University of Groningen
Carbohydrate-active enzymes that modify the cell wall of Aspergillus niger
van Munster, Jolanda
DOI:
10.1016/j.carres.2015.01.014
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van Munster, J. (2014). Carbohydrate-active enzymes that modify the cell wall of Aspergillus niger:
Biochemical properties and physiological functions during autolysis and differentiation [S.n.] DOI:
10.1016/j.carres.2015.01.014
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Chapter
2
Chapter
1
General Introduction
9
Chapter 1
Filamentous fungi are common in nature and various species are of significant industrial or medical importance. These fungi encounter carbon starvation in nature
as well as during industrial fermentations. In response, the fungi recycle part of their
mycelium to gain building blocks and energy to survive by e.g. formation of stress
resistant spores. We currently lack insight in the processes that take place in fungi
during carbon starvation. Enzymes that act on carbohydrates in the cell wall are key
effectors in the degradation of the cell wall during carbon starvation as well as in cell
wall changes that take place during the formation of spores. This thesis focuses on
the biochemical properties and physiological roles of carbohydrate-active enzymes
under carbon starvation conditions in the industrially important fungus Aspergillus
niger.
The Genus Aspergillus
Ubiquitous saprophytes
The kingdom of the Fungi harbors members as diverse as yeasts, molds and
mushrooms (Stajich et al., 2009). The fungi of the genus Aspergillus are common
saprophytes, they decay dead organic matter to release nutrients for their growth.
These fungi have adapted to living on a wide range of substrates and can efficiently
degrade plant polymers by secreting hydrolytic enzymes. Aspergillus spores, reproductive cells that are highly resistant to adverse environmental conditions, are
spread through the air to new habitats, where the spores germinate when nutrient
and water conditions are appropriate. The name Aspergillus originates from the
resemblance of the spore forming structures to the aspergill, a device used in the
Roman Catholic church to sprinkle holy water (Bennett, 2010). The Aspergilli belong
to the phylum Ascomycota, which also includes fungi such as the penicillin producer Penicillium chrysogenum, the laboratory model organism Neurospora crassa
as well as Trichoderma reesei, a source of cellulases used for biofuel production.
These fungi are molds, or filamentous fungi, which grow by generating elongated
cellular structures called hyphae, that form a network by branching and fusion.
The genus Aspergillus contains both medically and industrially important species.
The large number of Aspergillus genomes that have been sequenced in the last
decade illustrates the interest in this genus. After publication of the genome sequence of A. fumigatus (Denning et al., 2002) over ten years ago, the genomes of
A. niger (Andersen et al., 2011;Pel et al., 2007), A. oryzae, A. flavus (Machida et
10
Introduction
Aspergillus niger as producer of organic acids and proteins
A. niger is industrially important as producer of organic acids. As much as 900 kilotons of citric acid is produced annually for use in the food -, cleaning - and pharmaceutical industry, 95% of which is generated by A. niger fermentations (Karaffa &
Kubicek, 2003;Kubicek & Karaffa, 2006). Fermentations of A. niger are also used to
produce gluconic acid, which worldwide amounts to 60 kilotons annually. Gluconic
acid is a weak organic acid that is used as a food additive and has applications in
cleaning products and the pharmaceutical industry (Ramachandran et al., 2006).
Because of its capacity to secrete large amounts of proteins, A. niger is used as a
cell factory for production of heterologous and homologous enzymes. Traditional,
11
Chapter 1
Chapter 1
al., 2005;Payne et al., 2006), and A. nidulans (Galagan et al., 2005) have become
available. Many others have been added more recently and as a result the genome
sequences of 19 species are currently available in comparative databases such as
the Aspergillus Genome Database (AspGD ) (Arnaud et al., 2010) and the Central
Aspergillus Data Repository (CADRE) (Mabey Gilsenan et al., 2008). A. nidulans is
one of the model organisms in fungal genetics and cell biology. While the majority
of Aspergilli reproduce only asexually, A. nidulans has a sexual reproductive cycle
that been investigated in detail (Dyer & O’Gorman, 2012). This species produces the toxin sterigmatocystin, which makes it unsuitable for industrial applications
(Ward, 2011). A. flavus and A. parasiticus are also known to produce potent toxic
and carcinogenic metabolites such as aflatoxins. This is particularly problematic as
these species are commonly found contaminating grain and nut crops (Georgianna
& Payne, 2009). A. fumigatus is an opportunistic human pathogen. Its spores, with
a diameter of 2-3 µm, are small enough to enter into the lower pulmonary system
and can cause an allergy to fungal spores (Latgé, 1999;Moss, 2005). In patients
with severely compromised immune systems, A. fumigatus can cause invasive aspergillosis, cases of which have a mortality rate of between 50-90% (Latgé, 1999).
