klis2002

FEMS Microbiology Reviews 26 (2002) 239^256
www.fems-microbiology.org
Dynamics of cell wall structure in Saccharomyces cerevisiae
Frans M. Klis
a
a;
, Pieternella Mol a , Klaas Hellingwerf a , Stanley Brul
b
Swammerdam Institute for Life Sciences, University of Amsterdam, Nieuwe Achtergracht 166, 1018 WV Amsterdam, Netherlands
b
Food Processing Group, Unilever Research and Development, Olivier van Noortlaan 120, 3133 AT Vlaardingen, Netherlands
Received 3 January 2002; accepted 14 February 2002
First published online 21 March 2002
Abstract
The cell wall of Saccharomyces cerevisiae is an elastic structure that provides osmotic and physical protection and determines the shape
of the cell. The inner layer of the wall is largely responsible for the mechanical strength of the wall and also provides the attachment sites
for the proteins that form the outer layer of the wall. Here we find among others the sexual agglutinins and the flocculins. The outer
protein layer also limits the permeability of the cell wall, thus shielding the plasma membrane from attack by foreign enzymes and
membrane-perturbing compounds. The main features of the molecular organization of the yeast cell wall are now known. Importantly, the
molecular composition and organization of the cell wall may vary considerably. For example, the incorporation of many cell wall proteins
is temporally and spatially controlled and depends strongly on environmental conditions. Similarly, the formation of specific cell wall
protein^polysaccharide complexes is strongly affected by external conditions. This points to a tight regulation of cell wall construction.
Indeed, all five mitogen-activated protein kinase pathways in bakers’ yeast affect the cell wall, and additional cell wall-related signaling
routes have been identified. Finally, some potential targets for new antifungal compounds related to cell wall construction are
discussed. 0 2002 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved.
Keywords : Yeast; Cell wall ; Cell wall protein ; Stress; Candida albicans
Contents
1.
2.
3.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Cell wall architecture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.1. Composition and properties of the cell wall . . . . . . . . . . . . . . . . . . . .
2.2. The L1,3-glucan network . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.3. Chitin as an intrinsic part of the lateral walls . . . . . . . . . . . . . . . . . .
2.4. L1,6-Glucan . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.5. Cell wall proteins (CWPs) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.6. CWP^polysaccharide complexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.7. Molecular organization of the cell wall . . . . . . . . . . . . . . . . . . . . . . .
2.8. Predictive value of the cell wall model for ascomycotinous yeasts . . . .
2.9. Identi¢cation of cell wall mutants . . . . . . . . . . . . . . . . . . . . . . . . . . .
Cell wall dynamics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.1. Cell cycle-related changes in the cell wall . . . . . . . . . . . . . . . . . . . . . .
3.2. Stationary phase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.3. Activation of the Slt2/Mpk1 MAP kinase pathway by cell wall stress .
3.4. Ultrastructural changes in the wall in response to cell wall defects . . .
3.5. Control of GSC2/FKS2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.6. The Hog1 MAP kinase pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.7. Responses to anaerobic conditions . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.8. Responses to extreme environmental pH values . . . . . . . . . . . . . . . . .
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* Corresponding author. Tel. : +31 (20) 525 7834; Fax: +31 (20) 525 7056.
E-mail address : klis@science.uva.nl (F.M. Klis).
0168-6445 / 02 / $22.00 0 2002 Federation of European Microbiological Societies. Published by Elsevier Science B.V. All rights reserved.
PII : S 0 1 6 8 - 6 4 4 5 ( 0 2 ) 0 0 0 8 7 - 6
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4.
Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
250
Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
250
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
250
1. Introduction
This review focuses on recent advances in our understanding of the molecular organization of the cell wall of
Saccharomyces cerevisiae and on the various protein^polysaccharide complexes found in its cell wall. We further
discuss changes in composition and organization of the
wall in response to cell wall stress and to various environmental conditions. Cell wall construction is tightly regulated, but with a few exceptions the underlying control
mechanisms are still fragmentarily known. We will explore
some important ¢ndings in this area. For earlier general
reviews we refer to [1^3]. Other more specialized reviews
will be mentioned in the relevant sections.
are transferred to hypertonic solution, they rapidly shrink,
and depending on the osmotic stress they may lose more
than 60% of their initial volume. This process is reversible.
