Chapter 1 Introduction - Shodhgangashodhganga.inflibnet.ac.in/bitstream/10603/11668/10/11_chapter1.pdf · Chapter 1 Introduction 1 Organization of the chapter The work reported in

Chapter 1

Introduction

Chapter1 Introduction

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Organization of the chapter The work reported in this thesis involves delineation of the roles for

glycosylphosphatidylinositol (GPI)-anchored aspartyl proteases, CgYps1-11, in the

physiology of a human opportunistic fungal pathogen Candida glabrata. These

proteases, also referred as yapsins, are pivotal virulence determinants of C. glabrata.

Novel findings reported in this thesis demonstrate that one of the eleven aspartyl

proteases, CgYps1, is vital for regulation of intracellular pH homeostasis under acidic

environmental conditions. The work also uncovers pivotal roles for yapsins in

maintenance of cell wall architecture, energy, ion and vacuole homeostasis and

processing and secretion of vacuolar lumenal enzyme, carboxypeptidase Y. This chapter

provides a comprehensive review of literature on topics related to the work and is

organized in following sections:

(1.1) Candida and candidiasis

(1.2) Candida glabrata and its pathogenesis

(1.3) Aspartyl proteases: structure, regulation and function

(1.4) Yeast cell architecture and physiology

Last part of this chapter describes the objectives addressed in this study.

Section 1.1: Candida and candidiasis

1.1.1 The Fungal Kingdom

The Kingdom Fungi is comprised of a large, highly diverse group of non-

vascular eukaryotic organisms which are found everywhere i.e., in air, soil and water and

in and on plants, animals and humans (Walker and White, 2011). Fungi play pivotal

roles in ecology and human economy. They recycle nutrients through ecosystems by

decomposing non-living organic material and supply essential nutrients to plants by

growing symbiotically with their roots (Walker and White, 2011). Common fungi

encompass yeasts, molds, lichens, mildews and mushrooms. The role of fungi in shaping

human history, due to their wide-spread use in baking, fermentation and pharmaceutical

industries, has universally been acknowledged (Kendrick, 2011). Economically

important fungi include both yeasts (Saccharomyces species, Pichia pastoris and

Hansenula polymorpha) and molds (Penicillium spp. and Aspergillus spp.) (Kendrick,

2011).


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Fungi are both microscopic (yeasts and molds) and macroscopic (mushrooms and

puff balls) in nature (Walker and White, 2011). They absorb nutrients from the

environment via mutualism, parasitism or saprophytism (Kendrick, 2011). Fungi can

reproduce both sexually and asexually. The sexual and asexual fungal stages are referred

as telomorph and anamorph, respectively (Whiteway and Bachewich, 2011). The fungal

vegetative stage is either single-celled (yeast) or multi-cellular composed of long,

microscopic filaments (hyphae) (Walker and White, 2011). While unicellular yeast cells

divide by budding, hyphae branch and grow via tip elongation (Walker and White,

2011). Most of fungi exist in one of these two growth forms, however, some fungi,

referred as dimorphic, can switch between yeast and filamentous forms (van Burik and

Magee, 2001; Walker and White, 2011).

Of 1.5 million fungal species identified, less than 500 are pathogenic to animals

and humans (Warnock, 2007; Heitman, 2011). Medically important fungi include

Aspergillus spp., Cryptococcus neoformans, Histoplasma capsulatum and members of

the genus Candida (Warnock, 2007). Notably, species of Candida, Cryptococcus,

Fusarium and Ustilago can cause diseases in both humans and plants (Sullivan et al.,

2005; Ma and May, 2009; Doohan, 2011; McNeil and Palazzi, 2012).

Recent classification, based on molecular phylogenetic analyses, has divided the

kingdom of fungi into seven phyla, viz., Ascomycota, Basidiomycota, Chytridiomycota,

Neocallimastigomycota, Blastocladiomycota, Glomeromycota and Microsporidia. The

first two have been grouped in a subkingdom dikarya (Hibbett et al., 2007). Importantly,

human pathogenic fungi mainly belong to four classes, ascomycetes, basidiomycetes,

zygomycetes and deuteromycetes (Hibbett et al., 2007; Warnock, 2007).

Infections caused by fungi, also commonly referred as mycoses, pose a serious

threat to human health. Based on their location, mycoses can be divided into three major

classes; superficial (hair, nails and skin), intermediate (mucosal surfaces of oral cavity

(generally referred as thrush), genital, respiratory and gastrointestinal tracts) and

systemic (blood and internal organs) (van Burik and Magee, 2001; Sullivan et al., 2005).

Mortality rates for invasive mycoses due to C. albicans, C. neoformans and A. fumigatus

are 20-40%, 20-70% and 50-90%, respectively (Lai et al., 2008; Park et al., 2009).

1.1.2 Candida species: general features, clinical manifestations and prevalence

With introduction and wide-spread use of antibiotics since 1940, incidence of

both mucosal and invasive candidiasis has risen enormously in past two decades (Pfaller


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and Diekema, 2007). Candida spp., now recognised as highly prevalent and potentially

belligerent pathogens, rank as the fourth most common cause of nosocomial (hospital-

acquired) bloodstream infections (BSIs) worldwide (Wisplinghoff et al., 2004).

Currently, Candida spp. are the predominant cause of mycoses followed by Aspergillus

and Cryptococcus spp. (Krcmery and Barnes, 2002; Messer et al., 2006; Pfaller et al.,

2006).

The genus Candida belongs to the phylum Ascomycota and contains over 200

spp. (de Hoog et al., 2000). The fungi Candida exists predominantly in a unicellular

form with oval-shape cells of 3-6 µm diameter (Larone, 2002). Most of Candida species

exist in the environment as saprotrophs (Larone, 2002). At least 17 Candida spp. are

known to cause bloodstream infections in humans which include C. albicans, C.

glabrata, C. tropicalis, C. dublienensis, C. parapsilosis and C. krusei (Pfaller and

Diekema, 2004). Of these, C. albicans, C. glabrata, and C. krusei normally exist as

commensals in humans and reside on skin, gastrointestinal and genitourinary tracts, but

become pathogenic under conditions of immuno-compromise (Sullivan et al., 2005).

Majority of clinically important Candida spp. are diploid and switch between

unicellular yeast and filamentous pseudohypal or hyphal form (Calderone and Fonzi,

2001). This morphological switching occurs in response to various environmental cues

(Calderone and Fonzi, 2001; Bahn et al., 2007). While yeast cells are round to ovoid-

shape and single-celled, pseudohyphae and hyphae represent branched chains of

elongated yeast cells that are attached to each other at septa, and highly polarized, non-

constricted, long filaments, respectively (Calderone and Fonzi, 2001). The ability to

exist in more than one morphological form is usually referred as polymorphism

(Calderone and Fonzi, 2001). An exception to fungal polymorphism is the haploid,

single-celled budding yeast C. glabrata, which does not form true hyphae under any

growth conditions (reviewed in Silva et al., 2012).

Increase in the frequency of BSI due to Candida spp., over the last two decades,

has been attributed to increase in the number of susceptible hosts i.e., immuno-

compromised and critically ill intensive care unit- (ICU) patients (Pfaller and Diekema,

2007). Other possible factors that contribute to high candidemia rates include advanced

surgical procedures, increasing use of large catheter devices (central venous catheters),

antifungal and antibacterial drugs, increased organ transplants and cancer chemotherapy

(Klevay et al., 2009). Furthermore, continuous and inappropriate usage of antifungal

drugs has been postulated to result in emergence of several drug resistant Candida spp.


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including C. glabrata and C. krusei in tertiary-care hospitals around the world (Krcmery

and Barnes, 2002). Ironically, despite the development and use of various potent

antifungal drugs, the mortality rate attributable to candidiasis remains relatively

unchanged at about 49% (Gudlaugsson et al., 2003).

Clinical manifestations of infections due to Candida spp. range from mild fever

to fatal sepsis with multi-organ failure. In immunocompetent hosts, Candida spp. can

cause superficial infections on skin and mucosa, however, they can successfully

disseminate throughout the body resulting in severe, life-threatening, deep-organ

infections in immuno-suppressed individuals (Odds, 1987). Candida infections other

than skin or mucosal infections are known as invasive candidiasis which involves spread

and colonization of Candida to multiple organs including kidneys, brain, myocardium

and eyes via bloodstream (Lim et al., 2012). Chronic disseminated candidiasis is

prevalent in patients with acute leukemia (Massod and Sallah, 2005). Candida infections

range from oropharyngeal candidiasis (also known as oral thrush) to severe systemic

infections in patients suffering from AIDS (acquired immune deficiency syndrome)

(Odds, 1987). Genitourinary candidiasis is a common clinical complication seen in

women and ~ 75% of all adult women encounter at least one episode of vulvovaginal

candidiasis (VVC) during their child-bearing years (Achkar and Fries, 2010). C. albicans

ranks as the top most species in ~ 90% of VVC cases worldwide (Achkar and Fries,

2010). Notably, C. glabrata is responsible for about 37% cases of VVC in India (Achkar

and Fries, 2010).

About 95% of all Candida BSIs are caused globally by four spp., C. albicans, C.

glabrata, C. tropicalis and C. parapsilosis (Pfaller, et al., 2010). The remainder 5%

infections are caused by 12-14 other spp. including C. krusei (2 to 3%), C. guilliermondii

(0.5 to 1%), C. lusitaniae (0.5 to 0.6%), C. rugosa (0.03 to 0.7%), C. famata (0.08 to

0.5%), C. dubliniensis (0.01 to 0.1%), and others (Pfaller et al., 2010). Among all

Candida spp., C. albicans is still the most frequently isolated yeast species from blood

cultures and tissue samples (Pfaller and Diekema, 2007). However, an epidemiological

change to non-albicans Candida spp., such as C. glabrata, C. parapsilosis, C. tropicalis,

C. dubliniensis, and C. krusei has been observed over the last two decades (Krcmery and

Barnes, 2002; Pfaller et al., 2010). Of, non-albicans Candida spp., C. glabrata has

emerged as the second most important fungal pathogen in recent years, partly, due to its

high intrinsic resistance to azole antifungals (Krcmery and Barnes, 2002; Pfaller et al.,

2010, 2011, 2012).


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1.1.3 Epidemiology of Candida infections

Candida spp. currently account for 8 to 10% of all hospital-acquired bloodstream

infections world-wide and are exceeded in frequency only by gram-negative

staphylococci, Staphylococcus aureus and enterococci (Wisplinghoff et al., 2004). In

ICU patients, Candida spp. can account for upto 15% of nosocomial infections with

crude mortality rate of 25 to 60% (Bassetti et al., 2010). A recent study from the

SENTRY Antifungal Surveillance Program has revealed species distribution in clinical

isolates collected from 79 medical centres distributed across the globe and reported that

37.5% cases of candida infections are community-onset (CO) while the rest of the 63.5%

cases are hospital-acquired (Pfaller et al., 2011). The highest proportion of CO Candida

infections is being detected in USA with a frequency as high as 50.8% of total BSI

(Pfaller et al., 2011).

Several global multicentre analyses have studied changes in species distribution

and antifungal resistance pattern over last two decades (Pfaller et al., 2007b, 2010, 2011,

2012). Since 1990, C. albicans has been the most commonly isolated fungal pathogen

from the blood till date (Pfaller and Diekema, 2007). However, prevalence of C.

albicans as the major cause of candidiasis has come down to ~ 50% during last 10 years

compared to ~ 75-80% reported earlier (Pfaller and Diekema, 2007). Notably,

distribution of the species responsible for invasive candidiasis has shifted from C.

albicans to non-albicans Candida spp. such as C. glabrata, C. tropicalis and C. krusei

(Bassetti et al., 2010; Pfaller et al., 2011). Incidence of infection by non-albicans

Candida spp. is higher in ICU patients because of their prolonged hospitalization, use of

broad-spectrum antibiotics, presence of intravascular catheters, parenteral nutrition, etc.

(Pfaller et al., 2011). Geographical distribution of C. albicans ranges from 37% in Latin-

America to 70% in Norway and Finland (Pfaller and Diekema, 2007; Falagas et al.,

2010). C. glabrata is typically the second most common isolated species (8% to 20% of

total isolates) in USA and several European countries including Norway, Spain and

Denmark (Pfaller and Diekema, 2007). In other geographical regions including Spain,

Latin-America and Asia, C. parapsilosis and C. tropicalis are the most frequently

isolated non-albicans Candida spp. (Pfaller and Diekema, 2007).

Studies from different parts of India have uncovered regional variations in the

prevalence of candidiasis. While candidemia incidence rate of 1.6 per 1,000 hospital


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admissions was reported in Lucknow (Verma et al., 2003), incidence was 0.71 episodes

per 1,000 patients in North Indian hospitals (Singh et al., 2011). Importantly, C.

tropicalis is the second most important Candida spp. in Indian clinical settings

(Chakrabarti et al., 2008) while C. glabrata and C. parapsilosis could either be third or

fourth most common blood isolate. Intriguingly, a recent study found C. tropicalis to be

the most prevalent in candidemia patients (Kothari and Sagar, 2009).

Besides geographic location, prevalence of Candida spp. also depends upon the

patient age. For example, C. albicans and C. parapsilosis are the most frequent

pathogens isolated from neonates and children with occurrence rate of 39.4 and 43.9%

and 42.4 and 38.3%, respectively while C. glabrata, C. krusei and C. tropicalis are more

frequent in adult population with 17.2, 4.9 and 4.8% prevalence rate, respectively (Blyth

et al., 2009).

1.1.4 Antifungal therapies

Emergence of fungal diseases in early 1970s and increase in mortality rate

associated with fungal infections led to the development and discovery of novel

antifungal agents (Cowen and Steinbach, 2008). Currently available drugs for treatment

of mycoses belong to four main classes, viz., polyenes, azoles, pyrimidines and

echinocandins, as described below.

1.1.4.1 Polyenes

Polyene antifungals bind to ergosterol in the plasma membrane and disrupt

membrane permeability via pore formation. Leakage of intracellular contents through

these pores eventually leads to cell death (Akins, 2005, Cowen and Steinbach, 2008). Of

polyenes, amphotericin B (AMB) deoxycholate is the only approved drug for systemic

use, however, its use is very limited due to adverse side effects including severe

nephrotoxicity to the host (Ellis, 2002). Lipid-based formulations of AMB are

considerably less toxic and an effective treatment for children and adults with invasive

candidiasis (Akins, 2005; Cowen and Steinbach, 2008).

1.1.4.2 Azoles

Azole compounds are the most frequently used antifungal agents for the

treatment of fungal infections worldwide and consist of two broad structurally-unrelated

classes: imidazoles (ketoconazole, clotrimazole and miconazole) and triazoles

(itraconazole, fluconazole, voriconazole and posaconazole) (Akins, 2005; Cowen and

Steinbach, 2008). Both classes selectively inhibit the fungal cytochrome P450 enzyme,


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that carry out sterol C-14α-demethylation, resulting in decreased ergosterol synthesis and

accumulation of toxic intermediate methylated sterols (Akins, 2005). Although azole

antifungals are fungistatic in nature, they are preferred therapeutic agents than AMB due

to their broad spectrum activity and less toxicity (Akins, 2005; Cowen and Steinbach,

2008).

1.1.4.3 Pyrimidines

Antifungal activity of pyrimidine compounds is via inhibition of DNA and

protein synthesis in the fungal cell (Akins, 2005). Among this class, only flucytosine (5-

fluorocytosine) is approved for systemic use (Akins, 2005). However, due to limited

activity spectrum, toxic effects and rapid resistance emergence, use of flucytosine for

treatment of invasive infections is restricted. It is primarily used in conjunction with

amphotericin B to treat cryptococcal meningitis and endocarditis, meningitis and

hepatosplenic candidiassis (Cowen and Steinbach, 2008).

1.1.4.4 Echinocandins

The newest class of antifungals, echinocandins, which inhibit synthesis of β-1,3-

D-glucan, a major component of fungal cell wall, are composed of three drugs,

caspofungin, anidulafungin and micafungin (Cowen and Steinbach, 2008).

