Biochemical Society Focused Meetings
Enzymology and ecology of the nitrogen
cycle 15—17 September 2010 University of Birmingham ,
UK
Nitric oxide
and nitrosative stress tolerance in yeast
Anna Tillmann*1, Neil A.R. Gow*
and Alistair J.P. Brown*2
Aberdeen
Fungal Group, School of Medical Sciences, Institute of Medical Sciences,
University of Aberdeen, Foresterhill, Aberdeen AB25 2ZD, UK
Key words: fungal stress
response, flavohaemoglobin, nitric oxide, reactive nitrogen intermediates.
Abbreviations used: GSNO, S-nitrosoglutathione; ROS, reactive oxygen species; RNI,
reactive nitrogen intermediates; NO, nitric oxide; FAD, flavin adenine dinucleotide; NADP, Nicotinamide adenine dinucleotide
phosphate;
1. Presenting author
2. To whom correspondence should be addressed.
(al.brown@abdn.ac.uk)
Abstract
The
opportunistic human fungal pathogen Candida
albicans encounters diverse environmental stresses when it is in contact
with its host. When colonising and invading human tissues C. albicans is exposed to reactive oxygen (ROS) and reactive
nitrogen intermediates (RNI). ROS and RNI are generated in the first line of
host defence by phagocytic cells such as macrophages and neutrophils. In order
to escape these host-induced oxidative and nitrosative stresses C. albicans has developed various
detoxification mechanisms. One such mechanism is the detoxification of nitric
oxide (NO) to nitrate by the flavohaemoglobin enzyme, CaYhb1. Members of the
haemoglobin superfamily are highly conserved and are found in archaea,
eukaryotes, and bacteria. Flavohemoglobins have a dioxygenase activity (NOD)
and contain three domains: a globin domain, an FAD-binding domain, and an
NAD(P)-binding domain. Here we examine the nitrosative stress response in three
fungal models: the pathogenic yeast C.
albicans, the benign budding yeast Saccharomyces
cerevisiae, and the benign fission yeast Schizosaccharomyces pombe.
We compare their enzymatic and non-enzymatic NO and RNI detoxification
mechanisms and summarise fungal responses to nitrosative stress.
Why study nitrosative stress responses in yeasts?
The
evolutionarily divergent yeasts Candida
albicans, Saccharomyces
cerevisiae and
Schizosaccharomyces
pombe provide ideal model systems in which
to compare nitrosative stress responses. These three yeasts, which diverged about 500 million years ago [1], exist in different
environmental niches and therefore have been exposed to different evolutionary
pressures. As a major fungal pathogen of humans, C. albicans has evolved robust stress responses that facilitate adaptation
to environmental challenges such as changes in ambient pH, osmolartity and
nutrient availability, as well as exposure to ROS and RNI [2] (the latter
challenges being of particular interest in our laboratory). These unicellular yeasts have short life
cycles, they can be grown on defined experimental conditions, their genomes
have been sequenced [3]. Furthermore extensive
molecular toolboxes that have facilitated the dissection of fundamental cellular
processes such as the cell cycle, signal transduction and stress responses [4-6]. The ability to
survive these stresses contributes to the pathogenicity of C. albicans as well as virulence factors such as adhesins,
extracellular hydrolytic enzymes and phenotypic switching [7-9]. In contrast, the benign yeasts, S. cerevisiae and S. pombe, which are associated with environmental niches, tend to
be more sensitive to stresses than C. albicans
[10].
Nitric
oxide, RNI and their impact within the cell
Nitric oxide is
an ‘ancient’ molecule and nitric oxide and its derivates were oxidizing
substrates in the archaeal world, driving the evolution of a pathway related to
modern dissimilatory-denitrification [1], It has been suggested that aerobic respiration
has emerged from this pathway by adaptation of the enzyme NO reductase to its
new substrate, oxygen [11]. Nitric oxide is a gaseous
radical that can have beneficial or unfavourable effects within cells depending
on the concentration. At low concentrations NO can act as a second messenger
controlling numerous physiological processes in animal cells [12]. At high concentrations NO is
cytotoxic and is exploited as a weapon in host-pathogen defences [12]. As mentioned above, fungal
pathogens are relatively resistant to such stresses, and it is likely that the
ability of pathogenic fungi to combat host-pathogen defences evolved through ancient
interactions between fungi and phagocytic amoeba [13].