When transferred back to the original medium, the cells
immediately expand to their initial volume. This elasticity
of the wall is probably due to the elastic properties of the
L1,3-glucan chains (see Section 2.2). The elasticity of the
wall explains why the wall of living cells is much more
permeable than isolated cell walls. Whereas isolated walls
are permeable only to molecules of molecular mass up to
760 Da [30], walls of living cells are permeable to much
larger molecules especially under hypotonic conditions
and also depending on growth conditions [22,31,32].
2.2. The L1,3-glucan network
2. Cell wall architecture
2.1. Composition and properties of the cell wall
The cell wall is a sturdy structure providing physical
protection and osmotic support [4]. Electron microscopic
analysis of the wall using negative staining reveals a layered structure with an electron-transparent internal layer
of about 70^100 nm thick depending on growth conditions
and genetic background, and an electron-dense outer layer
[5,6]. In brewing yeast the electron-transparent inner layer
may be as thick as 200 nm [7]. The mechanical strength of
the wall is mainly due to the inner layer, which consists of
L1,3-glucan and chitin, and represents about 50^60% of
the wall dry weight (Table 1). The outer layer, which consists of heavily glycosylated mannoproteins emanating
from the cell surface [5,18], is involved among others in
cell^cell recognition events [5,19^21]. It also limits the accessibility of the inner part of the wall and the plasma
membrane to foreign enzymes such as cell wall-degrading
enzymes in plant tissue [22^24]. The carbohydrate side
chains of the cell surface proteins contain multiple phosphodiester bridges, resulting in numerous negative charges
at the cell surface at physiological pH values [25]. These
side chains are responsible for the hydrophilic properties
of the wall, and may be involved in water retention and
drought protection. The outer protein layer accounts for
about one third of the wall dry weight. Cell wall proteins
are covalently linked to the L1,3-glucan^chitin network
either indirectly through a L1,6-glucan moiety or directly
(Sections 2.6 and 2.7). In addition, some proteins are disul¢de-bonded to other cell wall proteins [26,27].
The cell wall is highly elastic [28,29]. When yeast cells
The mechanical strength of the cell wall is mainly due to
L1,3-glucan, which can be speci¢cally stained with aniline
blue [33,34]. L1,3-Glucan chains belong to the so-called
hollow helix family; in other words, they have a shape
comparable to a £exible wire spring that can exist in various states of extension [35]. This property explains the
above-mentioned elasticity of the cell wall. Using magicangle spinning 13 C-NMR on living cells, Krainer and coworkers have found that a portion of the L1,3-glucan indeed assumes a helical structure [36].
L1,3-Glucan is only slightly crystalline in lateral walls
[37]. In stationary phase cells, L1,3-glucan molecules were
found to consist of about 1500 glucose monomers [8]. This
may, however, be an underestimation as a result of partial
hydrolysis of glucan chains during the extraction procedure. Considerably higher degrees of polymerization have
been reported, depending on the type of acid used for
extraction [38]. In addition, the degree of polymerization
may depend on environmental conditions, because yeast
uses two L1,3-glucan synthase complexes, to an extent that
depends on growth phase and carbon source [39]. In their
mature form, L1,3-glucan chains of stationary phase cells
are moderately branched and contain about 3^4% L1,6linked glucose residues [16]. Conceivably, the degree of
branching of L1,3-glucan may also depend on growth conditions.
The moderate degree of branching in mature L1,3-glucan molecules prevents the extensive crystallization seen
on the surface of regenerating spheroplasts [6,37,40].
This is also consistent with the X-ray di¡raction data of
isolated walls, which point to a low level of crystalline
L1,3-glucan [37]. On the other hand, the presence of uninterrupted chain segments of substantial length permits
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stable interchain association, thus allowing the formation
of a three-dimensional network [16,40]. Consistent with
this, electron microscopic studies of isolated walls using
negative staining reveal a ¢ne network with meshes of
about 20^60 nm wide [37]. As isolated walls are in a
non-extended state, the pores of the glucan layer in living
cells are expected to be wider, explaining why the permeability of the inner wall is only limiting for very large
proteins [41].