A. terreus is used to produce the cholesterol lowering drug Lovastatin in submerged
liquid fermentations (Barrios-Gonzalez & Miranda, 2010). A. oryzae is key in fermentations of soy sauce and sake (Kitamoto, 2002;Machida et al., 2005). A. niger
and A. oryzae are industrial workhorses for the production of enzymes and primary
and secondary metabolites. Their long history of safe use in food applications has
resulted in the acquisition of the generally regarded as safe (GRAS) status, granted
by the Food and Drug Administration (Ward, 2011).
Chapter 1
mutagenesis based, strain selection procedures raised production levels to more
than 30 g protein l-1 (Punt et al., 2002). Enzymes such as alpha-amylase, glucoamylase, glucose oxidase, lipases, proteases, xylanases are produced for use in the
food industry, starch processing industry, detergent production, textile processing
and the pharmaceutical industry (Fleissner & Dersch, 2010;Meyer, 2008). The availability of genetic tools for manipulation of Aspergilli has opened up the possibility
to use these fungi as production hosts for secretion of heterologous proteins. Several inducible and constitutive promoters have been identified (Fleissner & Dersch,
2010) and metabolism independent, tight promoter systems are available (Meyer
et al., 2011;Shoji et al., 2005;Vogt et al., 2005). Various strategies have been employed to improve translation, folding and secretion, including boosting the natural
stress response to unfolded proteins in the ER, and fusions to native, secreted proteins. The use of protease deficient strains and protease inhibiting growth conditions
can further improve yield of a secreted protein, as reviewed by (Fleissner & Dersch,
2010;Ward, 2011). In spite of these recent advances, production of heterologous
proteins is still more problematic than homologous protein production (Ward, 2011).
Nevertheless, A. niger is widely used in the industrial production of proteins and organic acids, and insights into its physiology may help industrial strain development.
Growth, morphology and differentiation
Growth conditions and fungal morphology differ quite substantially between largescale, industrial fermentations and the situation in nature, although some features
are common to both. Filamentous fungi, such as A. niger, grow using elongated
hyphae, where growth takes place at the tip. Within each hyphae, multiple compartments are found, which are separated by a septum (Fig. 1A). Septal pores allow
passage of cytoplasm and transport of organelles, and sustain tip growth by relocating nutrients and other resources (Riquelme et al., 2011). This mode of growth
allows areas with low nutrient levels to be bridged and exploratory hyphae to extent
in search of new nutrients. Upon injury of hyphae, leakage of cytoplasm is minimized by plugging septa with specialized organelles called woronin bodies (Jedd,
2011;Markham & Collinge, 1987). Hyphae are surrounded by a cell wall, which is
discussed in more detail below.
The morphology of fungal colonies of a single strain can differ dramatically depending on growth conditions. Growth in submerged liquid cultures results in dispersed
12
Introduction
A
B
C
Chapter 1
Chapter 1
Fig. 1. Morphology of filamentous fungi (e.g. A. niger) under different conditions. A: a hyphae
with cell organelles, divided into connected compartments by septa, surrounded by a cell
wall. B: clumps of mycelium (pellets) as may be found in submerged liquid cultures. C: colony
growing on solid media such as an agar plate, with vegetative - and aerial hyphae and spore
forming structures.
mycelium or in balls of aggregated mycelium (pellets) (Fig. 1B), while growth on
agar plates and other surfaces results in flat, radial macro-colonies (Fig. 1C). When
considering a single one of these overall morphologies, the colonies are highly differentiated. In both pellets and flat macro-colonies, the center of the colony consists
of old mycelium and the colonies’ edge consists of newly formed mycelium. The total mRNA levels, gene expression and protein secretion differ going from the colony
center to the periphery (de Bekker et al., 2011b;Gordon et al., 2000;Krijgsheld et
al., 2012;Levin et al., 2007;Wösten et al., 1991). Even within one zone in a colony,
hyphae may be heterogeneous with respect to gene expression as well as levels of
transcription and translation (de Bekker et al., 2011a;Vinck et al., 2011;Wösten et
al., 2013). The closure of septa, resulting in the limitation of cytoplasmic streaming,
plays a role in maintaining this hyphal heterogeneity in A. oryzae (Bleichrodt et al.,
2012).