Nitrosative stress is mainly caused
by three forms of NO: the nitric oxide radical, the nitrosonium
cation and the nitroxyl anion. The NO
radical is a signalling molecule
that plays a regulatory role in cell proliferation, antimicrobial defence and
inflammatory responses [14-17].
Within the cell
NO reacts with oxygen species, with
thiol-containing proteins and with metalloproteins [18]. The
NO radical also reacts with oxygen to generate nitrogen dioxide which is
converted to the nitrite anion and further to the nitrate anion. Intermediates
of this oxidation include dinitrogen trioxide and the nitrite anion which
contributes to the nitric oxide toxicity by oxidising thiols and amines within
the cell. Due to its stability the nitrate anion is thought to be the end
metabolite of this NO pathway [19]. The nitrosonium cation is generated when one electron of NO is
released. In this reaction, the iron atom of Fe3+ containing
metalloproteins acts as the electron acceptor. The Fe2+-NO+
complex serves as a NO carrier which releases NO at specific target sites.
Additionally the nitrosonium cation reacts with nucleophilic centres and is
responsible for nitrosation generating nitroso-compounds including
nitrosamines, alkyl or aryl nitrite and S-nitrosothiols [20]. It has been proposed that NO is stored and carried
as S-nitrosoglutathione (GSNO), and that GNSO is used as an NO pool within
cells [21]. The nitroxyl anion is generated when one
electron is added to NO. This reduction is supported by the Fe2+ ion
and by Fe2+ containing metalloproteins which act as electron donors.
The nitroxyl anion is believed to mediate sulfhydryl oxidation of target
proteins. This process leads to the formation of nitrous oxide which is also
the result of nitroxyl
anion dimerisation [20].
In mammalian cells NO
biosynthesis is catalysed by three isoforms of NO synthase (NOS): the inducible (iNOS), the constitutive neuronal (nNOS) and endothelial isoforms (eNOS). All nitric oxide synthases use L-arginine and
NADPH to generate NO and citrulline [22]. As
mentioned above, macrophages that have taken up microbial cells release RNS and
RNI into the phagolysosome. Macrophages can produce up to 57 μM nitric oxide
and up to 14 mM of hydrogen peroxide [23]. ROS
such as superoxide anions (O2.-) and hydrogen peroxide (H2O2)
are generated with the help of NADPH oxidase as by-products of the respiratory
chain [24]. Furthermore the superoxide anion can also be converted with the help
of the myeloperoxidase to hypochlorous acid (HClO). Parallel to the production
of ROS, macrophages generate nitric oxide and nitrite with the help of iNOS.
Furthermore NO reacts with the superoxide anion to create the strong oxidant
peroxynitrite (ONOO-) which has fungicidal activity and is more
stable and a stronger oxidant than NO [12].
Due to the physical and chemical properties of NO it is more accurate to
imagine dynamic, temporary and local NO gradients within the cells. Hence NO
has a short-half life which varies depending on the intracellular and
extracellular redox state [25], the NO concentration, the
partial oxygen pressure, the presence of bivalent metals and thiol groups [12].
Nitrosative
stress responses in the model yeasts
C. albicans is exposed to NO and RNI, which are
generated during host-defence by phagocytic cells, and to non-enzymatically
generated NO from nitrates and nitrites of dietary products in the digestive
system [26].
Alternatively, NO can be generated by bacteria in the oral cavity or gut
[27-28]. On the other hand S. cerevisiae is exposed to endogenous NO
under hypoxic conditions, since the mitochondrial respiratory chain of S. cerevisiae can use endogenous nitrite
instead of oxygen as an electron acceptor, thereby generating NO within the
cell [29]. Several mechanisms exist to counteract these nitrosative stresses:
(1) the active detoxification of NO via flavohemoglobins; (2) the antioxidant
system for scavenging NO via GSH or trehalose; and (3) the up-regulation of
repair systems to counteract the caused damage. The systems that repair RNI
damage are poorly understood in yeasts [30].
