Molecular Microbiology (2011) 79(4), 968–989 䊏
doi:10.1111/j.1365-2958.2010.07503.x
First published online 30 December 2010
Cell wall integrity is linked to mitochondria and phospholipid
homeostasis in Candida albicans through the activity of the
post-transcriptional regulator Ccr4-Pop2
mmi_7503 968..989
Michael J. Dagley,1 Ian E. Gentle,2
Traude H. Beilharz,1 Filomena A. Pettolino,3
Julianne T. Djordjevic,4 Tricia L. Lo,1
Nathalie Uwamahoro,1 Thusitha Rupasinghe,5
Dedreja L. Tull,5 Malcolm McConville,5
Cecile Beaurepaire,6 André Nantel,6,7
Trevor Lithgow,1 Aaron P. Mitchell8 and Ana Traven1*
1
Department of Biochemistry and Molecular Biology,
Monash University, Melbourne, Australia.
2
Department of Biochemistry, La Trobe University,
Melbourne, Australia.
3
School of Botany, University of Melbourne, Melbourne,
Australia.
4
Centre for Infectious Diseases and Microbiology,
University of Sydney at Westmead Hospital, Sydney,
Australia.
5
Metabolomics Australia, The Bio21 Institute and the
University of Melbourne, Melbourne, Australia.
6
Biotechnology Research Institute, National Research
Council of Canada, Montreal, QC, Canada.
7
Department of Anatomy and Cell Biology, McGill
University, Montreal, QC, Canada.
8
Department of Biological Sciences, Carnegie Mellon
University, Pittsburgh, PA, USA.
Summary
The cell wall is essential for viability of fungi and is an
effective drug target in pathogens such as Candida
albicans. The contribution of post-transcriptional
gene regulators to cell wall integrity in C. albicans is
unknown. We show that the C. albicans Ccr4-Pop2
mRNA deadenylase, a regulator of mRNA stability and
translation, is required for cell wall integrity. The ccr4/
pop2 mutants display reduced wall b-glucans and
sensitivity to the echinocandin caspofungin. Moreover, the deadenylase mutants are compromised for
filamentation and virulence. We demonstrate that
defective cell walls in the ccr4/pop2 mutants are
Accepted 4 December, 2010. *For correspondence. E-mail ana.
[email protected]; Tel. (+61) 3 9902 9219; Fax (+61) 3 9905 3726.
© 2010 Blackwell Publishing Ltd
linked to dysfunctional mitochondria and phospholipid imbalance. To further understand mitochondrial
function in cell wall integrity, we screened a Saccharomyces cerevisiae collection of mitochondrial
mutants. We identify several mitochondrial proteins
required for caspofungin tolerance and find a connection between mitochondrial phospholipid homeostasis and caspofungin sensitivity. We focus on the
mitochondrial outer membrane SAM complex subunit
Sam37, demonstrating that it is required for both trafficking of phospholipids between the ER and mitochondria and cell wall integrity. Moreover, in C.
albicans also Sam37 is essential for caspofungin
tolerance. Our study provides the basis for an integrative view of mitochondrial function in fungal cell
wall biogenesis and resistance to echinocandin antifungal drugs.
Introduction
The yeast cell wall is essential for cell integrity and interactions with the environment. For human pathogens such
as Candida albicans, the cell wall governs key functions
including adherence and pathogenicity, and serves as a
pathogen-specific antifungal drug target (Ruiz-Herrera
et al., 2005; Walker et al., 2010).
The yeast cell wall is composed of a basal layer of
chitin, a body of b-glucans (1,3-b-glucan and 1,6-bglucan) and mannosylated proteins that decorate the wall
surface (Ruiz-Herrera et al., 2005; Klis et al., 2006). Cell
wall synthesis and remodelling is a dynamic process,
which responds to growth conditions, cell cycle progression, changes in cell morphology, and cell wall stress
(Staab et al., 1996; Ruiz-Herrera et al., 2005; Klis et al.,
2006; 2009; Bahn et al., 2007; Walker et al., 2008; Côte
et al., 2009). Several signalling pathways act to ensure
cell wall integrity, for example the PKC and HOG MAP
kinase pathways (Paravicini et al., 1996; Eisman et al.,
2006; Munro et al., 2007) and the Ca2+-calcineurin
pathway (Wiederhold et al., 2005; Munro et al., 2007;
Singh et al., 2009). Recent genetic screens with a kinase
mutant library in C. albicans indicate that the cell wall
Cell walls, mitochondria and phospholipids 969
regulatory network may include several additional pathways (Blankenship et al., 2010).
Cell wall biogenesis is disrupted by the echinocandin
antifungal drugs, which inhibit 1,3-b-glucan synthase
(Douglas et al., 1997). Although the echinocandins have
only recently been introduced into clinical practice (in
2001), already resistant clinical isolates with mutations in
the echinocandin target FKS1 have been identified,
raising concerns about emergence of drug resistance
(Perlin, 2007; Walker et al., 2010). Identifying pathways
governing cell wall integrity and tolerance to echinocandins is of critical importance for designing more efficient
antifungal therapies. These approaches have the potential to provide avenues for combinatorial therapy (Walker
et al., 2008; Sandowsky-Losica et al., 2008; Singh et al.,
2009; Walker et al., 2010).
The cell wall integrity pathways commonly regulate
the expression of genes required for cell wall biogenesis
and repair. Consequently, studies in C. albicans, as
well as in the model yeast Saccharomyces cerevisiae,
have focused on deciphering the roles of transcription
factors in cell wall biogenesis (García et al., 2004; Bruno
et al., 2006; Rauceo et al., 2008). Much less is known
about post-transcriptional gene expression in cell wall
integrity. In C. albicans post-transcriptional gene expression regulators have not been studied at all. In S. cerevisiae, mutants in some post-transcriptional regulators
have phenotypes consistent with a cell wall integrity
defect (Hata et al., 1998; Kaeberlein and Guarente,
2002; Markovich et al., 2004). For example, S. cerevisiae mutants in the Ccr4-Pop2-NOT mRNA deadenylase, an exonuclease that shortens mRNA poly(A) tails
to control mRNA decay and translation, are sensitive to
agents that perturb cell walls (Hata et al., 1998; Markovich et al., 2004). The nature of the cell wall integrity
defects in the S. cerevisiae ccr4/pop2 mutants, as well
as the molecular basis for these phenotypes, are
unknown.
Here we performed the first study of post-transcriptional
regulators in cell wall integrity in C. albicans and show
that the Ccr4-Pop2 mRNA deadenylase is required for
cell wall integrity, tolerance of the echinocandin caspofungin and virulence in the mouse model. Moreover,
we provide evidence that the cell wall defect of the
deadenylase mutants is linked to dysfunctional mitochondria and defective phosholipid homeostasis. To broaden
our understanding of the roles of mitochondria in
fungal cell wall biogenesis, we screened a collection
of mitochondrial morphology mutants in S. cerevisiae,
identifying several new genes required for tolerance
of cell wall inhibitors, and demonstrating that proteins, which have roles in mitochondrial phospholipid
homeostasis, have prominent roles in caspofungin
tolerance.
Results
The Ccr4-Pop2 mRNA deadenylase is required for cell
wall biogenesis in C. albicans
To address whether the Ccr4-Pop2 mRNA deadenylase is
required for cell wall biogenesis in C. albicans, we constructed homozygous deletion mutants in the genes
encoding the Ccr4 and Pop2 proteins. BLAST searches
were used to identify the orthologues of Ccr4 and Pop2 in
the C. albicans genome. Orf19.5101 and orf19.5734 show
high homology to S. cerevisiae Ccr4 and Pop2 respectively (43% identity for Ccr4, and 32.8% identity for Pop2),
and the C. albicans proteins contain the characteristic
Ccr4 and Pop2 domains: the leucine rich repeat LRR and
C-terminal exonuclease domain in Ccr4, and the RNaseD
Caf1 domain in Pop2 (Fig. 1A).
The C. albicans ccr4DD and pop2DD mutants were highly
sensitive to the cell wall perturbing agents Congo red and
the echinocandin caspofungin, and were moderately sensitive to the chitin-binding dye Calcofluor white (Fig. 1B).
These phenotypes were observed in two independently
constructed homozygous mutants for each of the genes
and could be complemented by re-introduction of the
CCR4 or POP2 genes into the mutant genomes (Fig. 1B).
Moreover, caspofungin sensitivity of the deadenylase
mutants could be rescued by addition of the osmostabilizer
sorbitol (Fig. 1B, right panel; of note, the mutants grew
slightly slower even in the presence of sorbitol at higher
doses of caspofungin. This could be because of additional
effects of Ccr4 and Pop2 or due to an inhibitory combinatorial effect of cell wall stress and higher osmolarity).
Consistent with a cell wall defect, the ccr4DD and
pop2DD mutants flocculated, microscopy showed that the
cells from mutant cultures were rounder, formed clumps,
lysed frequently and were somewhat bigger, and the
mutants hyper-activated the PKC-dependent cell wall
integrity pathway in response to treatment with Calcofluor
white (Fig. S1).
To further characterize the cell wall defect in the ccr4DD
and pop2DD mutants, we analysed the glycan composition of cell walls in wild-type, mutant and complemented
strains (Fig. 1C and Table S1). The ccr4DD and pop2DD
mutants displayed lower levels of 1,3-b-glucans and 1,6b-glucans in their walls. The levels of mannan were higher
in the mutants, likely reflecting a compensatory response.
Chitin levels were also on average somewhat lower in the
mutants (Fig. 1C and Table S1), although this was not
statistically significant in the pop2DD strain.
The Ccr4-Pop2 mRNA deadenylase is required for
wild-type filamentous growth and virulence
The ability to exist in different morphological forms, as
ovoid yeast-form cells and as elongated filaments, is
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
970 M. J. Dagley et al. 䊏
Fig. 1. The Ccr4-Pop2 mRNA deadenylase
is required for cell wall integrity in C. albicans.
A. Schematic representation of the domain
structure of the Ccr4 and Pop2 exonucleases
from C. albicans.