Differentiation within a colony plays a role in submerged fermentations as fungi
grown in pellets do not behave uniformly. This differentiation is minimized when the
fungus grows as dispersed mycelium. Differentiation is also important during the
response to carbon starvation. For example, during autolytic turnover of the mycelium, only a part of the hyphae is degraded and during sporulation only a number of
the compartments generate spore forming structures.
13
Chapter 1
The fungal cell wall
The fungal cell is surrounded by a cell wall which forms the interface of the fungus
with the environment. The cell wall not only protects the cells against environmental
influences, but the strength of the cell wall also enables functions such as substrate
penetration. Most of our current knowledge of the cell wall composition and structure, as well as of the enzymes involved in modification of the cell wall, originates
from research on the filamentous fungus A. fumigatus and the yeast Saccaromyces
cerevisiae.
Cell wall composition and structure
The fungal cell wall is composed of a network of carbohydrate polymers, with which
cell wall proteins are associated. The carbohydrate fraction of the cell wall of A. fumigatus is composed of 72 % glucose, 3 % mannose, 4 % galactose, 13 % N-acetylglucosamine, 4 % glucosamine and 6 % galactosamine (Maubon et al., 2006). The
A. niger cell wall has a similar composition, but with slighty less (N-acetyl)glucosamine (9-12 %) and more galactose (10-14 %) (Crook & Johnston, 1962;Johnston,
1963;Johnston, 1965a). The cell wall carbohydrates can be divided into an alkalineinsoluble backbone structure, which is embedded in alkaline-soluble polymers. The
alkaline-insoluble fraction of the A. fumigatus cell wall (Fig. 2A) consists of a β-1,3glucan backbone with β-1,6 branches. Other polysaccharides such as chitin (composed of β-1,4 linked N-acetylglucosamine units [GlcNAc]), β-1,3-1,4-linked glucan
and galactomannan (α-1,2/α-1,6-linked mannan with short β-1,5-galactofuranose
side chains) are attached to the non-reducing ends of this backbone (Fontaine et
al., 2000). The alkaline soluble part of the A. fumigatus cell wall (Fig. 2B) consists
mainly of α-1,3(/α-1,4)-linked glucose polymers (Latgé, 2007) and similar polymers
have been identified in A. niger. Nigeran has alternating α-1,3- and α-1,4-linkages
(Johnston, 1965a) while pseudonigeran mainly has α-1,3-linkages (Johnston,
1965b). Small amounts of other polymers can be found associated with the cell
wall. The galactomannan polymer (Bardalaye & Nordin, 1977;Baretto-Bergter et al.,
1980;Latgé et al., 1994) is not only found covalently attached to glucans but may
also be released in the culture filtrate (Fontaine et al., 2000) or may be present as
GPI-anchored polymer in the cell wall (Costachel et al., 2005). A linear heterogenous
α-1,4-galactose/galactosamine polymer (galactosaminogalactan) was found associated
with the alkali soluble part of the A. niger and A. fumigatus cell wall, as well as being secreted in the culture filtrate (Bardalaye & Nordin, 1976;Fontaine et al., 2011).
14
Introduction
A
B
β-1,6
β-1,3-glucan
with β-1,6-branches
α-1,3/1,4-glucan (nigeran)
α-1,3
α-1,3
α-1,4
Chapter 1
α-1,4
Chapter 1
β-1,3
β-1,4
β-1,4
β-1,3
β-1,3
β-1,3/1,4-glucan
α-1,3-glucan
β-1,4
β-1,4
α-1,6
galactomannan
α-1,2
chitin
β-1,5
galactomannan
Fig. 2. Structure of the carbohydrate network of the fungal cell wall. A: alkaline insoluble fraction, consisting of a backbone of β-1,3-glucan, to which polymers as chitin and galactomannan are attached. B: alkaline soluble fraction, consisting mainly of α-glucans. Monomers are
depicted as circle, glucose; square, N-acetyl-glucosamine; striped square, glucosamine; triangle, mannose; star, galactofuranose. Figure based on (Fontaine et al., 2000;Latgé, 2007).
The cell wall of spores differs from that of the vegetative mycelium. In dormant
spores, the cell wall is covered with a hydrophobic layer composed of hydrophobins.
