester bonds without apparent dissociation of the elongation complex, since it does not influence nucleoside triphosphate affinity and does not abolish the addition of multiple nucleotides (Brueckner and Cramer, 2008; Chafin et al., 1995; Rudd and Luse, 1996). In fact, upon α-amanitin induced RNAP II stalling, three elongation complex conformations with high, medium or no sensitivity to α-amanitin inhibition were identified, which correspond to the intermediate, the pre- and the post-translocation state, respectively (Brueckner and Cramer, 2008; Gong et al., 2004). Since α-amanitin stays attached to the active elongation complex, the slow elongation may occur when the elongation complex briefly escapes suppression (Brueckner and Cramer, 2008). In fact, α-amanitin bounds with RNAP II, in particular with the trigger loop and bridge helix, apparently get broken during the nucleotide addition cycle, allowing RNAP II translocation; however, in the next cycle, inhibition is again restored (Brueckner and Cramer, 2008; Gong et al., 2004). Thus, the reduction in the elongation rate in the presence of this toxin can be due to the energy required to break the bounds of α-amanitin with RNAP II and not to irreversible links (Brueckner and Cramer, 2008).

In response to the strong reduction of the elongation rate caused by α-amanitin transcriptional blockage, Rpb1, the largest subunit of RNAP II, is degraded (Nguyen et al., 1996). In accordance, exposure of murine fibroblasts to α-amanitin caused a dose-dependent degradation of Rpb1. This degradation was dependent on the interaction between α-amanitin and the Rpb1 subunit, as α-amanitin resistant Rpb1 was not degraded (Nguyen et al., 1996). Notably, Rpb1 degradation was not perceptibly accompanied by the degradation of other subunits of RNAP II (Nguyen et al., 1996). Even though upon α-amanitin treatment the other RNAP II subunits remain intact, α-amanitin induces enzyme disassembly and causes cytoplasmic accumulation of Rpb3, due to an active export mechanism from nuclei that occurs when Rpb1 is degraded (Boulon et al., 2010).

Rpb1 degradation was then shown to be mediated by the ubiquitin pathway, as upon α-amanitin treatment, Rpb1 is ubiquitinated (Lee et al., 2002; Mitsui and Sharp, 1999). This ubiquitination can aim to target Rpb1 for degradation to relieve transcription blockage (Lee and Sharp, 2004). RNAP II ubiquitination induced by α-amanitin occurs in transcriptionally engaged RNAP II as demonstrated by the reactivity of ubiquitinated RNAP II with antibodies targeting CTD Ser5 and Ser2 phosphorylation (Lee et al., 2002). Additionally, Boulon et al., showed that when nuclear import of Rpb1 is impaired, Rpb1 is insensitive to amanitin, reinforcing that

(32)

α-11

amanitin specifically promotes degradation of the elongating form of RNAP (Boulon et al., 2010). Despite the degradative ubiquitin signaling, a nondegradative signaling was also observed following α-amanitin-dependent transcription arrest, through RNAP II polyubiquitination on Lys 63 (Lee and Sharp, 2004). This signaling has also been shown to be involved in DNA damage, recruiting downstream factors to alleviate the transcriptional blockage, such as repair proteins (Lee and Sharp, 2004). In fact, transcription inhibition by α-amanitin stimulated polyubiquitination of RNAP II in a similar pattern to the one triggered by either UV-radiation (Anindya et al., 2007) or cisplatin (Lee et al., 2002), which also induce transcriptional stalling. Nonetheless, stalled RNAP II complexes seem to present different stabilities depending on the transcriptional blockage inducing agent, as cisplatin lesion complexes are much more stable that RNAP II complexes stalled by α-amanitin (Jung and Lippard, 2006).

1.2.1.3 Apoptosis induction following transcriptional blockage

Blockage of RNAP II by α-amanitin in Chinese hamster ovary cells, following exposure to 10mg/mL of α-amanitin for 27h, was coordinately related to nonrandom degradation of cellular chromatin, in a similar pattern to internucleosomal DNA cleavage, a marker associated with apoptosis (Damgaard et al., 1996).