B. Ten-fold serial dilutions of the indicated
strains were dropped on YPD plates with or
without Congo red, Calcofluor white and
caspofungin at the indicated doses. The
plates were incubated at 30°C for three days
and photographed. For the mutant strains in
the left panel, the original homozygous
deletant (URA3+ ARG4+ his1-, lane 2) was
tested, alongside two colonies of the mutants
complemented with vector only (which
restored histidine prototrophy, lanes 3 and 4)
or with a wild-type copy of the CCR4 or POP2
genes (lanes 5 and 6). The wild-type strain
was DAY185 (See Table S3). The same
strains were also tested on caspofungin
plates, except that the original homozygous
deletants were omitted (therefore all strains
are URA3+ ARG4+ HIS1+).
C. Cell walls were isolated as described in
Experimental procedures and the glycan
composition of the walls was determined by
mass spectrometry. Shown are averages of
three independent experiments performed
with two technical replicates. For chitin two
independent experiments were performed.
The error bar is the standard error of the
mean. **P < 0.01, *P < 0.05. See also
Table S1.
important for virulence of C. albicans (Lo et al., 1997;
Saville et al., 2003; Carlisle et al., 2009). Changes in the
cell wall accompany the morphological yeast-to-filaments
transition and we tested the ability of the ccr4DD and
pop2DD mutants to differentiate to filamentous form in
response to a variety of inducers.
As shown in Fig. 2A, the ccr4DD and pop2DD mutants
could not filament on 10% serum or Spider plates. The
mutants were able to undergo filamentous differentiation
in liquid cultures in response to 10% serum, albeit with
slower kinetics, possibly due to slower growth rates
(Fig. 2B; unlike in the wild-type, after 1 h in serum many
yeast-form cells were observed in the mutants, but after
2.5 h the mutants were filamentous). In contrast to this, in
Spider media the ccr4DD and pop2DD mutants were fully
defective for filamentous growth, and remained in yeast
form (right panel of Fig. 2B). The ccr4DD and pop2DD
mutants were also defective in differentiating to filamentous form in M199 media and in the presence of
N-acetylglucosamine (Fig. S2).
A defective cell wall and an inability to filament correlate
with reduced virulence of C. albicans (Lo et al., 1997;
Saville et al., 2003; Munro et al., 2005; Norice et al., 2007;
Carlisle et al., 2009). The virulence of the ccr4DD mutant
was compared with that of the complemented ccr4DD/
CCR4 strain using the tail-vein murine infection model of
candidaemia (Fig. 3). Only the ccr4DD mutant was tested
for virulence in the animal model, as the virulence of the
two mutants was not expected to be different: the cell wall
and filamentation phenotypes of the two mutants were
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
Cell walls, mitochondria and phospholipids 971
similar and both genes contribute the same molecular
function (exonuclease activity) to the Ccr4-Pop2-NOT
complex.
In two separate studies, using different inoculation
doses (2 ¥ 105 or 1 ¥ 106 yeast cells), mice infected with
ccr4DD showed no clinical signs of infection over a 20-day
period (Fig. 3A, the experiment using the higher inoculation dose is shown). This is in contrast to mice infected
with ccr4DD/CCR4 that exhibited a higher rate of mortality,
and this was more pronounced at the higher inoculation
dose (Fig. 3A). The reduced virulence of ccr4DD cells
coincided with a statistically significant reduction in
ccr4DD kidney burdens quantified by counting colonyforming units on day 1 (Fig. 3B), and a similar trend was
observed on day 2, although statistical significance could
not be obtained (Fig. 3B). Histopathology results were
fully consistent with kidney burden analysis, demonstrating that on day 1 and day 2 post infection hardly any
ccr4DD cells could be detected in the kidney tissue
(Fig. 3C, the arrows indicate the only mutant cells that
could be found in any of the sections analysed), while
abundant filamentous cells were detected in kidneys from
animals infected with the complemented strain (Fig. 3C).
Collectively, the results indicate that Ccr4 is required for
virulence of C. albicans in mice.
The transcriptome of the C. albicans ccr4DD mutants
indicates dysfunctional mitochondria
Fig. 2. The Ccr4-Pop2 mRNA deadenylase is required for
wild-type filamentous growth.
A. Cells from wild-type (DAY286, see Table S3) or homozygous
deletion mutants were streaked on YPD+10% calf serum or Spider
plates and incubated at 37°C for four days. The colonies on
serum-containing plates were photographed using a digital camera
and the scale bar represents 1 mm. The colonies on Spider plates
were photographed under 10 ¥ magnification, using the Olympus
fluorescent microscope and the scale bar represent 100 mm.
B. Cells from wild-type (DAY185), mutants or complemented strains
(all strains are URA3+ ARG4+ HIS1+) were grown over night in YPD
at 30°C and then diluted into pre-warmed YPD+10% calf serum or
Spider media and incubated at 37°C for the indicated times. The
cells were observed with the 100 ¥ objective. The scale bar is
10 mm.
To understand what cellular processes are affected by the
absence of Ccr4 in C. albicans and uncover pathways that
are linked to cell wall biogenesis, we performed microarray analysis of the transcriptome in ccr4DD strains compared with complemented ccr4DD /CCR4 cells.
In agreement with the fact that Ccr4 is a general regulator of gene expression and consistent with what has
been observed in S. cerevisiae (Cui et al., 2008; Azzouz
et al., 2009), the expression of a relatively large number of
genes was affected in the absence of Ccr4: 362 genes
were up- or downregulated at least 1.9-fold (with a
P-value below 0.05), which represents 5.6% of the
genome (Fig. 4A and Supporting Dataset 1; the complete
list of differentially expressed genes in the ccr4DD mutant
is shown in Supporting Dataset 2). Differential expression
of candidate transcripts was confirmed by qPCR analysis
(Supporting Dataset 3). Semi-quantitative PCR used for
analysis of poly(A) tail lengths of mRNAs also demonstrated good correlation with changes in mRNA levels
observed in the microarray (Fig. 4B).
Among the differentially expressed genes, three functional groups were prominent. First, genes related to mitochondrial biogenesis and function were differentially
expressed in cells lacking Ccr4 (30.5% of the genes that
were either up- or downregulated in ccr4DD mutants were
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
972 M. J. Dagley et al. 䊏
Fig. 3. CCR4 is required for pathogenicity in
a murine intravenous model of candidaemia.
A. Mice were inoculated intravenously with
either ccr4DD or ccr4DD/CCR4 and
euthanized after showing clinical signs of
infection. Mice not showing symptoms of
illness at 20 days post infection were also
euthanized. An estimation of differences in
survival (log-rank test) using the
Kaplan–Meier method was performed and a
P-value of < 0.05 was considered statistically
significant (as indicated by an asterisk ‘*’,
n = 10).
B. Mice were inoculated with the indicated
strains as described above and were
euthanized either on day 1 or day 2 post
inoculation. Kidneys were harvested and
assessed for infection burden by determining
colony-forming units (n ⱖ 3). The asterisk ‘*’
indicates that kidney burdens were statistically
lower in ccr4DD-infected kidneys collected on
day 1 post infection (P < 0.05 one-way
ANOVA), relative to ccr4DD/CCR4-infected
mice. On day 2, a similar trend was observed,
but there was no statistical significance.
C. Kidney histopathology was performed
using PAS-staining as described in
Experimental procedures. C. albicans hyphae
(pinkish purple) were detected in abundance
in ccr4DD/CCR4 and on day 1 and day 2 post
infection, but were rarely observed in
ccr4DD-infected kidneys. Arrows indicate the
only sign of ccr4DD infection detected in any
of the sections analysed. All other ccr4DD
sections analysed looked like the non-infected
control. In contrast, all sections collected from
ccr4DD/CCR4-infected kidneys on day 1 and
day 2 post infection contained abundant
infection foci. The scale bar represents
20 mm.
mitochondria-related in their function, which is 2.3-fold
enrichment over the background, P = 2.05E-15, see Supporting Dataset 1). Second, genes encoding rRNA and
ribosome biogenesis functions were upregulated in
ccr4DD cells. For example, 12.6% of genes in the upregulated group were classified in the GO term ‘rRNA metabolic process’, which is a 4.2-fold enrichment over the
background (P = 1.07E-07). Third, genes related to amino
acid metabolism were downregulated in the absence of
Ccr4 (14% of downregulated genes were classified to the
GO term ‘cellular amino acid metabolic process’, which is
a 4.6-fold enrichment over the background, P = 9.55E-07)
(Fig. 4A and Supporting Dataset 1).
To further understand Ccr4-dependent mRNA poly(A)
tail length control in C. albicans, we analysed the poly(A)
tail lengths on selected transcripts from the upregulated
group using LM-PAT (Ligation-Mediated Polyadenylation
Test) assays. This assay is based on reverse transcription
followed by PCR, with the sizes of the obtained products
reflecting the distribution of the mRNA poly(A) tail lengths
(see also Experimental procedures). In S. cerevisiae and
S. pombe, mRNAs display distinct poly(A) tail length at
steady-state levels in the wild-type, fractionating into ‘longtailed’ and ‘short-tailed’ groups (Beilharz and Preiss, 2007;
Lackner et al., 2007). The ‘short-tailed’ transcripts are
deadenylated in a gene-specific manner by Ccr4 (with the
alternative yeast deadenylase Pan2 playing only a minor
role), and this likely serves to achieve translational inhibition (Beilharz and Preiss, 2007). In C. albicans, mRNAs
were also preferentially ‘short-’ or ‘long-tailed’ in the wildtype (Fig. 4B, see also Fig. S3 for quantification of the
“short-” and “long-tailed” forms). The mRNAs encoding
cyclin Cln3 and the lipid metabolism protein Taz1 were
‘short-tailed’, whereas the mRNAs encoding the rRNA and
ribosome biogenesis proteins Pop3 and Hbr3, the phospholipid biosynthesis enzymes Ino1 and Cho2, the mitochondrial proteins Tim9 and Mrpl10, the adhesin Als1 and
Phr1, a protein required for cell wall biogenesis, were
‘long-tailed’ (Fig. 4B). In the ccr4DD and pop2DD mutants
the mRNAs stabilized with longer tails (Fig. 4B), with the
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
Cell walls, mitochondria and phospholipids 973
Fig. 4. Transcriptome analysis in the mRNA
deadenylase mutants.
