ROLE OF Ncr1p IN ENDOPLASMIC RETICULUM STRESS RESPONSE IN Saccharomyces ... · descontraídos. À Sara Silva e ao João Ferreira, que partilharam comigo a experiência de um ano de

ROLE OF Ncr1p IN ENDOPLASMIC RETICULUM STRESS

RESPONSE IN Saccharomyces cerevisiae

JOANA FILIPA MADUREIRA GAIFEM

Dissertação de Mestrado em Contaminação e Toxicologia Ambientais

2011

JOANA FILIPA MADUREIRA GAIFEM

ROLE OF Ncr1p IN ENDOPLASMIC RETICULUM STRESS

RESPONSE IN Saccharomyces cerevisiae

Dissertação de Candidatura ao grau de

Mestre em Contaminação e Toxicologia

Ambientais submetida ao Instituto de

Ciências Biomédicas de Abel Salazar da

Universidade do Porto.

Orientador – Doutor Vítor Costa

Categoria – Professor Associado

Afiliação – Instituto de Biologia Molecular e

Celular; Instituto de Ciências Biomédicas de

Abel Salazar

Role of Ncr1p in endoplasmic reticulum stress response in Saccharomyces cerevisiae

1

Acknowledgements

O trabalho apresentado nesta tese contou com o contributo importante de várias

pessoas, sem as quais a sua realização seria impossível ou estaria severamente

condicionada. Como tal, gostaria de lhes expressar os meus agradecimentos.

Em primeiro lugar, gostaria de agradecer ao meu orientador, o Professor Doutor Vítor

Costa, por ter aceitado orientar-me neste trabalho e por todos os contributos que deu

para a elaboração desta tese. Todos os conselhos, ensinamentos e correcções foram

essenciais para o desenvolvimento deste trabalho e para o meu crescimento profissional.

Muito obrigado Professor!

Agradeço à Rita Vilaça, que co-orientou este trabalho e que contribuiu de forma

fundamental para a realização do mesmo. Todas as interacções tiveram o seu papel na

minha aprendizagem, na criação de dinâmica de trabalho e na evolução do projecto. Por

todos estes motivos e pela disponibilidade demonstrada ao longo de todo o ano de

trabalho, o meu obrigado.

Porque o companheirismo e bom ambiente contribuem de forma notória para a

elaboração de um bom trabalho, quero expressar a minha gratidão a todos os elementos

do laboratório com quem pude conviver ao longo deste ano de trabalho. Ao Daniel,

agradeço por toda a disponibilidade em ajudar em diversos aspectos do meu trabalho,

assim como pelos conselhos dados na elaboração desta tese, e, não menos importante,

agradeço pela amizade espontânea e pelas conversas diárias, no laboratório ou fora

dele. Quero agradecer à Vanda, não só pelo convívio, mas também pelos contributos que

deu ao meu trabalho. Agradeço à Catarina Santos, que se revelou uma boa amiga,

sempre com uma palavra certa no momento certo, para além dos momentos de

descontracção. À Sílvia, agradeço toda a alegria que transmite no laboratório, bem como

a disponibilidade para ajudar no necessário. Quero agradecer à Catarina Pacheco e à

Maria João pelas conversas diárias que proporcionaram momentos agradáveis e

descontraídos. À Sara Silva e ao João Ferreira, que partilharam comigo a experiência de

um ano de trabalho para a elaboração de uma tese de mestrado, o meu obrigado.

Agradeço ao Prof. Dr. Pedro Moradas-Ferreira pelo interesse demonstrado no trabalho.

Não posso deixar de agradecer a todos os elementos dos grupos de Redox Cell

Signalling, Cellular and Applied Microbiology e Bioengineering and Synthetic Microbiology


2

pelo companheirismo e pelo excelente ambiente de trabalho. À Ângela, Filipe, Gabriel,

Ivan, Kalina, Marta, Miguel, Nuno, Prof. Dr.ª Paula Tamagnini, Prof. Dr.ª Arlete Santos,

Pedro, Rodrigo, Rita Mota, Sara Pereira, Tiago, Vítor e Zille, muito obrigado. Deixo

também os meus agradecimentos à Liliana Correia e à Helena Pinho, pela simpatia e

pela disponibilidade em ajudar no que fosse necessário.

A todos os meus amigos, o meu muito obrigado! Todos contribuíram para a

realização deste trabalho, pelo apoio e confiança que sempre me transmitiram. Um

agradecimento especial à Ana Luísa e ao Tiago Miguel, que mesmo à distância sempre

me apoiaram e de quem me orgulho de ser amiga. Aos meus colegas de mestrado,

agradeço a amizade e os bons momentos partilhados neste ciclo novo para todos nós, e

em particular, à Alexandra, com quem construí uma bela amizade e com quem partilhei

momentos de grande alegria e companheirismo.

Como não pode deixar de ser, quero agradecer a toda a minha família por todo o

apoio e confiança que me deram, e em particular aos meus avós António e Ana. Graças

ao seu apoio, não só durante este ano, mas ao longo de toda a minha vida, e a todos os

ensinamentos que me transmitiram, pude crescer como pessoa e chegar mais longe.

Teria sido muito mais difícil sem eles. Muito obrigado por tudo!

Por fim, agradeço às pessoas que convivem comigo diariamente e que

acompanharam mais de perto todos os meus passos. Aos meus pais, agradeço-lhes do

fundo do coração todo o carinho e confiança que sempre me transmitiram. Não há

palavras suficientes para descrever a gratidão que sinto por tudo o que fizeram por mim.

Em todos os momentos, não só este ano, mas ao longo de toda a minha vida, estiveram

presentes para me dar a mão sempre que precisei, para me mostrar o que está certo e o

que está errado, para me tornarem numa pessoa digna de chegar cada vez mais longe.

Devo-vos tudo o que sou. Ao meu irmão Bruno, dedico-lhe esta tese, pois desde o dia

que nasceu que é a minha maior inspiração. Todos os momentos de brincadeira, de

amizade e de apoio foram preponderantes para me dar a força necessária para os

momentos mais importantes. É o melhor irmão do mundo e tenho muito orgulho na

pessoa maravilhosa que é. Por último, quero agradecer ao Filipe, por todas as razões.

Pela confiança, pela amizade, por estar sempre ao meu lado, em momentos bons e

menos bons. É uma honra poder caminhar ao seu lado e com o seu apoio sei que

chegarei mais longe. Muito obrigado por tudo!


3

Table of Contents

Acknowledgements ........................................................................................................... 1

Table of Contents ............................................................................................................. 3

Figure Index ...................................................................................................................... 5

Table Index ....................................................................................................................... 5

Abbreviations .................................................................................................................... 7

Abstract ............................................................................................................................ 9

Resumo ...........................................................................................................................11

Chapter I - Introduction ....................................................................................................13

I-1. Niemann-Pick type C disease ................................................................................15

I-1.1. Epidemiology ...................................................................................................16

I-1.2. Clinical description and diagnosis ....................................................................16

I-1.3. Lipid-trafficking defects in NPC ........................................................................17

I-1.4. NPC genes and proteins ..................................................................................18

I-1.5. Conservation during evolution – NCR1, the yeast orthologue of hNPC1 ..........22

I-2. Endoplasmic reticulum stress .................................................................................23

I-2.1. Mechanism of protein folding ...........................................................................24

I-2.2. Recognition of unfolded proteins ......................................................................25

I-2.3. Transduction of the unfolded protein signal across the ER membrane .............25

I-2.4. Activation of protective responses by the UPR – IRE1 .....................................26

I-2.5. Role of IRE1 and HAC1 in membrane proliferation control ..............................28

I-2.6. Endoplasmic reticulum-associated degradation (ERAD) ..................................29

I-2.7. Cell signaling pathways related with ER stress conditions – the High Osmolarity Glycerol (HOG) pathway ...........................................................................................30

I-3. Saccharomyces cerevisiae as biological model ......................................................32

Chapter II – Aim of the work .............................................................................................33

Chapter III – Material and Methods ..................................................................................37

III-1. Yeast strains and growth conditions .....................................................................39

III-2. Genomic DNA extraction ......................................................................................40

III-3. Colony PCR and genomic DNA PCR ...................................................................40

III-4. Gene disruption ....................................................................................................41


4

III-5. Yeast electroporation ........................................................................................... 42

III-5.1. Preparation of electrocompetent cells ............................................................ 42

III-5.2. Electro-transformation and plating ................................................................. 42

III-6. Stress resistance ................................................................................................. 43

III-7. Oxidative stress markers ..................................................................................... 43

III-7.1. Quantification of reactive oxygen species (ROS) ........................................... 43

III-7.2. Quantification of glutathione levels ................................................................ 44

III-8. β-Galactosidase activity ....................................................................................... 44

III-9. ERAD assay ........................................................................................................ 45

III-10. Induction of the HOG pathway ........................................................................... 46

III-11. Statistical analysis ............................................................................................. 47

Chapter IV – Results and Discussion .............................................................................. 49

IV-1. Ncr1p deficiency decreases tunicamycin-induced growth arrest .......................... 51

IV-2. Ncr1p deficiency decreases tunicamycin-induced intracellular oxidation and glutathione depletion .................................................................................................... 53

IV-3. UPR activation in ncr1Δ cells ............................................................................... 56

III-4. ERAD is not inhibited by tunicamycin in ncr1Δ cells ............................................. 58

IV-5. Activation of Hog1p is compromised in ncr1Δ cells .............................................. 60

Chapter V – General Discussion and Future Perspectives .............................................. 63

References ...................................................................................................................... 69


5

Figure Index

Figure I.1. Lipid trafficking defects in NPC disease.. ........................................................18

Figure I.2. Topology of NPC1 ...........................................................................................19

Figure I.3. The Ire1p cascade of the UPR pathway ..........................................................27

Figure I.4. The High Osmolarity Glycerol (HOG) pathway.. ..............................................31

Figure IV.1. The ncr1Δ cells are resistant to tunicamycin.. ...............................................51

Figure IV.2. Effect of tunicamycin on cell growth. .............................................................52

Figure IV.3. Analysis of tunicamycin resistance. ..............................................................53

Figure IV.4. Analysis of ROS levels. ................................................................................54

Figure IV.5. Effect of tunicamycin on glutathione levels. ..................................................55

Figure IV.6. Structure of UPRE-lacZ gene reporter. .........................................................56

Figure IV.7. Hac1p activation by ER stress ......................................................................57

Figure IV.8. HAC1 deletion increases the sensitivity of ncr1Δ cells to tunicamycin ..........58

Figure IV.9. Analysis of CPY* stability ..............................................................................59

Figure IV.10. Quantification of CPY* decay. .....................................................................59

Figure IV.11. Hog1p phosphorylation is decreased in ncr1Δ cells ....................................60

Table Index

Table III.1. Yeast strains used in this work. ......................................................................39

Table III.2. Primers used in this work. ..............................................................................41

Table III.3. Reagents used in the preparation of a polyacrylamide gel. ............................46


7

Abbreviations

ATF – Activating Transcription Factors

ATF6 – Activating Transcription Factor 6

ATP – Adenosine Triphosphate

BSA – Bovine Serum Albumin

bZIP – Basic Leucine Zipper Domain

cAMP – Cyclic Adenosine Monophosphate

CDP – Cytidine Diphosphate

CFU –Colony Forming Units

CH - Cycloheximide

CNS – Central Nervous System

CPY* – Carboxypeptidase Y

CREB – cAMP Response Element Binding

DMSO – Dimethylsulfoxide

DNA – Deoxyribonucleic Acid

dNTP – Deoxyribonucleotides

DTNB – 5,5'-Dithiobis-(2-Nitrobenzoic Acid)

DTT – Dithiothreitol

EDEM – ER Degradation-Enhancing α-Mannosidase-like Protein

EDTA – Ethylenediaminetetraacetic Acid

ER – Endoplasmic Reticulum

ERAD – Endoplasmic Reticulum-Associated Degradation

GFP – Green Fluorescent Protein

GLS – Golgi Localization Sequences

GSH – Glutathione (reduced form)

GSSH – Glutathione (oxidized form)

GST – Glutathione S-Transferases

HOG – High Osmolarity Glycerol

H2DCF-DA – 2’-7’-Dichlorodihydrofluorescein diacetate

IRE1 – Inositol-Requiring Protein 1

LDL – Low-Density Lipoprotein

MAPK – Mitogen-Activated Protein Kinase

MOPS – 4-Morpholinepropanesulfonic Acid

mRNA – Messenger RNA

NADPH – Nicotinamide Adenine Dinucleotide Phosphate

NPC – Niemann-Pick type C


8

OD – Optical Density

ONPG – o-nitrophenylgalactopyranosyde

PAGE – Polyacrylamide Gel Electrophoresis

PBS – Phosphate Buffered Saline

PCR – Polymerase Chain Reaction

PDI – Protein Disulfide Isomerases

PERK – Protein kinase RNA-like ER kinase

PMSF – Phenylmethylsulfonyl Fluoride

RNA – Ribonucleic Acid

rpm – Revolutions Per Minute

ROS – Reactive Oxygen Species

SAPK – Stress-Activated Protein Kinase

SC – Synthetic Complete

SD – Standard Deviation

SDS – Sodium Dodecyl Sulfate

SSD – Sterol Sensing Domain

TEMED – N,N,N,N-Tetramethylethylenediamine

TOR – Target of Rapamycin

TPBS – Tween Phosphate Buffered Saline

tRNA – Transfer RNA

TBS – Tris Buffered Saline

TTBS – Tris-Tween Buffered Saline

TUN - Tunicamycin

UPR – Unfolded Protein Response

UPRE – Unfolded Protein Response Element

UPS – Ubiquitin-Proteasome System

UTR – Untranslated Region

wt – wild-type

XBP1 – X-box Binding Protein-1

YPD – Yeast extract Peptone Glycerol


9

Abstract

Niemann-Pick Type C (NPC) is an autossomal recessive lipid storage disease

characterized by abnormal cholesterol trafficking and intracellular accumulation in late

endosomes and lysosomes. NPC disease is related to a progressive neurodegenerative

phenotype and is caused by loss-of-function point mutations in either NPC1 or NPC2.