Afterwards, it was shown that prolonged inhibition of transcription seems to induce apoptosis through a p53-dependent mechanism (Blagosklonny et al., 2002). In fact, blockage of RNAP II and inhibition of mRNA synthesis trigger p53 accumulation and activation (Ljungman et al., 1999). Accordingly, in human fibroblasts a strong correlation between the dose required to inhibit mRNA synthesis by α-amanitin and p53 accumulation was observed (Ljungman et al., 1999). Regarding the mechanisms of α-amanitin p53-induced apoptosis, since transcription is blocked, transcription of p53-induced proapoptotic genes, such as Bax and Noxa, is also blocked (Arima et al., 2005). Nevertheless, inhibition of transcription also eliminates the expression of other p53-dependent genes, such as Mdm-2, which target p53 for degradation, thus leading to p53 accumulation (An et al., 1998). In fact, incubation with α-amanitin 10µM for 16h caused a significant reduction in Mdm-2 protein levels in MCF-7 cells, a breast cancer cell line, which accompanied p53 accumulation (Blagosklonny et al., 2002). Additionally, the CDK inhibitor, p21, presents altered protein levels following incubation with α-amanitin. p21 is a

(33)

12

major target of p53 and is often responsible for linking DNA damage to cell cycle arrest (Oren, 2003). Arima et al. reported that treatment of the colon cancer cell line HCT116 with 10µg/mL of α-amanitin for 24h caused a reduction in p21 protein levels, achieved by both transcriptional suppression and increased protein degradation (Arima et al., 2005). This downregulation of p21 allows cells to progress into S phase and undergo apoptosis (Arima et al., 2005). Moreover, downregulation of p21 may also contribute to the increased p53 levels, in accordance to Blagosklonny et al. who showed that p21 is required for normal p53 degradation in HCT166 cells (Blagosklonny et al., 2002). Additionally, the increase in p53 levels after α-amanitin exposure can also be owed to p53 phosphorylation, which favors its stabilization and activation (Arima et al., 2005).

While the transcription of p53-induced proapoptotic genes is blocked by α-amanitin (Arima et al., 2005), a transcription independent extranuclear function of p53 in apoptosis induction has been described. The extranuclear and transcription independent pro-apoptotic function of p53 is related with its accumulation in the cytosol and mitochondria, where p53 appears to induce apoptosis by interacting with members of the Bcl-2 family (Chipuk and Green, 2006; Mihara et al., 2003; Speidel, 2010). In fact, following α-amanitin treatment, p53 has been shown to accumulate in the mitochondria and to induce apoptosis (Arima et al., 2005; Leu and George, 2007). Accordingly, treatment of HepG2 cells with 10µg/mL of α-amanitin resulted in p53 mitochondrial localization and apoptosis induction, and both aspects were correlated with the interaction of p53 with Bak (Leu and George, 2007). Bak is a Bcl-2 protein - Bcl-2 antagonist/killer - that exerts a pro-apoptotic function leading to mitochondrial outer-membrane permeabilization, consequently causing the release of cytochrome c (Chipuk and Green, 2006). To further understand the requirement of p53 and its binding partner Bak to apoptosis induction in α-amanitin poisoning, 5µg/g of α-amanitin was intraperitoneally injected to p53-null and Bak-null animals and liver damage and apoptosis induction were analyzed (Leu and George, 2007). p53 and Bak deficient animals presented reduced liver damage and a lower proportion of TUNEL (terminal deoxynucleotidyl transferase-mediated biotin-dUTP nick-end labelling)-positive and caspase-3 positive apoptotic cells, when compared to wild type animals administered to the same dose of α-amanitin, leading to the conclusion that absence of Bak or absence of p53 provides protection against α-amanitin induced liver damage (Leu and George, 2007). Nevertheless, exposure of HepG2 cells to α-amanitin 10µg/mL also caused a decrease in the anti-apoptotic Bcl-xL and led to activation of caspase-8 and caspase-3 (Leu and George,

(34)

α-13

amanitin was blocked with 20µM of the broad-spectrum caspase inhibitor Z-Val-Ala-Asp(OMe)-fmk (Arima et al., 2005).