The two L1,3-glucan synthase complexes found in yeast
contain either Fks1 or Gsc2/Fks2 depending on environmental conditions. Both are multiple-spanning transmembrane proteins that are essential for the synthesis of L1,3glucan [39] and the corresponding genes are well conserved
among fungi [42,43]. It is generally assumed that Fks1 and
Gsc2 represent the catalytic subunits, but this has still to
be validated experimentally, because, for example, they
lack the known UDP-Glc binding site (K/RXGG) found
in glycogen synthase [44] and in an K1,3-glucan synthase
in Schizosaccharomyces pombe [45]. Alternatively, they
may represent a pore-forming protein that guides the
newly formed chain through the plasma membrane to
the outside. An elegant electron microscopic study, using
snap-frozen freeze-etched cells, has identi¢ed particles in
the plasma membrane, from which, on close examination,
¢brils with a diameter of 5 nm can be seen emerging [46].
These ¢brils disappear into the inner regions of the cell
wall, suggesting that they may represent L1,3-glucan
¢brils. A follow-up of this older work in combination
with the use of molecular genetic tools promises a more
complete understanding of in vivo L1,3-glucan synthesis.
Interestingly, a plant homologue of FKS1 has been identi¢ed in cotton ¢bers [47].
It is unknown whether growing L1,3-glucan chains in
yeast are extended at the reducing or non-reducing end.
However, L1,3-glucan from the fungus Sclerotium rolfsii is
extended at the non-reducing end [48]. Also, bacterial cellulose, diatom chitin, and plant homogalacturonan seem
to be extended at their non-reducing ends [49^51], suggesting that this is generally the case for processive glycosyl-
241
transferases such as L1,3-glucan synthase. This would
make the reducing end of the growing chain immediately
available for coupling to Pir cell wall proteins (see Section
2.5) and for processing enzymes such as Bgl2 [52,53].
As L1,3-glucan chains are synthesized as linear chains
[54], the presence of branches in the mature molecules
implies the existence of L1,3-glucan processing enzymes.
Additional enzymes may be needed for integrating newly
synthesized L1,3-glucan molecules into the growing wall
during isotropic growth. Enzymes potentially involved in
these steps are Gas1, an endotransglycosylase that may be
involved in extending and rearranging L1,3-glucan chains
[55,56], Bgl2, an endotransglycosylase that introduces intrachain L1,6-linkages [52,57], and, possibly, also the Crh1
[58] and the SUN family [57,59]. Their exact function in
vivo is, however, still incompletely understood.
2.3. Chitin as an intrinsic part of the lateral walls
Chitin is believed to occur as linear chains in the chitin
ring, in and around the bud scars and also to a minor
extent uniformly dispersed in the lateral walls of the mother cell [60^62]. Chitin isolated from bud scars consists of
about 190 N-acetylglucosamine monomers [17], but it is
unknown whether this is also a valid estimate for chitin in
the lateral walls. Calco£uor white is a £uorescent, anionic
dye that preferentially binds to L1,4-glucans such as chitin,
chitosan, and cellulose. It reacts to a lesser extent with
L1,3-glucan [63,64]. In S. cerevisiae, Calco£uor white is
frequently used to visualize chitin.
Chitin synthesis in S. cerevisiae involves three chitin
synthases and is tightly regulated [1,62]. For example,
under normal conditions the deposition of chitin in the
lateral walls takes place after cytokinesis [61]. This results
in a relatively low chitin level in the lateral walls: 0.1^0.2%
excluding the bud scar(s) and 1^2% including the bud
scars (see also Table 1). In cells with a (genetically) weakened wall, however, chitin synthesis is activated as part of
a salvage mechanism and the levels in the lateral walls of
those cells may become as high as 20% of the wall dry
Table 1
Cell wall macromolecules in S. cerevisiae
Macromolecule
Wall dry wt. (%)
Site of synthesis
Mannoproteins
L1,6-Glucan
L1,3-Glucan
Chitin
35^40
5^10
50^55
1^2
Secretory pathway
(PM)
PM
PM
Mature form
Refs.