Echinocandins possess fungicidal activity against most of the Candida and Aspergillus

spp. but are ineffective against Cryptococcus and Fusarium spp. (Cowen and Steinbach,

2008). Although echinocandins have been accepted as a common treatment option for

fungal infections, emergence of resistance against caspofungin is now being documented

(Pfaller et al., 2012)

Section 1.2: Candida glabrata and its pathogenesis

1.2.1 General biology, genome and classification

C. glabrata is an asexual haploid fungus found as a commensal in normal flora of

skin, mouth, gastrointestinal and urogenital tracts in humans (Fidel et al., 1999; Silva et

al., 2012). However, under immuno-compromised conditions, it can cause mild mucosal

as well as severe life-threatening systemic infections in host (Fidel et al., 1999; Silva et

al., 2012). C. glabrata belongs to the largest taxonomic class of fungal kingdom,

Ascomycetes, and is classified in subphyla, Saccharomycotina (Fidel et al., 1999). C.

glabrata is a non-dimorphic yeast which exists in small blastoconidia (~ 1-4 µm) form

under both commensal and pathogenic lifestyles (Fidel et al., 1999). Daughter cells are

formed exclusively by budding process in C. glabrata (Fidel et al., 1999) and are shown


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in Figure 1.1. On synthetic low-ammonium-dextrose agar medium, C. glabrata has been

observed to form pseudohyphae (reviewed in Kaur et al., 2005). Further, inactivation of

the transcription factor Ace2 results in cell-separation defect and hypervirulence

(reviewed in Kaur et al., 2005).

Figure 1.1 DIC (Differential interference contrast) image of logarithmic-phase C. glabrata wild-type (BG2) cells depicting bud formation (Magnification, 100X).

Based on 18s rRNA sequence, Kaur et al., showed that C. glabrata is

evolutionarily more closely related to Saccharomyces cerevisiae than to other pathogenic

Candida spp. including C. albicans (Kaur et al., 2005; Figure 1.2). Despite this

phylogenetic conservation with S. cerevisiae, C. glabrata does not undergo a life cycle

of diploid and haploid stages and remains exclusively as a haploid asexual organism

(Silva et al., 2012). Orthologs of genes associated with mating in S. cerevisiae have been

identified in C. glabrata. C. glabrata possesses three mating type-like (MTL) loci

(MTL1, MTL2, and MTL3) and can maintain two distinct ‘a’ and ‘alpha’ haploid mating

types (Srikantha et al., 2003; Muller et al., 2008), however, mating is yet to be reported

in C. glabrata (Tscherner et al., 2011). Similar to S. cerevisiae, C. glabrata is also a

petite-positive yeast and loss of mitochondria or its function is not lethal for its growth.

In fact, these “petite-positive” isolates of C. glabrata are more resistant to antifungals

than wild-type strain (Kaur et al., 2004; 2005).

Genome of a type strain of C. glabrata, CBS138 (ATCC 2001), originally

isolated from human faeces, was sequenced in 2004. Its genome, composed of 13

chromosomes is 12.3 million basepairs (Mbp) in length and encodes ~ 5283 protein-

encoding and 207 tRNA genes (Dujon et al., 2004, http://www.genolevures.org/). The

shortest chromosome of C. glabrata genome is 0.5 Mbp in length while the largest one is


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1.5 Mbp. Average G+C content of protein coding sequences (CDS) in C. glabrata

genome is 41%, while average length of CDS is 1479 bp (Dujon et al., 2004). Further,

only 1.5% of total CDS sequences in C. glabrata genome contain introns (Dujon et al.,

2004).

Figure 1.2 18S phylogeny of Candida spp. and other hemiascomycetes. C. glabrata is phylogenetically closer to S. cerevisiae than to other Candida spp. (Kaur et al., 2005).

C. glabrata and S. cerevisiae genome share significant similarity and synteny

(Dujon et al., 2004). However, probably owing to commensalism with humans, C.

glabrata has lost many genes related to metabolic processes, namely, genes involved in

galactose (GAL1,7,10) assimilation, phosphate (PHO3,5,11-12), nitrogen (DAL1-2) and

sulphur metabolism (SAM4), nicotinic acid, thiamine and pyridoxine biosynthesis

(SNO1-3) (Dujon et al., 2004; Jandric and Schuller, 2011). A recent study has shown

that C. glabrata cannot utilize phytic acid, which is one of the most common sources of

phosphate in plant materials such as fruits and seeds, while S. cerevisiae can easily grow

on phytic acid (Orkwis et al., 2010). Further, inability of C. glabrata to use phytic acid

as a sole phosphate source has been attributed to limited phosphatase activity (Orkwis et

al., 2010). It has also been speculated that lack of phytic acid abundance in mammalian

tissues may have invoked an evolutionary selection pressure on C. glabrata to lose

phytase activity in order to adapt to human host (Orkwis et al., 2010).


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1.2.2 Epidemiology and antifungal resistance

Although C. glabrata is phylogenetically closer to non-pathogenic S. cerevisiae,

it has the ability to cause infections in immunosuppressed individuals (Roetzer et al.,

2011). However, unlike C. albicans hyphal form that helps in tissue penetration,

pathogenicity of C. glabrata is entirely mediated by yeast/blastoconidia form (Fidel et

al., 1999; Brunke and Hube, 2013). Over the last two decades, C. glabrata has emerged

as the second most commonly isolated fungal pathogen in USA and several European

countries and now accounts for ~ 18-25% of total yeast BSI isolates in USA (Pfaller et

al., 2010). A significant increase in the incidence of C. glabrata BSI among ICU patients

in USA, since 1993, has been reported (Pfaller et al., 2011). Prolonged hospital stay (~

18.8 days) and frequent prior antimicrobial use were found to be associated with

increased C. glabrata infections in a multivariate prospective case-control study

(Vazquez et al., 1998). Similarly, fluconazole prophylaxis was reported as a

predisposing risk factor for C. glabrata BSI in cancer patients (Ray et al., 2008). Horn et

al. showed that old-age patients and solid organ transplant recipients are more likely to

be infected with C. glabrata than their younger counterparts (Horn et al., 2009).

However, recent reports suggest that previous use of fluconazole may not be a

significant risk factor for appearance of the fluconazole-resistant C. glabrata in BSI (Lee

et al., 2010; Garnacho-Montero et al., 2010).

Candidemia, an important nosocomial infection, is a major cause of mortality in

Indian hospitals. Although C. albicans and C. tropicalis are the predominant species

found in clinical settings, recent surveys from different Indian hospitals have shown

emergence of C. glabrata as a major pathogen in neonates and burn-patients (Gupta et

al., 2001; 2004; Chakrabarti et al., 2008; Giri and Kindo, 2012). In fact, in one of the

neonatal intensive care unit, C. glabrata was the most common species (42.1%)

responsible for candidemia followed by C. tropicalis (31.6%) and C. albicans (21.1%)

(Gupta et al., 2001).

C. glabrata can cause both mucosal and systemic infections depending upon the

host condition and commonly infects old-age patients, immune-compromised individuals

and patients with diabetes mellitus (Fidel et al., 1999). High mortality rates associated

with C. glabrata infection have partially been attributed to its intrinsic low susceptibility

and ability to acquire resistance to a widely used antifungal drug, fluconazole (Hitchcock

et al., 1993; Vanden-Bossche et al., 1998). In accord, recent studies in USA have shown

an increase in fluconazole resistance among C. glabrata isolates from 2001 to 2007


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compared to 1992-2001 period (Pfaller et al., 2009). In addition, MIC90 (minimum

inhibitory concentration) values for fluconazole for C. glabrata have been found to be as

high as 128 µg/ml in some regions of USA (Pfaller et al., 2009).

Overexpression of multidrug ABC (ATP-binding cassette) class of transporters,

CgCdr1, CgCdr2 and CgSnq2, and fluconazole target, lanosterol 14-α-demethylase

(Erg11), are the most common mechanisms of acquired resistance towards azole

antifungals in C. glabrata (reviewed in Tscherner et al., 2011). Additionally, gain-of-

function mutations in the zinc-cluster transcription factor, CgPdr1 (an orthologue of S.

cerevisiae Pdr1), which regulates expression of genes encoding multidrug efflux pumps,

have been reported to result in increased drug efflux and hyper virulence (Caudle et al.,

2011; Ferrari et al., 2009).

Further, loss of mitochondrial function and defective anionic phospholipid

biosynthesis has been linked with increased expression of CgCDR1, CgCDR2 and

CgSNQ2 (reviewed in Tscherner et al., 2011). Recent reports have demonstrated an

essential role for cell signalling pathways, viz., calcineurin-regulated, heat shock protein

Hsp90-dependent and protein kinase C (PKC)-mediated cell wall integrity pathways in

survival of azole and echinocandin drug stress in C. glabrata (Kaur et al., 2004; Borah et

al., 2011; Singh-Babak et al., 2012), thus, raising the possibility of combinatorial

therapy to treat fungal infections.

1.2.3 Virulence factors

C. glabrata, a harmless resident of the human microflora, has a propensity to

cause both mucosal and life-threatening systemic infections under immuno-

compromised conditions (Roetzer et al., 2011). Despite being the second most important

fungal pathogen in world population, pathobiology of C. glabrata including its virulence

factors, survival and adaptation mechanisms in human host and host defense against this

pathogen remains poorly-defined. Compared to other Candida spp., C. glabrata is

relatively less pathogenic and possesses limited virulence attributes (Roetzer et al., 2011;

Tscherner et al., 2011). Of common fungal virulence factors, C. glabrata displays

phenotypic switching, biofilm formation and ability to adhere to host tissues (Kaur et al.,

2005). However, it lacks some key fungal virulence traits including hyphae formation,

secreted proteolytic activity and mating (Kaur et al., 2005; Tscherner, et al., 2011). Key

virulence attributes of C. glabrata are discussed below.


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1.2.3.1 Adherence: role for Epa1-related adhesin proteins

Adherence has been recognized as one of the major virulence factor of human

microbial pathogens. C. glabrata shows comparable adherence ability to C. albicans

(Klotz et al., 1985). In vitro adherence of C. glabrata to epithelial cells has been

attributed to the expression of a glycosylphosphatidylicositol (GPI)-anchored cell

surface adhesin, Epa1 (Epithelial adhesin 1), which also recognizes host glycans

(Cormack et al., 1999). This adhesin belongs to a family of ∼ 23 cell wall proteins,

majority of which are sub-telomerically-encoded (Castaño et al., 2006). Of these 23

proteins, Epa1, Epa6 and Epa7 have been implicated in adherence of C. glabrata to

epithelial and endothelial cells (Cormack et al., 1999; Castano et al., 2006). Structural

studies of Epa1 suggest that its N-terminal domain is distantly related to S. cerevisiae

flocculins and PA14 like domain-containing proteins (anthrax protective antigen) (Ielasi

et al., 2012). Epa proteins usually have a Ca2+-dependent-carbohydrate (ligand)-binding

site at N-terminus and heavily glycosylated serine/threonine-rich region at C-terminus

(Ielasi et al., 2012).

Expression of telomere-associated EPA1-related genes is regulated in response to

environmental cues via chromatin-based transcriptional silencing (Castano et al., 2006).

Under in vitro culture conditions, Epa1 is the major adhesin expressed on the cell surface

while Epa6 and Epa7 are transcriptionally silenced due to their telomeric localization

(Castano et al., 2006). In urinary tract of murine model, limitation of nicotinic acid, a

precursor for NAD+ (Nicotinamide adenine dinucleotide), relieves NAD+-dependent

histone deacetylase Sir2-mediated silencing which results in the derepression of EPA6

(Domergue et al., 2005). In accord, mutations in telomeric silencing factors CgSir2,

CgSir3 and CgSir4 result in induced expression of EPA1, EPA6 and EPA7 causing

hyper-adherence of C. glabrata cells to cultured epithelial cells and increased ability to

colonize murine kidneys (Domergue et al., 2005; Castano et al., 2006). Further, using

glycan microarrays, Zupancic et al. showed that the carbohydate-binding specificity of

Epa6 and Epa7 is dependent on a five-amino acid region within their N-terminal ligand-

binding domain (Zupancic et al., 2008). They also reported that while Epa1 and Epa7

exclusively bind to galactose residue in β-1-3 or β-1-4 linked glycosides, Epa6 could

bind to either α-linked or β-linked terminal galactose residues, implying a broader

substrate specificity for Epa6 (Zupancic et al., 2008). Recently, a high-resolution crystal

structure of Epa1 N-terminal domain complexed with disaccharide ligands revealed that


13

two inner loops, CBL1 and CBL2, involved in calcium binding, and three outer loops

L1, L2 and L3, constituting the main carbohydrate attachment site, determine ligand

specificity (Maestre-Reyna et al., 2012).

1.2.3.2 Phenotypic switching

A heritable trait of high-frequency reversible phenotypic switching, which results

in a variety of colony morphologies, is an important determinant of C. albicans virulence

and occurs at sites of Candida infection (Calderone and Fonzi, 2001). Highly

spontaneous and reversible phenotypic switching has also been reported in C. glabrata

on solid medium containing copper sulphate (CuSO4) (Lachke et al., 2002). Four colony

phenotypes observed under these conditions were white (W), light brown (LB), dark

brown (DB) and very dark brown (vDB) (Lachke et al., 2002). Besides color, these

colonies displayed varied expression of metallothionein gene, MTII,

(Wh<LB<DB>vDB) and different morphologies i.e, budding cells, psuedohyphae and

tubular forms (Lachke et al., 2002). C. glabrata is also known to switch reversibly

between smooth and irregular wrinkled colonies (Lachke et al., 2002). Natural C.

glabrata isolates are predominantly DB and higher colonization of DB form has recently

been reported in spleen and liver during systemic murine infection (Brockert et al., 2003;

Srikantha et al., 2008). Importantly, it has also been postulated that different C. glabrata

switch phenotypes may colonize different host niches (Brockert et al., 2003).

1.2.3.3 Biofilm formation

An increase in the incidence of Candida infections in the last few decades has

largely been associated with biofilm formation on surgical implants or artificial devices

including catheters, stents, shunts, prostheses and pacemakers wherein C. albicans is the

most commonly found Candida spp. (Silva et al., 2010). C. glabrata has also been

shown to form thin biofilms on plastic surfaces in vitro (Iraqui et al., 2005; Silva et al.,

2010; Kraneveld et al., 2011). In contrast to C. albicans biofilm which is composed of

yeast cells, hyphae and pseudohyphae, C. glabrata biofilm exists as a multilayer

structure of only yeast cells embedded in the extracellular matrix (Silva et al., 2010). C.

glabrata has recently been shown to exhibit increased biofilm formation on the silicon

surface in the presence of urine (Silva et al., 2010). Four genes, CgRIF1, CgSIR4,

CgYAK1 and EPA6, were found to be pivotal for biofilm formation in C. glabrata and

lack of Epa6 significantly abolished biofilm formation in vitro (Iraqui et al., 2005).

EPA6 expression was induced in Flo8 and Mss11 transcriptional factor-dependent

manner upon exposure to weak acid-related chemical preservatives sorbic acid and


14

parabens and resulted in an increased adherence to the human vaginal epithelium

(Mundy and Cormack, 2009). Expression of other EPA genes, EPA1, EPA3, EPA7 and

EPA22, has also been shown to be upregulated in biofilms grown in semi-defined

medium (Kraneveld et al., 2011).

1.2.3.4 Pigment formation

Pigments are known to protect fungal pathogens from oxidative stress (reviewed

in van Burik and Magee, 2001). C. glabrata was earlier thought to be a non-pigmented

yeast, however, recently, indole-derived pigments, whose generation is dependent on the

presence of tryptophan as the sole nitrogen source in the medium and occurs via the

Ehrlich pathway, have been identified (Mayser et al., 2007; Brunke et al., 2010).

Furthermore, pigmented C. glabrata cells were better equipped to survive antifungal

response of human neutrophils and damage human epithelial cells (Brunke et al., 2010),

indicating a potential role for pigmentation during interaction with host cells.

1.2.3.5 Hydrolases

Many pathogenic fungi including C. albicans have the ability to secrete various

hydrolytic enzymes such as proteinases, lipases and phospholipases. Major role for these

hydrolytic enzymes is to provide nutrients for fungal cell proliferation, evade the host

immune defense system, and facilitate further tissue penetration and invasion (reviewed

in van Burik and Magee, 2001). Information available regarding these hydrolases in C.

glabrata is briefly summarized below.