Both proteins seem to regulate intracellular lipid transport through lysosomes and

endosomes. Several lipid disorders display evidences of endoplasmic reticulum (ER)

stress. Cell adaptation to ER stress is mediated by the unfolded protein response (UPR).

This signal transduction pathway detects unfolded proteins in the lumen of ER and

reduces stress by increasing the folding capacity of ER or triggers apoptosis of irreversibly

damaged cells.

In Saccharomyces cerevisiae, the vacuolar proteins Ncr1p and Npc2p are

orthologues of human NPC1 and NPC2, respectively. Yeast Ncr1p and Npc2p are

involved in ergosterol trafficking and can functionally complement the loss of function of

human NPC1 and NPC2, being able to suppress lipid trafficking defects associated with

NPC1 and NPC2 mutations. Therefore, studies using yeast as an eukaryotic model may

be useful to uncover the function of these proteins and to characterize molecular

mechanisms associated with NPC disease.

In this study, S. cerevisiae ncr1Δ mutant cells were used as a model system to study

the role of Ncr1p in ER stress response. Parental and ncr1Δ cells were treated with

tunicamycin, a drug that inhibits protein glycosylation and consequently activates UPR.

The results showed an increased resistance of ncr1Δ cells to tunicamycin and that Ncr1p

deficiency seems to have protective effects to yeast cells from tunicamycin-induced

growth arrest. The analysis of oxidative stress markers showed that tunicamycin

specifically decreased glutathione levels in parental cells, but not in ncr1Δ mutants.

However, ncr1Δ cells exhibited higher levels of reactive oxygen species. Notably, the

induction of a reporter gene controlled by Hac1p, the transcription factor involved in the

UPR, was suppressed in ncr1Δ cells exposed to tunicamycin. This effect is specific for this

drug since UPR was induced in both parental and ncr1Δ cells treated with dithiothreitol

(DTT), a compound that impairs disulfide bond formation. The hac1Δncr1Δ double mutant

displayed a higher sensitivity to tunicamycin when compared to ncr1Δ mutant, but was

more resistant than hac1Δ cells, suggesting that, for tunicamycin exposure, lack of NCR1

has protective effects by a Hac1p-independent mechanism. The analysis of endoplasmic

reticulum-associated degradation (ERAD) showed that after 1 h of tunicamycin exposure

this system is induced in ncr1Δ mutant cells but not in parental cells, in which tunicamycin

inhibits ERAD system by saturation of its capacity. The study of the High Osmolarity


10

Glycerol (HOG) pathway showed that Hog1p and phospho-Hog1p levels increased with

tunicamycin exposure in parental cells, but not in ncr1Δ mutant cells, indicating that the

resistance of ncr1Δ cells to tunicamycin is Hog1p-independent. These data suggest that

Ncr1p deficiency increases ER stress resistance induced by tunicamycin exposure via an

uncharacterized mechanism.


11

Resumo

Niemann-Pick tipo C (NPC) é uma doença lipídica autossómica recessiva,

caracterizada por disfunções no tráfego de colesterol e a sua acumulação a nível

intracelular nos endossomas e lisossomas. A degeneração neurológica está associada a

esta doença, sendo provocada por mutações pontuais nos genes NPC1 ou NPC2. Estas

proteínas aparentam regular o transporte intracelular de lípidos através dos lisossomas e

endossomas. Diversas doenças relacionadas com distúrbios lipídicos apresentam

evidências de stress do retículo endoplasmático. A adaptação celular ao stress do

retículo é mediada pela resposta à acumulação de proteínas mal conformacionadas,

designada por “unfolded protein response” (UPR). Esta via de transdução do sinal detecta

proteínas com conformações incorrectas no lúmen do retículo e, através do aumento da

capacidade de conformação do retículo, reduz o stress ou, caso os danos celulares

sejam irreversíveis, inicia o processo de apoptose.

Na levedura Saccharomyces cerevisiae, as proteínas vacuolares Ncr1p e Npc2p são

ortólogas da NPC1 e NPC2 humanas, respectivamente, e estão envolvidas no transporte

e tráfego de ergosterol. Ambas as proteínas Ncr1p e Npc2p podem complementar a

perda de função das respectivas proteínas humanas, suprimindo as anomalias ao nível

do tráfego lipídico associadas às mutações em NPC1 e NPC2. Como tal, a utilização da

levedura como modelo eucariótico pode ser vantajosa para o estudo das funções dessas

proteínas e para a caracterização de mecanismos moleculares relacionados com a

doença de NPC.

Neste trabalho, foram usados mutantes de S. cerevisiae ncr1Δ como modelo para o

estudo do papel da Ncr1p na resposta ao stress do retículo. Células parentais e do

mutante ncr1Δ foram tratadas com tunicamicina, um composto que inibe a glicosilação de

proteínas e consequentemente activa a UPR. Os resultados demonstraram uma maior

resistência do mutante ncr1Δ à tunicamicina e a deficiência em Ncr1p diminui a inibição

do crescimento induzida por este composto. A análise de marcadores de stress oxidativo

mostrou uma diminuição nos níveis de glutationa induzida pela tunicamicina nas células

parentais, contrariamente ao observado nos mutantes ncr1Δ. Todavia, estes mutantes

apresentaram níveis de espécies reactivas de oxigénio superiores aos das células

parentais. A indução de um gene repórter controlado pela proteína Hac1p, factor de

transcrição associado à UPR, através da exposição com tunicamicina, foi suprimida nos

mutantes ncr1Δ. Este efeito é específico para a tunicamicina uma vez que a UPR foi

induzida nas células parentais e no mutante após tratamento com ditiotreitol (DTT), um

composto que compromete a formação de ligações dissulfureto. O duplo mutante

ncr1Δhac1Δ apresentou uma maior sensibilidade à tunicamicina comparativamente ao


12

mutante ncr1Δ, mas também uma maior resistência que o mutante hac1Δ, o que sugere

que a delecção do gene NCR1 surte efeitos protectores por um mecanismo independente

do HAC1. Através da análise do sistema de degradação associada ao retículo

endoplasmático (ERAD) verificou-se que após uma hora de exposição à tunicamicina,

este sistema é induzido nos mutantes ncr1Δ, ao contrário do que sucede nas células

parentais, nas quais a tunicamicina inibe o sistema de ERAD devido à saturação da sua

capacidade. O estudo da via de alta osmolaridade do glicerol (HOG) demonstrou que os

níveis de Hog1p e de Hog1p na forma fosforilada aumentaram com exposição à

tunicamicina nas células parentais, mas não nos mutantes ncr1Δ, o que parece indicar

que a resistência dos mutantes ncr1Δ é independente da proteína Hog1p. Estes dados

sugerem que a deficiência na proteína Ncr1p aumenta a resistência ao stress do retículo

endoplasmático induzido pela exposição à tunicamicina através de um mecanismo não

identificado.


13

Chapter I

Introduction


15

I-1. Niemann-Pick type C disease

Sphingolipid storage diseases are a group of approximately forty genetic disorders,

caused by inherited defects of lysosomal hydrolytic processes or lipid transport that leads

to intracellular accumulations of cholesterol and lipids in the endosomal-lysosomal system

(Pacheco & Lieberman, 2008). Among this group is Niemann-Pick disease.

Niemann-Pick type C (NPC) disease, along with types A and B, belongs to the

Niemann-Pick group of lipidoses (Ikonen & Holtta-Vuori, 2004). This group of diseases

was first described in the late 1920’s by Albert Niemann and Ludwig Pick, as a

heterogeneous group of autossomal recessive lysosomal lipid storage disorders, with or

without neurological involvement, with regular features of hepatosplenomegaly and

sphingomyelin storage in reticuloendothelial and parenchymal tissues. It was later

demonstrated that there is a broad variability in age of onset, clinical expression and in the

level of sphingomyelin storage in tissues (Crocker & Farber, 1958), which led to a

classification of the disease into different groups (Crocker, 1961). Types A and B are

caused by loss-of-function mutations in the acid sphingomyelinase gene (Vanier & Millat,

2003). Type C was described as having a sub acute nervous system involvement, with

moderate/slower course and a mild visceral storage; however, later work led to a

reclassification of type C as a cellular lipid trafficking disorder, involving more specifically

endocytosed cholesterol (Pentchev et al., 1994). In NPC disease, cells fail to esterify

exogenously added cholesterol. This disorder is characterized by unique abnormalities of

intracellular transport of endocytosed cholesterol with accumulation of unesterified

cholesterol in endosomal/lysosomal compartment and the Golgi complex (Ikonen & Holtta-

Vuori, 2004; Vanier, 2010). Besides cholesterol sequestration, NPC cells can also

accumulate other lipids, in particular sphingolipids (Lusa et al., 2001; Puri et al., 1999;

Vanier, 1999; Zhang et al., 2001b). NPC is related to a progressive neurodegenerative

phenotype and in most cases is fatal (Patterson et al., 2001).

Advances in the knowledge of the disease led to the description of two genetic

complementation groups and the subsequent isolation of the two underlying genes: NPC1

and NPC2. They are represented in different proportions in the population – NPC1 is

involved in 95% of the cases (Patterson et al., 2001), while NPC2 is related to rare cases

(Vanier, 2010). NPC is caused by loss-of-function mutations in either NPC1 or NPC2

proteins, which mediate proper intracellular lipid transport through pathways that remain

unclear (Pacheco & Lieberman, 2008).

Introduction

16

I-1.1. Epidemiology

The prevalence of NPC disease is difficult to assess, due to insufficient clinical

awareness and difficult diagnosis. Estimates of birth prevalence for Western Europe have

been predictable to be 1 per 150,000 (Patterson et al., 2001). In Australia (Meikle et al.,

1999), The Netherlands (Poorthuis et al., 1999) and Portugal (Pinto et al., 2004) the

prevalence is 0.71, 0.53 and 3.3 per 150,000 births, respectively.

I-1.2. Clinical description and diagnosis

The clinical presentation of NPC is extremely heterogeneous, with patients

developing symptoms over a wide range of ages (Patterson et al., 2001). There is no

exact correlation between disease-causing mutations and the degree of severity of the

clinical phenotype (Vanier & Millat, 2003; Yamamoto et al., 2000). Similarly, the lifespan of

the patients varies between a few days (Spiegel et al., 2009) until over 60 years old

(Trendelenburg et al., 2006). This disease can be subdivided in four groups concerning to

the age of onset: early infantile, late infantile, juvenile and adult form of the disease.

However, the classic form of NPC, which encompasses approximately 70% of the cases,

presents between the ages of 3 and 15 years (Patterson et al., 2001). NPC severely

targets internal organs (mostly liver and spleen) and the first symptoms usually described

are hepatosplenomegaly (that seems to fluctuate and decrease with time) or obstructive

jaundice. The systemic involvement is usually severe, except for the perinatal period,

which is well tolerated (Vanier, 2010). Nevertheless, patients eventually develop

neurological and/or psychiatric symptoms, the severity of which is inversely associated

with lifespan (Imrie et al., 2002; Turpin et al., 1991).

The diagnosis of NPC disease is based on the analysis of dermal fibroblasts, with two

different approaches: a morphological approach, by filipin staining to detect the

accumulation of free cholesterol; and a biochemical approach, to monitor defective

cholesterol esterification in low density lipoprotein (LDL)-challenged cells (Vanier et al.,

1991). Currently, there are no effective treatments available to patients with this disorder

(Pacheco & Lieberman, 2008).

Despite the heterogeneity of the clinical symptoms of NPC disease, it is not totally

observed when it comes to the biochemical level of the disease. The majority of cases

present prominent accumulations of unesterified cholesterol, sphingolipids and complex

gangliosides in late endosomes and lysosomes, but a subset of patients with specific

mutations reveals less lipid storage (Millat et al., 2001a).


17

The development of NPC is characterized by a liver and spleen enlargement, caused

by the presence of lipid-laden macrophages. Kupffer cells in the liver and splenic

macrophages display clear cytoplasmic vacuolization that results from the accumulation of

cholesterol, phospholipids and glycolipids (Pacheco & Lieberman, 2008). Impairment of

lipid trafficking also has severe consequences in the central nervous system (CNS),

leading to neuron loss throughout the brain (Walkley & Suzuki, 2004). The presence of

swollen neuronal cell bodies in many regions in the brain is also a feature of NPC and

reflects lipid accumulation in late endosomes and lysosomes. It is observed also in NPC

an intracellular aggregation of the microtubule-binding protein tau, which is biochemically

similar to aggregates in Alzheimer’s disease (Auer et al., 1995).

I-1.3. Lipid-trafficking defects in NPC

Lipid-trafficking defects within the NPC brain reflect deficiencies in the pathway by

which cholesterol and other lipids reach neurons and are sorted intracellularly (Pacheco &

Lieberman, 2008). Neurons and other CNS cell types get cholesterol they need through

endogenous synthesis or by uptake of lipoprotein cholesterol particles produced and

released within the nervous system (Mauch et al., 2001). Cells internalize these particles

and unesterified cholesterol and other lipids are trafficked from the endosomal-lysosomal

system to organelles which they are destined, such as Golgi complex and endoplasmic

reticulum (ER). In NPC cells, lipoprotein cholesterol particles are internalized without

disruption, but stay entrapped in endosomal-lysosomal system, creating an insufficient

efflux of these particles (Figure I.1). It leads to an accumulation of unesterified cholesterol,

sphingolipids and complex gangliosides in cytoplasmic vesicles and a simultaneous

scarcity of these lipids in organelles where they are required (Pacheco & Lieberman,

2008).