DP
Branching
200a
140
1500
190b
Highly branched
Highly branched
Moderately branched
Linear
[13]
[12,14,15]
[8,16]
[17]
The data regarding the cell wall composition have been compiled from [8^10]. The mannoproteins are often heavily glycosylated; the protein content
per se of isolated SDS-extracted walls accounts for only a small percentage of cell wall dry weight. The cell wall components are presented in the order
in which they occur in the cell wall from the outside to the inside. The cell wall of C. albicans has a similar cell wall composition [11]. DP, degree of
polymerization; PM, plasma membrane. The site of synthesis of L1,6-glucan is uncertain [12]. Note that the data presented here may vary depending
on growth conditions.
a
N-chains.
b
Chitin from bud scars.
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synthases behave similarly, the reducing end of a chitin
chain is directly available for coupling to the acceptor sites
of L1,3- and L1,6-glucan molecules [70,71], when it
emerges from the pore through the plasma membrane.
2.4. L1,6-Glucan
Fig. 1. Schematic representation of the GPI-CWP Ccw12. The mature
form of Ccw12 is estimated to consist of less than 100 amino acids.
Ccw12 has three predicted attachment sites for N-chains. As N-chains
may contain up to 200 mannose residues each [13], they may together
have a molecular mass of about 90 kDa. Interestingly, CCW12 has a
codon adaptation index (CAI) of 0.87, indicating that it encodes a very
abundant protein. Deletion of CCW12 results in a reduced growth rate
of about 40% and increased sensitivity to Calco£uor white and Congo
red [128]. SP, signal peptide ; grey box, GPI anchor addition signal.
weight (see also Section 3.4) [10,66]. As to be expected for
a salvage mechanism, chitin is then also deposited in the
wall of the growing bud [66]. The responsible synthase is
encoded by CHS3 [67]. Chs3 is a multi-spanning membrane protein with its active site at the cytosolic side of
the plasma membrane [68]. Like other fungal chitin synthases, it is a processive enzyme with the diagnostic motif
D,D,D,QRRRW [69]. Diatom chitin synthase extends the
growing chitin chain at the non-reducing end using UDPGlcNAc as substrate [50]. Assuming that fungal chitin
L1,6-Glucan is in its mature form a highly branched,
water-soluble polymer consisting on average of about
130 glucose monomers [14]. It is unknown whether L1,6glucan is synthesized as a water-insoluble linear molecule
similar to L1,3-glucan [72], or as a water-soluble branched
molecule similar to the mixed L1,3-/L1,6-glucan produced
by S. rolfsii [48]. This is particularly relevant regarding the
development of an in vitro assay for L1,6-glucan synthase.
L1,6-Glucan is used in the cell wall to connect GPI-dependent cell wall proteins to the L1,3-glucan network (see
Fig. 4, Section 2.7). It may also function as acceptor site
for chitin, particularly in case of cell wall stress (see Section 2.6 and Fig. 2) [71,73]. The pioneering studies by
Bussey and co-workers have identi¢ed several genes that
a¡ect L1,6-glucan levels in the cell wall [74]. From these
studies it has emerged that various ER-resident proteins,
Golgi-resident proteins, and cell surface proteins strongly
a¡ect L1,6-glucan levels in the cell wall. This might be
interpreted in various ways: (i) the biogenesis of L1,6-glucan is a stepwise process that begins in the ER; (ii) the
synthesis of L1,6-glucan takes place at the plasma membrane but requires the stepwise synthesis of a primer; or
(iii) the synthesis of L1,6-glucan takes place at the plasma
membrane but the activity of the synthesizing complex is
highly sensitive to various defects in the secretory path-
Table 2
Known or putative functions of CWPs
Function
Genes
Refs.