1.2.3.5.1 Phospholipases

Phospholipase activity (ability to hydrolyze phospholipids) has been reported for

C. glabrata isolates (Kantarciolu and Yücel, 2002). Consistent with this, C. glabrata

genome harbours three genes, which are orthologs of S. cerevisiae phospholipase

encoding ORFs (www.candidagenome.org), however, role for these phospholipases in C.

glabrata virulence is yet to be examined.

1.2.3.5.2 Proteinases

Proteases are key virulence determinants of fungal pathogens and role for

secreted aspartyl proteases (Saps) in the virulence of C. albicans has extensively been

studied (reviewed in Naglik et al., 2003). In vitro production of proteinase in C. glabrata

was reported for the first time in 1991 (Chakrabarti et al., 1991) and a family of eleven

GPI-linked aspartyl proteases (CgYps1-11) has recently been identified in C. glabrata

(Kaur et al., 2007). However, despite the presence of these eleven proteases, secreted

proteolytic activity has not been observed in C. glabrata (Kaur et al., 2005; Silva et al.,


15

2012). Since the current study is aimed at deciphering the functions of aspartyl proteases

in the pathobiology of C. glabrata, literature related to the role for aspartyl proteases in

fungal cell physiology and virulence is discussed, in detail, in Section 1.3.

1.2.4 Interaction with host cells

Phagocytic cells of host innate immune system represent the first line of defense

against systemic candidiasis and play important roles in clearance of fungal infections

via phagocytosis and anti-inflammatory and ROS response (Netea et al., 2008; Romani,

2011). β-glucan and mannan present in the cell wall constitute major common pathogen-

associated molecular patterns for C. albicans (Klis et al., 2001; Poulain and Jouault,

2004). In accord, fungicidal antibodies and immune receptors have been shown to target

the inner β-glucan layer (Netea et al., 2008; Romani, 2011). Furthermore, dendritic cells

are known to recognize, phagocytose and process C. albicans for antigen presentation to

initiate the T-helper cellular immune response, which is central to mammalian host’s

long term resistance to candidiasis (Netea et al., 2008; Romani, 2011).

C. glabrata has recently been shown to replicate in murine and human monocyte-

derived macrophages (Kaur et al., 2007; Seider et al., 2011; Rai et al., 2012). This is in

contrary to the no intracellular proliferation of non pathogenic yeast S. cerevisiae (Kaur

et al., 2007). Survival of C. glabrata in macrophages is dependent upon metabolic

reprogramming, chromatin remodelling and maintenance of energy homeostasis (Kaur et

al., 2007; Rai et al., 2012). Similar to C. albicans, C. glabrata reconfigures its carbon

metabolism via up-regulation of genes involved in gluconeogenesis, glyoxylate

pathways and β-oxidation of fatty acids and repression of genes implicated in glycolysis

(Lorenz et al., 2004; Kaur et al., 2007). Although C. glabrata can successfully survive

and replicate in macrophages, its internalization does not lead to macrophage apoptosis

(Seider et al., 2011). Prevention of phagosome acidification appears to be a key

mechanism that C. glabrata employs to proliferate in the macrophage internal milieu

(Seider et al., 2011; Rai et al., 2012). Genes implicated in several physiological

processes, viz., chromatin remodelling, DNA repair, Golgi vesicle transport and cell wall

organization, have recently been identified to be required for survival and/or replication

of C. glabrata in human THP-1 macrophages (Rai et al., 2012). Notably, C. glabrata is

also known to display high intrinsic tolerance to several stresses including oxidative

stress (Cuéllar-Cruz et al., 2008; Roetzer et al., 2011).


16

1.2.5 Animal model systems to study C. glabrata pathogenesis

Owing to the relative low pathogenicity of C. glabrata, development of animal

model systems to study virulence of C. glabrata has not achieved the desired pace. Few

established murine models of C. glabrata infections include systemic, gastrointestinal

and vaginal candidiasis models (Fidel et al., 1996; Calcagno et al., 2003; Jacobsen et al.,

2010). Unlike C. albicans, systemic C. glabrata infection does not lead to clear clinical

manifestations even if mice are mildly immunocompromised (Calcagno et al., 2003).

However, prior treatment with immunosuppressive drug, cyclophosphamide, and high C.

glabrata inoculum (2 x 108 cells) do result in successful establishment of the disease

(Calcagno et al., 2003). Nevertheless, mortality of infected mice cannot be a sole

virulence evaluation criterion as, even with 100% mice mortality, evidence of necrosis or

inflammation around the site of C. glabrata infections are usually absent (Calcagno et

al., 2003). Notably, immunocompetent Balb/C mice have successfully been utilized to

measure the relative fitness of different C. glabrata strains via assessment of fungal

burden in different tissues in the disseminated model of candidiasis (Kaur et al., 2007;

Ferrari et al., 2009; Jacobsen et al., 2010).

Recently, Drosophila melanogaster infection model has been established to

study C. glabrata virulence (Roetzer et al., 2008). In this system, mutant MyD88 flies

defective in the humoral arm (Toll signalling) of the antifungal response were generated

which showed significantly increased mortality upon C. glabrata infection (Roetzer et

al., 2008). Other animal model systems including wax moth and silk worm larval

models, Caenorhabditis elegans and Zebrafish, have also been developed to study C.

glabrata pathogenesis (reviewed in MacCallum, 2012), however, their suitability as an

alternative to animal model systems need to be further investigated.

Section 1.3: Aspartyl proteases: structure, regulation and function

1.3.1 Structural features and catalytic mechanism

Proteases are a group of enzymes that catalyze the cleavage of peptide bonds

(CO-NH) in proteins. Based on the nature of the functional group at the active site and

the mechanisms of catalysis, they are broadly classified into eight classes, viz.,

asparagine, aspartic, cysteine, glutamic, metallo, serine, threonine, and unknown

(reviewed in Bairwa et al., 2013). While aspartic (aspartyl) proteases constitute a small

group of the protease family, they are nevertheless ubiquitous in nature and involved in

numerous physiological processes (Dunn et al., 2002). Aspartyl proteases with


17

molecular weight between 35 and 40 kDa are characterized by the presence of two

active-site aspartate (Asp) residues at their catalytic centre. They are optimally active at

acidic pH and inhibited by pepstatin A (reviewed in Bairwa et al., 2013).

The MEROPS database (http://merops.sanger.ac.uk) classifies aspartyl proteases

into 16 families (A1, A2, A3, A5, A8, A9, A11, A22, A24, A25, A26, A28, A31, A32,

A33 and A36) based on the structural and/or sequence similarities. Most of the known

fungal aspartyl proteases belong to the A1 family (pepsin A) of aspartyl proteases and

include aspergillopepsin I, candidapepsins, endothiapepsin, and rhizopuspepsin from

fungi and yapsins from S. cerevisiae (reviewed in Bairwa et al., 2013).

Aspartyl proteases of A1 family are usually monomeric, bilobal proteins, which

are synthesized as inactive zymogens (Dunn et al., 2002). One catalytic Asp residue is

contributed by each lobe with the active site located between two lobes of the enzyme.

An extended active-site cleft, which accommodates at least 7 amino acids, is a

characteristic feature of aspartyl proteases. Binding of at least 7 amino acids at the active

site facilitates tight interaction between the enzyme and the substrate, resulting in

maximum cleavage efficiency (Dunn et al., 2002). The catalytic Asp residues in most

members of the pepsin family are contained in an Asp-Thr-Gly-Xaa motif in both N- and

C-terminal lobes of the protease wherein Xaa is usually either Ser or Thr (Dunn et al.,

2002).

Structural and kinetic studies have shown that aspartyl proteases hydrolyze the

peptide bond by general acid-base catalysis mechanism instead of forming covalent

intermediates (Pearl and Taylor, 1987). In this mechanism, substrate’s peptide carbonyl

group undergoes a nucleophilic attack by aspartate-activated water molecule resulting in

the formation of a tetrahedral transition state intermediate (Polgar, 1987). This

tetrahedral transition intermediate is electrostatically stabilized by proximal tyrosine

residue present in the catalytic center of the enzyme. The second catalytic aspartate

residue of the enzyme then resolves this tetrahedral adduct by donating a proton to the

amino group produced by the hydrolysis of the peptide bond. Thus, one Asp (Asp-COO-

) acts as a general base to deprotonate the attacking H-OH while the second Asp (Asp-

COOH) acts as an acid by releasing a proton to the carbonyl oxygen of the resultant

amine product (Polgar, 1987). The catalytic rate of the enzyme depends upon the amino-

acid composition and length of the substrate (Pearl and Taylor, 1987).


18

1.3.2 Aspartyl proteases in pathogenic fungi

Several species of filamentous fungi and yeasts have been shown to harbor

aspartyl proteases in their proteome (reviewed in Bairwa et al., 2013) and aspartyl

proteases constitute one of the most common virulence trait of pathogenic fungi.

Aspartyl proteases from fungal spp. show significant similarity to each other (reviewed

in Bairwa et al., 2013). A typical fungal aspartyl protease contains a N-terminal signal

peptide, a pro-peptide region of approximately 17-65 amino acids, two catalytic domains

(β-structures) harboring two catalytic aspartic-acid residues within the hydrogen-

bonding distance of each other, a conserved C-terminus region (reviewed in Bairwa et

al., 2013 and shown in Figure 1.3). A GPI signal at the C-terminal is a unique trait of

yapsin protease family (reviewed in Bairwa et al., 2013 and shown in Figure 1.3). Based

on their location, they are of three types: secreted (Saps), destined to the cell

membrane/cell wall via a GPI anchor (Yaps) or destined to the vacuole (reviewed in

Bairwa et al., 2013)

1.3.2.1 Secreted aspartyl proteases

Expression and activity of extracellular proteolytic enzymes is tightly regulated

during individual stages of the fungal infection process. In C. albicans, extracellular

proteolytic activity is largely attributed to a family of ten secreted aspartyl proteases

(Sap1-10) which are isoenzymes with similar functions but different biochemical

properties (Naglik et al., 2003; Albrecht et al., 2006). They are encoded by the SAP gene

family (Naglik et al., 2003). Of ten Saps, eight are secreted (Sap1-Sap8) while Sap9 and

Sap10 belong to the class of GPI-anchored aspartyl proteases (Naglik et al., 2003;

Albrecht et al., 2006; Figure 1.3A and Table 1.1). Although Sap9 and Sap10 show 30-

36% similarity to S. cerevisiae asaprtyl proteases, they are phylogenetically more closely

related to other Saps than to their S. cerevisiae counterparts (Parra-Ortega et al., 2009).

Among SAP gene family, gene sequences of SAP2 and SAP3 show ~ 76% identity to

each other and to SAP1 while SAP4, SAP5 and SAP6 exhibit 87-94% identity to each

other. SAP4, SAP5 and SAP6 represent a distinct subgroup displaying ~ 70% identity to

SAP1, SAP2 and SAP3. SAP7 is the most diverged member of the group which is only

44% identical to SAP1-6. SAP8 sequence is 73% similar to SAP1 and 70% to SAP3.

Orthologs of Saps have been reported in other closely related spp. including C.

tropicalis, C. parapsilosis, and C. lusitaniae (Parra-Ortega et al., 2009). Notably, genes


19

coding for Saps have not been identified in C. glabrata and homologues of C. albicans

SAP5 and SAP6 are absent in C. dubliniensis (reviewed in Bairwa et al., 2013).

Figure 1.3: Schematic representation of the protein domain structures, modeled from Pfam database (http://pfam.sanger.ac.uk/family/pf00026), of aspartyl proteases in C. albicans (A) and C. glabrata and S. cerevisiae (B). Images are not drawn to scale.

1.3.2.1.1 Regulated expression of SAP genes

Individual members of the SAP gene family are differentially regulated under

various environmental conditions, viz., temperature, pH, nitrogen source and glucose

concentration, in different in vivo models and candidosis patients (reviewed in Bairwa et

al., 2013). Expression of Saps is also specific for the morphological form of C. albicans.

Saps 1–3 are secreted only by the yeast cells while Saps 4–6 are expressed only by the

hyphal forms of C. albicans (White and Agabian, 1995; Naglik et al., 2004).

Intriguingly, Sap 4-6 contain an integrin binding RGD motif that is postulated to

contribute to tissue invasion by hyphal cells (Parra-Ortega et al., 2009).


20

Table 1.1: Characteristics of known and putative fungal aspartyl proteases.

Aspartyl protease Length (amino acids)

Molecularweight (kDa)

Uniprot ID

Peptidase family

Sub-cellular localization pI pH

optimum

Active site

location (amino acids)

Signal peptide(amino acids)

Saccharomyces cerevisiae ScYps1 (YLR120C)¶ 569 60.0 P32329 A1 Cell surface/GPI 4.5 4.0 101, 371 1-21 ScYps2 (YDR144C) 596 64.2 P53379 A1 Cell surface/GPI 4.4 4.0 99, 360 1-22 ScYps3 (YLR121C) 508 54.5 Q12303 A1 Cell surface/GPI 9.3 4.0 81, 288 1-20 ScYps6 (YIR039C) 537 58.2 P40583 A1 Cell surface/GPI 3.9 ND 95, 324 1-24 ScYps7 (YDR349C)¶ 596 64.4 Q06325 A1 Cell surface/GPI 4.6 ND 74, 321 1-25 ScBar1 (YIL015W) 587 63.7 P12630 A1 Secreted 4.5 ND 63, 287 1-24 ScPep4 (YPL154C) 405 44.5 P07267 A1 Vacuolar 4.5 ND 109, 294 1-22

Candida albicansCaSap1 (orf19.5714)¶ 391 41.6 P0CY27 A1 Secreted 4.9 5.0 82, 267 1-18 CaSap2 (orf19.3708)¶ 398 42.3 P0DJ06 A1 Secreted 4.2 4.0 88, 274 1-18 CaSap3 (orf19.6001)¶ 398 42.8 P0CY29 A1 Secreted 4.2 3.0 90, 274 1-18 CaSap4 (orf.19.5716)¶ 417 45.3 Q5A8N2 A1 Secreted 5.0 5.0 107, 293 1-18 CaSap5 (orf19.5585)¶ 418 45.6 P43094 A1 Secreted 6.1 5.0 108, 294 1-18 CaSap6 (orf19.5542)¶ 418 45.4 Q5AC08 A1 Secreted 8.2 5.0 108, 294 1-18 CaSap7 (orf.19.756)¶ 588 62.5 Q59VH7 A1 Secreted 4.3 6.5 244, 464 1-16 CaSap8 (orf19.242)¶ 405 43.0 Q5AEM6 A1 Secreted 6.3 2.5 107, 292 1-17 CaSap9 (orf19.6928)¶ 544 58.5 O42779 A1 Cell surface/GPI 4.8 5.5 83, 371 1-17 CaSap10 (orf19.3839)¶ 453 49.3 Q5A651 A1 Cell surface/GPI 4.0 6.0 70, 266 1-20 CaYps7 (orf19.6481) 701 75.8 Q5AH56 A1 Cell surface/GPI 4.6 ND ND, 571 ND CaSap30 (orf19.2082) 435 47.9 Q5ACY5 A1 Secreted 3.9 ND 56, 232 1-16 CaSap98 (orf19.852) 364 39.6 Q5AHE4 A1 Secreted 5.6 ND 76, 248 1-19 CaSap99 (orf19.853) 363 39.1 Q5AHE3 A1 Secreted 5.9 ND 75, 247 1-17 CaApr1 (orf19.9447) 419 45.4 Q59U59 A1 Vacuolar 4.4 ND 122, 307 1-22

Candida glabrata CgYps1 (CAGL0M04191g) 601 63.7 Q6FJR5 A1 Cell surface/GPI 5.0 ND 91, 378 1-18

CgYps2 (CAGL0E01419g) 591 63.2 Q6FVJ4 A1 Cell surface/GPI 4.4 ND 85, 369 1-18

CgYps3 (CAGL0E01727g) 531 58.9 Q6FVI0 A1 Cell surface/GPI 6.4 ND 71, 308 1-14

CgYps4 (CAGL0E01749g) 482 53.2 Q6FVH9 A1 Cell surface/GPI 8.4 ND 68, 306 1-15



CgYps7 (CAGL0A02431g) 587 63.4 Q6FY32 A1 Cell surface/GPI 4.7 ND ND 1-18




CgYps11 (CAGL0E01881g) 508 55.5 Q6FVH3 A1 Cell surface/GPI 5.0 ND 66, 313 ND

CgYps12 (CAGL0J02288g) 541 59.5 Q76IP5 A1 ND 4.6 ND 71, 282 ND

CgPep4 (CAGL0M02211g) 415 45.4 Q6FK02 A1 Vacuolar 4.6 ND 109, 301 1-22

ND = Not determined, kDa = kilodalton, pI = Isoelectric point. ¶ Various parameters have been experimentally demonstrated.