Introduction

18

Figure I.1. Lipid trafficking defects in NPC disease. Under normal conditions, cholesterol particles enter the

cell and are trafficked from endosomal-lysosomal system to the endoplasmic reticulum, Golgi complex and

other intracellular organelles. In cells lacking NPC1 or NPC2, lipid trafficking is inhibited, leading to their

accumulation in endosomes and lysosomes and no efflux to other intracellular compartments. Adapted from

Pacheco & Lieberman, 2008.

The transport of sphingolipids from endosomes to Golgi complex can also be blocked

by high levels of cholesterol (Vanier & Millat, 2003). Lower levels of cholesterol in the

Golgi complex and ER result in deleterious effects in processes dependent on proper

membrane composition and also in a scarcity of substrate for further synthetic reactions

(Wojtanik & Liscum, 2003). Due to cholesterol sequestration, the subsequent induction of

all low-density lipoprotein cholesterol-mediated homeostatic responses is retarded in NPC

cells. Studies in patients cells demonstrated that lysosomal storage of unesterified

cholesterol may show a changeable intensity; however, fibroblasts from a large amount of

heterozygotes display mild but definitive changes (Argoff et al., 1991; Vanier et al., 1991).

This impairment in the process of endocytosed cholesterol is essential for the

pathogenesis of NPC disease and can clarify a more general dysfunction of intracellular

lipid metabolism (Walkley & Vanier, 2009).

I-1.4. NPC genes and proteins

NPC disease is genetically heterogeneous, and it is possible to distinguish two

complementation groups. Genes responsible for the disease have already been

described. The NPC1 gene was identified in 1997, by positional cloning, as the gene

mutated in the major complementation group (Carstea et al., 1997). This gene encodes a

large membrane glycoprotein that is mainly localized in late endosomes-lysosomes


19

(Higgins et al., 1999). In 2000, it was shown that the gene defective in the minor group

was HE1/NPC2 (Naureckiene et al., 2000), which encodes a small soluble lysosomal

protein with high affinity to cholesterol (Storch & Xu, 2009). Both genes are conserved

during evolution, even in organisms in which cholesterol is not one of the most important

components of membranes. The two genetic groups are biochemically and clinically

impossible to differentiate and due to the resemblance in the disease phenotypes, it is

believed that NPC1 and NPC2 may share the same metabolic pathway (Naureckiene et

al., 2000; Sleat et al., 2004). Nonetheless, it was never verified any direct interaction

between these two proteins (Ikonen & Holtta-Vuori, 2004).

The NPC1 gene, localized in chromosome 18q11-q12, encodes a 1278 amino acid

integral membrane protein with 13 transmembrane domains. The NPC1 domain is a

highly conserved region (amino acids 55-165) with a leucine zipper motif. The large

cysteine-rich luminal loop (amino acids 855-1098) includes a ring-finger motif and is a

likely site for protein-protein interaction. A sterol sensing domain (SSD) (amino-acids 615-

797) displays high homology to SSD of other integral membrane proteins that act in

response to ER cholesterol (Vanier & Millat, 2003) (Figure I.2).

Figure I.2. Topology of NPC1. LE/Lys – Late endosomes/Lysosomes. TM = Transmembrane region.

Adapted from Lloyd-Evans & Platt, 2010.

The current number for identified NPC1 disease-causing mutations is close to 300,

with a large majority of missense mutations, and more than 60 polymorphisms of the gene

have also been described (Vanier, 2010). These missense mutations are scattered

through the NPC1 gene and influence all functional domains, except the leucine zipper

motif. While more than one-third of the mutations are located in the cysteine-rich luminal

loop, there is a hot spot between amino acids 927 and 958, which harbors the three most

frequent mutations (Vanier & Millat, 2003). The most common, in allele p.I1061T, is

particularly frequent and is related with prominent cellular cholesterol trafficking

disturbances in fibroblasts of patients and it is correlated with a juvenile neurological onset

of NPC (Millat et al., 1999). The I1061T mutant was shown to be a functional protein

targeted for endoplasmic reticulum-associated degradation (ERAD), due to protein

Introduction

20

misfolding (Gelsthorpe et al., 2008). Curiously, mutations corresponding to a less brutal

impairment of cellular trafficking are located in this loop (Millat et al., 1999; Ribeiro et al.,

2001; Sun et al., 2001).

Along with cysteine-rich luminal loop, SSD also reveals mutations that emphasized

the functional significance of both domains. Homozygous mutations in SSD seem to be

very deleterious, corresponding to a lack of mature NPC1 protein and to a very severe

phenotype at both clinical and biochemical levels (Millat et al., 2001b).

The NPC2 gene, located in chromosome 14q24.3, is connected with very severe

clinical phenotypes. Missense mutations in NPC2 have been associated to more diverse

phenotypes, including juvenile and adult onset patients (Millat et al., 2001a; Verot et al.,

2007). The mature NPC2 is a glycoprotein with a ubiquitous expression in several tissues

(Naureckiene et al., 2000). Studies demonstrated a higher affinity binding and identified a

hydrophobic cholesterol-binding pocket around amino acid K97 (Ko et al., 2003). There

are few cases of NPC2 disease, but all present striking abnormalities of cellular

cholesterol processing. It was suggested that NPC1 could be a regulator of NPC2

transport, but it was not confirmed. With the increase of NPC2 cases, it is clear that it has

high heterogeneity as NPC1 (Vanier & Millat, 2003).

The exact functions of NPC1 and NPC2 have not yet been described (Vanier & Millat,

2003). The loss of function of both genes results in a versatile cellular pathology and,

contrary to several other lipidoses that result from defects in enzyme activity, NPC seems

to represent a primary transport defect. The failure of cholesterol homeostasis in NPC

cells is known but whether the cholesterol transport defect is the main problem or

potentially a consequence of some other malfunction remains unclear (Ikonen & Holtta-

Vuori, 2004).

The majority of cell biological studies about NPC pathology are referent to cells

defective in NPC1. NPC2 patients are rare and there are less models of study – a knock-

out mouse model has only become available recently, contrary to NPC1 models, such as

fibroblasts from affected patients, cells from the natural NPC1 -/- mouse (Loftus et al.,

1997) and several cell lines in which NPC1 has been mutated (Millard et al., 2000). In the

first studies, only biochemical assays were made concerning cholesterol trafficking, but

actually there are already tools for morphological analysis of defective sterols, such as

filipin staining (Coxey et al., 1993).

Some proteins related with cholesterol trafficking can also be involved in the

generation of impairments in NPC cholesterol trafficking. ABCA1, a protein involved in the

removal of cholesterol to apolipoprotein A-I, is suggested to resort to the endosomal-

lysosomal pool to get cholesterol for its efflux (Chen et al., 2001). In fact, ABCA1-

mediated cholesterol efflux is decreased in NPC1-deficient cells and the level of ABCA1


21

protein is also lower in these cells. In contrast, sequestration of NPC1 in the cholesterol-

laden late endocytic organelles in ABCA1-deficient (Tangier disease) fibroblasts was

recently observed, suggesting that the functions of both proteins are associated (Neufeld

et al., 2004).

The endosomal-lysosomal cholesterol sequestration in cells lacking NPC1 is

paralleled by the failure of distribution of cholesterol to several intracellular compartments,

such as the Golgi complex (Coxey et al., 1993), mitochondria (Frolov et al., 2003) and the

ER (Neufeld et al., 1996). A defective cholesterol esterification and an impaired

downregulation of cholesterol synthesis under cholesterol loading conditions indicate that

probably there is a cholesterol deprivation in the ER of NPC cells (Brown & Goldstein,

1999; Liscum & Faust, 1987; Neufeld et al., 1996). Nevertheless, the analysis of

cholesterol esterification in vitro using cell homogenates indicates that the ER cholesterol

level in NPC cells is more or less normal, and the only impairment observed was the

response to LDL-loading (Frolov et al., 2003; Lange et al., 2000).

The function of vesicular transport in endocytic cholesterol trafficking has been deeply

studied and it is well established the role of endocytic pathway in the transport of LDL

particles to organelles. The exit of cholesterol from late endosomes and lysosomes needs

functional vesicular machinery, and vesicular trafficking defects are involved in NPC.

Studies with green fluorescent protein (GFP)-fusion NPC1 protein in living fibroblast

cultures have shown that this compartment undergoes rapid movements that are strikingly

impaired in NPC1-mutant cells. These observations suggest that NPC1 is required for the

production of tubulovesicular structures that show loss of flexibility and slower rate of

movement in cells lacking NPC1. The NPC1-containing vesicles carry cholesterol from the

perinuclear regions to the cell periphery. These structures were also observed to interact

with the ER NPC1 (Ko et al., 2001; Zhang et al., 2001a).

It is possible to put forward two different scenarios concerning to vesicular transport

and cholesterol accumulation: loss of NPC1 activity leads to an impaired motility and

subsequently accumulation of cholesterol; instead, the accumulation of cholesterol

accounts for the lack of motility – and it is a secondary defect. Since the available

knowledge is limitative for conclusions, it seems plausible that both scenarios contribute to

the phenotype (Ikonen & Holtta-Vuori, 2004).

Despite some ambiguous results, some experimental data suggest that NPC1 plays a

role in regulation or mediation of retrograde transport of lysosomal cargo in the late

endosomal-lysosomal pathway. NPC1 seems to be also an intervenient in transport or

internalization of some compounds (Vanier & Millat, 2003).

It is believed that both NPC1 and NPC2 proteins function in a closely related fashion,

since it was not found any qualitative difference in their ability to respond to exogenous

Introduction

22

LDL cholesterol loading and in their tissue lipid storage (Vanier et al., 1996). Ioannou

(2001) suggested that NPC1 activity may be dependent on prior action of NPC2 to insert

sterol into the endosomal-lysosomal membrane. However, more information about both

proteins, such as structure and specific localization, may be useful to further understand

these processes.

I-1.5. Conservation during evolution – NCR1, the yeast orthologue of hNPC1

NPC proteins are ubiquitously expressed and present homology with proteins in

several organisms, indicating that NPC can play an important role in basic cellular

processes. The yeast Saccharomyces cerevisiae has a single copy of NCR1, a NPC1

orthologue. Ncr1p contains multiple transmembrane domains, such as NPC1 domain, a

conserved SSD domain, whose mutations highlight the importance of this domain for

proper Ncr1p function (Malathi et al., 2004), and a cysteine-rich domain (Berger et al.,

2005). Moreover, NPC1 proteins have an extremely high functional conservation among

species. Indeed, Ncr1p is able to suppress cholesterol and ganglioside accumulation

when expressed in NPC1-deficient Chinese hamster ovary cells (Malathi et al., 2004).

Many of the NPC1 patient mutations are in amino acids that are conserved in yeast

proteins. Indeed, Ncr1p presents a rate of identity and similarity of 34% and 57%,

respectively, when compared with human NPC1 (Carstea et al., 1997; Zhang et al., 2004).

Furthermore, of 105 identified miscoding patient mutations, 66% of the affected amino

acids are conserved in yeast and of these, 50% are identical between Ncr1p and human

NPC1 (Berger et al., 2005).

The first phenotype related with NCR1 deletion in yeast was the resistance to the

ether lipid drug, edelfosine (1-O-octadecyl-2-O-methyl-rac-glycero-3-phosphocoline). This

resistance seems to be due to the inability to move the compound out of the vacuole,

which probably provide protection to the cells from the inherent toxicity of edelfosine and

allow cell growth in its presence. Despite all these features, previous work established that

NCR1 is not essential for cell viability (Berger et al., 2005).

Other experiments were made to unravel the function of Ncr1p and subsequent

relationship with NPC disease. Malathi and co-workers studied the effect of mutations in

the SSD of Ncr1p and showed that Ncr1p plays an elemental role in subcellular

sphingolipid distribution, by recycling sphingolipids, and that defects in this process result

in sterols accumulation (Malathi et al., 2004). Despite these data, new studies are

required to understand the role of Ncr1p at cellular levels.


23

I-2. Endoplasmic reticulum stress

The endoplasmic reticulum (ER) is a specialized organelle with an important role in

the biology of the cell (Guerin et al., 2008). This is one of the largest intracellular

organelles, represented by a continuous membranous network that extends throughout

the cytoplasm and is contiguous with the nucleus (Kaufman, 1999). In eukaryotic cells,

the ER is the first compartment in the secretory pathway and the site of synthesis, folding

and delivery of secreted, membrane-bound and organelle-targeted proteins that are

correctly assembled to their proper targets within the secretory pathway and the

extracellular space. Only proteins with a correct folding are exported to the Golgi complex,

whereas incompletely folded proteins are retained in the ER to complete the folding

procedure or marked for degradation, in a process called quality-control. The ER is the

major site for synthesis of sterols and lipids and even a major part of the cell wall of lower

eukaryotes is synthesized in the ER (Cid et al., 1995).