A. For microarray analysis, ccr4DD mutants
were compared with ccr4DD/CCR4
complemented strains (both strains are
URA3+ ARG4+ HIS1+). Shown are selected
gene groups up- or downregulated in the
ccr4DD mutant at least 1.9-fold. The gene
functions were obtained from the Candida
Genome Database
(http://www.candidagenome.org) and from
literature searches.
B. mRNA poly(A) tail analysis was performed
using the LM-PAT assay as described in
Experimental procedures. TVN-PAT samples
were obtained using a reverse primer which
binds to the junction of the 3′ UTR and the
poly(A) tail and thus represent the shortest
poly(A) tails detected with this assay. The
TVN-PAT samples were used as controls for
comparison to the tail lengths in wild-type,
mutants and complemented strains. To
determine whether an mRNA presents with
preferentially short or long poly(A) tails, the
distribution of the poly(A) tail lengths for the
indicated genes was quantified in the
ccr4DD/CCR4 complemented strain- (see
Fig. S3). The ratio of long versus short tails
was 0.6 and 0.4 for CLN3 and TAZ1,
respectively, indicating these mRNAs are
‘short-tailed’, whereas the other mRNAs
presented with preferentially long poly(A) tails
and the ratios were as follows: 2.4 for POP3,
2.5 for HBR3, 1.6 for INO1, 1.5 for CHO2, 2.7
for TIM9, 1.5 for MRL10, 2.5 for PHR1 and
1.6 for ALS1 (see also Fig. S3). The asterisk
‘*’ indicates alternate 3′ untranslated regions
(UTR) usage in the HBR3 transcript.
tail sizes comparable to that observed in S. cerevisiae
(Beilharz and Preiss, 2007; Traven et al., 2009). We also
noticed that, similar to what we observed in S. cerevisiae
(Traven et al., 2009), for some genes such as CHO2, the
distribution of mRNA poly(A) tail lengths somewhat differed
between the ccr4DD and pop2DD mutants, with more
“shorter-tailed” transcripts present in the pop2DD cells
(Fig. 4B). In conclusion, these data demonstrate that these
Ccr4-Pop2 represent the major cytoplasmic deadenylase
in C. albicans and that in C. albicans also the ‘short-tailed’
transcripts are deadenylated by Ccr4-Pop2.
In terms of physiological changes in the absence of
Ccr4 activity, the strongest conclusion that we could make
from the transcriptome analysis was that the mutants
appeared to have dysfunctional mitochondria. This conclusion is based on the fact that the ccr4DD mutant
transcriptome showed upregulation of mitochondrial biogenesis, which is known to occur in response to mitochondrial dysfunction in yeast (Traven et al., 2001). For
example, genes encoding mitochondrial ribosomal subunits, mitochondrial proteins required for expression and
maintenance of the mitochondrial genome, as well as
some subunits of the mitochondrial protein import apparatus, were all upregulated in ccr4DD cells (Fig. 4A and
Supporting Dataset 1).
To further address the functional status of the mitochondria in ccr4DD and pop2DD mutants, we assayed their
growth in the presence of the non-fermentable carbon
source glycerol, as well as assessing mitochondrial morphology upon staining with MitoTracker red. The ccr4DD
and pop2DD mutants were able to grow on glycerol, both at
30°C and 37°C (Fig. 5A), showing that mitochondrial respiration is not majorly compromised; this might reflect
successful compensatory changes assisting mitochondrial
biogenesis. However, mitochondrial morphology was compromised in ccr4DD and pop2DD mutants. Wild-type cells
have an extensive tubular network of mitochondria, but in
both mutants the tubules were shorter and extended less
from the periphery into the cell centre (Fig. 5B).
The roles of mitochondria in cell wall integrity in yeast
The link between mitochondria and cell wall integrity in
fungi is largely unexplored. To our knowledge, only one
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
974 M. J. Dagley et al. 䊏
Fig. 5. The mRNA deadenylase mutants display altered
mitochondrial morphology.
A. Cells from the indicated strains were streaked on either
fermentable (glucose) or non-fermentable (glycerol) carbon source
containing plates and grown for 3 days at 30 or 37°C.
B. Mitochondria were strained with MitoTracker red and viewed by
confocal microscopy. The scale bar represents 5 mm.
gene from S. cerevisiae, PGS1 encoding the mitochondrial phosphatidylglycerol (PG) phosphate synthase, has
been firmly linked with cell wall biogenesis (Zhong et al.,
2005; 2007).
In order to understand if dysfunctional mitochondria
might be linked to defective cell walls in the C. albicans
ccr4/pop2 mutants, and what the underlying mechanism
might be, we took advantage of the availability of the S.
cerevisiae deletion mutant collection to further probe
these connections. Because the C. albicans ccr4/pop2
have mitochondrial morphology defects, we focused on
genes known to affect mitochondrial morphology in S.
cerevisiae (Dimmer et al., 2002; Meisinger et al., 2004;
Altmann and Westermann, 2005; Okamoto and Shaw,
2005; Sesaki et al., 2006; Tamura et al., 2009) (Table 1).
In addition to those mutants, we screened the cox11D
mutant defective in mitochondrial respiration. As a readout in the screen we assayed sensitivity to the antifungal
drug caspofungin, using agar serial dilution tests and
dose–response sensitivity assays in liquid media (see
Experimental procedures).
We found that a large number of mitochondrial morphology mutants were sensitive to caspofungin, indicating that
mitochondrial function generally contributes to caspofungin tolerance (Table 1, Table S2 and Fig. S4). However,
the degree of sensitivity differed markedly between
strains, with some mutants showing sensitivity only at
high doses (indicated as ‘+/-’ for mildly sensitive in
Table 1) and others showing more pronounced sensitivity
(strong sensitivity is indicated with ‘++’ and moderate sensitivity with ‘+’ in Table 1, see also Table S2 and Fig. S4).
The genes that we found to be strongly or moderately
required for tolerance of caspofungin are SAM37 encoding a subunit of the mitochondrial outer membrane SAM
(Sorting and Assembly Machinery) complex involved in
outer membrane biogenesis (Wiedemann et al., 2003),
MDM10 which encodes a subunit shared between the
SAM and the ERMES (ER–Mitochondria Encounter
Structure) complexes (ERMES is involved in physically
bridging the endoplasmic reticulum (ER) and mitochondrial membrane systems) (Meisinger et al., 2004; Kornmann et al., 2009), MDM31 encoding a mitochondrial
inner membrane protein, which displays genetic interactions with ERMES subunits (Dimmer et al., 2005), the
F-box protein genes MDM30 and MFB1 (Dürr et al.,
2006), MDM35 encoding a mitochondrial intermembrane
space protein involved in mitochondrial phospholipid
homeostasis (Osman et al., 2009), UPS2 that interacts
functionally and physically with MDM35 (Osman et al.,
2009; Tamura et al., 2009; 2010; Potting et al., 2010), the
mitochondrial fusion gene UGO1 (Okamoto and Shaw,
2005), the iron-sulphur cluster assembly chaperone
SSQ1, the mitochondrial rhomboid protease MDM37, as
well as genes which encode non-mitochondrial proteins
with likely indirect roles in mitochondrial morphology
establishment (Dimmer et al., 2002), MDM39, PTC1,
VPS45, NUP170, RIM9, CDC73 and REF2 (Table 1).
Next we tested the mutants for sensitivity to another cell
wall damaging drug, the chitin-binding dye Calcofluor
white. Out of the 40 mutants tested, seven were at least
moderately sensitive to both caspofungin and Calcofluor
white: sam37D, mdm10D, mdm35D, mdm39D, ref2D,
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
Cell walls, mitochondria and phospholipids 975
Table 1. Screen of the S. cerevisiae collection of mitochondrial morphology mutants for cell wall integrity phenotypes.
Gene
Function
Caspofungin
Calcofluor
white
Fluconazole
SAM37
MDM10
MDM12
MMM1
MDM34
TOM7
MDM31
FZO1
MGM1
UGO1
MDM30
MDM37
MFB1
DNM1
SAM complex
ERMES and SAM complexes
ERMES complex
ERMES complex
ERMES complex
TOM complex
IM protein, Interacts genetically with ERMES genes
Mitochondrial fusion, GTPase
Mitochondrial fusion, GTPase
Mitochondrial fusion, OM protein
Mitochondrial fusion, F box protein
Mitochondrial rhomboid protease, processing of Mgm1
Mitochondrial morphology, F box protein
Mitochondrial fission, Dynamin GTPase, Interacts with Fis1 and
Mdv1
Mitochondrial fission, Interacts with Dnm1 and Mdv1
Mitochondrial fission, Interacts with Fis1 and Dnm1
Mitochondrial fission, possibly involved in interactions with
Dnm1 and Num1
Mitochondrial fission, cell cortex protein
IMS protein, control mitochondrial phospholipid levels, Interacts
with Ups1 and Ups2
IMS protein, controls CL levels in mitochondria
IMS protein, controls CL and PE levels in mitochondria
ER GET complex subunit, insertion of proteins into ER
membrane
IM protein, possible involved in fusion of inner membranes
IM protein
OM GTPase
Mitochondrial transmission to bud, Intermediate filament protein
Mitochondrial membrane protein, function unknown
Component of translation initiation factor eIF3
RNA binding protein, 3′ end of mRNA maturation
Type 2C protein phosphatase
Component of the PAF1 complex, gene expression
Subunit of the NatB N-terminal protein acetylase
Vacuolar protein sorting
Inositol hexakisphosphate (IP6) and inositol heptakisphosphate
(IP7) kinase
Subunit of the nuclear pore complex
Proteolytic activation of Rim101
Mitochondrial DEAD-box RNA helicase
Chaperone, Fe-S cluster assembly
Cytochrome C oxidase activity
Phosphatidylserine decarboxylase
++
++
+/+/+/++
+/+/++
++
+
++
+/-
++
++
+/+/+/+
+/++
+/+/+/+
+/++
++
++
-
+/+/+/-
+
+/+/-
-
+/++
+
+
+
YES
YES
+/+
+
++
+
-
YES
YES
YES
+/+/+/+/+/++
+
+/++
+/-
+/+/++
++
+
++
-
+/++
-
YES
YES
YES
++
++
++
+/-
+/+
+/-
+/+/++
+/-
FIS1
MDV1
MDM36
NUM1
MDM35
UPS1
UPS2
MDM39
MDM33
MDM38
GEM1
MDM1
MPM1
CLU1
REF2
PTC1
CDC73
MDM20
VPS45
KCS1
NUP170
RIM9
MRH4
SSQ1
COX11
PSD1
Compromised
respiratory growth
YES at 37°C
YES
YES
YES
YES
YES at 37°C
YES
YES
YES
YES
YES
YES
YES
YES
YES
YES
YES
YES
YES
++, strong sensitivity; +, moderate sensitivity; +/-, mild sensitivity; -, not sensitive.