The cell wall underneath the hydrophobins consists of two layers, the outer of which
is pigmented (Bernard & Latgé, 2001;Latgé et al., 2005). Compared to vegetative
mycelium, the cell walls of A. fumigatus conidia contain less N-acetyl-glucosamine
(2 % of total carbohydrates instead of 13 %) and more mannose and galactose (25
% and 14 % instead of 4 %) (Maubon et al., 2006).
Cell wall synthesis, remodeling and degradation
The polymers that make up the carbohydrate network of the cell wall are synthesized individually and subsequently connected to the network of the cell wall. Carbohydrate-Active enZymes (CAZymes) are responsible for synthesis, remodeling,
cross-linking and degradation of these polymers.
The β-1,3-glucan that makes up the backbone of the cell wall, is synthesized at the
plasma membrane by a β-1,3-glucan synthase. The catalytic subunit of the synthase, encoded by the single fksA gene, uses UDP-glucose to generate and extrude
a linear polymer to the periplasm. This newly synthesized β-1,3-glucan is subsequently modified and attached to the existing cell wall carbohydrate network (Doug15
Chapter 1
las, 2001). The role of β-glucan modifying enzymes in A.fumigatus has recently
been reviewed (Mouyna et al., 2013). Members of family GH72, which are attached
to the cell wall by a GPI-anchor, can elongate β-1,3-chains. They use an endoacting mechanism to cleave a linear β-1,3-glucan, and transfer the donor to the nonreducing end of an acceptor β-1,3-glucan by creating a β-1,3 linkage (de MedinaRedondo et al., 2010;Hartland et al., 1996;Mouyna et al., 2000). Enzymes in family
GH17 can form β-1,6 branch points in β-1,3-glucan, either at the end (Mouyna et
al., 1998) or predominantly in the interior of the acceptor β-1,3-chain (Gastebois et
al., 2010). Evidence for a role of endo- and exo-β-1,3-glucanases in cell wall β-1,3glucan modification is also accumulating. Biochemical characterization showed that
the family GH81 A. fumigatus ENG1 is an endo-β-1,3-glucanase (Fontaine et al.,
1997b;Mouyna et al., 2002). Its ortholog in A. nidulans, EngA, causes fungal biomass reduction under carbon starvation conditions (Szilagyi et al., 2010), and is
thus likely to act on the cell wall. A. niger harbours three GH55 members; exo-β-1,3glucanases specific to filamentous fungi that release glucose from the non-reducing
end of β-1,3-glucan oligosaccharides. These enzymes may release gentobiose indicating they can hydrolyze β-1,3-glucan with a short β-1,6 branch as found in the cell
wall (Ishida et al., 2009;Oda et al., 2002) and thus complement the activity of cell
wall acting endo-β-1,3-glucanases. The recently identified GH132 family harbors
enzymes that can hydrolyze β-(1,3)-oligosaccharides ranging in size from dimer up
to insoluble polymeric β-(1,3)-glucan. Deletion of the gene encoding one of these
A. fumigatus enzymes results in cell wall defects, suggesting that the protein is involved in cell wall biogenesis (Gastebois et al., 2013).
Chitin is synthesized at the plasma membrane from UDP-GlcNAc by chitin synthases. The A. niger genome has 9 chitin synthases, with at least one representative of
the 7 different classes in which chitin synthases are divided based on their domain
structure (Pel et al., 2007;Rogg et al., 2012). These chitin synthases play distinct
roles in cell wall synthesis, with some of them being redundant (Rogg et al., 2012).
For example, the A. nidulans Class I ChsC and Class II ChsA have partially overlapping roles in hyphal wall integrity, conidiophore development and septum formation
(Fujiwara et al., 2000;Ichinomiya et al., 2005;Ichinomiya et al., 2002). The chsA
gene is expressed specifically during asexual reproduction while the chsC gene has
a much broader expression pattern (Lee et al., 2004). Once the new chitin chain
is synthesized, it can be connected through its reducing end to the β-1,3-glucan
backbone (Fontaine et al., 2000). The enzymes responsible for generating this con16
Introduction
The mechanism behind cell wall galactomannan synthesis in filamentous fungi is
largely unknown. In S. cerevisae the α-1,6-mannosyl-transferase Och1p initiates
synthesis of mannan and the Golgi enzyme-complexes containing Hoc1p, Van1p,
Anp1p Mnn9p, Mnn10p and Mnn11p synthesize α-mannan chains (Lambou et al.,
2010). Multiple homologs of Hoc1p and Och1p and unique homologs of the other
complex members are present in Aspergillus spp (Gastebois et al., 2009;Geysens
et al., 2009). The enzymes involved in synthesis of β-1,5-galactofuranose side
chains remain to be identified. A possible substrate, UDP-galactofuranose, is generated from UDP-galactopyranose by a UDP-galactomutase (Damveld et al., 2008).