Presentation of mannan chains
Cell wall permeability
Retention of water
Adhesion
Mating
Flocculation
Invasive/pseudohyphal growth
Bio¢lm formation
Cell wall processing
Glucan remodeling
Cell separation
Cell wall strengthening
Isotropic growth
Growth conditions
Sporulation
Stationary phase
Anaerobic growth
Low temperature
Metabolism
Esterase
Iron uptake
CCW12, YDR134C
Fig. 1
[22,24,41]
Section 2.1
SAG1, AGA1, AGA2, FIG2
FLO1, FLO5, FLO9, FLO10
FLO11
FLO11
[5,77,107^109]
[20,109^111]
[109]
[21]
CRH1, CRH2, CRR1, FIG2 (speculative), BGL2
EGT2, PRY3, CTS1, DSE2, DSE4, SCW11
CWP1, PIR1, PIR2/HSP150, PIR3, PIR4/CIS3
PIR1, PIR2/HSP150, PIR3, PIR4/CIS3
[52,58,106,108]
[57,100,112]
[65,113^116]
Section 3.1; 117
SPO1
SED1, SPI1
DAN1, DAN4, TIP1, TIR1, TIR2, TIR3, TIR4
TIP1, TIR1, TIR2, TIR4
[118]
[119,120]
[121^123]
[121,122,125]
TIP1
FIT1, FIT2, FIT3
[126]
[127]
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Fig. 2. Overview of the CWP^polysaccharide complexes in yeast. GPICWP, cell wall protein linked through a GPI remnant to L1,6-glucan
(L1,6-Glc); Pir-CWP, a cell wall protein from the PIR family linked
through an alkali-sensitive linkage to L1,3-glucan (L1,3-Glc). Kollar and
co-workers have identi¢ed several of the interconnecting linkages between cell wall macromolecules [70,71], but the interpolymer linkages
between GPI-CWP and Pir-CWP on the one hand and L1,3-glucan on
the other hand are unknown.
way. Using immunogold labeling, Montijn and co-workers
could not detect intracellular L1,6-glucan, not even in a
temperature-sensitive mutant that had accumulated secretory vesicles at the restrictive temperature for 2 h [12]. In
contrast, plasma membrane-derived vesicles and the cell
walls reacted strongly. This seems to exclude the ¢rst hypothesis. Because an in vitro assay for L1,6-glucan is not
yet available, di¡erentiation between the two other hypotheses is not simple, the more so since they are not
mutually exclusive. Summarizing, a clear picture of how
L1,6-glucan is synthesized is still lacking.
2.5. Cell wall proteins (CWPs)
The mannoproteins that form the outer cell wall layer
are highly glycosylated with a carbohydrate fraction that
often amounts to over 90% (w/w) [1]. Biotinylation of
intact cells using a sulfonated derivative of biotin that
does not cross the plasma membrane is a convenient
tool to distinguish between authentic cell wall proteins
and adventitiously bound proteins [75]. The outer layer
of mannoproteins is much less permeable to macromolecules than the internal ¢brillar layer [24]. This is largely
due to the presence of the long and highly branched carbohydrate side chains linked to asparagine residues [1,22]
and to the presence of disul¢de bridges [32]. Serine and
threonine residues, which may carry short oligomannosyl
chains, are often clustered, resulting in relatively rigid rodlike regions of the polypeptide backbone [76,77]. Finally,
due to phosphodiester bridges in both N- and O-linked
mannosyl side chains the cell surface of yeast contains
numerous negative charges [1,25]. Interestingly, the sensitivity of yeast cells to the antifungal plant protein osmotin,
which has a high isoelectric point and thus is positively
charged at physiological pH values, depends on the presence of mannosyl phosphate groups at the cell surface [78].
In addition, the cell wall may bind positively charged,
cytosolic proteins originating from lysed cells. Phosphodiester groups can be visualized with Alcian blue [25].
There are two main classes of proteins covalently
coupled to cell wall polysaccharides (Fig. 4, see Section
2.7).
(a) GPI-dependent cell wall proteins (GPI-CWPs). They
243
are generally indirectly linked to L1,3-glucan through a
connecting L1,6-glucan moiety. In the genome of S. cerevisiae about 60^70 GPI proteins have been identi¢ed ([79];
J.G. De Nobel, unpublished data). About 40 of them are
destined for the plasma membrane whereas the others become covalently linked to L1,6-glucan [71,80^84]. They
often contain repeats and serine- and threonine-rich regions. The most extensively studied GPI-CWP is Sag1,
which is involved in sexual agglutination [5,77,84,85]. Mature proteins only have a remnant of the original GPI
anchor, that links them to L1,6-glucan [71,81]. The core
structure of this remnant [71] is formed by ^ethanolamine^
Pi ^(Man)4 ^, and is probably substituted with additional
ethanolamine phosphate groups [86^88]. Interestingly,
L1,6-glucan extracted from cell walls by hot acetic acid
may contain a minor amount of galactose [14]. Conceivably, in some genetic backgrounds this is part of the GPI
anchor remnant of GPI-CWPs [89]. The presence of galactose in the cell wall of S. cerevisiae is consistent with the
evidence for a UDP-galactose transporter in bakers’ yeast
[90].