Information on respiratory growth defects was obtained from the Saccharomyces genome database.
SAM, sorting and assembly machinery; ERMES, ER–mitochondria encounter structure; GET, Golgi-ER trafficking; OM, mitochondrial outer
membrane;, IM mitochondrial inner membrane; IMS, mitochondrial intermembrane space.
ptc1D and vps45D (Table 1 and Fig. S5). We also found
that some mitochondrial morphology mutants that showed
only mild sensitivity to caspofungin in our assay, such as
tom7D and mdm20D, were clearly sensitive to Calcofluor
white (Table 1 and Fig. S5).
Mitochondrial defects are expected to lead to pleiotropic
stress response phenotypes. To address specificity, we
tested sensitivity to the azole fluconazole (which inhibits
ergosterol biosynthesis, affecting membrane biogenesis)
(Table 1 and Fig. S4). Several mitochondrial mutants
were sensitive to fluconazole; however, generally we did
not find a correlation between strong sensitivity to cell wall
damaging drugs and fluconazole, for example, sam37D
and mdm10D were strongly sensitive to cell wall inhibitors,
but not to fluconazole (Table 1).
Next we considered whether there are any commonalities between the mitochondrial morphology mutants that
we found to be strongly or moderately sensitive to cell wall
stress. We focused specifically on genes encoding mitochondrial proteins, which are thus more likely to have a
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
976 M. J. Dagley et al. 䊏
direct and specific role in the functional connections
between the mitochondrial and cell wall integrity.
Mitochondrial respiration might be contributing to cell
wall integrity, because the majority of mitochondrial morphology mutants, many of which are reported to be respiratory deficient, were at least mildly sensitive to
caspofungin (Table 1). However, there was not a strong
correlation between respiratory defects and the severity of
cell wall stress phenotypes, indicating that another function of the mitochondria has a more prominent role
(Table 1). Out of 10 mutants that displayed the strongest
sensitivity to caspofungin, five of these mitochondrial proteins have a connection to phospholipid homeostasis:
mdm10D, mdm35D, mdm31D, and ups2D have each been
reported to have altered levels of the phospholipid phosphatidylethanolamine (PE) and mdm10D, and mdm31D
also of cardiolipin (CL) (Kornmann et al., 2009; Osman
et al., 2009; Tamura et al., 2009). The sam37 mutant was
originally identified as a byproduct of a screen for phospholipid homeostasis mutants (Gratzer et al., 1995). In
fact, the only mitochondrial proteins that were moderately
or strongly required for tolerance of both cell wall inhibitors
in our study (caspofungin and Calcofluor white) are
Mdm10, Mdm35 and Sam37, and as explained above,
they all have been connected to maintenance of phospholipid homeostasis. That the sensitivity of these mutants to
caspofungin and Calcofluor white is due to a cell wall
defect is supported by rescue of the phenotypes by
osmotic stabilization (Fig. S6).
Based on these considerations, we suggest that sensitivity to cell wall inhibitors observed for a mitochondrial
morphology mutant is indicative of roles in phospholipid
homeostasis. We sought to explore these connections
further by using the sam37D mutant.
Although the sam37 mutant has been implicated in
phospholipid biosynthesis due to a positive hit in a screen
for phospholipid defects (Gratzer et al., 1995), a direct role
for Sam37 in phospholipid homeostasis has never been
established or explained. The majority of cellular phospholipid biosynthesis occurs in the ER, with the exception of
the synthesis of PE. PE biosynthesis occurs predominantly
in the mitochondria, by decarboxylation of ER-synthesized
phosphatidylserine (PS) by the mitochondrial PS decarboxylase Psd1 (Voelker, 2000; Birner and Daum, 2003).
These reactions necessitate transport between the ER and
the mitochondria: PS is synthesized in the ER and transported to mitochondria for decarboxylation to PE. PE then
travels back to ER for distribution to membranes and to
serve as precursor for synthesis of phosphatidylcholine
(PC) (Voelker, 2000; Birner and Daum, 2003). A recent
study identified the ERMES complex (Mmm1-Mdm10Mdm12-Mdm34) as being critical for physically bridging the
ER and mitochondrial membranes, enabling phospholipid
trafficking (Kornmann et al., 2009).
Fig. 6. Sam37 is required for transport-dependent decarboxylation
of phosphatidylserine.
A. Cells from wild-type and mutant cultures were labelled with
[3H]-L-serine as described in Experimental procedures, lipids
extracted and phospholipid species detected using thin layer
chromatography. Phospholipid species were identified by
comparison to standards and quantified. To determine the rate of
transport, levels of phosphatidylserine (PS),
decarboxylation-derived phospholipids, phosphatidylethanolamine
(PE) and phosphatidylcholine (PC) were plotted. Averages of three
independent experiments are shown and the error bar is the
standard error of the mean.
B. Psd1 activity was determined in isolated mitochondria, by
monitoring the conversion of [3H]-PS to [3H]-PE as described in the
Experimental procedures. Averages of three independent
experiments are shown and the error bar is the standard deviation.
To test how Sam37 affects phospholipid biosynthesis,
wild-type and sam37D cells were labelled with [3H]-L-serine
and the pool of newly made [3H]-PS was chased over time
to monitor conversion to PE and PC (Fig. 6A). Conversion
of PS to PE and PC was clearly slower in the absence of
Sam37 (approximately 50% less PS was converted in the
sam37D mutant, Fig. 6A). This defect could be due to lower
Psd1 activity in the absence of Sam37, or lower phospholipid trafficking between the ER and the mitochondria. To
discriminate between these two possibilities, we measured
Psd1 activity in isolated mitochondria in the presence or
absence of Sam37. The sam37D mutant did not display
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
Cell walls, mitochondria and phospholipids 977
Fig. 7. Sam37 is required for maturation of the GPI-anchored
protein Gas1.
A. Activation of the cell wall integrity gene CWP1 was monitored in
response to treatment with Calcofluor white by qPCR. Levels of
CWP1 were normalized to ACT1 levels. Shown are averages of
two independent experiments and the standard error.
B. Activation of the cell wall integrity pathway upon treatment with
125 ng ml-1 caspofungin was tested by monitoring the appearance
of the phosphorlyated form of the downstream kinase Slt2.
C. Total cell extracts were made from wild-type and sam37D
mutants and proteins visualized with antibodies directed against the
GPI-anchored protein Gas1. Cytosolic proteins Ssa1 and
hexokinase are shown as loading controls.
lower Psd1 activity, but rather higher activity was detected,
suggesting a compensatory response (Fig. 6B). We conclude that Sam37 is required for ER–mitochondria phospholipid trafficking and this is causing slower conversion of
PS to PE and PC in sam37D cells.
The S. cerevisiae pgs1D mutant, which is defective in
the biosynthesis of the mitochondrial phospholipid CL,
has a defect in activation of the PKC-dependent cell wall
integrity pathway (Zhong et al., 2007). In contrast, the
sam37D mutant activated the PKC pathway normally in
response to caspofungin and Calcofluor white (Fig. 7A
and B). Many cell wall proteins, as well as enzymes
required for cell wall synthesis are GPI-anchored (RuizHerrera et al., 2005; Klis et al., 2006; Plaine et al., 2008)
and PE is required for GPI anchor synthesis (Imhof et al.,
2000; Birner et al., 2001). We therefore considered
whether sam37D cells have a defect in maturation of
GPI-anchored proteins. In the absence of Sam37, the
steady-state protein levels of Gas1, a membrane and cell
wall localized GPI-anchored b-1,3-glucanosyltransferase
required for cell wall b-glucan remodelling, were lower
(Fig. 7C). The mRNA levels for GAS1 were not altered in
sam37D cells, as measured by qPCR. The average ratio
of GAS1 mRNA levels in the wild-type versus sam37D
mutant from five biological replicates was 1.18 (⫾ 0.28
standard error).
Ethanolamine can serve as a precursor for PE biosynthesis by the non-mitochondrial Kennedy pathway
(Voelker, 2000). The defect in Gas1 levels in sam37D cells
was partially suppressible by addition of exogenous ethanolamine (Fig. 7C), indicating it is, at least in part, due to
PE deficiency. We conclude that defective maturation of
GPI-anchored proteins could be contributing to the cell
wall defect in sam37D mutants.
To address whether the function of Sam37 in caspofungin tolerance is conserved in C. albicans, we constructed
a homozygous deletion mutant in the gene encoding the
Sam37 orthologue, orf19.1532. We found that Sam37
plays a bigger role in cell growth in C. albicans than in S.
cerevisiae, as the homozygous deletion mutant sam37DD
displayed substantially slower growth in the absence of
exogenous stress (Fig. 8). The C. albicans sam37DD
mutant was dramatically hypersensitive to caspofungin at
low doses of 35ng ml-1 and was also mildly sensitive to
Calcofluor white (Fig. 8). These data show that the function of Sam37 in tolerance of cell wall inhibitors is conserved in C. albicans.
The Ccr4-Pop2 deadenylase is required for
phospholipid homeostasis in C. albicans and this is
linked to its function in cell wall integrity
Based on the results of our S. cerevisiae caspofungin
sensitivity screen, we reasoned that Ccr4-Pop2 might be
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
978 M. J. Dagley et al. 䊏
almost control levels at lower caspofungin doses of
75 ng ml-1 (Fig. 9B; of note, at earlier time points we
observed that ethanolamine also somewhat improved the
growth of wild-type cells in the presence of caspofungin).
At a higher caspofungin dose of 125 ng ml-1, ethanolamine could not rescue the mutants (Fig. 9B), suggesting
that other pathways related to cell wall biogenesis are
also affected in the absence of Ccr4-Pop2.