A Golgi GDP-mannose transporter and UDP-galactofuranose transporters are required for the presence of galactomannan and galactofuran, respectively, in the cell
walls of A. fumigatus and A. nidulans (Afroz et al., 2011;Engel et al., 2012). This
suggests that galactomannan, in contrast to β-1,3-glucan and chitin, is produced in
the Golgi organel (Engel et al., 2012). The role of galactofuranose in A. niger was
recently reviewed (Tefsen et al., 2012).
Cell wall α-1,3-glucan is produced by α-glucan synthases, large proteins with an
extracellular GH13 domain and intracellular GT5 domain. A. nidulans has 2 genes
encoding α-glucan synthases, while A. fumigatus has 3 and A. niger even has 5 of
these genes. The involvement of multiple synthases may be related to functional
diversification. In A. nidulans, AgsB is the main α-glucan synthases during vegetative growth while AgsA is expressed during asexual sporulation (He et al., 2014).
Functional differences between the α-glucan synthases were also found in A. niger
17
Chapter 1
Chapter 1
nection have not been identified in filamentous fungi. However, in S. cerevisiae the
presence of either of the GH16 proteins Chr1 or Chr2 is required in-vivo to couple
chitin to β-1,6-glucan and β-1,3 glucan (Cabib et al., 2007;Cabib, 2009;Cabib et al.,
2008). The enzymes are induced during cell wall stress, and their activity is part
of the cell wall compensatory mechanism of the cell (Cabib et al., 2007;Garcia et
al., 2004). The A. niger genome encodes 3, respectively 2 GPI-anchored homologs of S. cerevisiae Chr1 and Chr2, which may be involved in cross-linking chitin
to β-1,3-glucan. Chitin can be modified through the action of chitinases and chitin
deacetylases. A. niger harbors a large number of chitinases, which all belong to GH
family 18. These chitinases vary in their domain structure, expression pattern and
biochemical activities and evidence for distinct physiological functions is accumulating. The Aspergillus chitinases are discussed in detail in Chapter 5 of this thesis.
Chapter 1
(Damveld et al., 2005;van der Kaaij et al., 2007a) and A. fumigatus (Beauvais et al.,
2005;Maubon et al., 2006). Deletion of all α-glucan synthase genes in A. fumigatus
and A.nidulans indicated that these mutant strains are viable while lacking α-1,3glucan (Henry et al., 2012;Yoshimi et al., 2013). The exact mechanism of action and
substrate used by the α-1,3-glucan synthases is not known. However, the enzyme
is thought to use a short primer of α-1,4-glucan to which glucose units are subsequently added (Grün et al., 2005;van der Kaaij et al., 2007a). Intracellular amylases
have been identified that contribute to α-1,3-glucan synthesis, perhaps by generating such as primer (He et al., 2014;Marion et al., 2006;van der Kaaij et al., 2007a).
Fungal enzymes that modify α-1,3-glucan can be found in family GH71. A number
of such enzymes from filamentous fungi have been characterized. Most of them are
secreted proteins that release mainly glucose from α-1,3-glucan (Fuglsang et al.,
2000;Grün et al., 2006;Sanz et al., 2005;Shalom et al., 2008;Wiater et al., 2001).
In addition some enzymes have been identified to be associated with the cell wall.
The A. nidulans MutA is localized in Hülle cells, which are present during sexual
sporulation (Wei et al., 2001;Zonneveld, 1972a;Zonneveld, 1972b). The enzymes
Agn1 and Agn2 from Schizosaccharomyces pombe release small oligosaccharides
from α-1,3-glucan and hydrolyze the cell wall during cell separation or sporulation
(Dekker et al., 2004;Dekker et al., 2007;Garcia et al., 2005;Grün et al., 2006). Furthermore, GPI-anchored enzymes from GH13 have been identified, such as A. niger AgtA, that are capable of transglycosylation reactions using α-1,4-glucan and
short α-1,3-glucan oligosaccharides. These enzymes may have a role in modification of cell wall α-glucan as deletion of agtA results in increased sensitivity towards
a cell wall disrupting compound (van der Kaaij et al., 2007b). It is currently unknown
how nigeran, with alternating α-1,3/ α-1,4-linkages, is produced. T.reesei secretes
an enzyme capable of degrading both α-1,3-glucan and nigeran,but the corresponding gene has not yet been cloned and it is not known to which enzyme family this
activity belongs (Hasegawa & Nordin, 1969).