Figure
1: Simplified RNI response network in yeasts.
Yeasts
cope with RNI stresses in different ways: they can enzymatically detoxify NO
via flavohemoglobins, they can scavenge NO through antioxidant systems or they
can repair the caused damage.
A number of antioxidant systems
contribute to nitrosative stress resistance, one of which is S-nitrosoglutathione reductase (GSNO reductase). Interestingly, compared with S. cerevisiae, S. pombe is particularly
sensitive to low concentrations of GSNO [31]. This might be due
to the inactivation of S. pombe GSNO
reductase by peroxynitrite since GSNO reductase activity is essentially required for the growth of S.
pombe, unlike in S. cerevisiae [32-33]. This observation
emphasises the importance of repair functions such as GSNO reductase that are
capable of reducing GSNO to ammonia and glutathione disulfide (GSSG)
[34]. However, other enzymes
such as thioredoxin
peroxidase (Tsa1) contribute to resistance to
endogenous RNI, and Tsa1 has also been shown to contribute to fungal virulence [30]. In addition, several
non-enzymatic antioxidants help to counteract the effects of RNI in yeasts,
such as GSH and metalloporphyrins. For C.
albicans the antioxidant trehalose is essentially linked to stress
adaptation [35]. The non-enzymatic antioxidant
systems and NO scavenger mechanisms are thought to have evolved a long time ago
when cells first were exposed to an aerobic environment. Since then gene
duplication events and the redundancy of stress resistance pathways and
antioxidant systems have facilitated the environmental adaptation of different
yeast species.
Flavohemoglobins are characterised by an NO
dioxygenase domain (NOD) which is highly conserved in bacteria and yeast and
converts nitric oxide to nitrate [36]. As the name suggests, flavohemoglobins contain a N-terminal heme group
followed by the C-terminal FAD (flavin adenine dinucleotide) domain and a
NAD(P) domain
[37]. S. cerevisiae and S. pombe
each have a single flavohemoglobin gene: ScYHB1
and the predicted SPAC869.02c
respectively (see Figure 1) [38]. In
contrast,
the C. albicans genome contains three
flavohemoglobin-like genes, namely CaYHB1,
CaYHB4 and CaYHB5 [39]. The sequence identity between ScYhb1 and the three C. albicans
flavohemoglobins ranges from 31% to 25% [40].
Figure 2: Flavohemoglobins of S. cerevisiae (ScYHB1), C. albicans (CaYHB1) and S. pombe (SPAC869.02c).
Flavohemoglobins consist of
three highly conserved domains an N-terminal globin domain (HEME), followed by
a FAD-binding domain (FAD) and an C-terminal NAD(P)-binding domain (NAD).
The flavohemoglobins in S. cerevisiae and S. pombe
are fully functional and the deletion of ScYHB1
leads to growth inhibition and the loss of the NOD function in S. cerevisiae [40]. In S. cerevisiae the ScYhb1 protein
is translocated to the mitochondria under hypoxic conditions where it
detoxifies NO [41]. This suggests
that flavohemoglobins are able to both protect yeasts against external as well
as internal sources of NO and RNI. In C. albicans only the deletion of CaYHB1 deletion attenuates virulence
slightly [39, 42]. Inactivation
of CaYHB4 or CaYHB5 did not inhibit NO
consumption under the experimental conditions tested or attenuate virulence in
the mouse model of systemic candidiasis [39], but this does not exclude the possibility that these gene products are
important under other conditions or at specific stages of infection.
These differences in YHB gene copy number and flavohemoglobin functionality might relate
to the different environmental niches of these yeasts and thus their individual
adaptation requirements. In vivo, CaYHB1 is expressed in C. albicans cells on epithelial surfaces
during oral infection [43] and in
cells infecting the mouse gastrointestinal tract [44]. However, CaYHB1 is
not up-regulated in deep tissue infections of liver, for example [45].