(b) Pir proteins (Pir-CWPs). They are presumably directly linked to L1,3-glucan through an alkali-sensitive
linkage. In S. cerevisiae a family of four such proteins
has been found [65,91,92]. They are all similarly organized
(SP^Kex2^repeat(s)^CX(66)CX(16)CX(12)C), consisting
of an N-terminal signal peptide, a Kex2 site, followed by
a repeat-containing region with up to 11 repeats, and a
highly conserved carboxy-terminal region with four cysteine residues in a conserved spacing pattern. Pir1, Pir2/
Hsp150, Pir3, and Pir4/Cis3 all have been localized to the
cell wall immunologically [27,93,94]. Several additional
proteins, such as Pau1 and its homologues, and Sps100,
which is believed to contribute to maturation of the spore
wall [95], and also Ygp1, which is induced by nutrient
limitation [96], are predicted to have an N-terminal signal
peptide, but not an addition signal for a GPI anchor.
Possibly, their ¢nal destination is not the medium, but
the cell wall, to which they may become linked in a PirCWP-like fashion.
In addition, several cell wall proteins such as Bar1, a
protease [27], Aga2 [5,97], the active subunit of the sexual
agglutinin complex in MATa cells, Pir4/Cis3 [27], and
some known or potential cell wall glycanases such as
Fig. 3. Hypothetical linkage between a Pir-CWP and a L1,3-glucan
polymer. According to this scheme a L1,3-glucan molecule is linked with
its reducing end to an oligomannoside, O-linked to a serine or threonine
residue of a Pir-CWP. The glycopeptide linkage is alkali-sensitive.
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Fig. 4. Putative molecular organization of the cell wall of S. cerevisiae.
The mechanical strength of the wall is due to an elastic three-dimensional network of L1,3-glucan molecules kept together by hydrogen bonding
between locally aligned chains [40]. As the arrows in the model indicate,
the terminal non-reducing ends of the L1,3-glucan side chains are believed to function as acceptor sites for L1,6-glucan and chitin, whereas
the reducing end of the L1,3-glucan molecules may be involved in the
linkage to Pir-CWPs. The L1,3-glucan layer forms the inner layer of the
cell wall, whereas cell wall proteins form the outer layer. The two most
abundant CWP^polysaccharide complexes are shown here : GPI-CWP^
L1,6-glucan^L1,3-glucan and Pir-CWP^L1,3-glucan. The L1,6-glucan
molecules are highly branched and thus water-soluble, tethering GPICWPs to the L1,3-glucan network. GPI-CWPs represent the major component of the outer protein layer. The Pir-CWPs are directly linked to
the L1,3-glucan network through an alkali-sensitive linkage and are distributed throughout the L1,3-glucan network. This model is based on
[2,65,70,71,73,80,115,137]. The model is believed to be valid for C. albicans as well [11]. GPI-CWP, cell wall protein linked through a GPI
remnant to L1,6-glucan (L1,6-Glc); Pir-CWP, a cell wall protein from
the PIR family linked through an alkali-sensitive linkage to L1,3-glucan.
Sun4/Scw3 [57], can be released from intact cells using a
reducing agent. This suggests that they might be disul¢delinked to other cell wall proteins. Reducing agents are also
expected to release soluble, intermediate forms of GPICWPs [98]. Finally, SDS extraction of isolated walls releases many proteins. With a few exceptions, like the
transglucosylase Bgl2 [52] and the chitinase Cts1
[99,100], they are not authentic cell wall proteins and their
presence is due to contamination with membrane fragments [11,57, 101].
Members of the Hsp (heat-shock protein) family [102]
and abundant glycolytic enzymes such as Tdh1, Tdh2,
and Tdh3 [103] are often found at the cell surface. They
can be extracted from intact cells with a reducing agent
such as mercaptoethanol under slightly alkaline conditions, suggesting that they are either trapped inside the
wall or are ionically bound to cell surface proteins. It is
not clear whether these proteins originate from lysed cells
or as frequently claimed are exported by a non-conventional secretory mechanism [102^104]. Heat-shock proteins
and glycolytic enzymes have also been found in the medium of regenerating spheroplasts, which raises the same
issue [104^106].