Importantly, rescue of the caspofungin sensitivity of the
ccr4DD and pop2DD mutants by ethanolamine was
specific. Caspofungin sensitivity of mutants in the cell
wall integrity PKC pathway (bck1-/- and mkc1-/-) was
not suppressed by ethanolamine (Fig. S7). Furthermore,
upregulation of cell wall-related genes in the ccr4DD
mutant (likely a compensatory response to cell wall
defects, see Fig. 4 and Supporting Dataset 1) was
rescued by ethanolamine (Fig. 9C). This shows that ethanolamine is repairing the cell wall defects of the ccr4DD
mutant. Upregulation of the lipid biosynthesis genes INO1
and TAZ1 and the mitochondrial ribosomal gene MRPL10
in ccr4DD mutants was also reverted by ethanolamine
(Fig. 9C), indicating a rescue of phospholipid and mitochondrial defects.
Discussion
Fig. 8. Sam37 is required for caspofungin tolerance in C.
albicans. Cells from the indicated strains were grown to log phase.
Ten-fold serial dilutions were dropped on plates supplemented with
caspofungin or Calcofluor white at the doses indicated on the left
side of the figure. The plates were photographed after three days
of growth at 30°C. Two independent homozygous deletion mutants
were tested (lane 2 and 5), alongside complemented strains (lanes
3–4 and 6–7). The wild-type was DAY185.
affecting phospholipid homeostasis in C. albicans, and
that this links its roles in cell wall integrity and mitochondrial morphology.
To test this hypothesis, we isolated total lipids from
wild-type, ccr4DD and ccr4DD/CCR4 strains and determined the levels of phospholipids by mass spectrometry.
As shown in Fig. 9A, the ccr4DD mutant had lower levels
of phospholipids, including PE (Fig. 9A). To test directly
whether the cell wall, phospholipid and mitochondrial
defects of the deadenylase mutants are linked, we
assayed suppression of cell wall defects in the mutants by
exogenous ethanolamine supplementation. Supplementing the growth media with ethanolamine suppressed
caspofungin sensitivity of ccr4DD and pop2DD cells to
In this report, we describe the first characterization of
post-transcriptional factors in regulating cell wall integrity
in C. albicans, focusing on the Ccr4-Pop2 mRNA
deadenylase. We show that Ccr4-Pop2 is required for cell
wall biogenesis in C. albicans: its activity affects cell wall
b-glucan levels and thereby mediates basal resistance to
the antifungal drug caspofungin. Our data in C. albicans
are likely to have general relevance, explaining the observation that Ccr4-Pop2 is required for caspofungin tolerance in S. cerevisiae (Markovich et al., 2004) and the
fungal pathogen Cryptococcus neoformans (Panepinto
et al., 2007). Data we gathered in S. cerevisiae show that
it is the exonuclease activity of this complex that is
required for growth in the presence of caspofungin (Fig.
S8). The catalytic activity of this complex thus represents
a promising antifungal drug target.
We demonstrate that the cell wall integrity defects of the
C. albicans ccr4 and pop2 mutants are functionally linked
with defects in mitochondrial morphology and phospholipid homeostasis, providing for the first time an explanation for the cell wall integrity defects in the Ccr4-Pop2
deadenylase mutants. We extended the functional links
between mitochondrial and cell wall defects by screening
a collection of mitochondrial morphology mutants in S.
cerevisiae for caspofungin sensitivity. This screen identified several new genes with roles in cell wall integrity and
it pointed to a link between cell wall integrity and the role
of mitochondria in phospholipid homeostasis. We show
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
Cell walls, mitochondria and phospholipids 979
Fig. 9. The C. albicans ccr4DD mutant
displays lower phospholipid levels and this is
functionally linked to defects in cell wall
integrity.
A. Lipids were extracted from wild-type,
ccr4DD and complemented strains and
phospholipid levels determined by mass
spectrometry as described in Experimental
procedures. Two independent experiments
were performed and the averages and
standard errors are shown. PE,
phosphatidylethanolamine; PC,
phosphatidylcholine; PI, phosphatidylinositol;
PS, phosphatidylserine; PG,
phosphatidylglycerol; CL, cardiolipin.
B. Ten-fold serial dilutions of cells from the
indicated strains were dropped on synthetic
complete plates with or without caspofungin.
Ethanolamine was supplemented to one set
of plates as indicated.
C. Strains from ccr4DD mutants or
complemented ccr4DD/CCR4 strains were
grown in synthetic complete media with or
without 1 mM ethanolamine. Total RNA was
isolated and qPCRs performed to assess
upregulation of the indicated cell wall integrity,
lipid biosynthesis and mitochondrial
biogenesis genes in the ccr4DD mutant. ACT1
levels were used for normalization. The graph
shows the average and standard error from
two biological replicates.
that this role is conserved in C. albicans, by demonstrating that the homozygous deletion mutant in SAM37 is
sensitive to caspofungin, providing the foundation for
studying the roles of mitochondria in echinocandin resistance in this major fungal pathogen.
Conservation of Ccr4-Pop2 roles at the molecular level
In S. cerevisiae and S. pombe, Ccr4-Pop2 represents the
major mRNA deadenylase, post-transcriptionally regulating mRNA stability and translation (Tucker et al., 2001;
Grigull et al., 2004; Beilharz and Preiss, 2007; Jonstrup
et al., 2007; Lackner et al., 2007; Goldstrohm and
Wickens, 2008). Our work shows that the molecular role
of Ccr4-Pop2 in C. albicans is conserved with these other
fungal species. In C. albicans, mRNAs preferentially
display a short or a long poly(A) tail at steady state in
wild-type cells, and the ‘short-tailed’ mRNAs are deadenylated by Ccr4-Pop2. This is analogous to what has been
observed in S. cerevisiae and S. pombe (Beilharz and
Preiss, 2007; Lackner et al., 2007) and suggests that in C.
albicans also, gene-specific mRNA poly(A) tail length
control by the Ccr4-Pop2 deadenylase modulates gene
expression. In S. cerevisiae, mRNAs involved in rRNA
and ribosome biogenesis are short-lived and their stability
is regulated by Ccr4 (Grigull et al., 2004). This is a conserved role for Ccr4, as we found that in C. albicans the
levels of the rRNA and ribosome biogenesis genes were
elevated in the absence of Ccr4.
At a cellular level, we showed that a major function of
Ccr4-Pop2 in C. albicans is to maintain cell wall integrity.
The ccr4DD and pop2DD mutants display lower levels of
cell wall b-glucans, are hypersensitive to cell wall targeting drugs, and show hyperactivation of the PKCdependent cell wall integrity pathway and a cell lysis
phenotype. The identity of the mRNA targets of Ccr4Pop2 responsible for these cell wall defects is presently
unknown. Microarray analysis of the ccr4DD mutant transcriptome revealed extensive compensatory mechanisms
at play and the complexity of this regulatory network
makes it difficult to distinguish, at a transcriptome level,
between direct and indirect effects from loss of Ccr4
activity. For example, deadenylation regulates not only
mRNA stability, but also the efficiency of translation, which
cannot be determined from a transcriptome analysis.
However, our result that TAZ1 is a ‘short-tailed’ gene and
overexpressed in the C. albicans ccr4DD mutant, suggests that Ccr4-dependent deadenylation could be regulating the expression of genes required for phospholipid
homeostasis. Future experiments, using CCR4 shut-
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
980 M. J. Dagley et al. 䊏
down strains (to potentially avoid large compensatory
effects present in homozygous deletants that have
adapted to loss of Ccr4), and a proteomics approach will
be required to find the mRNAs regulated by Ccr4 which
impact on cell wall biogenesis.
Ccr4-Pop2, phospholipid homeostasis and cell wall
biogenesis
Taken together, our data suggest that Ccr4-Pop2 controls
the expression of genes required for ER and mitochondrial function, thereby affecting phospholipid biosynthesis
and consequently cell wall integrity (Figs. 9 and 10). The
experiment demonstrating a direct link between the phospholipid and cell wall defects is the rescue of caspofungin
sensitivity of the ccr4DD and pop2DD mutants by ethanolamine supplementation.
Our conclusion that phospholipid imbalance is causative of cell wall defects in the Ccr4-Pop2 mRNA deadenylase mutants is further supported by a recent report on
the effects of PS synthase (CHO1) and PS decarboxylase
(PSD1 and PSD2) in C. albicans (Chen et al., 2010). The
parallels between the cho1, psd1 psd2 mutants and the
mRNA deadenylase mutants include defects in cell walls,
as well as compromised filamentation and virulence. The
nature of the filamentation defects in the deadenylase
mutants is similar to those of the phospholipid biosynthesis mutants in that filamentation is strongly defective in
Spider and M199 media and only partially defective in
response to serum (Fig. 2, Fig. S2 and Chen et al., 2010).
An important difference between the phospholipid biosynthesis mutants (Chen et al., 2010) and the Ccr4-Pop2
mRNA deadenylase mutants (this study) is the extent of
sensitivity to the echinocandin caspofungin and other
cell wall inhibitors. The deadenylase mutants are highly
sensitive to caspofungin, whereas the phospholipid biosynthesis mutants are only sensitive at a very high concentration of 25 mg ml-1 caspofungin (Chen et al., 2010).
Moreover, the deadenylase mutants are sensitive to
Congo red and Calcofluor white, whereas the cho1 and
psd1 psd2 mutants are not (Chen et al., 2010). We
suggest that this difference reflects a global role of Ccr4Pop2 in promoting cell wall integrity.
How exactly phospholipids affect cell wall integrity is
unknown. Effects of PE on GPI anchor synthesis and
compromised signalling through the PKC or calcineurin
pathways in the phospholipid biosynthesis mutants have
been proposed as mechanisms (Chen et al., 2010). The
ccr4DD and pop2DD mutants hyper-accumulate phosphorylated Mkc1 (Fig. S1), suggesting that signalling through
the PKC pathway is intact. Our data with the S. cerevisiae
sam37D mutant also show that PKC pathway activation is
not defective. Based on (i) caspofungin sensitivity of both
the deadenylase mutants (our study) and the cho1 and
psd1 psd2 mutants (Chen et al., 2010), (ii) our data
showing a defect in cell wall b-glucans in the absence of
the Ccr4-Pop2 mRNA deadenylase and (iii) ethanolamine
rescue of caspofungin sensitivity of the deadenylase
mutants, we now propose that phospholipid imbalance
affects the activity of b-glucan synthase. b-Glucan synthase is an integral membrane protein and earlier biochemical work indicates a critical role for the phospholipid
environment in ensuring proper b-glucan synthase activity
(Wasserman and McCarthy, 1986; Sloan et al., 1987;
Saugy et al., 1988). Future experiments will test this
hypothesis.