Carbon starvation
The industrial production of enzymes, organic acids, or secondary metabolites by
filamentous fungi typically takes place in large scale fermentors. Both in nature
and under laboratory or industrial conditions, fungi encounter carbon starvation the
moment when substrates run out. Our current understanding of the processes that
take place under carbon starvation conditions is limited, while the effects can be
18
Introduction
Fungi undergo morphological changes during carbon starvation. Long and thin hyphae are formed, and a large number of vacuoles appear in the cytoplasm of older
hyphae. In addition, empty hyphal compartments may be observed as well as, after
prolonged starvation, spore forming structures (McIntyre et al., 2001;Nitsche et al.,
2012;White et al., 2002). Fungal programmed cell death and fragmentation of the
fungal cell wall have been reported during carbon starvation.
Programmed cell death can occur through necrosis, apoptosis or be associated with
autophagy. Necrosis is associated with rapid loss of plasma membrane integrity, organelle swelling and mitochondrial dysfunction (Hitomi et al., 2008), resulting in the
uncontrolled release of all kinds of compounds. Typical apoptotic features include
the externalization of phosphatidylserine to the membrane surface, DNA cleavage
and mitochondrial damage (Hamann et al., 2008). Apoptotic markers have been
found associated with carbon starving and aging fungi (Emri et al., 2005a;Mousavi
& Robson, 2003). Autophagy is a non-selective process in which cytoplasm and cell
organelles are degraded in the vacuole and their components recycled (Pollack et
al., 2009). Autophagy is not only associated with cell death but is also important in
normal cell maintenance. It was recently shown that autophagy plays a role in the
direct reponse of A. niger to carbon starvation. Carbon starvation induces genes associated with autophagy in A. niger (Nitsche et al., 2012) and A. nidulans (Szilagyi
et al., 2013). Deletion of genes atg1 and atg8, which are required for autophagy,
resulted in reduced mitochondrial turnover and a prolonged viability during carbon
starvation in A. niger (Nitsche et al., 2013).
19
Chapter 1
Chapter 1
substantial (Emri et al., 2008;White et al., 2002). Aspergillus may respond to carbon
starvation by recycling part of its own mycelium. This process is usually referred to
as autolysis, and is accompanied by the disintegration and degradation of fungal
biomass, the production of hydrolytic enzymes such as proteases and chitinases,
and the release of ammonia (White et al., 2002). Autolysis may affect industrial fermentations as cultures are losing productive power and biomass fragmentation can
cause downstream processing problems. The induction of hydrolytic enzymes such
as proteases can damage or even destroy the products of interest, especially in the
case of enzymes (White et al., 2002). On the other hand, insights into the process
of autolysis could provide tools to assist in the recovery of intracellular fermentation
products (Emri et al., 2008;White et al., 2002).
Chapter 1
During carbon starvation the fungal cell wall is thought to undergo drastic remodelling or degradation. Fungal carbohydrate-active enzymes are key effectors in these
cell wall changes. The production of such hydrolytic activities in response to carbon
starvation has been observed in a number of fungi (Jaroszuk-Scisel et al., 2011;Lahoz et al., 1976;McIntyre et al., 2000;Perez-Leblic et al., 1982;White et al., 2002).
Recent investigations of the transcriptome and proteome of carbon starving A. niger
(Nitsche et al., 2012) and A. nidulans (Szilagyi et al., 2013) cultures have provided
an overview of the up-regulated fungal cell wall active carbohydrate hydrolases.
However, the roles of only a number of these enzymes have been confirmed and investigated in more detail. Chitinase ChiB is rate limiting in biomass degradation during carbon starvation in A. nidulans. Deletion of the corresponding gene results in
less biomass loss and pellet disintegration (Shin et al., 2009;Yamazaki et al., 2007).