The ER provides the conditions required for protein folding, such as ATP, Ca2+ and an

oxidizing environment that allows disulfide-bond formation and protein folding (Guerin et

al., 2008; Shen et al., 2004). An appropriate ER function is essential for some cell

physiological aspects, such as vesicle and lipid trafficking and protein targeting and

secretion (Guerin et al., 2008). Environmental perturbations in these parameters, such as

disruption of Ca2+ homeostasis, inhibition of protein glycosylation or disulfide bond

formation, hypoxia and virus or bacteria infection, compromise the normal functioning of

ER, leading to an accumulation and aggregation of unfolded proteins in the ER lumen that

induces ER stress (Banhegyi et al., 2007; Shen et al., 2004). When the capability to

process the protein folding is compromised, the ER activates a signal transduction

pathway, known as unfolded protein response (UPR), in order to decrease the

accumulation of these proteins in the ER (Kaneko & Nomura, 2003).

The UPR is the biochemical basis for several ER storage diseases, such as

Huntington’s, Parkinson’s and Alzheimer’s disease, in which unfolded or misfolded

proteins form aggregates (Rutishauser & Spiess, 2002; Vembar & Brodsky, 2008). This

pathway was first described in the budding yeast S. cerevisiae (Back et al., 2005; Shen et

al., 2004), after the identification of an unfolded protein response element (UPRE) in the

yeast KAR2 promoter, by Sambrook and co-workers (Mori et al., 1993) and Walter and

co-workers (Cox et al., 1993). The UPRE is essential and sufficient to confer ER stress

inducibility on a heterologous reporter gene (Mori et al., 1992).

The ER has evolved mechanisms to sense the stress in the lumen and to induce an

adaptive response that aims to reestablish its normal physiological state, either by up-

regulating its folding capacity (by the induction of ER-resident molecular chaperones and

Introduction

24

foldases) and increasing its size, or down-regulating the biosynthetic load of the ER

through shut-off of protein synthesis, at the transcriptional (Martinez & Chrispeels, 2003;

Pakula et al., 2003) and translational level (Harding et al., 1999). In addition, the induction

of ER associated degradation (ERAD) increases the degradation of unfolded proteins

(Friedlander et al., 2000; Travers et al., 2000). When these mechanisms fail to restore

normal ER homeostasis, apoptosis is activated to eliminate damaged cells (Shen et al.,

2004).

I-2.1. Mechanism of protein folding

The ER holds specific characteristics, from its chemical composition to its machinery,

that are different from those of other organelles and significantly influence protein folding

processes. The ratio of glutathione forms (the major redox buffer in the cell) in the ER is

also different from that in the cytosol. Levels of reduced (GSH) to oxidized glutathione

(GSSG) in the ER is 1:1 to 3:1, against 30:1 to 100:1 in the cytosol (Hwang et al., 1992).

The ER presents a neutral pH and high concentration of Ca2+ that can reach 5 mM,

against 0.1 µM in the cytosol (Orrenius et al., 2003). Since the majority of ER-resident

molecular chaperones and foldases have high affinity to Ca2+, perturbations of the ER

Ca2+ pool can severely affect their folding and interactions with other chaperones (Corbett

et al., 1999; Lloyd-Evans & Platt, 2010).

There are numerous post-translational alterations in the ER, such as disulfide bond

formation and N-linked glycosylation. Glycosylation plays a key role in protein folding. This

process starts at the ER, during protein synthesis in ribosomes and it is suggest that there

is a higher thermodynamic stability of glycoproteins in the glycosylated form (Shental-

Bechor & Levy, 2008). Disulfide bond formation is one of the most relevant parameters

required for maturation of proteins in the ER. It is catalyzed by protein disulfide

isomerases (PDI), which in turn is reoxidised by the FAD-dependent oxidase Ero1p.

Ero1p is extremely important in yeast under anaerobic conditions and an uncoupling of

Ero1p from its electron acceptor during ER stress, may lead to the formation of reactive

oxygen species (ROS) (Tu & Weissman, 2002).

As a way to monitor if ER is assembling its products correctly, secreted proteins are

targeted to ER quality control. Primary mediators of ER quality-control are molecular

chaperones, which besides the sampling of correctly assembled proteins, help

polypeptides to fold and evaluate the conformation of their substrates (Vembar & Brodsky,

2008). One of the ER quality-control machinery is the calnexin/calreticulin cycle, that

analyzes protein conformations and defines if a molecule is exported to the Golgi complex

or if is targeted for ERAD (Ellgaard et al., 1999).


25

The protein folding machinery of the ER comprises three distinct groups of proteins:

foldases, molecular chaperones and the lectins calnexin, calreticulin and EDEM (ER

degradation-enhancing α-mannosidase-like protein). Foldases are enzymes whose role is

to catalyze stages in protein folding. Chaperones increase the efficiency of protein folding

by recognizing and stabilizing the partially folded intermediates during the folding process.

They can be classified into several groups according to their cytosolic counterparts. One

of these molecular chaperones is BiP, which takes part in the translocation of nascent

polypeptide chains into the ER (Gupta & Tuteja, 2011). In fact, the interaction between

unfolded proteins and ER-resident molecular chaperones represents a second quality-

control checkpoint of the ER machinery (Ellgaard & Helenius, 2003).

I-2.2. Recognition of unfolded proteins

Biochemically, unfolded proteins present a conformation that interacts with molecular

chaperones. However, different chaperones recognize and make possible the folding of

different proteins. BiP also plays a role in recognition of unfolded proteins. This chaperone

has high affinity to protein substrates. When unfolded proteins bind to BiP and become

locked in their conformation, the ATPase activity of the chaperone is induced. BiP exists in

equilibrium between monomeric and oligomeric forms. Only the monomeric form of BiP

can bind unfolded proteins, and this association increases the monomeric, unmodified BiP

pool (Freiden et al., 1992). Hence, it was suggested that BiP is recruited to the monomeric

pool from a modified oligomeric BiP storage pool, by interaction with unfolded proteins

(Gething, 1999). Furthermore, the UPR may respond to changes in the protein folding

demand reflected by the available pool of free BiP (Shen et al., 2004). These mechanisms

seem to be the first events in signal transduction, in response to the accumulation of

unfolded proteins in the ER.

I-2.3. Transduction of the unfolded protein signal across the ER membrane

In higher eukaryotes there are three transmembrane proteins that transduce the

unfolded protein signal across the ER membrane. Two of them belong to the ER luminal

domains of the type I, IRE1 (inositol-requiring protein 1) and PERK (protein kinase RNA-

like ER kinase), and the third is the type II transmembrane protein activating transcription

factor 6 (ATF6) (Pineau & Ferreira, 2010; Shen et al., 2002). ATF6 holds two independent

ER stress regulated Golgi localization sequences (GLS). Nonetheless, only PERK and

IRE1 display a degree of conservation throughout all eukaryotes, and no ATF6 orthologue

has been discovered in yeast until now (Liu et al., 2000; Torres-Quiroz et al., 2010).

Introduction

26

Experiments in yeast showed that the ER luminal domains of PERK and IRE1 are similar

and their functions are evolutionarily conserved (Liu et al., 2000).

In an active state, there is a relationship between the luminal domains, PERK and

IRE1, and BiP (Bertolotti et al., 2000). In fact, BiP can be considered as a master negative

regulator of the UPR, because in unstressed cells, BiP binds to the luminal domains,

keeping them inactive. However, when ER stress occurs and unfolded proteins

accumulate in the lumen, BiP disassociates from these ER stress sensors to take part in

protein folding attempt. Subsequently, oligomerization of luminal domains is initiated, as

well as activation of these proximal signal transducers (Bertolotti et al., 2000; Shen et al.,

2004).

The calnexin/calreticulin cycle and recognition of unfolded proteins by BiP play an

important role in the regulation of activity of the proximal stress transducer ATF6 (Hong et

al., 2004). Nevertheless, the conserved N-linked glycosylation site in yeast Ire1p is not

essential for its function (Liu et al., 2000), which suggests that differential regulation of

these three sections of UPR (IRE1, PERK and ATF6) exists to improve UPR signaling to

specific folding requirements in the ER (Yoshida et al., 2003).

I-2.4. Activation of protective responses by the UPR – IRE1

Yeast and plants lack ATF6 and PERK. In yeast, the UPR is rather a simple linear

pathway, with transcriptional regulation exclusively mediated by the IRE1 pathway,

through the induction of chaperones and ERAD (Shen et al., 2004). This pathway is

characterized by exclusive features in stress signal transduction and is observed in all

eukaryotes. The IRE1 gene was identified in a forward genetic screen for mutations

related with the activation of an UPRE::LACZ reporter by ER stress (Cox et al., 1993; Mori

et al., 1993). IRE1 encodes a type I transmembrane ER resident protein, with an N-

terminal luminal domain that senses the ER stress and a C-terminal cytoplasmic domain

required for KAR2 (the yeast orthologue of BiP) expression (Shen et al., 2004).

The substrate for the Ire1p endoribonuclease is the mRNA for the bZIP transcription

factor Hac1p. HAC1 contains a large intron of 252 bp, located in the 3’-end of the mRNA.

The presence of unfolded proteins in the ER lumen, induced by agents such as

tunicamycin, a natural inhibitor of N-linked glycosylation, or dithiothreitol (DTT), which

impairs disulfide bond formation, leads to the dimerization and trans-autophosphorylation

of Ire1p (Back et al., 2005; Fei et al., 2009; Shen et al., 2004). This activates its RNase

activity and induces the cleavage of both 5’- and 3’-exon-intron junctions in HAC1 mRNA,

leading to the formation of 5’-OH and 3’-cyclic PO4 ends (exons), that are joined by tRNA

ligase (Gonzalez et al., 1999; Sidrauski et al., 1996) (Figure I.3). This transcription factor


27

was first described in yeast as an immediate downstream substrate for the RNase activity

of Ire1p, and showed ability to bind with KAR2 UPRE (Mori et al., 1996). Notwithstanding,

there are no readily recognizable UPREs in the promoters of the genes associated with

ERAD (Back et al., 2005).

The mechanism of HAC1 mRNA splicing is similar to pre-tRNA splicing, but differs in

the localization, since HAC1 mRNA splicing is likely to be cytoplasmic. Despite the

resemblance of both splicing mechanisms, it is unknown how the ligase differentiates

exons and introns. In vitro assays showed that HAC1 exons remain associated after the

cleavage induced by Ire1p (Abelson et al., 1998; Gonzalez et al., 1999). The translation of

unspliced mRNA is suppressed by the base pairing between the 5’-UTR (untranslated

region) of unspliced HAC1 mRNA and the intron (Ruegsegger et al., 2001). However, the

increased transcriptional activation potential that is observed in the spliced forms opposed

to the unspliced ones is not yet fully explained.

The HAC1 mRNA splicing leads to the expression of an alternative C-terminus with

high transcriptional activation potential and to the removal of a translational attenuator

from HAC1 mRNA (Mori et al., 2000; Ruegsegger et al., 2001). Then, Hac1p binds to the

UPRE (CAGCGTG) (Mori et al., 1998). After suppression of protein synthesis in

ribosomes, ER chaperones are induced to correct protein conformation, by refolding

unfolded proteins. The remaining unfolded proteins are then eliminated from the ER to the

cytosol through retrograde transport, and degraded by the proteasome (ERAD) (Kaneko &

Nomura, 2003).

Figure I.3. The Ire1p cascade of the UPR pathway. Ire1p detects high levels of misfolded proteins in the ER

lumen and promotes HAC1 mRNA splicing reaction, removing the intron from the precursor mRNA which

encodes Hac1p. Hac1p binds to UPR related genes and upregulate the expression of chaperones and ERAD

proteins in the nucleus.

Introduction

28

I-2.5. Role of IRE1 and HAC1 in membrane proliferation control

The UPR may have a role in the regulation of membrane proliferation. In cells

lacking Ire1p and Hac1p (that are inositol auxotrophs), ER stress triggered by tunicamycin

(TUN) upregulate INO1 gene, which encodes for inositol-1-phosphate synthase, a key

enzyme in phospholipid biosynthesis (Cox et al., 1997; Travers et al., 2000). In addition,

the induction of membrane proliferation is in some cases dependent on a functional UPR

pathway (Cox et al., 1997; Takewaka et al., 1999). Therefore, UPR seems to hold a

specialized function in the increment of phospholipid biosynthesis and ER proliferation

when it comes to an acute and/or severe ER stress. On the other hand, studies with ire1Δ

and hac1Δ mutants revealed that the activation of INO1 by inositol starvation was only

moderately defective in these strains (Chang et al., 2002; Cox et al., 1997). These mutant

strains presented, after 4h inositol starvation, increased values of CDP-diacylglycerol,

compared to wild-type, and decreased levels of phosphatidic acid and

phosphatidylinositol. These alterations were reversed and INO1 induction was not

compromised by HAC1 deletion in a strain with an overexpression of inositol phenotype

(Opi-). The changes in phospholipid levels in ire1Δ and hac1Δ strains indicate that UPR

plays a role in the regulation of metabolic reactions in phospholipid metabolism at the ER

membrane (Chang et al., 2002).

The bZIP transcription factor downstream of IRE1 presents a high degree of

divergence, even in organisms evolutionarily close, such as yeasts and filamentous fungi

(Saloheimo et al., 2003). In metazoans, XBP1 (X-box binding protein-1) is a bZIP

transcription factor of the ATF/CREB family and is the functional homologue for Hac1p. It

plays a key role in the regulation of a subset of ER-resident molecular chaperones. XBP1

splicing also introduces a frame-shift and an alternative C-terminus with increased

transcriptional activation potential. The mechanism of XBP1 splicing is still unclear.

Nevertheless, despite the divergence, the splice junctions in both XBP1 and HAC1 mRNA

are conserved (Lee et al., 2002).