Cell wall proteins may have various functions, which are
summarized in Table 2. In many cases their precise function is unknown. Some proteins, like the very small but
presumably abundant GPI-CWPs Ccw12 and Ydr134c
with a predicted unprocessed size of 133 and 66 amino
acids, respectively, and a codon adaptation index of
0.870 and 0.646, respectively, may be used as a means to
present mannan to the cell surface (Fig. 1). Various GPICWPs are involved in adhesion events like sexual agglutination and £occulation of yeast cells. Others appear to
have an enzymatic function like Crh1, Crh2, and Crr1
[58]. Fig. 2 is required for the normal width of the conjugation tube [108], suggesting that it may be involved in
remodeling of L-glucan in the conjugation tube.
The expression of PIR genes is also up-regulated in case
of cell wall stress (see Section 3.3) [113,114], consistent
with the idea that their gene products might be involved
in cell wall strengthening. Disruption of all four genes
results in swollen cells that grow slightly slower and are
more sensitive to Calco£uor white and Congo red, indicating that their cell wall is indeed weakened [92]. Interestingly, cells that overexpress PIR2 are more resistant to the
plant antifungal protein osmotin whereas cells that lack
PIR1, PIR2, and PIR3 are more sensitive to it [93]. Di¡erences in tolerance to osmotin depend on the presence of an
intact cell wall, because after spheroplasting the cells are
equally sensitive [88]. Possibly, Pir-CWPs make the cell
wall less permeable to osmotin and other proteins.
2.6. CWP^polysaccharide complexes
Analysis of CWP^polysaccharide complexes requires a
relatively small set of reagents. Recombinant L1,3-glucanase and L1,6-glucanase are now commercially available.
For detection, antibodies speci¢c for L1,3-glucan are commercially available, whereas antibodies speci¢c for L1,6glucan are easy to raise [129]. The lectins concanavalin
A for mannoproteins and wheat germ agglutinin for chitin
oligomers further complement this list.
Cell wall proteins may be linked to the L1,3-glucan network in various ways (Fig. 2). In cells grown in rich medium the GPI-CWP^L1,6-glucan^L1,3-glucan complex is
the most abundant one (Fig. 2A). It consists of cell wall
proteins such as Ccw12, Tip1, Ssr1, Cwp1, or Sag1 covalently linked through a GPI remnant to a non-reducing
end of L1,6-glucan, which in turn is linked to a non-reducing end of L1,3-glucan [2,71,80,81]. Because the GPI
remnant (core structure: ^ethanolamine^Pi ^(Man)4 ^) contains a phosphodiester bridge, GPI-CWPs in this complex
can be speci¢cally released by using aqueous hydro£uoric
acid [80]. Pir-CWP^L1,3-glucan represents another, abundant CWP^polysaccharide complex in which a protein
from the PIR family is linked through an alkali-sensitive
linkage to L1,3-glucan (Fig. 2D) [65,75,92]. The nature of
this linkage is not known, but in view of its sensitivity to
mild alkali it is tempting to postulate that it involves an
O-linked side chain. Interestingly, immunogold labeling of
Pir-CWPs shows that the signal is uniformly dispersed
over the chitin^L1,3-glucan layer of the cell wall [94]. In
contrast, immunogold labeling of the GPI-CWPs Sag1 and
Flo1 results in strong labeling of the outer ¢brillar wall
layer [5,110]. This indicates that Pir-CWPs are part of the
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inner layer. A possible interpretation of these observations
is presented in Fig. 3, in which it is proposed that a L1,3glucan molecule is covalently linked through its reducing
end to an O-linked side chain of a Pir-CWP, possibly
present in the latter’s internal repeats.
Some GPI-CWPs such as Cwp1 may also be bound
directly to L1,3-glucan through an alkali-sensitive bond
presumably in a Pir-CWP-like fashion [115]. As a result,
two additional GPI-CWP^polysaccharide complexes can
be distinguished, which are either single- or double-linked
(Fig. 2B,C) [115]. Importantly, Cwp1 and possibly also
other potentially double-linked GPI-CWPs play an important role in the response of cells towards cell wall stress
[113,116]. Finally, a ¢fth complex has been identi¢ed in
which a GPI-CWP is linked to a L1,6-glucan molecule,
which is directly interconnected to chitin (Fig. 2E).