Specific aspects of mitochondrial function impact on cell
wall integrity
In addition to rescuing caspofungin sensitivity, ethanolamine supplementation also rescued the compensatory
upregulation of mitochondrial biogenesis and phospholipid biosynthesis genes observed in the ccr4DD mutants.
This provides a link between mitochondrial morphology
defects, phospholipid defects and cell wall integrity
defects arising from the absence of Ccr4-Pop2 in C.
albicans. Mitochondrial morphology has been wellcharacterized in S. cerevisiae, with a number of mutants
having been identified that show mitochondrial morphology defects (Dimmer et al., 2002; Meisinger et al., 2004;
Altmann and Westermann, 2005, Okamoto and Shaw,
2005; Sesaki et al., 2006; Tamura et al., 2009). Our
screen of the collection of mitochondrial morphology
mutants identified that mitochondrial function is required
for caspofungin tolerance, as the majority of mutants were
at least mildly sensitive to caspofungin in our assay. A
previous whole-genome study of caspofungin sensitivity
did not report a connection to mitochondrial function
(Markovich et al., 2004), possibly due to the fact that the
cut-off used was a fourfold decrease in minimum inhibitor
concentration and the defects of the mutants in our study
were milder. Our conclusion that mitochondrial function
modulates caspofungin sensitivity is supported by a large
chemical genomics screen of the S. cerevisiae deletion
collection performed by Hillenmeyer et al. in which the
heterozygous deletion mutant in COX17, which is
required for cytochrome C oxidase activity and respiration, was among the most sensitive to caspofungin (Hillenmeyer et al., 2008).
In our assay, however, respiratory deficiency did not
correlate with the severity of hypersensitivity to cell wall
stress (Table 1), and thus a function of the mitochondria
different to respiration appears to be critical for cell wall
integrity. Our screen suggested that mitochondrial phospholipid homeostasis is important for cell wall biogenesis.
Previously, the pgs1D mutant affected in synthesis of the
mitochondrial phospholipid CL has been shown to have a
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
Cell walls, mitochondria and phospholipids 981
cell wall defect (Zhong et al., 2005; 2007). Our study
significantly expands the list of mitochondrial mutants with
phospholipid defects that are sensitive to cell wall stress,
as we show that mdm10D, mdm35D, mdm31D, ups2D and
sam37D are sensitive to caspofungin and a subset of
these mutants also to Calcofluor white. In support of our
results, we found that mdm35, mdm31 and sam37 heterozygous mutants were listed in the group of mutants
with a fitness defect in the presence of caspofungin in the
large-scale chemical genomic screen of more than 400
compounds and conditions performed by Hillenmeyer
et al. (2008).
We further characterized the sam37D mutant to probe
the connections between cell wall integrity and the role
of mitochondria in phospholipid homeostasis. Cell wall
defects arising in the absence of SAM components are
suggested by the fact that cells inactivated for the
Sam37 subunit are sensitive to caspofungin and Calcofluor white, and that this can be suppressed by osmotic
stabilization. Previous genome-wide screens list sam37D
mutants among the mutants that are sensitive to Calcofluor white (Lesage et al., 2005) and to caspofungin in a
heterozygote mutant situation (Hillenmeyer et al., 2008).
Furthermore, the sam37D mutant was noted to hyperaccumulate chitin in the wall, a known response to cell
wall stress (Lesage et al., 2005), is known to flocculate
(I. Gentle, unpubl. obs.) and exhibits synthetic genetic
interactions with mutants in several cell wall biogenesis
genes (e.g. KNH1, KTR3, KRE1, KRE11, HKR1, CHS3)
(Tong et al., 2004). In the context of our study, these
various observations can now be rationalized with
Sam37 functioning to promote cell wall integrity, via its
role in establishing contacts between the ER and mitochondrial membranes that are necessary for phosholipid
biosynthesis. We demonstrate that Sam37 is required
for wild-type rates of PS decarboxylation to PE in the
mitochondria and that this is affecting maturation of the
GPI-anchored b-glucan remodelling enzyme Gas1, suggesting that cell wall defects of sam37D cells arise, at
least in part, from lower PE levels. Importantly, we demonstrate that the role of Sam37 in caspofungin tolerance
is conserved in C. albicans, demonstrating for the first
time directly that mitochondrial function affects echinocandin resistance in this pathogen.
Mdm10 is another subunit of the SAM complex that we
found to be required for cell wall integrity. Mdm10 is
also a subunit of the ERMES complex, involved in
ER–mitochondria contacts and phospholipid trafficking
(Meisinger et al., 2007; Kornmann et al., 2009). However,
in our assays, mutants in the other ERMES subunit
(mmm1D, mdm12D and mdm34D), displayed a milder
sensitivity to cell wall stress, suggesting that it is the role
of Mdm10 within the SAM complex that dominates its
impact on cell wall integrity. Consistent with this, Tom7, a
subunit of the TOM (Translocase of the Outer Membrane)
complex that functions to diminish association of Mdm10
with the SAM complex and to promote its interactions with
ERMES (Meisinger et al., 2006; Yamano et al., 2010;
Becker et al., 2011), was not required for caspofungin
sensitivity. However, we found that tom7D was sensitive to
Calcofluor white, supporting a distinct role for Tom7 in cell
wall integrity.
Our data provide the first direct evidence of, and an
explanation for, the requirement for Sam37 in phospholipid
biosynthesis. PE biosynthesis from the ER-synthesized
precursor PS is compromised in sam37D cells, but Psd1
activity is not lower, indicating that the function of Sam37 is
in phospholipid trafficking between the ER and the mitochondria. We therefore propose that, in addition to the
recently discovered ERMES complex (Kornmann et al.,
2009), the SAM complex is required for establishing functionally important connections between the ER and mitochondrial membranes. Functional interactions between
the ERMES and SAM complexes have been previously
proposed, based on a shared subunit (Mdm10) and the
requirement of Mdm12 and Mmm1 in assembly of mitochondrial outer-membrane proteins mediated by the SAM
complex (Meisinger et al., 2004; 2007). That other factors
would be involved in ER–mitochondria phospholipid trafficking has been suggested based on the moderate phospholipid defects of ERMES mutants (Kornmann et al.,
2009; Kornmann and Walter, 2010). The ERMES complex
mutants displayed a much milder cell wall integrity phenotype in our assay, suggesting that in individual ERMES
mutants, phospholipid transfer is less affected than it is in
the sam37D mutant. This notion is supported by wild-type
total steady-state levels of the majority of phospholipids in
mdm12D cells (Kornmann et al., 2009). A recent report
showed that Mmm1, Mdm10 and Mdm34 have equivalent
TULIP domains for binding phospholipids, perhaps
explaining a functional redundancy (Kopec et al., 2010).
Until more is known about the structure and function of
ERMES, it is difficult to discern the precise role in phospholipid homeostasis, cell wall integrity and mitochondrial
function.
How the SAM complex controls phospholipid transport
between the two membranes is not yet clear. The SAM
complex might directly regulate the number of membrane
contacts, being required for the assembly of b-barrel proteins such as Mdm10 into the mitochondrial outer membrane (Pfanner et al., 2004). Alternatively, the SAM
complex could itself be involved in the physical interactions between the ER and mitochondria: the Sam37 and
Sam35 subunits of the SAM complex are peripheral membrane proteins on the outer face of the mitochondria
(Gratzer et al., 1995; Ishikawa et al., 2004; Milenkovic
et al., 2004; Waizenegger et al., 2004), in a position to
make protein–protein or protein–lipid contacts with the
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
982 M. J. Dagley et al. 䊏
Fig. 10. The factors connecting cell wall biogenesis to mitochondrial function and phospholipid homeostasis. The blue arrow represents the
overall process of cell wall biogenesis, which requires delivery of phospholipids to the plasma membrane, the targeting and activity of
enzymes responsible for synthesis of wall components in the plasma membrane, and the targeting of mannoproteins and modifying enzymes
into the wall. Phospholipid biosynthesis depends on both the endoplasmic reticulum and mitochondria and co-ordinated function is ensured
through the ERMES and SAM complexes required for interactions between the mitochondria and the endoplasmic reticulum membrane
systems, and Ccr4-Pop2-dependent deadenylation of mRNAs encoding proteins destined for either organelle. The mitochondrial proteins
Mdm31, Mdm35, Ups2, Mdm10 and Sam37 are required for phospholipid homeostasis and have prominent roles in tolerance of the
echinocandin caspofungin.
ER. Future experiments will need to distinguish these
potential mechanisms.
A network of gene products regulating cell wall integrity
and drug sensitivity in fungi
Several genetic interactions have been observed
between the genes that we found to contribute to cell wall
integrity in S. cerevisiae and C. albicans (Table 1, Figs 1,
8 and 10). For example, both CCR4 and POP2 show
synthetic genetic interactions with the GET complex
subunit MDM39/GET1 and the F-box protein MDM30,
and POP2 also displays synthetic interactions with
another GET complex subunit, GET2 (Pan et al., 2006).
CCR4 also shows synthetic genetic interactions with the
ERMES subunit MDM12 (Costanzo et al., 2010). In addition to its genetic interactions with factors involved in cell
wall biogenesis (Tong et al., 2004), SAM37 displays
genetic interactions with the MDM39/GET1, GET2 and
GET3 subunits of the GET complex (Pan et al., 2006;
Collins et al., 2007; Costanzo et al., 2010). MDM31
genetically interacts with MDM10 and the other ERMES
subunits (Dimmer et al., 2005), as well as with UPS2
(Costanzo et al., 2010), TOM7 with MFB1, and interestingly also with the mitochondrial phosphatydylserine
decarboxylase PSD1 (Costanzo et al., 2010) and MDM35
with the GET complex subunit GET3 and with PSD1
(Costanzo et al., 2010). We suggest that the reported
genetic interactions reflect the roles of these genes in
phospholipid homeostasis and cell wall integrity (Fig. 10).