Being the only major pathway in yeast, the IRE1 pathway coordinates several

features of the secretory pathway, such as membrane biogenesis, chaperone induction,

upregulation of ERAD genes and ER quality-control (Friedlander et al., 2000; Travers et

al., 2000; Yoshida et al., 2003). Previous experiments have shown that moderate IRE1-

and HAC1-independent transcriptional induction from a core promoter occurs in response

to ER stress in yeast. Thus, a second signal transduction pathway that modulates and

stimulates activation of ER chaperone genes by a Ire1p-hac1p independent pathway, in

response to stress, may exist in this organism (Schroder et al., 2003).


29

I-2.6. Endoplasmic reticulum-associated degradation (ERAD)

ER stress conditions can also induce the proteasome-dependent ERAD system to

counteract the high levels of unfolded or misfolded proteins present in the lumen of the ER

and restore ER homeostasis (Guerin et al., 2008).

ERAD systems are ubiquitous among eukaryotes and have been well studied in S.

cerevisiae (Xie & Ng, 2010). These systems aim to remove unfolded proteins by

retrograde transport from the ER to the cytosol via the translocon and, consequently, sort

them for degradation by the ubiquitin-proteasome system (UPS) (Kaneko & Nomura,

2003). Unfolded proteins in the lumen of the ER are recognized through the detection of

specific domains, such as unpaired cysteines or exposed hydrophobic regions.

Retrotranslocation of these proteins from the ER to the cytoplasm may use the same core

protein complex Sec61p that provides the conducting channel in the translocon through

which proteins are imported into the ER lumen. Then, a cascade of enzymatic reactions

leads to a formation of a polyubiquitinated protein that will be recognized by the

proteasome subunits and subsequently degraded. Initially, the UPS was connected with

an ER quality control mechanism. Several studies indicated later that the UPS is able to

degrade proteins with anomalous conformations, leading to discover of some proteins that

are targeted to degradation by UPS, such as the yeast vacuolar protease

carboxypeptidase Y (CPY*), which is unfolded, retained and later degraded by ERAD

(Vembar & Brodsky, 2008; Xie & Ng, 2010).

In yeast, many components of ERAD pathway are induced by the UPR, such as

DER1, HRD3, HRD1/DER3 and UBC7 (Travers et al., 2000). Hrd1p is an ER type I

transmembrane protein that has E2 ubiquitin ligase activity (Bays et al., 2001);

nevertheless, it prefers an unfolded protein as an ubiquitination substrate and uses only

E2 ubiquitin-conjugating enzymes to mediate ubiquitination of ERAD substrates. Although

UBC1 mRNA levels are not affected by DTT treatment, UBC7 and HRD1 genes are

induced upon ER stress by a Hac1p- and Ire1p-dependent mechanism, implying that the

UPR may regulate some parts of ERAD system in yeast. The UPR is not essential for

basal expression of these proteins, suggesting that there is a basal level of ERAD,

sufficient for elimination of unfolded proteins under normal physiological conditions.

However, upon ER stress, the UPR is induced to increase ERAD activity to face the new

conditions (Friedlander et al., 2000).

The association between UPR and ERAD is not totally understood. It is known that

cells unable to perform ERAD are more sensitive to stress, as observed by a constitutive

activation of the UPR and a requirement for the UPR for normal growth and survival under

Introduction

30

mild stress conditions (Friedlander et al., 2000). Some unfolded protein substrates of the

ERAD pathway were characterized in cells lacking IRE1 (Casagrande et al., 2000). The

identification in mammalian cells of EDEM also supports the idea of an interaction

between the UPR and ERAD. The induction of EDEM on ER stress conditions is mediated

by IRE1/XBP1, but not by ATF6, suggesting that one of the functions of IRE1/XBP1 is to

upregulate ERAD (Yoshida et al., 2003).

I-2.7. Cell signaling pathways related with ER stress conditions – the High

Osmolarity Glycerol (HOG) pathway

When environmental conditions change, organisms evolve responses in order to

survive that include the induction of cell signaling pathways. These comprise Mitogen-

Activated Protein Kinase (MAPK) cascades that consist of a three component signaling

system, namely a MAPK Kinase Kinase, a MAPK Kinase and a MAPK that are

sequentially activated by phosphorylation (Robinson & Cobb, 1997).

Many of these MAPK cascades are evolutionarily conserved in eukaryotes (Chen &

Thorner, 2007). In S. cerevisiae, there are five MAPK pathways: pheromone response

pathway mediates cellular responses to pheromones; filamentous growth pathway leads

to a regulation to nutrient limiting conditions; spore wall assembly pathway acts during

meiosis and sporulation; cell wall integrity pathway is involved in conditions of cell wall

stress, such as hypo-osmotic shock; and the High Osmolarity Glycerol (HOG) pathway

plays a key role in survival under hyperosmotic conditions.

The HOG pathway is activated in response to an increase of osmolarity in

extracellular medium, leading to higher glycerol production. In order to maintain osmotic

balance, cells also increase glycerol uptake and, therefore, intracellular osmolyte

concentration. Several studies indicate the HOG pathway as an essential cascade for

regulating adaptation to severe conditions, such as heat stress and citric acid. This

pathway consists of two branches that encompass putative osmosensors coupled to a

MAPK cascade that, by phosphorylation, may lead to the activation of the Hog1p MAPK,

the orthologue to mammalian p38 stress-activated protein kinase (SAPK) (Schroeter et

al., 2002; Torres-Quiroz et al., 2010) (Figure I.4).


31

Figure I.4. The High Osmolarity Glycerol (HOG) pathway. During osmotic stress, the HOG pathway is

induced and the Hog1p MAPK phosphorylates and activates transcription factors that mediate stress

response.

It was recently shown that the HOG pathway also has a major role in ER stress

resistance. The mechanism of Hog1p phosphorylation during ER stress is divergent from

that associated with cellular response to other stress conditions and only Sln1p branch of

the HOG pathway is required, along with both Ire1p and Hac1p (Bicknell et al., 2010;

Torres-Quiroz et al., 2010). Strains lacking Hog1p present sensitivity to tunicamycin or β-

mercaptoethanol (Torres-Quiroz et al., 2010), a reducing agent that also induces ER

stress by preventing disulfide bond formation. These results indicate that Hog1p is vital to

deal with chemical agents that form unfolded protein aggregates in the ER. In contrast,

when a hyperactivation of this pathway occurs, cells reveal resistance to tunicamycin,

indicating that kinase activity of Hog1p is necessary to deal with N-glycosylation defects

promoted by tunicamycin exposure (Torres-Quiroz et al., 2010).

The HOG pathway is also involved in late phases of ER stress (Bicknell et al.,

2010). Hog1p translocates into the nucleus and controls the expression of genes that are

exclusively activated in late points of ER stress. Hog1p also induces autophagy

components (Prick et al., 2006), indicating that the HOG pathway takes part in several

aspects of cellular response to long term ER stress.

Introduction

32

I-3. Saccharomyces cerevisiae as biological model

The yeast Saccharomyces cerevisiae is a unicellular eukaryotic fungus encompassed

in the Ascomycete family. It is frequently used as a model organism for the study of the

eukaryotic cell and biological processes conserved during evolution (Jazwinski, 2005). It is

established that S. cerevisiae is appropriate to the study of fundamental cellular

mechanisms and correlations with those in higher eukaryotes, including humans. Due to

the high degree of similarity among eukaryotes – from the organization and function of

molecules, organelles, genes, to signaling pathways necessary for the regulation of cell

growth (Botstein et al., 1997), stress responses (Gasch & Werner-Washburne, 2002) and

intracellular transport (Kucharczyk & Rytka, 2001) – it is possible to study all of these

mechanisms in such a simple organism as yeast.

S. cerevisiae was the first organism to have its genome fully sequenced and a major

part is functionally characterized. Indeed, the development of genomic and proteomic

tools, combined with the several online databases that contain information about yeast

genes and proteins (Pena-Castillo & Hughes, 2007), provides a wide range of knowledge

about several aspects of the organism. It is also a microorganism easy to genetically

manipulate and techniques for its manipulation and harvesting are strongly optimized

(Amberg et al., 2005).

Since yeast presents orthologues of human genes, it has been used for the study of

several diseases, including cancer (Hartwell, 2002) and neurological disorders like

Huntington’s (Giorgini et al., 2005). S. cerevisiae has also been used as a model for the

study of NPC disease, due to the existence of hNPC1 and hNPC2 orthologues, NCR1 and

NPC2, respectively. In fact, the identification of a phenotype for the ncr1Δ mutants allows

the use of conventional yeast genetics to define cell functions for NPC proteins (Berger et

al., 2005). It is also possible to use yeast to study mechanisms related with ER stress

response, due to the conservation during evolution of the IRE1 pathway, which controls

UPR, and its downstream target, the Hac1p transcription factor (orthologue of human

XBP1) (Shen et al., 2004). Therefore, we selected S. cerevisiae as our model organism to

study the role of Ncr1p in ER stress response.


33

Chapter II

Aim of the work


35

Previous studies propose neurodegenerative diseases as one of the main causes of

accumulation of unfolded proteins and subsequent ER stress (Kaneko & Nomura, 2003).

Thereby, we decided to unravel the role of Ncr1p in endoplasmic reticulum stress

response in S. cerevisiae. This work aimed to:

i. characterize the sensitivity of ncr1Δ cells to stress conditions, by exposure to

compounds that induce ER stress;

ii. establish the correlation between ER stress sensitivity and oxidative stress

markers, by measuring ROS and glutathione levels;

iii. uncover putative alterations in cell signaling pathways of ncr1Δ cells, related

to ER stress response


37

Chapter III

Material and Methods


39

III-1. Yeast strains and growth conditions

The Saccharomyces cerevisiae strains used in this work are described in Table III.1.

Table III.1. Yeast strains used in this work.

Strain Genotype Reference/Source

BY4741 Mata, his3Δ1, leu2Δ0, met15Δ0,

ura3Δ0

EUROSCARF

ncr1Δ BY4741 ncr1Δ::KanMx4 Vilaça, R.

hac1Δ BY4741 hac1Δ::KanMx4 EUROSCARF

BY4741 pRS316 BY4741 carrying pRS316 Vilaça, R.

BY4741 pJT30 BY4741 carrying pJT30 This work

ncr1Δ pJT30 ncr1Δ carrying pJT30 This work

BY4741 pRS315 BY4741 carrying pRS315 This work

ncr1Δ pRS315 ncr1Δ carrying pRS315 This work

BY4741 pCPY BY4741 carrying pCPY This work

ncr1Δ pCPY ncr1Δ carrying pCPY This work

W303a Mata, ura3Δ1, leu2Δ3, his3Δ11,

trp1Δ1, ade2Δ1, can1Δ100

EUROSCARF

SEC63-GFP VPH-cherry W303a SEC63-GFP::HIS3 VPH-

cherry::TRP1

Schuck et al., 2009

SEC63-GFP VPH-cherry hac1Δ W303a SEC63-GFP::HIS3 VPH-

cherry::TRP1 hac1Δ::LEU2

Schuck et al., 2009

SEC63-GFP VPH-cherry ncr1Δ W303a SEC63-GFP::HIS3 VPH-

cherry::TRP1 ncr1Δ::KanMx4

This work

SEC63-GFP VPH-cherry ncr1Δhac1Δ W303a SEC63-GFP::HIS3 VPH-

cherry::TRP1 hac1Δ::LEU2

ncr1Δ::KanMx4

This work

The growth media used were YPD (1% (w/v) yeast extract, 2% (w/v) bactopeptone,

2% (w/v) glucose), synthetic complete (SC)-glucose medium (2% (w/v) glucose, 0.67%

(w/v) yeast nitrogen base without aminoacids, 0.14% (w/v) drop-out medium lacking

histidine, leucine, tryptophan and uracil, 0.008% (w/v) histidine, 0.04% (w/v) leucine and

0.008% (w/v) tryptophan or minimal medium (0.67% (w/v) yeast nitrogen base without

aminoacids, 2% (w/v) glucose, 0.004% (w/v) histidine, 0.004% (w/v) methionine and

0.004% (w/v) uracil).

Cells were grown aerobically at 26 ºC to early exponential phase (OD600nm = 0.6), in

an orbital shaker at 140 rpm, with a ratio of flask volume/medium volume of 5:1.


40

III-2. Genomic DNA extraction

Cells (10 mL) were grown to stationary phase and harvested by centrifugation during

5 min at 4000 rpm. The cell pellet was collected and washed once with deionized water

and resuspended in 100 µL of lysis buffer (2% Triton X-100, 1% SDS, 100 mM NaCl, 10

mM Tris-HCl pH 8.0, 1 mM EDTA) and 100 µL of phenol:chloroform:isoamyl alcohol

(50:48:2). Cells were lysed by vigorous shacking of the cell suspension in the presence of

glass beads for 3 min (short pulses of 1 min were used, with 1 min intervals). The

aqueous phase was recovered after centrifugation at 4000 rpm 5 min, and 100 µL of

chloroform were added. The mixture was homogenized by vortexing 3 min (as described

previously), supplemented with 100 µL TE 10x (100 mM Tris, 10 mM EDTA pH 8.0) and

centrifuged (14000 rpm, 5 min). The aqueous phase was transferred to a new eppendorf

tube, added to 1 mL of 100% ethanol and mixed to wash. After centrifugation (14000 rpm,

3 minutes), the pellet was resuspended in 400 µL of TE 1x. It was added 30 µL of 1

mg/mL RNase A and the mixture was incubated 5 min at 37 ºC. Then, 10 µL of 4 M

ammonium acetate and 1 mL of 100% ethanol were added. A DNA pellet was collected by

centrifugation (14000 rpm, 3 min), washed twice with 70% ethanol, dried and

resuspended in sterile water. The genomic DNA was quantified using a BioPhotometer

(Eppendorf) and analyzed by agarose gel electrophoresis.