Deletion of the homologous A. fumigatus chiB1 - also encoding the main chitinase
induced during starvation - does not result in a change in biomass loss during starvation (Jaques et al., 2003): the physiological role of these enzymes thus appears
to be different. A. fumigatus ChiB1 hydrolyzes chitin and chitin oligosaccharides
by cleaving mainly chitobiose from the non-reducing substrate end, but can also
perform a transglycosylation reaction (Escott et al., 1998;Jaques et al., 2003;Lu
et al., 2009;Xia et al., 2001). The A. nidulans ChiB appears to have similar activity
(Erdei et al., 2008). We show in chapter 3 that the biochemical activity of A. niger
CfcA is similar to its homologs A. nidulans ChiB and A. fumigatus ChiB1, and that
the enzyme is responsible for cell wall degradation despite the fact that no changes
in biomass decrease were detected upon gene deletion. The A. nidulans EngA also
plays a role in biomass decrease during autolysis (Szilagyi et al., 2010). Its ortholog
in A. fumigatus has been characterized as an endo-acting β-1,3-glucanase, and
was found to be associated with the fungal cell wall (Fontaine et al., 1997a;Mouyna
et al., 2002). The β-N-acetyl-hexosaminidase encoded by nagA, which is expressed
during carbon starvation in A. nidulans, is thought to contribute to cell death but not
to autolysis (Shin et al., 2009). To identify the full repertoire of autolytic hydrolases
in A. niger, we investigated the regulation behind their expression as well as the
biochemical and physiological role of two of these enzymes in detail (Chapter 2, 3).
Upon carbon starvation in A. niger both apoptotic/necrotic cell death and/or autophagy may result in cytoplasm degradation and the appearance of empty hyphal
compartment. During prolonged starvation, spore forming structures are generated
and, under the right conditions, the cell walls can be fragmented and degraded
20
Introduction
During periods of stress, Aspergillus can survive by forming stress resistant spores.
The formation of these spores requires energy and building blocks. In case of nutrient limitation, fungal autolysis and autophagy may make these nutrients available
by recycling the cell wall or intracellular components. The regulatory machinery responsible for the initiation of sporulation is linked to the regulation of autolysis.
Sporulation
Asexual reproduction by means of stress resistant spores is an important part of
the life cycle of A. niger. The production of asexual spores (conidia) and the spore
bearing structures (conidiophores) requires a set of tightly regulated morphological
changes, which have been extensively investigated in A. nidulans. The conidiophore formation starts from a foot cell, a specialized cell with a thick cell wall from
which a stalk is formed that elongates into the air (Fig. 3). The top of the stalk swells
to form a vesicle, in which nuclei multiply and migrate towards the top of the vesicle.
Multiple buds will form, each harboring one of the nuclei. These buds, metulae,
produce two phialides, which are the cells that will form a chain of conidia. As each
vesicle harbors 60-70 nuclei and over 100 conidia are found in a chains extending
from the phialide, each conidophore generates over 10 000 conidia (Adams et al.,
1998;Krijgsheld et al., 2013). The produced conidia are highly resistant to adverse
environmental conditions. Their outer layer is covered by pigments, which renders
the spores of A. nidulans and A. fumigatus green, and those of A. niger black (Bennett, 2010).
The regulation of conidiation is divided in a global upstream response and a conidiophore specific regulatory pathway (Fig. 1 of Chapter 2). Sensors and regulators such as the VeA complex react to environmental stimuli such as light, carbon
depletion and osmotic stress (Bayram & Braus, 2012;Etxebeste et al., 2010). They
21
Chapter 1
Chapter 1
by hydrolytic activities (Nitsche et al., 2012),(Chapter 2,3). During carbon starvation, both cell death and fungal cell wall fragmentation occur but these processes
are regulated independently (Emri et al., 2006;Emri et al., 2005a;Nitsche et al.,
2012;Shin et al., 2009). For example, a low pH during carbon starvation may inhibit
cell wall hydrolysis in A. niger, resulting in the formation of a network of empty hyphal compartments (Nitsche et al., 2012). In A. nidulans, apoptosis can take place
in mutants that lack an autolytic phenotype (Emri et al., 2005a).
Chapter 1
Fig. 3. Morphological changes taking place during generation of spore forming structures and
spores. Figure modified with permission from (Etxebeste et al., 2010).
activate upstream developmental activators, which work as an intertwined network
that regulates secondary metabolism, autolysis and the initiation of early condiophore formation. As deletion of these genes results in a non-sporulating, fluffy phenotype with undifferentiated vegetative hyphae, they are designated ‘fluffy’ genes.
FluG produces an extracellular signal, accumulation of which is thought to signal
that hyphae grow into the atmosphere and form aerial mycelium (Lee & Adams,
1994b;Lee & Adams, 1996). Signal accumulation results in activation of the regulatory pathway composed of the ‘fluffy’ genes, which ultimately activates the conidiation specific pathway governed by brlA (Seo et al., 2006). As reviewed in (Adams et
al., 1998;Etxebeste et al., 2010;Krijgsheld et al., 2013), brlA expression is activated
when the conidiophore stalk is elongated. BrlA is important in further conidiophore
development, as well as for activation of transcriptional activators AbaA and WetA.