We further suggest that sensitivity of mitochondrial
mutants to cell wall inhibitors is predictive of roles in these
pathways. In future experiments, we will be testing this
prediction with the mitochondrial morphology mutants
identified in the current study.
We demonstrated that Sam37 is required for phospholipid trafficking between the mitochondria and the ER.
Phospholipid trafficking is centred on transport of the
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
Cell walls, mitochondria and phospholipids 983
Psd1 substrate PS to the mitochondria for its decarboxylation to PE, and transport of PE from the mitochondria to
the ER (Fig. 10). However, in both S. cerevisiae and C.
albicans, inactivation of PSD1 is not equivalent to inactivation of SAM37, with the sam37 mutant displaying a
considerably more pronounced hypersensitivity to cell
wall stress (our study and Chen et al., 2010). In a complex
genetic network as described above, it remains possible,
and perhaps likely, that molecules in addition to phospholipids might help co-ordinate the role of ER–mitochondria
connections in cell wall integrity. Trafficking of Ca2+
between the ER and mitochondria could also be mediated
by physical contacts between these two organelles (Kornmann et al., 2009; Kornmann and Walter, 2010). At least
one Ca2+-dependent pathway, the calcineurin pathway,
affects cell wall biogenesis in both C. albicans and S.
cerevisiae (Wiederhold et al., 2005; Munro et al., 2007;
Singh et al., 2009), and might provide for co-ordination
over mitochondrial biogenesis, phospholipid homeostasis
and cell wall biogenesis.
Experimental procedures
Yeast strains and growth conditions
The C. albicans strains used in this study are derivatives of
BWP17 (Wilson et al., 1999), and are listed in Table S3. The
ccr4DD, pop2DD and sam37DD mutants were constructed by
standard methods based on PCR and homologous recombination, using ARG4 and URA3 as selective markers. The
complemented strains were constructed by introducing a
wild-type copy of CCR4, POP2 or SAM37 under their own
promoter and terminator into the HIS1 locus of the respective
mutants, using the integrative plasmid pDDB78. Unless otherwise stated, fully prototrophic strains (URA+ ARG+ HIS+)
were used for all experiments.
The S. cerevisiae strains are listed in Table S3. The
BY4741 deletion collection was obtained from Open
Biosystems. The ccr4D, pop2D and the catalytic inactive
mutant ccr4-1 used in Fig. S8 are in the KY803 background
and are described in Traven et al. (2005).
Standard growth conditions were YPD (2% glucose, 2%
peptone, 1% yeast extract), at 30°C, 200 r.p.m. For the
C. albicans strains, the media were supplemented with
80 mg ml-1 uridine. The mutants were selected using minimal
media lacking the appropriate amino acids.
For testing drug sensitivity, 10-fold serial dilutions of logphase cells from wild-type, mutant and complemented strains
were dropped on plates with or without the drugs: 35 and
50 ng ml-1 caspofungin, 20 and 50 mg ml-1 Calcofluor white
and 10 and 20 mg ml-1 fluconazole. We noticed that the ability
of both wild-type and mutants to grow on caspofungin
declined with chronological age (i.e. when the cultures were
inoculated from plates kept at 4°C) and so strains were
freshly streaked from stocks for the experiments. The plates
were incubated 2–4 days at 30°C before photographing. For
caspofungin dose–response experiments in liquid media,
cells were grown to log phase and then inoculated into
96-well plates containing caspofungin at 0, 10, 20, 35, 50, 60
and 100 ng ml-1. Per well, 105 cells were used. Growth was
scored after 24 h at 30°C. Representative experiments are
shown in Figs S4 and S5 and Table S2.
Growth on glycerol was tested on YPG plates (2% glycerol,
2% peptone, 1% yeast extract). Ethanolamine supplementation experiments were done in synthetic complete media.
Ethanolamine was added at 1 mM concentration, as previously described in S. pombe (Matsuo et al., 2007).
Filamentous growth, mitochondrial morphology
determination and microscopy
For testing filamentous growth, cells from overnight cultures
grown in YPD at 30°C were diluted to OD600 = 0.2 into prewarmed YPD+10% calf serum, Spider media (1% nutrient
broth, 1% D-mannitol), M199 or N-acetylglucosamine
media (9 g NaCl, 6.7 g yeast nitrogen base and 0.56 g
N-acetylglucosamine per litre) and incubated at 37°C for the
times indicates in the figures. Before viewing, cells were
washed and resuspended in PBS (phosphate-buffered
saline). Cells were imaged using an Olympus IX81 microscope with the Olympus cell^M software.
For testing filamentation on plates, cells were re-streaked
on Spider or YPD+10% serum plates and incubated at 37°C
for 4 days.
For viewing mitochondrial morphology, strains were grown
at 30°C to mid-log phase. Mitochondria were stained with
1 mM MitoTracker Red for 30 min in the dark. After staining,
cells were washed and resuspended in water and mounted
1:1 with 1% low-melt agarose. Cells were imaged using a
Leica SP5 laser scanning confocal microscope, using a 100 ¥
oil-immersion objective. The confocal images were compiled
using the Leica LAS AF Lite software.
Cell wall carbohydrate analysis
Cell walls were prepared according to the method of François
(2006). Cell walls were isolated from 1.8 ¥ 109 cells. To
isolate cell walls, cells were washed twice in water, resuspended in TE (Tris-EDTA) with glass beads and disrupted by
three cycles of 2 min Beadbeater lysis. Lysates were centrifuged (1000 g, 1 min), and the supernatant kept. Beads were
washed five times with TE and pelleted at 500 g for 1 min
after each wash; all supernatants were pooled with lysate.
The pooled lysate/wash supernatant was centrifuged at
4800 g for 15 min, and the pellet resuspended in water. Cell
debris was removed by centrifugation at 500 g for 1 min, and
the supernatant centrifuged at 3000 g for 5 min to recover cell
walls. Cell walls were dried overnight in a 50°C oven.
Wall polysaccharides were analysed as partially methylated alditol acetates by mass spectrometry as described
(Van de Wouw et al., 2009). The mole percentage of each
polysaccharide was estimated from 1,4-linked glucosamine
for chitin, 1,3-linked glucopyranose for 1,3-b-glucan, 1,6linked glucopyranose for 1,6-b-glucan, and addition of all
mannopyranosyl derivatives for mannan. The average mole
percentage of the carbohydrates in the cell walls was calculated from three independent preparations analysed with two
technical replicates (for chitin, two independent preparations
were analysed in duplicate), and the error bar is the standard
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
984 M. J. Dagley et al. 䊏
error of the mean. The P-values were obtained by comparing
the mutants to the wild-type using the paired Student’s t-test.
Lipid analysis
Yeast cells were inoculated at OD600 = 0.1 into YPD (10 ml)
and grown to OD600 = 1. Lipids were extracted from 1 ml
aliquots of the cultures. Part of the culture was removed for
extraction of total proteins for normalization of lipid levels
from the same cultures. Proteins were extracted by trichloroacetic acid (TCA) precipitation and quantified using the Bradford assay. Normalization of lipid to protein levels is standard
for quantitative measurements of lipids in yeast (Nebauer
et al., 2007; Schuiki et al., 2010) and was chosen over normalization to cell numbers because of the difference in size
between wild-type and deadenylase mutant cells (Fig. S1).
Two independent experiments were performed, with four
technical repeats each. Cells were recovered by centrifugation for 5 min, and washed twice in 500 ml of chilled PBS. Cell
pellets were lysed by three 20 s cycles of freeze/thawing
between liquid nitrogen and a dry-ice-ethanol bath. Chloroform (200 ml) with 5 mM final of LPC standard was added to
the pellet, mixed, and then 560 ml of methanol : water (2:0.8,
v/v) was added. Lipids were extracted by vigorous vortexing,
followed by 5 min of centrifugation at maximum speed at 0°C.
The solvent phase (containing the extracted lipids) was
removed and stored at -20°C prior to analysis.
For liquid chromatography-mass spectrometry (LC-MS)
analysis, extracted lipids in chloroform/methanol/water were
dried and resuspended in (100 ml) butan-1-ol/ 10 mM ammonium formate in methanol (1:1, v/v). An aliquot (5 ml) was
loaded onto a 50 mm ¥ 2.1 mm ¥ 2.7 mm Ascentis Express
RP Amide column (Supelco) and the lipids separated with a
5 min gradient of water/methanol/tetrahydrofuran (50:20:30,
v/v/v) to water/methanol/tetrahydrofuran (5:20:75, v/v/v), with
the final buffer held for 3 min at a flow rate of 0.2 ml min-1 using
an Agilent LC 1200 system. The lipids were analysed in an
on-line Agilent Triple Quad 6410 mass spectrometer in either
the MS or MS/MS fragmentation mode. CL was identified in the
mass spectra from the retention time and accurate mass. PCs
were identified from MS/MS fragmentation data by scanning
for the precursor of m/z 184.1 in the positive ion mode while
phosphatidylinositols (PIs) and PGs were identified by scanning for the precursor of m/z 241 and 153, respectively, in the
negative ion mode. PEs and PSs were identified by scanning
for a neutral loss of m/z 141 in the positive mode and m/z 87 in
the negative mode respectively. The individual PC, PE, PG, PI,
PS and CL were quantified by multiple reaction monitoring
using fragmentor voltage range of 60–160 V and collision
energy range of 25–40 V and, respectively, using nitrogen as
the collision gas at 7 l min-1. LC-MS data was processed using
Agilent MassHunter quantitative software.
RNA preparation and microarray analysis
Overnight cultures of the ccr4DD mutant and the ccr4DD/
CCR4 complemented strain (both strains are URA+ ARG+
HIS+) were diluted to OD600 = 0.1 in 100 ml of YPD and grown
to OD600 = 1.0. Cells were recovered by centrifugation and the
pellets washed in water. The cell pellets were snap frozen in
liquid nitrogen and stored at -80°C. RNA was isolated using
the Ambion RiboPure Yeast RNA isolation kit according to
manufacturer’s instructions, with the exception that cell lysis
was performed by Beadbeater using three cycles of 2 min of
lysis with a 1 min break between cycles. The quality and
quantity of the RNA was determined using an Agilent 2100
Bioanalyzer.