III-3. Colony PCR and genomic DNA PCR

For colony PCR, a small portion of culture was peaked from agar plates, resuspended

in 25 µL NaOH and boiled at 100 ºC for 15 min. Samples were mixed by vortexing and

supernatant containing the DNA was collected after centrifugation (13000 rpm, 1 min).

For PCR, a mix of 20 µL containing 1 x Reaction Buffer (Thermo Scientific), 1.5 mM

MgCl2 (Thermo Scientific), 0.2 mM forward primer, 0.2 mM reverse primer, 0.2 µM dNTPs

(Fermentas), 1 U Taq Polymerase (Thermo Scientific), 1 µL of colony PCR supernatant or

genomic DNA was prepared. PCR were analyzed in agarose gels using TAE 1x as buffer,

and DNA bands were compared to Gene Ruler Ladder Mix (Fermentas).


41

III-4. Gene disruption

For NCR1 gene disruption in BY4741 and hac1Δ cells, a deletion fragment containing

HIS3 and the flanking regions of NCR1 was amplified by PCR using pRS313 and the

primers Ncr1_40bp_HIS3_S and Ncr1_40bp_HIS3_AS (presented in the Table III.2).

Purification of DNA from TAE agarose gel was performed with GFXTM PCR DNA and Gel

Band Purification Kit (GE Healthcare). Cells were transformed by electroporation (as

described below) and mutant cells were selected in minimal medium lacking histidine. To

confirm gene disruption, colony PCR was performed using primers Ext_NCR1_conf_S

and Int_HIS3_AS. Since all mutant strains tested were false-positive, we decided to

disrupt HAC1 in BY4741 ncr1Δ cells. For that, a fragment containing LEU2 and the

flanking regions of HAC1 was amplified by PCR using genomic DNA from W303a hac1Δ

strain and primers HAC1_amplif_S and HAC1_amplif_AS. The ncr1Δ cells were

transformed with this DNA fragment but all mutant cells obtained were also false-positive.

To disrupt NCR1 in W303a and W303a hac1Δ cells, a deletion fragment containing

KanMx4 and the flanking regions of NCR1 was amplified by PCR using genomic DNA

from the BY4741 ncr1Δ strain and the primers Ext_NCR1_S and Ext_NCR1_AS. Cells

were transformed by electroporation and mutants were selected in minimal medium

supplemented with geneticin 0.4 mg/mL. Gene disruption was confirmed by PCR using

primers Ext_NCR1_conf_S and Int_KAN_AS or Ext_NCR1_conf_S and Int_NCR1_AS.

Table III.2. Primers used in this work.

Primersa

Sequence

Ncr1_40bp_HIS3_S CTCCAAAAAGAACAAGAGCAGAACTTCAAT

TAGTAAAACCCGTTTTAAGAGCTTGGT

Ncr1_40bp_HIS3_AS TATTTTTTCACTACGTAAAATATAGTATAATCT

GCTATGGCTACATAAGAACACCTT

Ext_NCR1_conf_S AAGGTGCGAAATGACGGAAGA

Int_HIS3_AS AGAAAATGCGGGATCATCTCG

HAC1_amplif_S ATGAGGGTTGTAAGGCAAAGTGG

HAC1_amplif_AS TGTTCAGTGTCGCTGCCCAGT

Ext_NCR1_S CCGTGGCTAATGTCACAACA

Ext_NCR1_AS TTACGAGTGAAGCGTTCTGG

Int_NCR1_AS CGTCGTCCACAATCATTGCCC

Int_KAN_AS TGCTGTTTTGCCGGGGAT

aS - sense; AS – antisense.


42

III-5. Yeast electroporation

Yeast strains were transformed with plasmids by electroporation. This is a method

used to insert polar molecules into a host cell by electric pulses that produce transient

holes in the cell membrane. In this study, yeast strains were transformed with pRS316

and pRS315 (empty vectors), pJT30 (pRS316 harboring an UPRE-lacZ reporter) and

pCPY (pRS315 expressing an HA-tag version of CPY*). For gene disruption, deletion

fragments amplified by PCR were integrated in genomic DNA by homologous

recombination.

III-5.1. Preparation of electrocompetent cells

Cells were grown in 50 mL of YPD medium to an OD600nm = 1.3 – 1.5, harvested, and

resuspended in 10 mL of 10 mM Tris 1 mM EDTA 100 mM lithium acetate ph 7.5 and

gently shacked during 45 min at 30 ºC. Then, 250 µL of 1 M DTT was added and cells

were shacked 15 min at 30 ºC. Ice-cold sterile water was added for a final volume of 50

mL and cells were centrifuged at 4 ºC. Cells were washed first with 25 mL of ice-cold

sterile water and then with 2 mL of 1 M sorbitol (4 ºC), and resuspended in 50 µL of 1 M

sorbitol (4 ºC).

III-5.2. Electro-transformation and plating

Electrocompetent cells (40 µL) were mixed with 5 µL of plasmid DNA (± 0.1 µg) or

deletion fragment (± 0.5 µg) and incubated on ice for 5 minutes. The mixture was

transferred to prechilled sterile 2 mm electroporation cuvette. An electric pulse (1.5 kV, 25

µF and 200 Ω) was applied in parallel using an electroporation system (BioRad). After the

pulse delivery, 1 mL of selective minimal medium containing 1 M sorbitol was immediately

added and cells were gently shacked for 30 min (plasmid) or 4 h (deletion fragment) at 26

ºC. Cells were plated in selective minimal medium and grown at 26 ºC for 3 days.


43

III-6. Stress resistance

For the analysis of stress resistance, yeast cells were grown to early exponential

phase (OD600nm = 0.6) and exposed to DMSO (control) or 1 µg/µL tunicamycin (dissolved

in DMSO) for 1 h, 4 h and 18 h. Cell growth was followed spectrophotometrically. Cell

viability was determined by standard dilution plate counts on YPD medium containing

1.5% (w/v) agar. Cells were grown at 26 ºC for 3 days and then colonies were counted.

Viability was expressed as the percentage of the colony-forming units (CFU). For the

spotting assay, cells were diluted to OD600nm = 0.1 and fivefold serial dilutions were plated

on YPD solid media supplemented with DMSO (control) or 1 µg/µL tunicamycin (dissolved

in DMSO).

III-7. Oxidative stress markers

III-7.1. Quantification of reactive oxygen species (ROS)

This method is based on an in vivo intracellular oxidation of the oxidant-sensitive

probe 2’-7’-dichlorodihydrofluorescein diacetate (H2DCF-DA), as previously described by

Davidson et al. (1996).

Yeast cells (6 mL) were exposed to DMSO (control) or 1 µg/µL tunicamycin for 4 h or

18 h. A group of cells were exposed to H2O2 for 1 h as probe control. In the last hour of

treatment, these cultures were divided in two, and one part was incubated with 6 µL of 10

µM H2DCF-DA (Invitrogen; dissolved in DMSO) in the dark at 26 ºC.

Cells were cooled on ice, harvested by centrifugation and resuspended in 100 µL of

50 mM potassium phosphate buffer pH 6.4. A volume corresponding to 2 x 107 cells was

lysed by vortexing for 5 min in the presence of glass beads (short pulses of 1 min were

used, with 1 min intervals on ice). The supernatant was collected, diluted with 50 mM

potassium phosphate buffer pH 6.4 (42 µL of sample + 958 µL buffer) and the

fluorescence was measured using a spectrofluorimeter (Horiba Fluoromax-4) set at an

excitation wavelength of 504 nm and an emission wavelength of 524 nm.

Autofluorescence (measured using unlabeled control cells) was subtracted.


44

III-7.2. Quantification of glutathione levels

The preparation of yeast extracts was performed as described by Belinha et al.

(2007) and glutathione was assayed by the method of Tietze (1969). Yeast cells (50 mL)

were grown to early exponential phase (OD600nm = 0.6) and cultures were treated with

DMSO (control) or 1 µg/µL tunicamycin for 1 or 18 h. Yeast extracts were prepared by

combining equal volumes of 2 M perchloric acid and a cell suspension in 100 mM

potassium phosphate buffer 2 mM EDTA pH 7.4. The mixture was lysed by vortexing in

the presence of glass beads for 3 min – short pulses of 30 s were used, with 30 s intervals

on ice. Cell debris was removed by centrifugation at 13000 rpm for 5 min. The

supernatant was neutralized to pH 7.0 with 2 M KOH 0.3 M 4-morpholinepropanesulfonic

(MOPS) acid. Samples were frozen (-80 ºC) for 1 hour and centrifuged at 13000 rpm for 1

min. Protein content was determined by the method of Lowry, using bovine serum albumin

(BSA) as a standard. For oxidized glutathione determination, samples were treated with

2% (v/v) 2-vinylpiridine, and incubated for 1 hour at 4 ºC with agitation. A mix containing

750 µL of 100 mM KPi 1 mM EDTA pH 7.0, 0.133 mg/mL NADPH and 0.05 mg/mL DTNB

was added to 10 or 50 µL of sample (for GSH+GSSG or GSSG quantification,

respectively) and 80 µL of glutathione reductase 2 U/mL and the rate of color

development was monitored at 405 nm. The concentration was determined by reference

to a GSSG standard added to the assay cuvette (internal standard) and expressed as

nmol of glutathione (µg protein)-1.

III-8. β-Galactosidase activity

To analyze UPR activation after tunicamycin exposure, BY4741 and ncr1Δ cells (10

mL) containing pJT30 (harboring an UPRE-lacZ reporter) or pRS315 (empty vector) were

grown in SC-glucose medium lacking uracil to early exponential phase (OD600nm = 0.6) and

exposed to DMSO (control) or 1 µg/µL tunicamycin (dissolved in DMSO) for 1 h. Cells

were harvested by centrifugation, resuspended in 100 µL of Breaking Buffer (100 mM

Tris-HCl, 1 mM DTT, 10% glycerol) and protease inhibitors (1:20) (Complete, Mini, EDTA-

free Protease Cocktail Inhibitor Tablets; Boehringer Mannheim), lysed by vortexing for 5

min in the presence of glass beads (short pulses of 1 min were used, with 1 min intervals

on ice) and centrifuged at 13000 rpm for 15 min. The supernatant was collected and

protein content was determined by the method of Lowry, using BSA as a standard. For β-


45

galactosidase assay, samples (40 µg of protein) were mixed with LacZ buffer (60 mM

Na2HPO4, 40 mM NaH2PO4, 10 mM KCl, 1 mM MgSO4, 50 mM β-mercaptoethanol) to a

final volume of 800 µL. Then, samples were incubated at 32 ºC for 5 min and 200 µL of o-

nitrophenylgalactopyranosyde (ONPG) were added to initiate the reaction. When color

became yellow, the reaction was stopped by addition of 400 µL of Na2CO3. Absorbance

was measured at OD420nm.

III-9. ERAD assay

The analysis of protein degradation by ERAD was performed as described by Ellis et

al. (2004), with some alterations. BY4741 and Δncr1 cells, both harboring pRS315 (empty

vector) or pCPY, were grown in minimal medium lacking leucine to early exponential

phase (OD600nm = 0.6). Yeast cells were exposed to DMSO (control) or 1 µg/µL

tunicamycin (dissolved in DMSO) for 0, 1 or 6 h and subsequently treated with 100 µg µL-1

cycloheximide. Aliquots were removed at 0, 30 and 60 min, to tubes containing 10 mM

NaN3 (final concentration); and cells were collected by centrifugation at 4000 rpm and

washed once with cold buffer (10 mM NaN3, 1 mM EDTA). Cell pellets were resuspended

in 200 µL cold protein extraction buffer (10 mM Tris-HCl pH 8.0, 25 mM ammonium

acetate, 1 mM PMSF, 10% trichloroacetic) acid supplemented with protease inhibitors

(Complete, Mini, EDTA-free Protease Cocktail Inhibitor Tablets; Boehringer Mannheim)

and shacked in the presence of glass beads for 5 min, with short pulses of 1 min and

equal time of intervals on ice. Cell lysates were transferred to new tubes. Glass beads

were washed with protein extraction buffer (500 µL) by vortexing for 1 min and the new

lysates were added to previous one. Lysates were centrifuged at 14000 rpm for 10 min at

4 ºC and pellets were resuspended in 120 µL buffer I (100 mM Tris base, 3 % SDS, 1 mM

PMSF) and boiled for 5 min. Insoluble debris was removed by centrifugation at 14000 rpm

for 5 min. The supernatant (18 µL) was separated by electrophoresis using 10%

polyacrylamide gels (described in Table III.3). Electrophoresis was performed at 12 mA

during the stacking gel and 16 mA during the running gel, using a LMW Calibration Kit for

SDS Electrophoresis (GE Healthcare) as protein standards and the following buffer: 0.025

M Tris pH 8.3, 0.192 M glycine, 0.1% SDS. Proteins were blotted into a nitrocellulose

membrane (Hybond-C, GE Healthcare) at 0.8 mA/cm2 during 1 h, using a semi-dry system

and a transfer buffer (39 mM glycine, 48 mM Tris, 0.0375% SDS, 20% methanol). After

blotting, the nitrocellulose membranes were stained with Ponceau S (0.2% Ponceau S,

0.03% trichloroacetic acid, 0.03% sulfosalicylic acid) to visualize proteins.