21

In C. albicans, extracellular pH plays an important role in regulating the

expression of secreted aspartyl proteases (Naglik et al., 2003). Expression of Sap

isoenzymes in C. albicans increases in the late log and the stationary phase of growth

when surrounding medium pH is acidic (White and Agabian, 1995). Specifically,

expression of SAP2 mRNA had been shown to be maximal at acidic pH 4.0 while SAP3

was found to be optimally expressed at pH 3.2 (Hube et al., 1994; White and Agabian,

1995). Expression of C. albicans SAP1-7 was found to be induced in both oral and

vaginal cavity, which varies in their internal pH conditions (Hube et al., 1994; De

Bernardis et al., 1995; Naglik et al., 1999).

Further, expression of Sap2, Sap3 and Sap8 has been reported to be elevated

under low temperature conditions (Crandall and Edwards, 1987). An increased

expression of SAPs in the presence of complex nitrogen sources, viz., bovine serum

albumin (BSA), haemoglobin and collagen, in nitrogen-starved C. albicans cells has also

been observed (Naglik et al., 2003; reviewed in Bairwa et al., 2013).

In C. albicans WO-1 strain, expression of SAP1 is tightly regulated by colony

morphology and coupled to the white-opaque switch (Morrow et al., 1993). While log-

phase opaque cells show high expression of SAP1 mRNA, switching to the white colony

phenotype results in a reduction in the SAP1 expression (Morrow et al., 1993). In the

same strain, SAP3 expression is linked with opaque cells while SAP2 mRNA can be

detected in log phase of both white and opaque colonies (Morrow et al., 1993).

Expression of other SAPs, viz., SAP4, SAP5 and SAP6, is dependent upon pH and

is generally induced at pH 6.0 or above (Hube et al., 1994). Transcript levels of SAP4,

SAP5 and SAP6 are elevated during serum-induced hyphal development at neutral pH

with SAP6 being the most abundant transcript followed by SAP5 and SAP4 (Hube et al.,

1994; White and Agabian, 1995). SAP7 expression has not been observed in vitro,

although, few studies have shown induced expression of SAP7 in vaginal candidiasis

patients and mice (Naglik et al., 2003; Taylor et al., 2005). SAP8 mRNA is expressed

preferentially in yeast cells at early logarithmic phase at 25˚C and also detectable at 37˚C

at a lower level (Monod et al., 1998). In contrast, SAP9 is expressed preferentially in

later growth phases when SAP8 expression is reduced (Monod et al., 1998). Both SAP9

and SAP10 are expressed in commensal and infection stages and are responsible for

survival of C. albicans in oral cavity (Albrecht et al., 2006).

C. albicans SAP2 is generally induced in late stages of the infection when fungal

cells had spread into the deep tissue and SAP2 expression had also been observed in


22

vaginitis infection along with SAP1 (De Bernardis et al., 1995; Staib et al., 1999). Of all

10 SAPs in C. albicans, SAP5 and SAP9 are the most highly expressed SAPs in vivo

(Naglik et al., 2008). SAP5 expression has also been found to be induced during early

stage of the infection in a mouse model of vaginitis (Taylor et al., 2005). SAP8

expression has recently been found to be highly upregulated in mature biofilms produced

by C. albicans strains isolated from patients with denture stomatitis (Ramage et al.,

2012). Further, SAP9 and SAP10 expression has been detected in clinical patients with

oral Candida infections (Naglik et al., 2008).

1.3.2.1.2 Role for secreted aspartyl proteases in pathogenicity

C. albicans Saps exhibit broad substrate specificity and are active over a wide

range of pH, viz., pH 2.0 to pH 7.0. (reviewed in Bairwa et al., 2013). They cleave

various mammalian proteins including mucin, keratin, laminin, fibronectin, collagen,

albumin, hemoglobin, salivary lactoferin, interleukin-1β, cystatin A, and

immunoglobulin A (listed in Table 1.2) and facilitate processes of nutrient acquisition,

tissue invasion and evasion of immune responses (Naglik et al., 2003; Naglik et al.,

2004). In accord, Sap-mediated proteolysis in C. albicans regulates the host immune

system. C. albicans Saps degrade human host complement factors such as C3b, C4b and

C5 under in vitro growth conditions (Naglik et al., 2003; Gropp et al., 2009). Sap2 has

been shown to possess mucinolytic activity at low pH 3.5 in vitro and promote tissue

invasion and spread of the fungus (Colina et al., 1996). Sap-mediated degradation of

salivary protein, lactoferrin, and cathepsin D in the oral cavity has also been reported

(Naglik et al., 2003; Naglik et al., 2004). Furthermore, Sap5 has been implicated in

degradation of the human E-cadherin during oral mucosal tissue invasion (Villar et al.,

2007).

C. albicans mutants lacking Saps have been found to be attenuated for virulence

(reviewed in Naglik et al., 2003). Homozygous null C. albicans mutants deleted for

SAP1, SAP2 and SAP3 were first generated by Hube et al. using Ura (uracil)-blaster gene

disruption method (Hube et al., 1997). Among these, sap2 mutant showed considerably

reduced proteolytic activity and was significantly avirulent in a murine model of

disseminated oral candidiasis. sap1 and sap3 null mutants also showed moderate

virulence attenuation (Hube et al., 1997). Using similar gene-deletion strategy, Sanglard

et al. constructed a mutant deleted for SAP4, SAP5 and SAP6 and found significantly

attenuated virulence in murine models of disseminated candidiasis and peritonitis which


23

suggest an important role for Sap4-6 isoenzymes in C. albicans pathogenesis (Sanglard

et al., 1997).

Additionally, adherence of each of the sap1, sap2 and sap3 null mutants was

significantly reduced to human buccal epithelial cells while sap4-6 mutant was

hyperadherent under these conditions (Hube et al., 1997; Sanglard et al., 1997).

Recently, sap7 homozygous mutant has been created (Taylor et al., 2005). Despite a

persistent SAP7 expression in intravenous vaginal infection, sap7 null mutant was not

avirulent in a vaginal model of candidiasis (Taylor et al., 2005).

C. albicans Sap9 and Sap10, which show significant homology to S. cerevisiae

aspartyl proteases, have been implicated in cell wall integrity. C. albicans strains deleted

for either SAP9 or SAP10 were significantly sensitive to cell wall perturbing agents

including hygromycin B, amorolfine, calcofluor white and congo red and displayed

increased chitin content (3-4%) and abnormal budding phenotype (Albrecht et al., 2006).

Further, deletion of SAP9 and SAP10 rendered cells defective in invasion and damage of

epithelial cells in a reconstituted human epithelium (RHE) model of oral infection

(Albrecht et al., 2006).

1.3.2.2 GPI-linked aspartyl proteases (Yapsins)

Yapsins, which are non-secreted aspartic proteases, have been implicated in the

maintenance of cell wall integrity under environmental stress conditions (Krysan et al.,

2005). They belong to a family of cell surface-localised GPI-linked aspartyl proteases

with an average length of 500-600 amino acids (Gagnon-Arsenault et al., 2006). The

GPI anchor consists of a single phospholipid moiety attached to the plasma-membrane

and a complex head group of a phosphodiester-linked inositol, a glucosamine, a linear

chain of three mannose sugar and a phosphoethanolamine (Canivenc-Gansel et al., 1998;

Ferguson, 1999). An amide bond between the C-terminal residue of the protein and the

amino group of phosphoethnoalmine helps in the attachment of the protein to the GPI-

anchor (reviewed in Chatterjee and Mayor, 2001). The GPI glycolipid anchor, composed

of a conserved core structure of ethanolamine-P-Man3GlcN-PI, is added to the C-

terminus of the protease in the endoplasmic reticulum (Canivenc-Gansel et al., 1998;

Ferguson, 1999; Gagnon-Arsenault et al., 2006). The GPI attachment motif is comprised

of three sequence regions: an anchoring amino acid (the ω site) followed by two small

amino acids, a polar spacer region of 8–12 amino acids and a hydrophobic C-terminal

region of 11-20 amino acid residues (Canivene-Gansel et al., 1998; Ferguson, 1999).


24

S. cerevisiae genome harbours a gene family that codes for five GPI-linked

aspartyl proteases (Yps1-3, Yps6 and Yps7) which are also referred as yapsins (Krysan

et al., 2005; Gagnon-Arsenault et al., 2006 and shown in Figure 1.3B and Table 1.1).

Genes encoding Yps4 and Yps5 are pseudogenes. Yps1 and Yps2 have been shown to

cleave peptides and proteins C-terminal ends to basic residues both in vitro and in vivo

(reviewed in Gagnon-Arsenault et al., 2006). Homologs of yapsins are present in many

fungi including C. albicans, C. glabrata, A. oryzae and A. fumigatus (reviewed in

Bairwa et al., 2013).

In C. glabrata, GPI-linked aspartyl proteases are encoded by a family of eleven

genes (CgYPS1-11) and the CAGL0J02288g ORF (open reading frame) (Figure 1.3B,

Table 1.1). C. glabrata YPS1, YPS2 and YPS7 are orthologs of S. cerevisiae YPS1, YPS2

and YPS7, respectively, and are encoded at syntenic loci (Kaur et al., 2007; Figure 1.4).

A cluster of 8 YPS genes (CgYPS3-6 and CgYPS8-11), designated as ‘C’, is unique to C.

glabrata and is present on the chromosome E (Kaur et al., 2007). The ORF

CAGL0J02288g, which is syntenic to its ortholog, BAR1, in S. cerevisiae codes for

CgYps12 (Figure 1.4). Bar1 in S. cerevisiae is known to be localized to the periplasmic

space of mating type a cells and inactivate alpha factor by proteolytic cleavage (Manney,

1983). C. albicans contains only one YPS gene which encodes CaYps7 (Table 1.1).

The list of aspartyl proteases present in C. albicans, C. glabrata and S. cerevisiae

along with their known and/or predicted properties is presented in table 1.1.

Additionally, Pfam database-modeled domain organization of C. glabrata and S.

cerevisiae yapsins and C. albicans Saps is depicted in Figure 1.3. Presence of multiple

Saps and yapsins in the pathogenic fungi suggests that they probably arose owing to

duplication events in the ancestral sequence and may reflect their pivotal roles in

infection of the mammalian host.


25

Figure 1.4 Synteny of the genes encoding aspartyl proteases between S. cerevisiae (Sc) and C. glabrata (Cg). Orientation of the genes is depicted by the pointed end, the dotted box represents the absence of the corresponding genes in the organism and un-annotated genes are denoted by ORF numbers. White-filled box depicts the gene whose synteny is being presented while grey-filled boxes denote the neighboring genes. The synteny was mapped with the help of these databases: Candida genome data base (http://candidagenome.org/), Candida DB (http://genodb.pasteur.fr/cgi-bin/WebObjects/CandidaDB.woa/ wa/), Genolevures (http://genolevures.org/) and Candida Gene Order Browser (http://cgob.ucd.ie/). Figures are not drawn to scale. Sequence-based modelling of the S. cerevisiae Yps1 structure reveals that the mature

Yps1 enzyme is composed of two subunits, α and β, which are generated after the

removal of N-terminal propeptide of 46 amino acids followed by a proteolytic cleavage

in the serine/threonine (S/T) rich loop region of ~ 100 amino acid residues (reviewed in

Gagnon-Arsenault et al., 2006 and shown in Figure 1.5). Presence of this unique loop

insertion just after the first catalytic Asp residue is unique for yapsin family of aspartyl

proteases (reviewed in Gagnon-Arsenault et al., 2006). This loop region contains at least

one dibasic residue pair for further cleavage to generate two-subunit enzyme (Cawley et

al., 1998). The α- and β- subunits in Yps1 are linked to each other by a disulphide bridge

and contribute one catalytic aspartate residue each, which flank the exposed loop region

(Cawley et al., 1998; reviewed in Gagnon-Arsenault et al., 2006). Another characteristic

feature of yapsins is the presence of serine-threonine rich region, C-terminal to the

catalytic domain, which is thought to be heavily glycosylated (Ash et al., 1995).

Consistent with this, ScYps1, ScYps2 and ScYps3 have 10, 9 and 11 putative N-

glycosylation sites, respectively (Ash et al., 1995; Komano and Fuller, 1995). Finally,

the most important and unique characteristic of fungal yapsins is the presence of a C-

terminal sequence designated for the attachment of a GPI anchor (Komano and Fuller,

1995; Krysan et al., 2005 and shown in Figure 1.5).


26

Figure 1.5: Pictorial representation of the domain organization of pro-zymogen form of S. cerevisiae Yps1 with pro-peptide fragment of 67 amino acids. Pro-peptide is cleaved during maturation. Mature form of Yps1 enzyme has catalytic aspartate residues at 34 and 304 positions and the Cys 50, Cys 119 and Cys 339, Cys 371, correspond to Cys 117, Cys 186 and Cys 406, Cys 438 in the pro-zymogen Yps1, respectively. Asn 548 (Asn 481 in mature enzyme) at C-terminal denotes the GPI-anchor attachment site in Yps1. Figure is not drawn to scale.

A recent study has shown a pH-dependent proteolytic processing of S. cerevisiae

Yps1 protease, which contributes to the maturation, activation and shedding of the

enzyme from the cell surface and links the yapsin activity with the pH-dependent

reorganization of the yeast cell wall (Gagnon-Arsenault et al., 2008). GPI-Yps1 was

found to be active in vivo at pH 3.0 and pH 6.0. While the differential processing of

Yps1 at pH 3.0 was mostly autocatalytic, the Yps1 enzyme processing required

assistance of the other proteases at pH 6.0 (Gagnon-Arsenault et al., 2008). In addition,

autocatalytic processing of the loop region, to generate the mature two-subunit ScYps1

enzyme, was modulated by external pH via selection of the alternate cleavage sites

(Gagnon-Arsenault et al., 2008).

1.3.2.2.1 Regulated expression of YPS genes

Similar to SAPs, expression of yapsin encoding genes is regulated in response to

several environmental cues. S. cerevisiae YPS1 and YPS3 transcript levels are up

regulated upon exposure to cell wall stress agents including congo red and zymolyase

(Garcia et al., 2004; Gagnon-Arsenault et al., 2006). YPS1 expression is also known to


27

induce in response to thermal stress (Krysan et al., 2005). Notably, S. cerevisiae YPS1

gene contains a putative CDRE (calcineurin-dependent response element) motif

GTGGCTT at -304 to -298 position and its expression is under the regulation of the

calcineurin-dependent transcription factor Crz1 (Krysan et al., 2005). Notably, 1.5- to 8-

fold increased expression of CgYPS2, CgYPS4-5 and CgYPS8-11 was observed in

response to the macrophage internal milieu (Kaur et al., 2007). Additionally, transcript

levels of CgYPS1 were elevated during thermal stress and this heat-induced expression

of CgYPS1 was primarily dependent upon calcineurin-Crz1 pathway and partially on

Slt2 MAPK (mitogen-activated protein kinase) pathway (Miyazaki et al., 2011).

1.3.2.2.2 Role for yapsins in fungal physiology and virulence

(i) Role for S. cerevisiae yapsins

The fungal cell wall is a dynamic structure which undergoes continuous

remodelling to adapt to the environmental conditions. It is mainly a multilayer

meshwork of three essential components, viz., β-glucan, mannoproteins and chitin. β-

Glucan (comprised of β-1,3- and β-1,6-glucans) and the complex network of cell wall

mannoproteins are the major cell wall constituents and represent 40-50% and 30-50% of

total cell wall dry mass, respectively (Lipke and Ovalle, 1998). Chitin, a minor cell wall

component, accounts for 1-2% of total cell wall dry weight (Lipke and Ovalle, 1998).