Microarray analysis was performed essentially as
described (Nantel et al., 2006). Data normalization and
analysis was conducted in GeneSpring GX version 7.3
(Agilent Technologies). Genes that were up- or downregulated in the ccr4DD mutant by 1.9-fold or more (P ⱕ 0.05)
were selected from a Volcano Plot and considered to be
differentially expressed. Gene ontology analysis was performed using the tools at the Candida genome database
(CGD, http://www.candidagenome.org) (Skrzypek et al.,
2010).
Quantitative real-time PCR and ligation-mediated
polyadenylation test (LM-PAT)
Quantitative real-time PCR was performed on RNAs isolated
from the ccr4DD mutant and the ccr4DD/CCR4 complemented strain, using primers specific for the genes listed in
Supporting Dataset 3. For the ethanolamine supplementation
experiments, a single colony from mutant or complemented
strains was resuspended in 20 ml of water and then split
into either synthetic complete media without ethanolamine
or synthetic complete media supplemented with 1 mM
ethanolamine. Cultures were grown over night at 30°C, and
then diluted to OD600 = 0.2 and grown until reaching
OD600 = 1. For activation of CWP1 expression in S. cerevisiae
(Fig. 7), cells were grown to mid-log phase and then treated
with 40 mg ml-1 Calcofluor white for 2 h. RNA was isolated
using the hot-phenol method and DNase treated prior to
reverse transcription using the Transcriptor High Fidelity
cDNA synthesis kit from Roche. qPCR reactions were prepared using Fast-Start Sybr Green Master (Roche) on an
Eppendorf Realplex master cycler and analysed by absolute
quantification. The expression levels were normalized to the
level of ACT1.
LM-PAT was performed as described previously (Beilharz
and Preiss, 2007; Traven et al., 2009). The TVN-PAT
samples represent the shortest poly(A) tail length obtained by
the assay and were obtained from wild-type cDNA using a
PAT-T12-VN reverse transcription primer (5′-GCGAGCT
CCGCGGCCGCGTTTTTTTTTTTTVN; where V is any nucleotide except T and N is any nucleotide), which binds at the 3′
UTR and poly(A) junction. The LM-PAT reverse primer and
the TVN-PAT primer share 5′ sequence to enable parallel
PCR amplification from these control samples and the experimental cDNAs. Whether an mRNA is ‘short-tailed’ or ‘longtailed’ was determined by comparison to the TVN-PAT
sample after quantification of sections corresponding to short
or longer poly(A) tails. LM-PAT PCR amplicons were detected
by SYBR Safe DNA gel stain using Fujifilm Las-300 gel
documentation system and multiguage V3.0 software.
Western blots
For the phospho-Mkc1 Western blots, yeast cells were grown
to mid-log phase and then treated with 50 mM Calcofluor
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
Cell walls, mitochondria and phospholipids 985
white (Fig. S1) or 125 ng ml-1 caspofungin (Fig. 7). Samples
were taken before and after addition of the cell wall inhibitors
at the time points indicated in the figures. Proteins were
extracted by TCA precipitation and Western blots performed.
Phospho-Mkc1 was detected using the mouse monoclonal
anti-phospho p44/42 MAP kinase antibody (Cell signalling),
followed by the anti-actin antibody as a loading control.
For monitoring Gas1 protein levels, cells from wild-type
and sam37D mutants were grown in the presence or absence
of ethanolamine. The Gas1 antibody was a generous gift from
Howard Riezman (University of Geneva, Switzerland). Antibodies against hexokinase and Ssa1 were used as loading
controls.
In vivo phosphatidylserine transfer and in vitro Psd1
activity assays
The rate of conversion of PS to PE and PC in vivo was
determined by labelling the cells with [3H]-L-serine and then
following PS, PE and PC formation in a pulse chase
experiment. Cells were grown to mid-log phase in YPD at
25°C and labelled with 10 mCi ml-1 of [3H]-L-serine in PBS for
15 min at 25°C, after which cells were harvested, washed
and the 0 time point taken. The remainder of the culture was
resuspended in YPD and incubated at 25°C for the indicated
time points. At each time point, cells were pelleted and snap
frozen in liquid nitrogen.
Lipids were extracted in chloroform : methanol : water
(2:1:0.8; v/v/v) by bead beating 3 ¥ 1 min, followed by mixing
with beads for at least 1 h at 4°C. Supernatants were washed
three times by water-equilibrated butanol and phases
partitioned. The organic phase was dried under nitrogen.
Pellets were resuspended in 10 ml chloroform : methanol
(2:1; v/v) and applied to silica 60 HPTLC plates (Merck) and
developed. TLC phospholipid standards (Avanti) were run on
the same plates. Bands corresponding to the phospholipid
species were quantified with a Berthold TLC plate scanner.
Three independent experiments were performed and the
average ratio of [3H]-PE+PC/[3H]-PS calculated. The error bar
is the standard error of the mean of the calculated ratios.
For in vitro Psd1 activity assays mitochondria were isolated
as described (Gabriel et al., 2003) and resuspended in 400 ml
at a concentration of ª 2 mg ml-1 mitochondrial protein. The
amount of mitochondria was determined by immunodetection
with antibodies against various mitochondrial proteins
(Tom70, mtHsp70, Tom40, Tim23, porin, Cox2 and F1b). The
amount of protein used was adjusted to provide for the conversion of substrate linearly over time. The substrate
was [3H]-PS, which was synthesized from mitochondriaassociated membranes (MAMs) essentially as described
(Kuchler et al., 1986). MAMs were produced by isolating
mitochondria using breaking buffer at pH 7.4 (0.6 M sorbitol,
200 mM K+HEPES pH 7.4) to maintain the connection of
MAMs. MAMs were separated from mitochondria on a
sucrose gradient of 20–50% in 0.6 M sorbitol, 20 mM
K+HEPES pH 6.0) and pelleted at 100 000 g for 1 h. The
pellet was resuspended in breaking buffer at a protein concentration of 25 mg ml-1 and snap frozen in liquid nitrogen.
Psd1 activity assays were performed as described (Choi
et al., 2005). Reactions were started by addition of substrate,
incubated at 30°C for the indicated time points and stopped
by aliquoting into chloroform:methanol (2:1, v/v) and rapid
vortexing. Reaction products were separated on HPTLC, and
bands corresponding to [3H]-PS and [3H]-PE detected and
quantified as described above. Represented are averages of
three independent experiments and error bars show the standard deviation.
Virulence studies in mice
Virulence studies and organ burden analysis were conducted
in 8- to 9-week-old BALB/c mice (Animal Resource Centre,
Floreat Park, Western Australia). All animal experimentation
was done in accordance with the guidelines issued by the
Sydney West Area Health Service Animal Ethics Committee,
Department of Animal care, Westmead Hospital, Harkesbury
Road, Westmead, PO BOX 533, Wentworthville NSW 2145
(Animal Ethics Approval Protocol No. 5048). Animals were
anaesthetized by inhalation of methoxyflurane.
For the survival study, groups of 10 mice were inoculated
via tale vein injection with each C. albicans strain (2 ¥ 105 or
1 ¥ 106 yeast cells in 200 ml saline) and observed daily for
signs of ill-health. Mice which had lost 20% of their preinfection weight (Day 0), or which showed debilitating clinical
signs prior to losing 20% of their pre-infection weight, were
euthanized by CO2 inhalation followed by cervical dislocation.
Otherwise they were sacrificed 20 days post-infection.
For organ burdens, groups of six mice were inoculated via
tale vein injection with each C. albicans strain (1 ¥ 106 yeast
cells in 200 ml saline). Three mice from each group were
sacrificed on day 1 and day 2 post infection. Kidneys were
harvested and homogenized for determination of colonyforming units on SABD agar plates [Sabouraud Brain Heart
Infusion Agar Plus Drugs – 5 g peptone (Difco), 20 g glucose,
26 g Brain Heart Infusion Agar (Difco), 7.5 g agar, 0.025 g
Gentamicin and 0.25 g chloramphenicol per litre] after 2 days’
incubation at 30°C. Alternatively, kidneys were fixed in 10%
neutral buffered formalin (NBF) and selected tissue blocks
were placed into plastic cassettes and processed overnight
using a routine overnight cycle in a tissue processor. The
tissue blocks were then embedded in wax, serially sliced into
5 mm sections. Slides holding the sections were stained with
Periodic Acid Schiff (PAS) stain.
Statistical analysis was performed as follows. For the
murine intravenous model of candidemia, an estimation of
differences in survival (log-rank test) using the Kaplan–Meier
method was obtained and the survival curves plotted, using
the SPSS version 16 statistical software. In all cases, a
P-value of < 0.05 was considered statistically significant. Student’s t-test and one-way ANOVA were used to compare
means between groups using SPSS version 16 statistical
software.
Acknowledgements
We thank Jörg Heierhorst for his support and encouragement
in the initial phases of this work, Thomas Preiss for assistance with early experiments looking at mRNA poly(A) tail
lengths in C. albicans, Judy Callaghan for assistance with
confocal microscopy, Peter Boag for access to the Olympus
fluorescence microscope, Virginia James for histochemistry,
© 2010 Blackwell Publishing Ltd, Molecular Microbiology, 79, 968–989
986 M. J. Dagley et al. 䊏
Gary Martinic and Christabel F Wilson for technical assistance with animals and Karen Bythe for help with statistics.
We thank Mark Prescott and Dalibor Mijaljica for access to
the S. cerevisiae deletion collection and Kip Gabriel for
advice and discussions on the roles of the yeast mitochondrial proteins.
This work was supported by a Peter Doherty fellowship from
the Australian National Health and Medical Research Council
(NH&MRC), a short-term postdoctoral fellowship from the
Human Frontier Science Program Organization (HFSPO) and
a discovery project grant from the Australian Research Council
(ARC) (to A.T.), an ARC Federation fellowship (to T.L.), an ARC
Australian Research fellowship (to T.H.B.) and an NH&MRC
Principal Research Fellowship (to M.M.).
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