Chapter 1 Introduction -...
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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
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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
Chapter1 Introduction
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.,
Chapter1 Introduction
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).
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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
Chapter1 Introduction
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).
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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
Chapter1 Introduction
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).
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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
CgYps5 (CAGL0E01771g) 519 57.2 Q6FVH8 A1 Cell surface/GPI 5.5 ND 69, 307 1-17
CgYps6 (CAGL0E01793g) 516 55.9 Q6FVH7 A1 Cell surface/GPI 4.6 ND 67, 304 1-15
CgYps7 (CAGL0A02431g) 587 63.4 Q6FY32 A1 Cell surface/GPI 4.7 ND ND 1-18
CgYps8 (CAGL0E01815g) 519 56.6 Q6FVH6 A1 Cell surface/GPI 6.8 ND 68, 307 1-15
CgYps9 (CAGL0E01837g) 521 56.8 Q6FVH5 A1 Cell surface/GPI 5.1 ND 68, 303 1-16
CgYps10 (CAGL0E01859g) 505 55.3 Q6FVH4 A1 Cell surface/GPI 7.3 ND 64, 301 1-13
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.
Chapter1 Introduction
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
Chapter1 Introduction
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
Chapter1 Introduction
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).
Chapter1 Introduction
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.
Chapter1 Introduction
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).
Chapter1 Introduction
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
Chapter1 Introduction
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.,
Chapter1 Introduction
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).
Chapter1 Introduction
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
Chapter1 Introduction
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
Chapter1 Introduction
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
Chapter1 Introduction
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,
Chapter1 Introduction
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
Chapter1 Introduction
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).
Chapter1 Introduction
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
Chapter1 Introduction
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
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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).
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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
Chapter1 Introduction
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
Chapter1 Introduction
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.
Chapter1 Introduction
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
Chapter1 Introduction
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
Chapter1 Introduction
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
Chapter1 Introduction
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).
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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
Chapter1 Introduction
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
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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
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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).
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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).
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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.
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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.