During cell wall stress, genes associated with all the three components of cell wall show

differential expression to accommodate the changes (Lesage and Bussey, 2006). Some of

the well-studied examples of these genes in S. cerevisiae include genes involved in β-

glucan synthesis and modification (FKS2, GAS1 and BGL2), chitin synthesis (CHS3) and

a variety of cell wall mannoproteins encoding genes (CWP1, CCW14, ECM11

and PRH1) (Gagnon-Arsenault et al., 2006; Lesage and Bussey, 2006).

Localization of yapsins to the cell surface in S. cerevisiae suggested that these

proteases may play a role in shedding and processing of the cell surface proteins

(Komano and Fuller, 1995; Ash et al., 1995; Gagnon-Arsenault et al., 2006). In addition,

the presence of a dibasic residue just N-terminal to the ω-site for GPI-attachment in

several plasma membrane and cell wall-associated proteins provide support to the above

notion (Caro et al., 1997; Schild et al., 2011) Consistent with this, Yps1 appears to have

overlapping functions in the processing of cell wall-related enzymes with Kex2 which is

a subtilisin-related serine protease and has been implicated in the proteolytic processing

of Exg1 (β-glucanase) (Komano and Fuller, 1995; Larriba et al., 1995; Bader et al.,


28

2001; Gagnon-Arsenault et al., 2006, 2008) Recently, Yps1 has been shown to cleave

the extracellular inhibitory domain of mucin Msb2 resulting in the generation of active

signalling mucin which further activates the Cdc42-dependent MAPK pathway in

response to nutritional cues (Vadaie et al., 2008).

In S. cerevisiae, yapsins generally function at an optimum pH of 3.5 to 4.5 and

are stable between pH 2.0 to pH 7.0 (Gagnon-Arsenault et al., 2006). S. cerevisiae Yps1

has been reported to be enzymatically active at pH 4.0 to 4.5 and could cleave the paired

basic residues (Arg-Arg, Arg-Lys or Lys-Lys) of the mammalian

adrenocorticotropin/endorphin prohormone, pro-opiomelanocortin, anglerfish pro-

somatostatin I and II and pro-insulin in vitro (Azaryan et al., 1993; Zhang et al., 1997;

Gagnon-Arsenault et al., 2008; Table 1.2). Additionally, Yps1 has been reported to

function as a shedder for a subset of GPI-anchored enzymes, including itself and the

Gas1 glucanosyltransferase (Gagnon-Arsenault et al., 2008; Table 1.2). Recently, Yps1

has also been implicated in the processing of human proglucagon into glucagon in vitro

(Cawley et al., 2011; Table 1.2). Similar to Yps1, Yps2 has also been found to have pH

optima of 4.0 (Komano et al., 1998). Further, Yps2 (Mkc7) and Yps3 (Yap3) have been

shown to cleave human Alzheimer β-amyloid precursor protein under in vitro conditions

(Komano et al., 1998; Table 1.2).


29

Table 1.2: Known and putative substrates of aspartyl proteases in S. cerevisiae, C. albicans and C. glabrata.

Aspartyl protease Substrate in yeast Substrate in mammals Function/role of the substrate

Saccharomyces cerevisiae

Yps1

pro-alpha mating factor Yeast mating pheromone

Mucin; Msb2 Upstream protein in Cdc42

dependent-MAPK signalling, osmosensor

Vacuolar carboxypeptidase Y

Prc1

An exopeptidase involved in non-specific protein degradation in the

vacuole Aspartyl protease;

Yps1 Cell surface-associated GPI-linked aspartyl protease

Glucanosyl-transferase; Gas1

Cell wall protein involved in cell wall organization, and transcriptional

silencing Cell wall protein; Pir4 Glucan cross linking

Anglerfish pro-somatostatin I

and II Precursor of neuro-peptide hormones

Human 7B2 Regulate secretion

Pro-insulin

Precursor of insulin, fat and carbohydrate metabolism

Dynorphin A

Precursor of endogenous opoid peptide

Dynorphin B

Precursor of endogenous opoid peptide

Amidorphin Opoid peptide

Adrenocorticotropic hormone

Synthesis and secretion of androgenic steroids

Human pro-elafin; Trappin-2

Precursor of elafin, an elastase-

specific inhibitor

β-amyloid peptide

Human β-amyloid precursor protein; (APP)

Precursor of β-amyloid,

Synapse formation

Cholecystokinin 33

Role in digestion

Human Pro-glucagon Precursor of glucagon

Yps2

β-Amyloid peptide

Human β-amyloid

precursor protein (APP)

Precursor of β-amyloid

Synapse formation

Yps3

β-amyloid peptide Precursor of β-amyloid Cholecystokinin 33 Role in digestion

β-endorphin Endogenous opoid peptide neurotransmitter

Candida albicans

Sap1, Sap2, Sap3

Insulin B-chain, Structural component of insulin Bovine hemoglobin Oxygen carrier protein in RBCs

Complement C3b, C4b

and C5 Casein

Complement factors

Milk protein


30

Sap2

α2-macroglobulin Immune effectors Collagen Extracellular matrix protein Vimentin Cell organelles anchoring Cystatin A cysteine protease inhibitor Mucin A glycoconjugate Immunoglobulin-A Immune effectors Keratin Structural protein of mammalian skin Histatin-5 Salivary anti-microbial peptide

Sap4 Casein Milk protein

Sap5 Casein, E-cadherin Milk protein, Adhesive protein

Sap6 Casein, Hemoglobin Milk protein, oxygen carrier protein in RBCs

Sap9 Casein Milk protein

Sap9 and Sap10

Histatin-5 Salivary anti-microbial peptide Chitinase; Cht2 Chitin synthesis

Cell wall protein; Pir1 Glucan cross linking Cell wall protein;

Ywp1 Cell dispersal

Adhesin; Als2 and Als10 Cell adhesion

Cell wall protein; Rhd3 Unknown function

Cell wall protein; Rbt5 Hemoglobin utilization Cell wall protein;

Ecm33 Cell wall integrity

Cell wall protein; Pga4 β-1,3-glucanosyltransferase

Candida glabrataYps1 and Yps7 Adhesin; Epa1 Cell adhesion Yps1 to Yps11 Histatin-5 Salivary anti-microbial peptide

Further, various yapsin deleted strains in S. cerevisiae showed differential

susceptibility to cell wall damaging agents; yps1∆ mutant was hyper-sensitive to

caspofungin (a β-1,3-glucan synthase inhibitor) and caffeine (Krysan et al., 2005;

Gagnon-Arsenault et al., 2006). A S. cerevisiae strain deleted for all five YPS genes

(YPS1-3,6,7) displayed an enhanced propensity for cell lysis at 37˚C, reduced amount of

β-glucan in the cell wall and hypersensitivity to cell wall damaging agents (Krysan et al.,

2005; Gagnon-Arsenault et al., 2006). Activity of β-glucan synthetases in yps1-3∆6∆7∆

remained unperturbed indicating a possible role for yapsins either in the incorporation or

retention of β-glucan in the cell wall (Krysan et al., 2005; Gagnon-Arsenault et al.,

2006). In agreement with this, over-expression of YPS1 partially suppressed the

caspofungin sensitivity of wild type cells suggesting a role for Yps1 in the stabilization

of the cell wall glucan. YPS1 expression during cell wall stress was dually regulated by

classical protein kinase C (Pkc1)-MAPK signalling and Crz1-regulated calcineurin-

mediated signalling (Gagnon-Arsenault et al., 2006). Expression of YPS1 mirrored that

of the stress-induced FKS2 (encode subunit of β-glucan synthase) implying coordinated


31

functions of Yps1 and Fks1 in maintaining the β-glucan core of the cell wall during cell

wall stress (Gagnon-Arsenault et al., 2006).

(ii) Role for C. glabrata yapsins

Similar to S. cerevisiae, C. glabrata yapsins have also been implicated in

maintenance of cell wall integrity and survival of thermal stress (Kaur et al., 2007;

Miyazaki et al., 2011). Cgyps1∆ mutant showed highly attenuated growth at 41˚C and

this growth defect could easily be rescued by addition of an osmotic stabilizer, sorbitol

(Miyazaki et al., 2011). However, the sensitivity of Cgyps1∆ to thermal stress appears to

be a strain-dependent phenotype (Kaur et al., 2007; Miyazaki et al., 2011). In addition,

essential role for CgYps1 and CgYps7 in survival of stationary phase stress has also

been reported (Kaur et al., 2007). Disruption of CgYPS1 and CgYPS7 genes individually

rendered cells sensitive to salt stress and caffeine and calcofluor white, caffeine and

congo red, respectively (Kaur et al., 2007). Further, deletion of either YPS1 or YPS7

singly or in combination led to an enhanced resistance to zymolyase which digests β-

glucan (Kaur et al., 2007). Phenotypes of C. glabrata mutants lacking yapsins are

summarized in Table 1.3.

Moreover, yapsins in C. glabrata play an essential role in cell wall remodelling

by removal and release of the major GPI-anchored cell wall adhesin, Epa1 (Kaur et al.,

2007). Fluorescence-activated cell sorting (FACS) and Western blot analysis on Cgyps1-

11∆ mutant cells revealed Epa1 to be stabilized at the cell surface with mutant cells

displaying hyper adherence to Lec2 epithelial cells (Kaur et al., 2007). Further,

significantly reduced amounts of proteolyzed fragments of Epa1 were released into the

culture medium in Cgyps1-11∆ strain compared to the wild type strain indicative of a

defective Epa1 processing from the cell surface (Kaur et al., 2007).

Importantly, deletion of CgYPS1 and CgYPS7 in combination or deletion of all

eleven YPS genes led to reduced survival in murine macrophages by 2- and 33-fold,

respectively, compared to wild-type (wt) strain, and, simultaneously, resulted in

activation of macrophages leading to increased nitrite production (Kaur et al., 2007).

Notably, similar to C. albicans, disruption of a single yapsin encoding gene did not

significantly attenuate virulence (Naglik et al., 2003; Kaur et al., 2007) indicating

functional redundancy and/or synergistic effects of C. glabrata yapsins during systemic

infections of mice.

Collectively, these studies indicate a central role for GPI-linked aspartyl

proteases in maintenance of cell wall structure under stressful conditions, however, the

Chapter 1 Introduction

32

Table 1.3: Phenotypic summary of C. glabrata mutants lacking yapsins.

Mutant genotype

Environmental conditions Adherence to

Lec2 epithelial

cells

compared to

wild-type

Replication in

murine

macrophages

(J774A.1)

compared to

wild-type

Survival in

murine model of

disseminated

candidiasis

compared to

wild-type

CW CR Caffeine NaCl Temperature

(42˚C)

Zymolyase

treatment

β-glucan content in

YPD-grown cells

Cgyps1∆ W W S S W R Decreased Increased Decreased Decreased

Cgyps2∆ W W W W W W ND ND ND ND

Cgyps7∆ S S W W W R ND Increased No change No change

Cgyps1∆yps7∆ S S S S W R ND Increased Decreased Decreased

CgypsC∆

(Cgyps3-6∆, yps8-

11∆)

W W W W W W ND Increased No change No change

Cgyps1∆C∆ W W S S W R ND Increased Decreased Decreased

Cgyps2∆C∆ W W W W W W ND ND ND ND

Cgyps7∆C∆ S S W W W R ND ND Decreased Decreased

Cgyps1-11∆ S S S S W R Decreased Increased Decreased Decreased

CW = Calcofluor white, CR = Congo red, R = Resistant, S = Sensitive, ND = Not determined, W = Wild-type phenotype


33

precise mechanism, by which these proteases regulate cell wall remodelling,

remains to be identified. The reduced proteolytic cleavage of Epa1 in C. glabrata

Cgyps1-11∆ mutant (Kaur et al., 2007), identification of Gas1, Yps1 and Pir4 as

endogenous substrates for S. cerevisiae Yps1 (Gagnon-Arsenault et al., 2008) and the

recent illustration of an in vitro cleavage of covalently-linked cell wall proteins including

Cht2 and Pir1 by C. albicans Sap9 and Sap10 (Schild et al., 2011) suggest recycling

and/or processing of the cell wall mannoprotein layer as the major cell wall organization

mechanism.

1.3.2.3. Vacuolar aspartyl proteases

The yeast vacuole is a fluid-filled, single membrane-bound dynamic organelle

which acts as a storage compartment for amino acids and polyphosphates and plays an

important role in ion and pH homeostasis, osmoregulation, removal of toxic substances

and recycling of macronutrients (Li and Kane, 2009; Armstrong, 2010). The major yeast

vacuole processing protease, proteinase A (PrA), is an aspartic peptidase and is encoded

by the PEP4 gene in S. cerevisiae (Woolford et al., 1986). Mutants disrupted for PEP4

accumulate multiple vacuolar zymogen, indicative of Pep4’s pivotal role in processing

and activation of vacuolar hydrolases (Zubenko et al., 1983). Pep4 is required for

mitochondrial degradation during acetic acid-induced apoptosis, chronological aging,

sporulation, cellular response to starvation and microautophagy (Teichert et al., 1989;

Palmer et al., 2007; Pereira et al., 2010). In C. glabrata, the ORF CAGL0M02211g

codes for CgPep4 (Figure 1.4) which shows 63% identity with S. cerevisiae Pep4.

However, contrary to PEP4, CgPEP4 expression is known to be induced in response to

osmotic stress, pH stress and glucose starvation (Gasch et al., 2000; Roetzer et al.,

2008). Recently, CgPEP4 expression was also found to be up-regulated in the biofilm

mode of growth (Seneviratne et al., 2010). In C. albicans, the orf19.1891, expressed in

both yeast and hyphal forms, encodes Apr1, a Pep4 homolog (Figure 1.4), and is

predicted to play a central role in survival under starvation conditions (Niimi et al.,

1997; Kusch et al., 2008).

Section 1.4: Yeast cell architecture and physiology

1.4.1 Yeast cell wall

The fungal cell wall is a complex, dynamic and protective three-dimensional

multilayer meshwork of polysaccharide and glycoproteins which together provides

structural integrity to the cell (Lipke and Ovalle, 1998). Despite being a rigid structure,


34

architecture of the fungal cell wall continuously varies during different growth stages

and environmental conditions (Levin, 2011). In accord, biogenesis, assembly and

composition of the cell wall are influenced by cell cycle progression, growth and

different stress conditions (Lipke and Ovalle, 1998; Levin, 2011). Four major functions

of yeast cell wall include maintenance of osmotic balance via regulating the passage of

the water through the cell, cell shape maintenance for proper cell division and bud

formation, protection against mechanical stress and serving as scaffold for cell surface

proteins (Lipke and Ovalle, 1998; Levin, 2011). The polysaccharide component of the

cell wall provides an attachment site for heavily N- and O-glycosylated proteins which

help in restricting the permeability of the cell wall to foreign macromolecules (Klis et

al., 2002; Levin, 2011).

The fungal cell wall constitutes about 10-25% of total cell biomass (Klis et al.,

2002). Electron microscopy analysis has revealed the bilayer architecture of cell wall in

S. cerevisiae, C. albicans and C. glabrata wherein inner and outer layer are mostly

electron-transparent and electron-dense, respectively (Klis et al., 2002; de Groot et al.,

2008). General constituents of the cell wall in S. cerevisiae, C. albicans and C. glabrata

are the same and consist of β-1,3-glucan, β-1,6-glucan, chitin and mannoproteins (Klis et

al., 2002; Lesage and Bussey, 2006). The electron-transparent inner-most layer of the

cell wall consists of chitin and glucan polymers. Chitin is a linear polymer of β-1,4-N-

acetyl glucosamine synthesised by chitin synthases and mostly concentrated in bud neck,

septum, and bud scar areas (Lesage and Bussey, 2006). Total levels of chitin vary in

different fungal species with chitin constituting only 1.1-1.3% of total cell wall dry mass

in C. glabrata during logarithmic growth conditions (de Groot et al., 2008).