α-Amanitin induced changes in gene and protein expression

In addition, α-amanitin induces changes in gene expression. In fact, analysis of blood samples 24h following mice intravenous injection of α-amanitin (0.357mg/kg) caused alterations in the expression of 85 genes, of which, Catnβ was upregulated whereas the expression of Flt3-L, IL-7r and Rpo2-4 was downregulated (Chen et al., 2009). Also, in another study, mice intravenous injection of α-amanitin (0.357mg/kg) caused alterations in the expression of 146 genes, of which 66 had lower expression levels, whereas the expression of 80 genes was increased (Zhao et al., 2006). Genes which encoded proteins regulated by RNAP II were inhibited, as well as genes encoding proteins involved in the transcription process and mRNA splicing (Zhao et al., 2006). On the other hand, negative regulators of RNAP II, such as Zfpm1, and the transcription factor Fos increased after α-amanitin administration (Zhao et al., 2006). Furthermore, transcription inhibition with concentrations of α-amanitin up to 30 µg/mL enhanced transcription driven by human immunodeficiency virus (HIV) long terminal repeats (LTR) at the level of transcription elongation, as detected by an increase in luciferase expression in murine Ltk cells stably transfected with a plasmid pHIVLucA41, on which luciferase cDNA has been placed under the control of HIV LTR (Casse et al., 1999). This seems to be achieved by promoting major chromatin rearrangements that may increase the accessibility to CTD kinases or decrease its accessibility to phosphatases, therefore enhancing CTD phosphorylation (Casse et al., 1999). Moreover, stress activation of HIV LTR seems to be caused by general inhibition of transcription. In fact, other conditions, (e.g. UV radiation and DNA damage) that also partially inhibit the global transcriptional activity, activate HIV LTR transcription and promote CTD phosphorylation (Carrier et al., 1994; Casse et al., 1999; Kumar et al., 1996; Valerie et al., 1988).

Additionally, Tsao et al., showed that α-amanitin treatment causes selective protein loss (Tsao et al., 2012). In fact, the protein expression of DSIF160 and cyclin F was significantly reduced after a 14-h incubation of COS cells with α-amanitin 20 µg/mL (Tsao et al., 2012). The authors showed that the decrease in protein expression was a result of an induced accelerated degradation of the above-mentioned proteins by α-amanitin, as treatment with the proteasome

(35)

14

inhibitor MG132, rescued DSIF160 and cyclin F from α-amanitin induced loss (Tsao et al., 2012). Finally, the authors suggested that besides DSIF160, other Rpb1-interacting proteins could be at risk for continuous co-degradation in α-amanitin treated cells (Tsao et al., 2012).

Inflammation and oxidative stress

Inflammation and oxidative stress are pathophysiological events tightly linked with one another (Biswas, 2016). In fact, activation of inflammatory cells leads to the release of reactive species (RS) and causes oxidative stress, while these RS can also initiate an intracellular cascade that enhances the expression of proinflammatory genes (Biswas, 2016).

Tumor necrosis factor (TNF), a cytokine involved in systemic inflammation, has been implied in α-amanitin toxicity. In fact, TNF-α seems to play a pivotal role in α-amanitin hepatotoxicity since mice treated with anti-TNF-α antibodies and transgenic mice lacking the TNF-α receptor exhibited low hepatotoxicity (Leist et al., 1997). TNF-α appears to shorten the latency period associated with α-amanitin intoxications acting synergistically with this amatoxin (El-Bahay et al., 1999; Leist et al., 1997). Additionally, transcription blockage by α-amanitin sensitizes hepatocytes towards apoptosis induced by TNF-α (Leist et al., 1994; Leist et al., 1997). Moreover, TNF-α can activate the transcription factor nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB), a crucial transcription factor that participates in a number of physiological and pathological conditions, including inflammation and oxidative stress (Biswas, 2016). Accordingly, Garcia et al. showed that intraperitoneal administration of α-amanitin to mice induces activation NF-κB in hepatocytes and in macrophage-like cells in the liver and in the kidney (Garcia et al., 2015b).

Besides the induction of a pro-inflammatory state, α-amanitin administration also causes alterations in the activities of superoxide dismutase and catalase, crucial enzymes for the prevention of oxidative stress-related injury. In fact, exposure of mice or human hepatocytes to α-amanitin caused an increase in superoxide dismutase activity (Magdalan et al., 2011; Marciniak et al., 2013; Zheleva et al., 2007). Regarding catalase, administration of α-amanitin doses above 0.5mg/kg to mice caused a decrease in catalase activity in the liver (Zheleva et al., 2007), whereas administration of a lower dose (0.25mg/kg), significantly increased catalase activity in a putative adaptive state (Marciniak et al., 2013). Moreover, exposure to α-amanitin 2µM for 48h also caused a reduction in catalase activity in human hepatocytes (Magdalan et