BrlA and WetA activate downstream genes that are involved in early development
such as hydrophobins and pigment production. In addition to the ‘fluffy’ genes the
22
Introduction
Extensive morphological changes are taking place during sporulation, and the fungal cell wall needs to be adapted to enable these changes. This can be observed by
changes in appearance (Adams et al., 1998), as well as by the differences in carbohydrate composition of vegetative mycelium and spores (Maubon et al., 2006).
The cell wall consists mainly of carbohydrates, and CAZymes build and remodel
the cell wall during sporulation. Sporulation specific chitin synthases (Fujiwara et
al., 2000;Lee et al., 2004) have been identified that contribute to these cell wall
changes. However, it is likely that other CAZymes also change the cell wall during
sporulation in macro-colonies on plates and/or during submerged fermentations.
In this thesis we identified such a set of genes and characterized two sporulation
specific chitinases in detail (Chapter 2, 4).
Scope of this thesis
Fungal aging and stress conditions such as nutrient starvation, can lead to the initiation of sporulation and autolysis. To adapt the fungal cell wall during the struggle to
survive when nutrients become limiting, carbohydrate-active enzymes modify the cell
wall by acting on polysaccharides such as chitin, α-glucans and β-glucans. Although
it is recognized that several CAZymes play a role in remodeling the cell wall during
starvation and sporulation, detailed insights into the roles these enzymes play is far
from complete. This thesis focuses on the identity, biochemical activity and physiological roles of glycoside hydrolases during carbon starvation and sporulation.
Chapter two provides a systematic overview of the carbohydrate-active enzymes
produced by A. niger during prolonged carbon starvation, based on a detailed analysis of its transcriptome, proteome and enzyme activities. To differentiate between
genes and enzymes induced by starvation and those induced by conidiation, the
expression profiles of glycoside hydrolases in wild-type and strains carrying a deletion of the developmental regulators brlA and flbA are compared.
23
Chapter 1
Chapter 1
flbA is required for initiation of sporulation. FlbA is a regulator of G-protein signaling
that blocks vegetative growth by inhibiting heterotrimeric G-protein signaling. This
block is required to initiate conidiation (Seo et al., 2005;Yu, 2006). This regulatory
network is also linked to autolysis in A. nidulans, as for example the fluG deletion
strain shows reduced autolysis and flbA deletion results in a hyper-autolytic phenotype (Emri et al., 2005b;Lee & Adams, 1994a;Wieser et al., 1994).
Chapter 1
Chapter three focuses on the characterization of two glycoside hydrolases expressed strongly upon carbon starvation, AgnB and CfcA. AgnB was found to be
an alpha-1,3-glucanase which releases glucose from alpha-1,3-glucan substrates.
CfcA is an exo-chitinase, cleaving (GlcNAc)2 from the non-reducing end of chitin.
CfcA is linked to recycling of the cell wall during autolysis.
In chapter four, evidence is presented for the physiological role of chitinases CfcI
and CtcB in sporulation. Transcriptome analysis shows that these chitinases, together with other glycoside hydrolases, are strongly induced in sporulating aerial
mycelium compared to vegetative mycelium. Using promoter fusion strains, activities of the promoters of cfcI and ctcB were located to the conidophores. Deletion of
both genes resulted in accumulation of chitin in the spore cell wall, indicating both
enzymes have a function in modification of the cell wall during sporulation.
Chapter five focuses on the biochemical characterization of one of these sporulation specific chitinases. Chitinase CfcI belongs to a subgroup of glycoside hydrolase
family 18 of which until now no enzymes were characterized. CfcI is capable of hydrolysing chitotriose and longer chitin oligosaccharides by cleaving off monomers,
possibly in a processive mode. The enzyme acts on the reducing end of the oligosaccharide substrates.
In chapter six, the development and validation of an analytical method using
HPAEC-PAD for the separation and quantification of native chitin oligosaccharides
is described. To show the suitability of the method in chitinase activity assays, the
HPAEC-PAD method is used to determine the kinetic parameters of chitinase CfcI.
In addition, the function of the CBM18 inserted in the catalytic domain of CfcI is
assessed by comparing the kinetic parameters of CfcI with those of a CfcI mutant
where key aromatic acids in the CBM18 domain were mutated.
The results are summarized and discussed in Chapter seven.
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