Polysaccharide β-glucan forms the main structural constituent of the cell wall and

is divided into two different classes based on the type of the glycosidic bond between

two carbon moieties in monosaccharide units of the polysaccharide. These classes are β-

1,3-glucan (85% of total β-glucan) and β-1,6-glucan (15% of total β-glucan) (Klis et al.,

2006). A multi-layer meshwork of β-1,3-glucan (~ 1500 glucose residues) is branched

through short polymers of β-1,6 glucan (~ 150 glucose residues) via covalent bonds and

provides mechanical and tensile strength to the cell wall (Lipke and Ovale, 1998; Klis et

al., 2006). A recent study by de Groot et al. showed that S. cerevisiae and C. albicans

cell wall contains 26-27% of alkali-resistant β-1,3-glucan. Levels of alkali-insoluble β-

1,6-glucan were found to be 7 and 11% in S. cerevisiae and C. albicans cell wall,

respectively (de Groot et al., 2008). Interestingly, de Groot et al. also reported that


35

compared to S. cerevisiae and C. albicans, cell wall of C. glabrata had significantly

lesser amount of alkali-resistant β-1,3-glucan (17% of dried-cell wall mass) and β-1,6-

glucan (4% of dried-cell wall mass) suggesting fewer cross-link between glucan and

chitin in C. glabrata cell wall (de Groot et al., 2008).

The outer electron-dense layer of the cell wall consists of a fibrillar network of

cell wall mannoproteins (Klis et al., 2006). Both covalently- and non-covalently-bound

mannoproteins are embedded in the glucan network and form 30-50% of the cell wall

mass. Cell wall proteins (CWPs) are glycosylated in endoplasmic reticulum and/or Golgi

apparatus during transit to the cell wall (Klis et al., 2002). Three major groups of

covalently bound CWPs present in S. cerevisiae, C. albicans and C. glabrata are GPI-

anchored proteins, Pir-proteins (proteins with internal repeats) and proteins linked to

other proteins by disulfide bridges (Klis et al., 2006). The outer mannoprotein layer of

the cell wall not only acts as a barrier to foreign macromolecules but also plays

important roles in the regulation of cell surface hydrophobicity, adhesion and

antigenicity of fungal cell (Albrecht et al., 2006; Kaur et al., 2007; de Groot et al., 2008;

Netea et al., 2008).

Intriguingly, the cell wall in C. glabrata contains significantly higher mannose to

glucose ratio and 50% more proteins compared to the S. cerevisiae wall (de Groot et al.,

2008). In silico analysis has identified 106 putative GPI-anchored proteins of different

functional groups including 51 adhesin-like proteins, 17 enzymatic proteins (proteases

and lipases) and several other proteins of unknown function in C. glabrata (Weig et al.,

2004). Consistent with this, de Groot et al. predicted a total of 67 putative adhesin-like

GPI-proteins and identified 18 CWPs using mass spectrometry in C. glabrata (de Groot

et al., 2008).

Cell wall remodelling in response to environmental cues in S. cerevisiae is

regulated by a conserved PKC-mediated cell wall integrity signalling pathway which is

comprised of a linear array of MAPKs, Pkc1-Bck1-Mkk1/Mkk2-Slt2 (Levin, 2011).

Phosphorylation of the terminal MAPK, Slt2, leads to its translocation to the nucleus and

subsequent activation of transcription factors Swi4 and Rlm1, which stimulate

transcription of genes implicated in cell cycle and cell wall metabolism, respectively

(Levin, 2011). In Pkc1-mediated signalling, cell wall stress signal is first transmitted

through a family of cell surface sensors to the small Rho1 GTPase, which activates

several effector molecules including Pkc1 and results in the synthesis and the proper

delivery of β-glucan for appropriate remodelling of the cell wall (Levin, 2011).


36

1.4.2 Yeast stress responses

All types of cells regularly encounter varied environmental conditions which can

adversely impact cell growth. Ability to respond to such changes in the extracellular

environment (temperature, pH, presence of toxic chemicals and nutrient availability)

requires a coordinated cascade to sense the change, transduce the signal and mount a

response via appropriate reprogramming of transcriptional and metabolic pathways

(Bahn, 2008; Selvig and Alspaugh, 2011). For yeast cells, extracellular pH is an

important environmental signal that regulates cell growth, physiology, metabolism and

differentiation (Selvig and Alspaugh, 2011). Yeast cells grow more rapidly in acidic

medium than in neutral or alkaline medium (Peñalva and Arst, 2004). Non-pathogenic

yeast S. cerevisiae and several pathogenic fungi including C. albicans, A. nidulans and

C. neoformans possess efficient adaptation mechanisms to survive broad pH alterations

particularly in an alkaline pH environment (Peñalva and Arst, 2004; Bahn, 2008; Selvig

and Alspaugh, 2011). This part of the chapter describes fungal external pH adaptation

responses.

1.4.2.1 Environmental stress response

S. cerevisiae displays an extensive array of transcriptional responses to different

environmental stress conditions including heat shock, pH, oxidative, nutrient and

osmotic stresses (Gasch et al., 2000; Causton et al., 2001). Overall response of the cell

toward these environmental stress signals is referred as environmental stress response

(ESR) or common environmental response (CER) (Gasch et al., 2000; Causton et al.,

2001). Genes which are differentially expressed during ESR constitute about 14% of

total predicted genes in S. cerevisiae genome (Gasch et al., 2000). Major stress-induced

transcription factors regulating ESR in yeast are Hsf1 (heat shock), Skn7 and Yap1

(oxidative stress), Hog1 (osmotic) and Msn2 and Msn4 (general stress response). ESR

response has also been identified in other fungi including C. albicans, C. glabrata and

fission yeast Schizosaccharomyces pombe (Chen et al., 2003; Smith et al., 2004;

Enjalbert et al., 2006; Roetzer et al., 2008).

Genome expression profiling analyses in S. cerevisiae and C. glabrata revealed

that ~ 90% of up-regulated genes during ESR are targets of highly conserved Cys2His2

zinc-finger transcriptional factors, Msn2 and Msn4 (Gasch et al., 2000; Causton et al.,

2001; Roetzer et al., 2008). These ESR-induced genes are involved in carbohydrate

metabolism, heat shock response, protein degradation, vacuolar functions, DNA damage


37

repair, ion homeostasis and anti-oxidant mechanisms and signalling (Gasch et al., 2000;

Causton et al., 2001; Roetzer et al., 2008). Genes involved in RNA metabolism,

nucleotide biosynthesis, ribosomal biogenesis, rRNA processing, and translation

initiation constituted the repressed gene set in ESR (Gasch et al., 2000; Causton et al.,

2001). Exclusive gene targets of CgMsn2 and CgMsn4 in C. glabrata include MDH3

(encodes glyoxylate cycle enzyme), FBP26 (encodes glycolytic enzyme), PHM8

(encodes putative lysophosphatidic acid phosphatise, involved in phosphate metabolism)

and YCK1 (casein kinase, involved in cell morphogenesis) (Roetzer et al., 2008).

In contrast, ESR in C. albicans and S. pombe are mainly regulated by Hog1, Sty1

and Atf1 mediated stress-activated MAPK pathway and not by Msn2 and Msn4

(Enjalbert et al., 2006; Chen et al., 2003). Despite this difference, substantial overlap has

been observed in the ESRs of C. albicans, S. pombe and S. cerevisiae (Gasch et al.,

2000; Causton et al., 2001; Chen et al., 2003; Enjalbert et al., 2006).

1.4.2.1.1 Msn2 and Msn4 transcriptional factors

Msn2 and Msn4 constitute major regulators of ESR gene expression system in S.

cerevisiae and C. glabrata (Gasch et al., 2000; Causton et al., 2001; Roetzer et al.,

2008). MSN2 and MSN4 genes were originally identified as multi-copy suppressors of

SNF1 protein kinase-defective, temperature sensitive mutant of S. cerevisiae (Martinez-

Pastor et al., 1996; Schmitt et al., 1996). Msn2 and Msn4 are functionally redundant

transcriptional factors containing two Cys2His2 zinc-fingers at C-terminus. S. cerevisiae

Msn2 and Msn4 show 32% identity among themselves and share 28 and 27% identity

with their othrologs in C. glabrata, respectively. Both Msn2 and Msn4 bind to a

consensus DNA sequence, known as stress responsive element (STRE), in promoters of

their target genes. In accord, designated STRE sequence, AGGGG/CCCCT, is present in

promoter regions of several genes induced during stressful environmental conditions

(Gasch et al., 2000; Causton et al., 2001; Roetzer et al., 2008). Deletion of Msn2 and

Msn4 in both S. cerevisiae as well as in C. glabrata rendered cells hypersensitive to a

variety of stresses including temperature, osmotic and oxidative stress (Gasch et al.,

2000; Roetzer et al., 2008).

1.4.2.1.2 Regulation of Msn2 and Msn4 activity

Activity of Msn2 and Msn4 transcriptional factors has primarily been studied in

S. cerevisiae and is known to be dependent on their localization in the cell (Gorner et al.,

1998). Under normal growth conditions, Msn2 and Msn4 reside in cytoplasm in

deactivated forms. However, these transcription factors, in response to stress, are


38

activated and rapidly localize to the nucleus (Gorner et al., 1998). Both Msn2 and Msn4

contain nuclear localization signal (NLS) adjacent to the zinc-finger domain (Gorner et

al., 1998). Upon activation, Msn2 and Msn4 show repetitive shuttling between nucleus

and cytoplasm and have been shown to require an exportin, Msn5, for their nuclear

export (Gorner et al., 2002).

NLS of Msn2 and Msn4 is negatively controlled by cAMP-dependent protein

kinase A (PKA) signalling pathway which prevents localization of these transcription

factors to the nucleus. PKA was earlier thought to directly phosphorylate Msn2 and

Msn4 to initiate their cytoplasmic relocalization as several putative PKA-

phosphorylation sites were identified in Msn2 and Msn4 proteins (Gorner et al., 1998).

However, Msn2 and Msn4 were found to exist in phosphorylated states under normal

growth conditions and hyper-phosphorylated forms upon stress exposure (Garreau et al.,

2000). During PKA signalling activation, these hyper-phosphorylated stages were

reversed resulting in Msn2 and Msn4 re-localization to the cytoplasm (Gorner et al.,

2002). Notably, both, genes coding for two of the three PKA catalytic subunits, TPK1

and TPK2, as well as, negative regulators of PKA signalling, BCY1, PDE1 and YAK1,

showed elevated expression during ESR.

Another pathway that regulates Msn2 and Msn4 activation is TOR (target of

rapamycin) signalling which helps in sequestration of these transcription factors in the

cytosol by 14-3-3 adaptor protein, Bmh2 (Beck and Hall, 1999). In the presence of stress

or rapamycin-mediated inhibition of TOR signalling, association between Msn2 and

Bmh2 is lost leading to Msn2 localization to the nucleus (Beck and Hall, 1999).

Besides signal transduction pathways, degradation of Msn2 is also known to

regulate its activity. Stress-dependent proteosomal degradation of Msn2, which is

mediated by cyclin-dependent protein kinase Srb10, has been observed in the nucleus

(Lallet et al., 2004). During heat stress, Srb10 is involved in the hyper-phosphorylation

of Msn2 causing its localization to the nucleus and further ubiquitination by SCF (Skp,

Cullin, F-box containing complex) E3 ubiquitin ligase complex (Lallet et al., 2004).

Deletion of UMP1 which encodes proteosome maturation factor, increases Msn2 and

Msn4 activity corroborating the notion that proteosome degradation system regulates

Msn2 and Msn4 activity (Sadeh et al., 2011).


39

1.4.2.1.3 Targets of Msn2 and Msn4

Deletion of MSN2 and MSN4 in S. cerevisiae affected transcriptional activation

of ∼ 60% genes which were up-regulated during heat shock or oxidative stress,

indicating a substantial contribution of Msn2 and Msn4 to the ESR response (Gasch et

al., 2000). Similarly, > 90% of Msn2- and Msn4-regulated genes showed higher

expression when both MSN2 and MSN4 were over expressed (Gasch et al., 2000). Over

expression of MSN2 and MSN4 additionally induced ~ 80 new ESR genes whose

expression, upon heat and oxidative stress, remained unaffected by deletion of these

factors (Gasch et al., 2000). Msn2 and Msn4 transcription factors are also involved in

the induction of about 180 genes during hydrogen peroxide (H2O2) stress and the

msn2∆msn4∆ mutant is hypersensitive to H2O2 (Martinez-Pastor et al., 1996; Gasch et

al., 2000). Msn2 and Msn4 are also important for regulation of genes involved in

synthesis and degradation of disaccharide sugar trehalose (Zahringer et al., 2000).

Notably, trehalose is known to act as a thermo-protectant and deletion of TPS1 rendered

cells highly sensitive to thermal stress (De Virgilio et al., 1994). Further, trehalose is

postulated to serve as a chemical chaperone for protein folding during high temperature

stress (Simola et al., 2000).

1.4.2.2 Transcriptional responses to environmental pH changes

Transcriptional responses to external pH changes, especially shift towards

alkaline pH, have been well-characterized in S. cerevisiae, A. nidulans and C. albicans.

Alkaline pH is known to induce transcription of genes involved in ion transport,

metabolism and stress responses in S. cerevisiae (Viladeval et al., 2004). External pH

serves as a regulator of differentiation and development in C. albicans (Calderone and

Fonzi, 2001). While acidic external pH favours growth in the yeast form, alkaline

conditions induce the hyphal growth. Importantly, the yeast-to-hyphal transformation

has been shown to be essential for C. albicans virulence (Calderone and Fonzi, 2001).

1.4.2.2.1 Alkaline pH response in A. nidulans and S. cerevisiae

Major effectors of alkaline pH response, Rim101 in S. cerevisiae, and PacC in A.

nidulans, are Cys2His2 zinc-finger transcription factors, which bind to the consensus

DNA-binding sites, TGCCAAG and TGCCARG, respectively, in the promoter of their

target genes (reviewed in Selvig and Alspaugh, 2011). Both Rim101 and PacC are

activated in response to neutral and alkaline environment via proteolytic cleavage of an

inhibitory C-terminal domain. The Rim101/PacC pathway constitutes a highly conserved


40

cascade of (i) signalling complexes present on the plasma membrane, (ii) adaptor

molecules present in the cytosol and (iii) proteases present on the endosomal

membranes, where Rim101 and PacC processing and activation occur (Reviewed in

Penalva et al., 2008). Unlike PacC of A. nidulans, Rim101 in S. cerevisiae is also

proteolytically processed in acidic growth conditions (Li et al., 1997).

During pH signalling, environmental pH is sensed by a plasma membrane-bound

seven transmembrane-domain receptor, Rim21/PalH. The Rim21/PalH is subsequently

internalized in the cytosol by an arrestin, Rim8/PalF, and transported to endosomal

membranes via endocytic transport machinery composed of ESCRT (endosomal sorting

complex required for transport) complex. In neutral and alkaline pH, PalF is

phosphorylated and ubiquitinated resulting in its internalization in the cytosol by

ESCRT-I complex (Herranz et al., 2005). Unlike PalF, internalization of its ortholog,

Rim8 in S. cerevisiae is independent of its ubiquitinylation status (Herrador et al., 2010).

After internalization, the ESCRT-I complex recruits the ESCRT-III a heterodimeric

complex of Vps20 and Snf7 subunits via ESCRT-II complex for complete endocytosis

(Kullas et al., 2004). Once endocytosed, the Rim21-Rim8/PalH-PalF complex interacts

with downstream proteins, Rim20/PalA and calpain-like protease Rim13/PalB (Galindo

et al., 2007). Finally, the Rim21-Rim8-Rim20/PalH-PalF-PalA trimeric-complex

interacts with the C-terminus of Rim101/PacC and Rim13/PalB, leading to the

Rim13/PalB-mediated proteolysis and activation of Rim101/PacC transcription factor on

endosomal membrane. Activated Rim101 or PacC relocate to the nucleus and regulate

the expression of their target genes.