It is not clear how yeasts detect NO and which signalling
pathways mediate NO and RNI responses. In contrast to mammalian cells, yeasts do
not express an obvious NO receptor. However Chiranand et al. [46] recently found that in C. albicans, CaYHB1
expression is activated by the regulator CaCta4. By mutating the regulatory
region of CaYHB1 they identified a
nitric oxide-responsive element (NORE) which is crucial for CaYHB1 gene regulation in response to NO.
Once this NORE promoter element was identified, CaCta4 (a Zn(II)2-Cys6 transcription factor) was
then shown to bind directly to NORE.
Furthermore, Chiranand and coworkers demonstrated that inactivation of CaCTA4 inhibits CaYHB1 induction in response to NO [47]. Moreover C. albicans Dcta4 null mutant display attenuated virulence in the mouse model
of systemic candidiasis, reinforcing the idea that robust nitrosative stress
responses contribute to the pathogenicity of C. albicans. CaCTA also
up-regulates a putative sulphite transporter gene (CaSSU1) in response to RNI. Interestingly C. albicans Dssu1 cells are not sensitive to NO, unlike the situation in S. cerevisiae where SSU1 mediates NO resistance under certain environmental conditions [47].
Comparisons of NO-induced genes in S. cerevisiae and C. albicans are intriguing [42, 48]. For
instance catalase and iron
acquisition genes are up-regulated in both species. However, as illustrated by the case of SSU1, even where apparent orthologues
are highly expressed in both S.
cerevisiae and C. albicans, the
molecular activities and responses appear to be specific for each yeast species
and might only be explainable by their evolutionary adaptation to their
environmental requirements [46]. Furthermore, the transcription
factors that regulate the nitrosative stress responses in these yeasts are even
more divergent. The closest homologue to CaCTA4
in S. cerevisiae is ScOAF1 [49], an oleate receptor. The next
closest homologue of CaCTA4 in S. cerevisiae is ScHAP1 [50], a heme-responsive
transcription factor. Neither ScOAF1 nor
ScHAP1 appears to be involved in
nitrosative stress response in S.
cerevisiae. Instead, Sarver and DeRisi have shown that the C2H2
zinc finger transcription factor ScFZF1
is involved in NO sensing in S.
cerevisiae [46]. In S. pombe, the AP-1-like bZIP
transcription factor, SpPap1, regulates nitrosative as well as oxidative and
nutritional stress responses [51]. The orthologue of SpPap1, ScYap1, regulates the oxidative stress
response in S. cerevisiae. These observations illustrate the functional
reassignment of transcription factors across these evolutionarily divergent
yeasts, an observation that also holds between S. cerevisiae and C. albicans
[52-53].
Conclusions
and future perspectives
Our
understanding of nitrosative stress pathways in most organisms is rudimentary
at best, and much work remains to be done to elucidate fungal nitrosative
stress response mechanisms. This cannot be simply done by genome sequence comparisons
because fungi lack obvious homologues of many nitrosative stress functions that
are present in other organisms. Also there has been rewiring of nitrosative
stress regulators across the ascomycetes [54]. Nevertheless it is important to
study NO and RNI defence mechanisms in yeasts because they contribute to fungal
pathogenicity and presumably to the survival of yeasts in other environmental
niches. Understanding the crosstalk between nitrosative and oxidative stress
responses is likely to lead to a better understanding of host-pathogen
interactions and fungal virulence because pathogenic yeasts are exposed to both
ROS and RNI during contact with host immune defences. Finally new antifungal drug
targets might be revealed by a more complete understanding of the biochemical
detoxification pathways of pathogenic fungi.
Acknowledgments
AT is supported
by the CRISP project (Combinatorial Responses In Stress Pathways) funded by the
BBSRC (BB/F00513X/1) under the Systems Approaches to Biological Research (SABR)
Initiative, and by the University of Aberdeen.
AB and NG are also supported by the Wellcome Trust (080088)
and the European Commission (PITN-GA-2008-214004-FINSysB; ERC-2009-AdG-249793;
FP7-ITN-2008-237936-Ariadne).
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