46

Membranes were blocked for 1 h with 5% milk powder in TPBS (0.065% Tween 20,

0.4% NaCl, 0.01% KCl, 0.09% Na2HPO4.2H2O, 0.012% KH2PO4) and incubated overnight

with the primary antibody rabbit anti-HA (1:500; Sigma Aldrich) or rabbit anti-actin (1:200;

Sigma Aldrich). Membranes were washed twice for 15 min with TPBS and incubated for 1

h with the secondary antibody anti-rabbit IgG peroxidase (1:5000; Sigma Aldrich).

Membranes were washed twice with TPBS (15 min) and with PBS (0.4% NaCl, 0.01%

KCl, 0.09% Na2HPO4.2H2O, 0.012% KH2PO4) (15 min). Immunodetection was performed

by chemiluminescence, using ECLTM Western Blotting Detection Reagents (GE

Healthcare). Membranes were exposed to a Hybond-ECL film (GE Healthcare), and the

film was developed.

Table III.3. Reagents used in the preparation of a polyacrylamide gel.

Reagents Running gel Stacking gel

Acrylamide 30 % 1.8 mL 250 µL

Running Buffer (1.5 M Tris-HCl pH 8.8,

0.4 % SDS)

1.3 mL -

Stacking Buffer (0.5 M Tris-HCl pH 6.8,

0.4 % SDS)

- 625 µL

H2O 2.3 mL 1.6 mL

Ammonium Persulfate 10 % 41 µL 18.8 µL

TEMED 4.3 µL 2.5 µL

III-10. Induction of the HOG pathway

BY4741 and Δncr1 cells were grown in YPD medium to early exponential phase

(OD600nm = 0.6) and exposed to DMSO (control) or 1 µg/µL tunicamycin (dissolved in

DMSO) for 1 h. Alternatively, cells were grown in YPD medium supplemented with 1M

sorbitol (positive control). Cell pellets were collected by centrifugation at 4000 rpm for 5

min. Yeast extracts were prepared in 50 mM potassium phosphate buffer (pH 7.0)

containing protease inhibitors (Complete, Mini, EDTA-free Protease Cocktail Inhibitor

Tablets; Boehringer Mannheim) and phosphatase inhibitors (50 mM sodium fluoride, 5

mM sodium pyrophosphate, 1 mM sodium orthovanadate), by shaking of cell suspension

in the presence of glass beads for 5 minutes, with short pulses of 1 minute and equal time


47

of intervals on ice. Cell debris was removed by centrifugation (13000 rpm, 15 minutes, 4

ºC) and protein content was determined by the method of Lowry (using BSA as a

standard).

For SDS-PAGE, samples (20 µg for Hog1p and 50 µg for phospho-Hog1p detection)

were prepared in sample buffer (0.125 M Tris-HCl pH 6.8, 4% SDS, 20% glycerol, 10% β-

mercaptoethanol) and boiled for 3 minutes. Proteins were separated by electrophoresis as

described in III.9.

Immunodetection was performed as described in III.9, using rabbit anti-Hog1p MAPK

(1:1000; Santa Cruz Biotechnology, Inc, USA) or rabbit anti-phospho-p38 MAPK (1:500;

Cell Signaling Technology, Beverly, MA, USA) as primary antibodies to detect Hog1p and

phospho-Hog1p, respectively, and anti-rabbit IgG peroxidase (1:5000; Sigma Aldrich) as

secondary antibody. For blocking with milk powder and successive washes, it was used

TPBS (0.15% Tween 20, 0.4% NaCl, 0.01% KCl, 0.09% Na2HPO4.2H2O, 0.012%

KH2PO4) and PBS (0.4% NaCl, 0.01% KCl, 0.09% Na2HPO4.2H2O, 0.012% KH2PO4) for

Hog1p detection, and TTBS (0.1% Tween 20, 0.242% Tris, 0.8% NaCl) and TBS (0.242%

Tris, 0.8% NaCl) for phospho-Hog1p detection.

III-11. Statistical analysis

Data are expressed as mean values ± SD of at least three independent experiments.

Values were compared by Student’s t-test. The 0.05 probability level was chosen as the

point of statistical significance throughout.


49

Chapter IV

Results and Discussion


51

IV-1. Ncr1p deficiency decreases tunicamycin-induced growth arrest

Ncr1p deficient cells show an increased resistance to edelfosine, due to the

accumulation of this compound inside the vacuole that prevents its toxicity (Berger et al.,

2005). Nevertheless, not much is known about the effects of other drugs in cells lacking

Ncr1p. To study the role of Ncr1p in response to ER stress conditions, we analyzed the

resistance of ncr1Δ cells to tunicamycin. This compound inhibits protein glycosylation,

inducing the unfolded protein response (UPR), and it is often used to assess ER stress

(Back et al., 2005; Fei et al., 2009). Tunicamycin impairs bud formation and arrests the

cells in the G1/S phase of the cell cycle, leading to a disturbance in cell growth (Back et

al., 2005).

To test tunicamycin resistance, S. cerevisiae BY4741 (parental) and ncr1Δ mutant

cells were plated on solid medium containing tunicamycin or DMSO (vehicle). Parental

cells revealed sensitivity to tunicamycin whereas ncr1Δ cells showed a higher resistance

to this compound (Figure IV.1).

Figure IV.1. The ncr1Δ cells are resistant to tunicamycin. S. cerevisiae BY4741 and ncr1Δ cells were

grown on YPD media until the exponential phase. Cultures were diluted to O.D. 600 nm = 0.1 and a series of 1:5

dilution was plated in YPD media supplemented with DMSO or 1 µg/mL tunicamycin (TUN).

Tunicamycin resistance was also tested in liquid cultures. Parental and ncr1Δ cells

were treated with tunicamycin or DMSO and cell growth was followed overtime. The

results are shown in Figure IV.2. In both strains, tunicamycin decreased cell growth, but

this effect was more severe in parental cells than in ncr1Δ mutant cells. These results also

suggest that Ncr1p deficiency increases tunicamycin resistance. Tunicamycin exposure

for 1 h and 4 h did not significantly affect the growth of parental or ncr1Δ cells, suggesting

that its toxic effects are only observed after long-term exposure.


52

Figure IV.2. Effect of tunicamycin on cell growth. S. cerevisiae BY4741 and ncr1Δ cells were grown to

early exponential phase and treated with 1 µg/mL tunicamycin (TUN) or equal volume of DMSO for 1h, 4h and

18h. Values are fold changes relative to measure at T0, and correspond to means ± SD of at least three

experiments. *p<0.05.

The analysis of cell viability (Figure IV.3) suggests that parental and ncr1Δ cells show

similar resistance to tunicamycin: in both strains, 50% of the cells remained viable after 18

h of treatment. The overall results indicate that Ncr1p deficiency protects yeast cells from

tunicamycin-induced growth arrest but not from cell death.


53

Figure IV.3. Analysis of tunicamycin resistance. S. cerevisiae BY4741 and ncr1Δ cells were grown to

exponential phase and exposed to 1 µg/mL tunicamycin or equal volume of DMSO for 1 h, 4 h or 18 h.

Cellular viability was determined by standard dilution plate counts and expressed as the percentage of the

colony forming units of nonstressed cells. Data are means ± standard deviations of at least three independent

experiments.

IV-2. Ncr1p deficiency decreases tunicamycin-induced intracellular oxidation

and glutathione depletion

To assess if tunicamycin toxicity is associated with oxidative stress markers, we

analyzed the levels of reactive oxygen species (ROS) and glutathione. ROS are produced

as byproducts of cell metabolism and their effects are prevented by antioxidant defenses,

including superoxide dismutases, catalase and glutathione. When the balance between

antioxidant capacity and ROS levels is decreased, an oxidative stress occurs in the cell.

ROS production increases under stress conditions, like radiation and exposure to

xenobiotics (Pagano, 2002; Riley, 1994). ROS are considered stress biomarkers in yeast,

for instance, during induction of cell death by apoptosis (Madeo et al., 2004).

To quantify ROS levels, parental and ncr1Δ cells were exposed to DMSO or

tunicamycin for 4 h or 18h and labeled with H2DCF-DA, a molecular probe that is sensitive

to ROS. Parental cells treated with H2O2 during 1 h were used as positive control. After 4

hours of treatment with tunicamycin, both strains present similar low levels of ROS (Figure

IV.4). However, after 18 h exposure, ROS levels were 50% higher in parental cells treated

with tunicamycin, compared with DMSO-treated cells. It should be noticed that

intracellular oxidation increased during the growth of control cells (from 4 h to 18 h). This

is due to an increase in mitochondria function associated with the transition from a


54

fermentative (exponential phase) to a respiratory metabolism (post-diauxic phase) (Costa

& Moradas-Ferreira, 2001). Consistent with the protective effect of Ncr1p deficiency

against tunicamycin, ROS levels did not increase in ncr1Δ cells. Notably, this mutant

showed an increase of basal ROS levels, compared with that of parental cells. This is

probably due to a decrease in antioxidant defenses and mitochondrial dysfunctions

displayed by ncr1Δ cells at post-diauxic phase (Vilaça et al, unpublished).

Figure IV.4. Analysis of ROS levels. S. cerevisiae BY4741 and ncr1Δ cells were grown in YPD to

exponential phase and exposed to 1 µg/mL tunicamycin (TUN) or equal volume of DMSO (control) for 4 h and

18 h. Parental cells were exposed to 1.5 mM H2O2 for 1 h (positive control). ROS levels were quantified by

labeling cells with the molecular probe H2DCF-DA. Values are means ± SD of at least three experiments.

*p<0.05.

We hypothesized that the resistance of ncr1Δ cells could be correlated with higher

levels of glutathione, a tripeptide that plays a key role in detoxification processes. The

conjugation of glutathione (GSH) with the xenobiotics, catalyzed by glutathione S-

transferases (GST), leads to the production of glutathione S-conjugates and, therefore,

prevents the toxicity of those compounds (Pocsi et al., 2004). Glutathione also functions in

antioxidant protection, being required for glutathione peroxidases for the reduction of

hydrogen peroxide or lipid hydroperoxides. This reaction generates a glutathione disulfide

(GSSG) that is reduced to GSH by glutathione reductase, at the expense of NADPH


55

(Costa & Moradas-Ferreira, 2001; Pocsi et al., 2004). Therefore, we measured GSH and

GSSG levels in parental and ncr1Δ cells exposed to DMSO or tunicamycin for 1 h or 18h.

The results (Figure IV.5) have shown that tunicamycin treatment for 1h did not affect the

levels of either oxidized or reduced glutathione in both strains. However, a long-term

exposure to the drug led to GSH depletion in parental cells, an effect that was not

observed in ncr1Δ cells. This is consistent with the decrease in tunicamycin-induced

growth inhibition observed in this mutant strain. Notably, ncr1Δ cells showed lower basal

levels of glutathione. Although there is no evidence in the literature that glutathione is

involved in tunicamycin detoxification, our data suggest that glutathione does not mediate

the higher resistance of ncr1Δ cells to this compound.

Figure IV.5. Effect of tunicamycin on glutathione levels. S. cerevisiae BY4741 and ncr1Δ cells were grown

in YPD to exponential phase and exposed to 1 µg/mL tunicamycin (TUN) or equal volume of DMSO (control)

for 1 h or 18 h. The concentration of reduced (GSH) and oxidized (GSSG) glutathione was determined by

reference to a GSSG standard added to the assay cuvette and measured at 405 nm. Values are means ± SD

of at least three experiments. *p<0.05.


56

IV-3. UPR activation in ncr1Δ cells

The accumulation of unfolded protein in the ER activates the unfolded protein

response (UPR) that aims to maintain ER homeostasis, leading to the activation of Ire1p

pathway. Unfolded proteins lead to the dimerization of Ire1p and subsequent

autophosphorylation, which activates Hac1p, the substrate for Ire1p endonuclease. Hac1p

binds to unfolded protein response elements (UPREs) and induces chaperones to correct

protein conformation (Kaneko & Nomura, 2003; Pineau & Ferreira, 2010).

The protective effect of NCR1 deletion against tunicamycin led us to evaluate the

activation of Hac1p. Parental and ncr1Δ cells expressing an UPRE-lacZ gene reporter

(pJT30) (Figure IV.6) were treated with tunicamycin or DMSO and β-galactosidase (β-

GAL) activity was measured. The results obtained are presented in Figure IV.7.

Figure IV.6. Structure of UPRE-lacZ gene reporter. The plasmid pJT30 harbors a LacZ reporter under the

control of UPRE. Upon activation, the Hac1p transcription factor binds to the UPRE and β-galactosidase

activity increases.

Consistent with published data (Schroder et al., 2003), β-GAL activity increased 12-

fold in parental cells treated with tunicamycin, indicating that Hac1p was activated.

However, tunicamycin did not induce the UPR in ncr1Δ cells.

Aim to assess if this lack of response is a trait of ncr1Δ cells submitted to ER stress or

if it is specific for tunicamycin, a similar study was performed using DTT, another

compound that induces ER stress and subsequently the UPR by inhibiting disulfide bond

formation. In this assay, control cells were treated with H2O. The results of this control

were similar to those obtained in cells exposed to DMSO (data not shown).

As expected, β-galactosidase activity increased in parental cells exposed to DTT. In

ncr1Δ cells, β-galactosidase activity also increased, although to levels slightly lower to the

observed in parental cells. This result indicates that the UPR machinery is not

compromised in the ncr1Δ cells. Thus, the specific suppression of tunicamycin-induced

Hac1p activation is correlated with the attenuation of the growth-inhibitory effect observed

in this mutant strain.


57

Figure IV.7. Hac1p activation by ER stress. S. cerevisiae BY4741 and ncr1Δ cells harboring pJT30 (UPRE-

LacZ reporter) were grown in SC-glucose medium to exponential phase (O.D.600 nm = 0.6). β-galactosidase

activity was determined in cells untreated and treated with 1 µg/mL tunicamycin (TUN) or 10 mM DTT for 1 h.