Activated PacC in A. nidulans, regulates expression of both acid- and alkaline-

responsive genes by repressing the former and inducing the latter (Arst and Penalva,

2003). PacC pathway in A. nidulans also plays important role in virulence of this

pathogenic fungus (Hua et al., 2010).

In S. cerevisiae, Rim101 primarily acts as a repressor and downregulates the

expression of NRG1 (negative regulator of glucose-repressed genes). The transcriptional

repressor, Nrg1, is known to suppress the expression of ion transporter Ena1, which is

pivotal to ion homeostasis during elevated pH conditions (Lamb et al., 2001; Lamb and

Mitchell, 2003). However, as an inducer, Rim101 positively regulates the expression of

VMA4, which encodes a vacuolar H+-ATPase subunit and is required for growth under

alkaline pH conditions (Lamb et al., 2001). Other than NRG1 and VMA4, expression of

ARN4 (encodes a bacterial siderophore-iron-transpoter), FET4 (encodes low-affinity


41

Fe(II) transporter) and NRG2 (encodes a transcriptional repressor similar to Nrg1) is also

regulated by the Rim101 pathway (Lamb et al., 2001; Lamb and Mitchell, 2003).

Additionally, Rim101 is involved in invasive growth, sporulation and ion homeostasis in

S. cerevisiae (Lamb et al., 2001; Lamb and Mitchell, 2003).

1.4.2.2.2 Alkaline pH response in C. albicans

In C. albicans, the Rim101 pathway regulates several alkaline pH responses

including induction of alkaline pH-responsive genes, PRA1 and PHR1, and repression of

acidic responsive gene, PHR2, under alkaline pH growth conditions and transition from

yeast-to-hyphal stage (Davis, 2009). Notably, PRA1 and PHR1 encode cell surface

proteins and are involved in filamentation and cell growth at alkaline pH (reviewed in

Selvig and Alspaugh, 2011). Most of the signalling components of Rim101 pathway in

C. albicans are conserved with those in S. cerevisiae including homologs of Rim21,

Rim8, Rim20, Rim13 and Rim9.

A genome-wide expression profiling analysis on C. albicans cells, grown at

acidic pH 4.0 and alkaline pH 8.0, identified 514 pH-responsive genes (Bensen et al.,

2004). Of 514 genes, Rim101 was found to regulate expression of ∼ 118 genes (23%).

Genes induced at pH 8.0 were either involved in iron acquisition or encoded hyphal-

specific proteins. While expression of iron acquisition genes was mostly independent of

Rim101, hyphal-specific genes showed a significant Rim101-dependency but for the

SAP4 and the SAP6 genes. Overall, genes implicated in several processes, viz.,

carbohydrate, amino acid and lipid metabolism, signal transduction, electron and ion

transport, cell wall integrity, hyphal development and protein synthesis, folding and

degradation, were differentially regulated in response to ambient pH. Although Rim101

does not play any prominent role in regulating the transcriptional responses at acidic pH

in C. albicans (Bensen et al., 2004), it is still essential for virulence in the murine model

of systemic candidiasis (Davis et al., 2000). C. albicans Rim101 is also involved in

tissue invasion via positive regulation of the gene encoding secreted aspartyl protease,

Sap5 (Villar et al., 2007). Further, Rim101-dependent expression of Sap5 mediates

proteolytic degradation of the adhesin molecule, E-cadherin, during interaction of C.

albicans with oral epithelial cells (Villar et al., 2007). C. glabrata possesses orthologs of

S. cerevisiae Rim101, Rim21 and Rim8 (http://www.candidagenome.org), however,

their role/s in pH adaptation and virulence remain to be investigated.


42

Other signalling pathways involved in fungal alkaline pH response are

calcineurin-mediated signalling and MAPK signalling. Pkc1-mediated cascade in S.

cerevisiae upregulated expression of genes involved in cell wall biogenesis, metabolism

and transport, during pH adaptation response, in an alkaline stress-responsive Slt2

kinase- and Wsc1 membrane sensor-dependent manner (Serrano et al., 2006). Deletion

of several components of MAPK pathway rendered cells sensitive to alkaline pH

(Serrano et al., 2006). Msn2 and Msn4 have recently been implicated in alkaline pH

response, wherein owing to a transient reduction in cytosolic cAMP levels, PKA-

mediated negative regulation of Msn2 and Msn4 was lost which resulted in the nuclear

localization of Msn2 and Msn4 and the transcriptional activation of 331 genes upon

growth in alkaline pH medium (Casado et al., 2011).

1.4.2.2.3 Acidic pH response in S. cerevisiae

In S. cerevisiae, adaptation to low environmental pH conditions including

presence of weak acids involves four main regulatory systems which include (i) Msn2

and Msn4 transcription factors (Schuller et al., 2004), (ii) War1, a Zn2Cys6 transcription

factor (Schuller et al., 2004), (iii) Haa1, a transcription factor, required for adaptation

and resistance to acetic acid and propionic acid (Fernandes et al., 2005) and (iv)

Rim101, classical alkaline pH adaptive response pathway (Mira et al., 2009). PDR12,

coding for an ABC class membrane transporter, is induced under low pH conditions and

sorbic acid stress and effluxes benzoic and sorbic weak acids (Mira et al., 2010).

Transcription factor War1 regulates the expression of PDR12 and plays important role in

survival of S. cerevisiae during weak acid stress (Kren et al., 2003). Similarly, Rim101

pathway was also found to regulate the expression of several genes in response to

propionic acid (Mira et al., 2009). Deletion of RIM101 perturbed cytosolic pH

homeostasis, vacuolar acidification and cell wall structure during propionic acid stress

(Mira et al., 2009).

Msn2 and Msn4 transcription factors regulate expression of several pH

responsive genes including genes encoding small molecule transporters, carbohydrate

metabolism and cytochrome-c oxidase machinery in S. cerevisiae (Causton et al., 2001).

Of 147 up-regulated genes at pH 4.0, Msn2 and Msn4 were required for the induced

expression of 136 (93%) genes (Causton et al., 2001).

1.4.2.2.4 Acidic pH response in C. albicans

In contrast to alkaline pH response, fungal survival under acidic conditions has

not been well-studied. C. albicans PHR2, which encodes a cell wall β-glycosidase, is


43

expressed preferentially at acidic pH in a Rim101-dependent manner and required for

virulence in a vaginal model but not in a systemic model of candidiasis (Baek et al.,

2006). Another gene in C. albicans, RBR1, which codes for a GPI-linked cell wall

protein, shows induction at low pH 4.5 in an Nrg1-dependent manner and plays an

essential role in filamentation at low pH (Lotz et al., 2004). Importantly, Crz1 and Crz2-

dependent calcineurin signalling is required for survival of C. albicans and C. glabrata

in an acidic environment (Kullas et al., 2007; Chen et al., 2012).

1.4.2.2.5 Ambient pH response in C. glabrata

C. glabrata response to low or alkaline pH is largely unstudied. A recent whole

proteome analysis on C. glabrata cells grown under different pH environment, pH 4.0,

7.4 and 8.0, displayed distinct profiles of proteins implicated in protein synthesis,

folding and degradation, cytoskeleton organization and amine and carbon metabolism

(Schmidt et al., 2008). In general, protein expression pattern was more similar between

cells grown at pH 7.4 and pH 8.0 compared to cells grown at pH 4.0. Proteins involved

in organic acid and carbon metabolism, protein folding and protein-complex assembly

showed significantly lower expression at pH 7.4 and 8.0 compared with their expression

at pH 4.0. In contrast, proteins involved in cell signalling, endocytosis and protein

folding and turnover were expressed at significantly higher levels at pH 7.4 and 8.0

compared with those at pH 4.0 (Schmidt et al., 2008).

Interestingly, expression profiles of proteins involved in protein metabolism were

different in pH 8.0-grown C. glabrata and C. albicans cells. While genes involved in

protein synthesis and protein degradation are generally up-regulated and down-regulated,

respectively, at pH 8.0 in C. albicans, these proteins displayed the reverse expression

trend in C. glabrata. Schmidt et al. suggested that C. glabrata senses low pH as less

stressful than alkaline pH. These results were corroborated independently by

transcriptional profiling analysis on pH 4.5-grown C. glabrata cells wherein expression

of genes involved in cellular respiration, ribosome biogenesis and protein complex

biogenesis was up-regulated (Seider et al., 2011).

1.4.2.3 pH homeostatic mechanisms in S. cerevisiae

All cellular processes in biological systems are dependent on pH, and, thus,

intracellular pH (pHi) is a tightly regulated physiological parameter in the cell. pH is

defined by the negative logarithm of the hydrogen ion concentration in solution. In yeast

cells, pHi is tightly regulated and maintained at a homeostatic value of neutral pH in


44

response to extracellular conditions including any shifts in surrounding environmental

pH and nutrient availability (Orij et al., 2009). The master regulator of pHi in S.

cerervisiae is the plasma membrane-bound P2-type H+-ATPase, Pma1 (plasma

membrane ATPase) (Ferreira et al., 2001). P-type ATPases are located at the cell

membrane and hydrolyse ATP, which is coupled with transport of protons to the outside

of the cell. Pma1, a single 100-kDa polypeptide, is one of the most abundant proteins in

the plasma membrane and pumps protons out of the cell in a stoichiometry of 1 H+

extruded per ATP hydrolysed (Morsomme, et al., 2000). It is structurally and

functionally related to other P-type ATPases, viz., Na+, K+ and Ca2+-ATPases of animal

cells and H+-ATPase of plant cell. Studies from S. cerevisiae, S. pombe and Neurospora

crassa showed that Pma1 is firmly embedded in the lipid bilayer via 10 transmembrane

α-helices, with its N- and C-termini located in the cytoplasm (Ambesi et al., 2000).

Pma1 in S. cerevisiae is an essential gene and its expression is under the control

of an essential transcription factor, Rap1 (repressor activator protein), also known as

Tuf1 or Grf1 (Capieaux et al., 1989). PMA1 transcription and translation is known to be

modulated by external glucose concentration in the surrounding medium (Serrano, 1983;

Rao et al., 1993). Glucose has also been shown to induce phosphorylation of Pma1,

which leads to increased affinity for ATP and an increased maximum reaction rate

(Vmax) (Lecchi et al., 2007). Mass spectrometry and trypsin digestion analyses have

identified C-terminally located Ser-911 and Thr-912 as the phosphorylation sites in

Pma1 (Lecchi et al., 2007).

S. cerevisiae possesses a second P2-type H+-ATPase, Pma2, which is not

essential and expressed at 300-fold lower level than Pma1 under normal growth

conditions (Supply et al., 1993). PMA2 gene shows 89% identity to PMA1 and can

functionally replace Pma1, if expressed under the control of PMA1 promoter (Supply et

al., 1993). Besides Pma1 and Pma2, vacuole membrane-associated H+-ATPase also

plays important role in regulation of pHi homeostasis as discussed in Section.

1.4.3 Yeast vacuole

Vacuole, the most acidic organelle, in a yeast cell is functionally equivalent to

the lysosome of higher eukaryotes (Klionsky et al., 1990). Both organelles contain

several hydrolytic enzymes which are optimally active at acidic pH (Li and Kane, 2009).

Vacuoles perform myriad functions in cell physiology, viz., intracellular protein

degradation/turnover, pH and ion homeostasis, osmo-regulation, storage of


45

carbohydrates, amino acids and polyphosphate and detoxification of harmful substances

(Kane, 2007).

1.4.3.1 Morphological features

The yeast vacuole is a dynamic organelle surrounded by a single membrane

whose morphology constantly changes in response to intracellular and extracellular

environmental cues (Kane, 2007). Actively dividing yeast cells have 2-3 prominent

vacuoles in their cytoplasm. Based on growth conditions, vacuole undergoes fission and

fusion to alter their volume and size (Klionsky et al., 1990). While small vacuoles fuse

to form one large vacuole when a cell enters into the stationary phase, vacuole is also

known to fragment into many small vesicles during osmotic stress (Wickner et al.,

2002). The lipid profile of the vacuolar membrane is distinct from that of the plasma

membrane and contains reduced levels of sphingolipids and a very low level of

ergosterol to phospholipid ratio, which render vacuole membrane detergent soluble

(Lauwers et al., 2006). Characteristically, vacuolar membrane lacks sphingolipid-rich

lipid rafts which participate in signalling and membrane trafficking processes in plasma

membrane (Foster, 2003).

1.4.3.2 Protein constituents

The yeast vacuole contains several proteins in its lumen and membrane (Kane,

2007; Wiederhold et al., 2009). Owing to its role in macromolecule (protein and

carbohydrate) degradation, various hydrolytic enzymes are bonafide residents of vacuole

lumen and membrane. Vacuolar hydrolytic enzymes in S. cerevisiae are well-

characterized and include carboxypeptidase Y (CPY), carboxypeptidase S (CPS),

proteinase A (PrA), proteinase B (PrB), aminopeptidase I (AP-I), dipeptidyl

amimopeptidase B (DAPB), alkaline phosphatase (AP) and α- mannosidase (reviewed in

Kane, 2007). Additionally, another class of proteins, transporters, is enriched in the

vacuolar proteome and implicated in the transport of amino acids, metal ions and

glutathione conjugates (Wiederhold et al., 2009). Another well-studied vacuolar

membrane-associated enzyme is the highly conserved vacuolar H+-ATPase (V-ATPase)

which is evolutionarily related to the F1F0-ATPase of the mitochondria (reviewed in

Kane, 2006).


46

1.4.3.3 Vacuolar H+-ATPase: structure, function and regulation

Based on external growth conditions, pH of the yeast vacuole ranges from 5.0 to

6.5 (Martinez-Munoz et al., 2008). This pH homeostasis is achieved by highly regulated

proton pump activity of V-ATPase which couples ATP hydrolysis with proton transport

from the cytosol to the vacuolar lumen (Martinez-Munoz et al., 2008). Unlike plasma

membrane H+-ATPase, yeast V-ATPase is a large multi-subunit enzyme organized into

two domains.

(i) Structure of V-ATPase

Two domains of V-ATPase, V1 and V0, consist of several subunits (Zhang et al.,

2008). The soluble peripheral V1 domain is comprised of eight different subunits, named

as A to H, of approximately 10-70 kDa molecular mass (reviewed in Kane, 2006;

Beyenbach and Wieczorek, 2009; Pérez-Sayáns et al., 2012). The V1 domain is involved

in ATP hydrolysis and organized in stoichiometric arrangement of A3 B3 C1 D1 E2 F1 G2

H1-2. Three copies each of subunit A and B forms a hexameric head portion of V1

domain. This A3B3 hexameric complex contains ATP binding site and three catalytic

sites wherein catalytic residues are mainly contributed by subunit A, while subunit B

contributes three non-catalytic, regulatory sites (reviewed in Kane, 2006; Beyenbach and

Wieczorek, 2009; Pérez-Sayáns et al., 2012).

In S. cerevisiae, subunits A and B are encoded by VMA1 and VMA2 genes,

respectively (Drory et al., 2006). The A3B3 hexameric head portion of V1 domain is

connected to V0 domain via two types of stalks known as central stalk and peripheral

stalk. The V0 domain consists of 6 different subunits, viz., a, d, e, c, c’ and c’’ (reviewed

in Kane, 2006; Beyenbach and Wieczorek, 2009; Pérez-Sayáns et al., 2012). The central

stalk, composed of subunits D and F of V1 domain and d of V0 domain, extends from

proteolipid subunit (composed of c, c’ and c”) of V0 domain to the central core of A3B3

hexameric head of V1 domain. The peripheral stalk, composed of subunits C, E, G, and

H, connects A3B3 hexamer of V1 domain to N-terminal domain of subunit a of V0

domain (reviewed in Kane, 2006; Beyenbach and Wieczorek, 2009; Pérez-Sayáns et al.,

2012). ATP hydrolysis in A3B3 hexameric head of V1 domain drives rotation of the

central stalk and the proteolipid ring, relative to the subunit a of V0 domain. The subunit

a of V0 domain is held fixed relative to the A3B3 hexameric head of V1 by peripheral

stalk, which acts as a stator .This arrangement facilitates the translocation of proton from

the cytoplasmic side of membrane to the lumenal side, thereby, acidifying the vacuolar


47

lumen (reviewed in Kane, 2006; Beyenbach and Wieczorek, 2009; Pérez-Sayáns et al.,

2012).

One copy each of subunit c and subunit c’ and 4-5 copies of subunit c” form the

proteolipid ring of V0 domain. Central stalk’s subunits D and F bind to subunit d of V0

domain. The whole complex containing proteolipid ring, subunit d of V0 domain and

subunit D and F of V1 domain is referred as the rotary complex (reviewed in Kane, 2006;

Beyenbach and Wieczorek, 2009; Pérez-Sayáns et al., 2012). Structure of S. cerevisiae

V-ATPase is pictorially presented in Figure 1.6.

In S. cerevisiae, subunit a of V0 domain is present in two isoforms which are

encoded by VPH1 and STV1 genes (Manolson et al., 1992; 1994). The subunit a is

involved in targeting of V-ATPase to distinct cellular compartments with Vph1- and

Stv1-containing V-ATPases being sorted to the vacuolar membrane and the Golgi

network, respectively (Kawasaki-Nishi et al., 2001a). Other subunits of V0 domain i.e. d,

c, c’ and c” are encoded by VMA6, VMA3, VMA11 and VMA16 genes, respectively.

Assembly of the V0 domain occurs in the endoplasmic reticulum in the presence of five

assembly factors, viz., Vma21, Vma22, Vma12, Pkr1, and Voa1 (Ryan et al., 2008).

Figure 1.6: Diagrammatic representation of yeast vacuole H+-ATPase (V-ATPase). V-ATPase is composed of two domains, V1 and V0. The peripheral V1 domain consists of eight different subunits (A-H) in the stoichiometry shown in the figure. The integral V0 domain is composed of six subunits; a, e, d, c, c’ and c’’. The rotating central ‘rotor’ is composed of the subunit D, F, d and proteolipid ring (c, c’, c’’; shown in orange) while reminder subunits (A, B, C, H, G, E, a, e) form the stable ‘stator’ complex. (A) An assembled V-ATPase structure: The central stalk (composed of subunits D, F and d) and the peripheral stalk (composed of C, E, G and N-terminal cytoplasmic domain of subunit a) connects the V1 and V0 domain. ATP hydrolysis drives rotation of the central rotor complex resulting in H+ translocation into the vacuole lumen. (B) Disassembled V-ATPase: In yeast, glucose deprivation drives the reversible disassembly of the V-ATPase. During disassembly, the V1 domain dissociates from the subunit C and V0


48

domain. The subunit H holds rest of the V1 complex for further assembly process, while free subunit C binds with actin filaments. The disassembled V1 and V0 domain does not perform ATP hydrolysis and proton translocation, respectively (reviewed in Kane, 2006).

(ii) Functions of V-ATPase

V-ATPases are present in many cellular membranes such as endosomes,

lysosomes, Golgi-derived vesicles, secretory vesicles and plasma membrane (Klionsky

et al., 1990). V-ATPase-dependent acidification of cellular compartments is necessary

for protein degradation, zymogen activation, pH homeostasis, receptor-ligand

dissociation and uptake of small molecules such as metal ions in yeast vacuoles

(reviewed in Klionsky et al., 1990; Martinez-Munoz et al., 2008). Further, V-ATPase

activity is also essential for endocytosis and receptor cycling (Klionsky et al., 1990;

Martinez-Munoz et al., 2008).

Deletion of one of the subunits of V-ATPase results in cell death in most of the

eukaryotic cells including mammalian cells (Davies et al., 1996; Kane, 2007). Contrary

to this, S. cerevisiae and other yeast cells including C. albicans and C. glabrata can

survive mutations that impede V-ATPase activity (Kane, 2006). S. cerevisiae mutants

deleted for VMA (vacuolar membrane ATPase) genes, which encode different V-ATPase

subunits, exhibit distinct phenotypes, viz., inability to grow at high pH, in the presence of

high and low Ca2+ concentrations and non fermentable carbon souces such as glycerol

and ethanol (Kane, 2006). Moreover, deletion of genes encoding V-ATPase subunit A, B

and c also affects protein sorting leading to accumulation of precursor forms of

membrane vacuolar enzyme, AP, and soluble hydrolases, CPY and PrA, within the

secretory pathway, thereby, preventing their delivery to the vacuole (Yaver et al., 1993).

(iii) Regulation of V-ATPase activity

Cellular pH homeostasis is maintained by the collaborative efforts of plasma-

membrane H+-ATPase and vacuolar H+-ATPase. V-ATPase activity in S. cerevisiae is

primarily regulated by reversible assembly and disassembly of V1 and V0 domains in

response to different environmental cues including extracellular pH and glucose

availability (Kane, 2006; Beyenbach and Wieczorek, 2009). In yeast cells, V-ATPase

disassembles during glucose starvation and reassembles when glucose is added back to

the medium (Kane, 2006; Beyenbach and Wieczorek, 2009). Furthermore, reversible

disulphide bond formation between two conserved cysteines in the catalytic site of V-

ATPase subunit A, modulation of coupling between proton transport and ATP


49

hydrolysis and changes in the activity of vacuolar membrane ion transporters and

intracellular pH homeostasis system are known to regulate V-ATPase activity (Feng et

al., 1994; Kawasaki-Nishi et al., 2001a and 2001b; Brett et al., 2005).

1.4.3.4 Functions of vacuole

Vacuole is pivotal to cell physiology and its major cellular functions are protein

degradation, ion and metabolite storage and detoxification (Klionsky et al., 1990). Yeast

vacuole functions are briefly summarized below and schematically depicted in Figure

1.7.

1.4.3.4.1 Vacuole as a storage organelle

In yeast cells, vacuole plays a major role in storage of metal ions, phosphate, trehalose

and amino acids as discussed below.

(i) Storage of metal ions

Storage and release of metal ions, Ca²+, Zn²+, Ni²+, Cd²+, Fe3+, Mn2+, Cu2+ in vacuole is

controlled by different transporters including low affinity Ca2+/H+ exchanger, Vcx1, and

high affinity P-type Ca2+-ATPase, Pmc1. Vcx1 mediates Ca2+ uptake in a V-ATPase

dependent manner (Miseta et al., 1999) while Pmc1 is independent of the proton

gradient. Pmc1 plays an important role in Ca2+ storage in vacuoles under stress

conditions and is essential for survival of vma mutants in high extracellular Ca2+

concentrations (Cunningham and Fink, 1994). Overall, yeast vacuole is the major

intracellular calcium store with > 95% of total cellular Ca2+ ions. Vacuole is also a major

storage house for Zn2+, Mn2+ and Fe2+/Fe3+ ions (reviewed in Klionsky et al., 1990).

Owing to an essential role for vacuole in metal ion homeostasis, mutants defective in

vacuolar biogenesis (vps16, vps41 and pep12) and V-ATPase activity (vma5, vma7, cup5

and tfp1) displayed altered accumulation of manganese, calcium, sulfur, copper, cobalt,

selenium, magnesium, and nickel metal ions (Eide et al., 2005).


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Figure 1.7: Functions of the yeast vacuole: 1) Degradation: Several hydrolases (shown in green) in the vacuolar lumen helps in degradation of macromolecules delivered via multiple trafficking pathways to the vacuole. 2) Storage: Vacuole stores several ions, metals and amino acids imported by various transporters (shown in pink). Polyphosphate is also stored in vacuoles. 3) pH homeostasis and vacuolar acidification: The V-ATPase (depicted in orange) is a multi-subunit enzyme responsible for vacuolar acidification and maintenance of intracellular pH homeostasis along with the plasma membrane-bound Pma1 (shown in light blue). 4) Detoxification: vacuolar ABC transporters (shown in red) help in sequestration of toxic metals, drugs and harmful by-products of cellular metabolism.

(ii) Role for calcineurin signalling in ion homeostasis

In yeast cell, Ca2+ homeostasis is tightly regulated by calcineurin which is a

highly conserved, Ca2+/calmodulin-activated, serine/threonine protein phosphatase

(Rusnak and Mertz, 2000). It exists as a hetero-dimer of two subunits: (i) catalytic

subunit calcineurin A, and (ii) regulatory subunit calcineurin B. All eukaryotic

organisms possess one or more genes coding for each of these subunits. In S. cerevisiae

three genes CNA1, CNA2 and CNB1 encode catalytic and regulatory subunits of

calcineurin, respectively (Cyert and Thorner, 1992). Calcineurin signalling is known to

regulate several aspects of yeast cell physiology (Kraus and Heitman, 2003). In accord,

calcineurin-deficient cells (cna1∆cna2∆ or cnb1∆) show growth defects upon exposure

to high extracellular levels of Na+, Li+ and Mn2+, alkaline pH, elevated temperature and

prolonged incubation in the presence of α-factor (Mendoza et al., 1994).


51

In yeast, calcineurin-induced gene expression is dependent on activation of the

transcription factor, Crz1/Tcn1 (Stathopoulos et al., 1999). Calcineurin dephosphorylates

Crz1/Tcn1, leading to its translocation to the nucleus in a karyopherin Nmd5-dependent

manner (Polizotto and Cyert, 2001). Crz1 is known to bind to a consensus sequence, 5’-

GNGGC(G/T)CA-3’, known as CDRE (calcineurin dependent response element), in

promoters of its about 160 target genes which are mainly involved in ion homeostasis,

cell wall maintenance, vesicular transport, and protein modification (Yoshimoto et al.,

2002). Interestingly, expression of VCX1 and PMC1 is tightly regulated by

Ca2+/calmodulin-mediated calcineurin signalling (Pittman et al., 2004). Role for

calcineurin signalling in vacuolar ion homeostasis is supported by the findings that S.

cerevisiae vma mutants require functional calcineurin for vegetative growth (Garrette-

Engele et al., 1995). Hence, not surprisingly, calcineurin has been shown to be required

for virulence of C. glabrata and C. albicans (Miyazaki et al., 2010b; Chen et al., 2012).

(iii) Storage of amino acids

Vacuole is also known to store large amounts of basic and neutral amino acids in

its lumen. Basic amino acids are transported to the vacuolar lumen by Vba1, Vba2 and

Vba3 transporters (Shimazu et al., 2005) while transport of neutral amino acids is

facilitated by Avt1, a member of a family of seven transporters (Avt1-7) (Russnak et al.,

2001). Others members of this family, Avt3, Avt4 and Avt6 are implicated in amino acid

export from the vacuole (Russnak et al., 2001). Importantly, activity of all Avt family

transporters requires proton gradient across the vacuolar membrane for efficient

transport process (Russnak et al., 2001).

(iv) Storage of phosphate

The vacuole contains a very high concentration of phosphate in the form of

polyphosphate (Poly P) which is a polymer of tens to hundreds of phosphate residues

linked by phosphoanhydride bonds (Rao et al., 2009). In S. cerevisiae, 90-99% poly P

pool resides in the vacuole (Freimoser et. al., 2006). Poly P are synthesized and

accumulated in the vacuole by consorted action of four vacuolar transporter chaperone

(Vtc) proteins, Vtc1, Vtc2, Vtc3 and Vtc4, in response to phosphate availability (Ogawa

et al., 2000). In yeast cells, although polyphosphate mainly act as a major source of

phosphate and buffering system for positively charged ions including Ca2+ and Mg2+,

Poly P have been implicated in long-term stress survival, blood coagulation and

fibrinolysis, energy homeostasis and regulation of chromatin condensation and

translation in other organisms (reviewed in Rao et al., 2009; Pavlov et al., 2010).


52

Phosphate storage and utilization in yeast cells is tightly regulated by phosphate-

responsive signalling (PHO) pathway (Freimoser et al., 2006). During phosphate

starvation, phosphate is acquired by four cell membrane-bound phosphate transporters

(Wykoff and O’Shea, 2001). Another recently identified phosphate transporter, Pho91,

which resides in vacuole, plays an important role in Poly P accumulation (Hurlimann et

al., 2007). Under low-phosphate conditions, transcription factor, Pho4, gets

dephosphorylated and localized to the nucleus where it triggers activation of

transcription of phosphate responsive genes and these events occur in the reverse order

under phosphate-rich conditions (Springer et al., 2003).

Polyphosphate metabolism is tightly linked with phosphate availability. Poly P

are synthesized and accumulated in vacuoles under phosphate-surplus conditions while

they are rapidly utilized in phosphate-deficient environment (Kulaev et al., 1999). Poly P

are also known to buffer transient fluctuations in extracellular phosphate levels (Thomas

et al., 2005). A complex comprised of four vacuolar transporters, Vtc1, Vtc2, Vtc3 and

Vtc4, has been implicated in maintenance of poly P homeostasis in yeast (Ogawa et al.,

2000; Hothorn, 2009).

Some components of S. cerevisiae PHO signalling pathway including Pho4,

Pho81 and Pho84 are conserved in C. glabrata (Kerwin and Wykoff, 2009), although, C.

glabrata has lost the gene coding for a phosphate-repressible acid phosphatase, Pho5,

along with other genes, PHO3, PHO11, PHO12, that are required for phosphate

metabolism in S. cerevisiae (Orkwis, et al., 2010; Jandric and Schüller, 2011).

1.4.3.4.2 Vacuole as a proteolytic degradation system

Vacuole also functions as a cellular protein degradation system along with

proteosome-mediated protein degradation and harbors several proteolytic enzymes (van

Den Hazel et al., 1996). In S. cerevisiae, vacuole contains soluble vacuolar proteases,

PrA, PrB, CPY and CPS, and membrane bound proteases, AP-1 and DPAP-B (reviewed

in Klionsky et al., 1990). Several cellular proteins are targeted to the vacuole for

degradation either when not required or as a part of their recycling process (Springael

and Andre, 1998; Liu, Y et al., 2006). Additionally, breakdown of cargo containing

vesicles in the vacuole during autophagy is also reported to be dependent on vacuolar

proteases, PrA and PrB (Harding et al., 1995).


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1.4.3.4.3 Detoxification functions of vacuole

Yeast vacuole also serves as a detoxification house of the cell, wherein various

toxic molecules and catabolites are sequestered away from the cytosol in glutathione,

glucuronide or sulfate-conjugated forms (Sharma et al., 2002). Vacuolar transport of

these conjugated molecules is primarily mediated by multidrug resistance-related

proteins (MRPs) and transport activity of two such vacuolar membrane proteins, Ycf1

and Ybt1, has been reported to be independent of the V-ATPase activity (Sharma et al.,

2002). Expectedly, vacuolar function-defective mutants displayed elevated susceptibility

to multiple drugs (Parsons et al., 2004).

1.4.3.4.4 Role of vacuole in fungal virulence

Vacuole size and morphology are known to undergo dynamic changes with

emergence and extension of germ tube (Barelle et al., 2003). Consistently, disruption of

vacuolar proteins, Abg1 and Vac8, resulted in altered hyphal branching in C. albicans

(Veses et al., 2005; Barelle et al., 2006). Recently, Vma7, a putative V-ATPase subunit,

in C. albicans has been shown to play an essential role in vacuolar acidification and

virulence in a murine model of systemic candidiasis (Poltermann et al., 2005). Similarly,

a pivotal link between properly functioning vacuole and virulence has also been

established in C. neoformans (Erickson et al., 2001; Liu, X et al., 2006). However, role

for vacuole in pathogenesis of C. glabrata remains an unexplored niche till date.


54

Objectives of the present study

Fungal virulence is determined by a set of attributes wherein aspartyl proteases

occupy a coveted position. Despite the recent emergence of C. glabrata as a major

nosocomial pathogen, its virulence factors remain poorly-studied. The primary goal of

the current study was to decipher the traits which make C. glabrata a successful

pathogen. A family of eleven GPI-linked aspartyl proteases has recently been shown to

be required for virulence of C. glabrata, however, functions of individual members of

this protease family in pathobiology of C. glabrata are not known. Hence, the current

study is mainly aimed at delineating the role for eleven GPI-linked aspartyl proteases in

cell physiology of C. glabrata. Specific objectives of this work can be summarized as

follows.

1. To investigate the role for GPI-anchored aspartyl proteases in C. glabrata

survival under different environmental stresses.

2. To identify the environmental cues that regulate expression of yapsin-encoding

genes in C. glabrata.

3. To decipher the molecular basis underlying yapsin essentiality for C. glabrata

virulence.

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