Values are means ± SD of at least three experiments. *p<0.05.

To assess if the tunicamycin resistance of ncr1Δ cells was Hac1p-dependent, we

studied the effect of NCR1 deletion in a hac1Δ strain (W303a background), which is

known to be sensitive to tunicamycin (Schuck et al., 2009). Parental, hac1Δ, ncr1Δ and

ncr1Δhac1Δ cells were plated on YPD media supplemented with DMSO or tunicamycin.

Since the W303a strain displayed a higher sensitivity to tunicamycin, compared with the

BY4741 strain, a lower dose of this compound was used in this assay (0.05 µg/mL). The

results obtained are presented in Figure IV.8.


58

Figure IV.8. HAC1 deletion increases the sensitivity of ncr1Δ cells to tunicamycin. S. cerevisiae W303a

SEC63-GFP VPH-cherry (wt) cells and its isogenic strains hac1Δ, ncr1Δ and ncr1Δhac1Δ were grown on YPD

media until the exponential phase. Cultures were diluted to OD 600 nm = 0.1 and a series of 1:5 dilution was

plated in YPD media supplemented with DMSO or tunicamycin. Cells were grown at 26 ºC for 3 days.

As expected, hac1Δ cells showed a high sensitivity to tunicamycin (Schuck et al.,

2009). Under these conditions, the growth of ncr1Δ cells was similar to the observed in

parental cells. Notably, ncr1Δhac1Δ double mutants displayed a higher sensitivity to

tunicamycin, when compared with the ncr1Δ strain, but its resistance was higher to that of

hac1Δ cells. These results suggest that NCR1 deletion partially protects yeast cells from

tunicamycin by a Hac1p-independent mechanism.

III-4. ERAD is not inhibited by tunicamycin in ncr1Δ cells

Under ER stress conditions, cells can induce other mechanisms besides UPR to

restore ER homeostasis, such as ER associated protein degradation (ERAD) (Guerin et

al., 2008). This pathway acts by removing unfolded or misfolded proteins from the ER to

the cytosol and, subsequently, sorting them for degradation by the ubiquitin-proteasome

system (Kaneko & Nomura, 2003).

To assess if Ncr1p deficiency affects ERAD activation, parental and ncr1Δ cells

carrying pCPY (a plasmid harboring CPY*, that is targeted for degradation by ERAD) were

exposed to DMSO or 1 µg/µL tunicamycin for 0, 1 or 6 h, and subsequently treated with

cycloheximide to monitor CPY* stability. The results are presented in Figures IV.9 and

IV.10.


59

Figure IV.9. Analysis of CPY* stability. S. cerevisiae BY4741 and ncr1Δ cells harboring pCPY were grown

in minimal medium lacking leucine to exponential phase (OD 600 nm = 0.6) and treated with DMSO or 1 µg/µL

tunicamycin for 0, 1 and 6 h. Subsequently, cells were treated with 100 µg/µL cycloheximide (CH) for 0, 30

and 60 min. Cells were lysed and protein extracts were separated by SDS-PAGE and analyzed by

immunoblotting, using anti-HA and anti-actin (loading control) antibodies. A representative experiment is

shown.

Figure IV.10. Quantification of CPY* decay. Band intensities (Figure IV.9) were quantified by densitometry

and values were normalized for t0 of cycloheximide treatment.


60

In parental cells, tunicamycin exposure for 1 h inhibited ERAD. This is consistent with

published data and has been attributed to the accumulation of high levels of unfolded

proteins that severely decrease the rate of protein degradation due to a saturation of

ERAD capacity (Travers et al., 2000). After 6 h of tunicamycin exposure, CPY* decay

increased, suggesting that ERAD was activated. It is likely that the induction of UPR

increases the capacity of the ER to cope with the unfolded proteins, preventing ERAD

saturation.

In the absence of ER stress, the rate of CPY* degradation was similar in parental and

ncr1Δ cells. However, tunicamycin treatment for 1 h did not compromise ERAD in ncr1Δ

cells. Since this mutant strain is more resistant to tunicamycin, these results suggest that

probably there is no saturation of the ERAD system in cells lacking Ncr1p.

IV-5. Activation of Hog1p is compromised in ncr1Δ cells

The HOG signaling pathway plays a key role in osmotic and oxidative stress

responses, through the activation of proteins and transcription factors that control the

production of glycerol, with the purpose of maintaining the osmotic balance, and

antioxidant defenses (Bilsland et al., 2004; Rep et al., 2001; Schuller et al., 1994). This

pathway is also involved in response to ER stressors. Strains lacking the Hog1p MAPK

display sensitivity to tunicamycin and β-mercaptoethanol, whereas the activation of the

pathway enhances ER stress resistance (Torres-Quiroz et al., 2010). To evaluate

changes in this pathway associated with Ncr1p deficiency, parental and ncr1Δ cells were

treated with tunicamycin or DMSO and the levels of Hog1p and phospho-Hog1p were

analyzed by immunoblotting (Figure IV.11).

Figure IV.11. Hog1p phosphorylation is decreased in ncr1Δ cells. S. cerevisiae BY4741 and ncr1Δ cells

were grown in YPD medium and treated with DMSO or 1 µg/µL tunicamycin (TUN) for 1 h. Phospho-Hog1p

and Hog1p levels were analyzed by immunoblotting, as described in methods. The hog1Δ cells were used as

negative control. One representative experiment out of two is shown. *unspecific band.


61

In parental cells, Hog1p and phospho-Hog1p levels increased with tunicamycin

exposure. Hog1p activity is required for glycerol production in response to ER stress and

deletion of genes involved in glycerol synthesis increases the sensitivity of yeast cells to

tunicamycin (Torres-Quiroz et al., 2010). In ncr1Δ cells, Hog1p did not increase upon

tunicamycin treatment. In addition, the levels of phospho-Hog1p were very low in this

mutant strain, even after induction of ER stress. These results suggest that the resistance

of ncr1Δ cells to tunicamycin is Hog1p-independent. Nevertheless, a recent report

suggested that the basal activity of Hog1p, rather than its phosphorylated form, mediates

cellular protection against ER stress (Torres-Quiroz et al., 2010). However, basal levels of

Hog1p of both strains are similar and despite the increase in Hog1p levels in parental cells

after tunicamycin exposure, the same is not observed in ncr1Δ cells, which support the

hypothesis that resistance of ncr1Δ cells to tunicamycin is not dependent of Hog1p.


63

Chapter V

General Discussion and Future Perspectives


65

Sphingolipid storage diseases have been a main topic of research in the last years,

mainly due to its high prevalence worldwide. The molecular pathology of these disorders

has been subject of study in an attempt to understand the processes underlying these

diseases.

Niemann-Pick type C (NPC) is an autossomal recessive lipid storage disease

characterized by abnormal cholesterol trafficking and accumulation of unesterified

cholesterol in endosomal/lysosomal system (Ikonen & Holtta-Vuori, 2004; Pacheco &

Lieberman, 2008). NPC is associated with a progressive neurodegenerative phenotype

and is fatal in most of the cases (Patterson et al., 2001). The two complementation groups

and correspondent genes have already been described. Point mutations in either NPC1 or

NPC2, which mediates intracellular lipid transport through pathways that are not totally

understood, are the causes of this lipidoses (Pacheco & Lieberman, 2008).

Numerous lipid disorders display evidences of endoplasmic reticulum (ER) stress

(Klein et al., 2011). The ER plays an essential role in the biology of the cell, by regulating

protein export into the Golgi or retaining misfolded proteins to complete its folding

procedure or to target them for degradation. It is also the major site for synthesis of sterols

and lipids (Cid et al., 1995). The ER provides ideal conditions for several physiological

aspects, such as lipid trafficking and protein folding (Guerin et al., 2008). Environmental

alterations, such as inhibition of protein glycosylation or disulfide bond formation,

compromise ER normal functions, leading to an accumulation of unfolded proteins in the

ER lumen that induces ER stress (Banhegyi et al., 2007; Shen et al., 2004).

In this study, we investigated the role of Ncr1p, a vacuolar protein, in ER stress

response, using Saccharomyces cerevisiae cells as a model system. Ncr1p is an

orthologue of the human NPC1 protein (Berger et al., 2005). Tunicamycin, a compound

that inhibits protein glycosylation (Back et al., 2005), was used to induce ER stress. Our

results suggest that Ncr1p deficiency protects yeast cells from tunicamycin-induced

growth arrest, but not from cell death. It is known that ER is a potential source of ROS,

generating approximately 25% of its content. Furthermore, under stress conditions,

formation of ROS by the ER is known to increase (Tan et al., 2009). Reactive oxygen

species (ROS) and glutathione levels were measured to assess the redox state of the cell

(Tan et al., 2009; Zampieri et al., 2009). In parental cells, tunicamycin toxicity was

correlated with an increase of ROS levels and glutathione depletion. These oxidative

stress markers were suppressed in ncr1Δ mutant cells, which is consistent with the

protective effect of Ncr1p deficiency against tunicamycin. The basal levels of ROS were

increased and glutathione levels decreased in ncr1Δ cells, compared with those in

parental cells, which is probably due to a decrease in antioxidant defenses and

mitochondrial dysfunctions displayed by this mutant at post-diauxic phase (Vilaça et al,

General Discussion and Future Perspectives

66

unpublished). Although a previous study showed that tunicamycin-induced ER stress

affects glutathione levels (Tan et al., 2009), it is still unclear whether glutathione may take

part in its detoxification. However, the lower levels of glutathione observed in ncr1Δ cells

suggests that glutathione probably does not mediate the tunicamycin resistant phenotype

of ncr1Δ cells.

The unfolded protein response (UPR) is a mechanism that is induced during ER

stress in order to decrease the accumulation of unfolded proteins in the lumen of the ER

(Kaneko & Nomura, 2003; Schroder et al., 2003). The induction of this pathway leads to

the activation of Hac1p, a transcription factor that upregulates UPR target genes in an

attempt to restore ER homeostasis (Back et al., 2005). Therefore, hac1Δ cells are

sensitive to tunicamycin (Schuck et al., 2009). In parental cells, exposure to tunicamycin

led to the UPR induction, as assessed by measuring Hac1p activation, and a transient

inhibition of ERAD. This inhibition has been associated with the accumulation of unfolded

proteins to high levels that saturate ERAD capacity and decrease the rate of protein

degradation (Travers et al., 2000). In ncr1Δ cells, the activation of Hac1p and ERAD

inhibition were suppressed upon exposure to tunicamycin. These results indicate that the

resistance of ncr1Δ cells is not correlated with an increase in UPR and suggest that the

lack of UPR activation and ERAD inhibition may result from a higher capacity of this

mutant to detoxify tunicamycin. However, most of the tunicamycin resistance of ncr1Δ

cells is Hac1p-dependent, since it was severely affected by HAC1 deletion. It will be

important to characterize the mechanism by which Ncr1p deficiency exerts a protective

effect from tunicamycin in a Hac1p-independent manner. It should be noticed that the

UPR machinery of ncr1Δ cells is functional since Hac1p was activated by ER stress

induced by DTT.

The High Osmolarity Glycerol (HOG) pathway is also important to cell homeostasis

and it was recently implicated in the regulation of cellular responses to ER stress (Torres-

Quiroz et al., 2010). In ncr1Δ cells, Hog1p levels did not increase, and even phospho-

Hog1p levels were very low, indicating that resistance to tunicamycin is Hog1p-

independent. In parental cells, both Hog1p and phospho-Hog1p levels increased after

tunicamycin exposure. This is consistent with published data and aims to increase

glycerol production in response to ER stress (Torres-Quiroz et al., 2010).

Despite recent breakthroughs, unraveling the role of Ncr1p is a process that it is still

in a very inceptive stage. The first phenotype associated to ncr1Δ mutant cells is the

resistance to edelfosine, an anti-tumor ether lipid drug. Mutant cells were unable to export

the compound out of the vacuole and this confers protection against its toxicity and in turn

allows growth in its presence (Berger et al., 2005). Nevertheless, the molecular effects of

other compounds are still unclear. Therefore, some questions can be posed to understand


67

the intracellular effects of tunicamycin: if tunicamycin can enter the cell or if its

detoxification is somehow facilitated in ncr1Δ cells are points of interest in tunicamycin

study. A deeper study of the vacuole, since compounds can be retained in this organelle

and subsequently their effects can be suppressed, may contribute to the understanding of

the dynamics of tunicamycin in the cell. Since tunicamycin exposure differentially

influences cell growth in parental and ncr1Δ strains, the evaluation of signaling networks

related with cell growth should be considered, such as the Target of Rapamycin (TOR)

pathway, which is known to regulate cell growth in response to several stimulus, including

stress (Soulard et al., 2009). It is also known that basal activity of Hog1p, and not only its

phosphorylated form, plays a protective role against ER stress (Torres-Quiroz et al.,

2010). The importance of the HOG pathway should be clarified, to assess if other

components of this pathway could be involved in ncr1Δ phenotype.

In summary, these results suggest that Ncr1p deficiency increases ER stress

resistance induced by tunicamycin, a phenotype associated with a decrease in stress-

induced oxidative stress markers. Moreover, ERAD inhibition and ER stress responses

such as UPR and Hog1p activation are decreased in ncr1Δ cells, although a Hac1p-

independent mechanism plays a minor role in the resistance of this mutant strain to

tunicamycin. The characterization of the Hac1p-independent mechanisms that contribute

to the resistance of ncr1Δ cells to tunicamycin will contribute to our understanding of the

role of Ncr1p in the regulation of ER stress responses.


69

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