Although this type of complex is relatively rare in cells
growing on rich medium, under special conditions it can
become much more prominent (see Section 3.4 and Table
3) [73]. Interestingly, chitin in the lateral walls, which represents V10% of the total chitin, seems to be speci¢cally
associated with L1,6-glucan [60]. This may mean that chitin in the lateral walls is predominantly present in the form
of the complex GPI-CWP^L1,6-glucan^chitin. Finally, chitin is also directly linked to L1,3-glucan [70]. Chitin is,
however, omitted from the ¢rst four complexes shown in
Fig. 2, because (i) such complexes may exist without a
chitin molecule attached to it, for example, in the growing
bud, and (ii) a chitin molecule may become linked to another L1,3-glucan molecule than the one shown in the
various cell wall protein^polysaccharide complexes, i.e.
the linkage between L1,6-glucan and chitin may be quite
indirect.
As the synthesis of chitin and L1,3-glucan takes place at
the plasma membrane [54,68] and the coupling of GPICWPs to L1,6-glucan also occurs outside the plasma membrane [12,84], the interconnections between the cell wall
macromolecules also have to be made outside the plasma
membrane. This strongly points to the existence of various
classes of cell wall assembly enzymes ^ presumably transglycosylases ^ responsible for interconnecting (a) GPICWPs to L1,6-glucan, (b) L1,6-glucan to L1,3-glucan, (c)
chitin to L1,3-glucan, (d) chitin to L1,6-glucan, and (e) cell
wall proteins directly to L1,3-glucan through an alkali-sen-
245
sitive linkage. Interestingly, cells growing in the presence
of gentiobiose, which consists of two L1,6-linked glucose
monomers, incorporate a GPI-CWP reporter protein less
e⁄ciently into the cell wall than wild-type cells, and this is
accompanied by increased sensitivity to Zymolyase [136].
This suggests that gentiobiose inhibits a cell wall assembly
step involving L1,6-glucan. At present, none of the postulated assembly enzymes has been identi¢ed. Also, the sequence of coupling events is largely unknown (see Section
2.7).
2.7. Molecular organization of the cell wall
The cell wall forms a bilayered, supramolecular structure that surrounds the entire cell (Fig. 4). Its mechanical
strength is largely based on an internal layer of moderately
branched L1,3-glucan molecules that form a three-dimensional network that is kept together by hydrogen bonding
between laterally associated chains [40]. Methylation analysis has revealed that each L1,3-glucan polymer has multiple side chains [16]. Their terminal non-reducing ends are
believed to function as acceptor sites for L1,6-glucan and
chitin [70,71], whereas the reducing end of the L1,3-glucan
molecules may also be involved in the linkage to PirCWPs (see Section 2.6) [115]. Methylation analysis of mature L1,6-glucan has shown that it is a highly branched
molecule [14], which explains why it is water-soluble
[15,40]. Chitin is deposited in the lateral walls after cytokinesis has taken place, probably sti¡ening the cell wall
[61]. It represents less than 10% of the total chitin content
of the walls and may be coupled to either L1,3-glucan or
L1,6-glucan [70,71].
The biosynthetic pathway of the sexual agglutinin Sag1
has been studied in detail and several intermediate glycoforms have been identi¢ed including two extracellular glycoforms, namely, a plasma membrane-bound form and a
later, soluble form [84,98]. In contrast to the mature wallbound form, both extracellular intermediates are not yet
linked to L1,6-glucan. This suggests that the formation of
the GPI-CWP^L1,6-glucan^L1,3-glucan complex proceeds
as follows : (i) L1,6-glucan is connected to L1,3-glucan, and
(ii) a GPI-CWP is coupled to L1,6-glucan. Nothing is
known about how the formation of other CWP^polysaccharide complexes proceeds.
Table 3
Changes in cell wall composition and organization in response to genetically caused cell wall defects
Defect
Response
Refs.
Less L1,3-glucan
More chitin in the lateral walls
More GPI-CWP-L1,6-glucan-chitin
More GPI-CWPs in the medium
More Pir-CWPs in the wall
More L1,3-glucan and chitin in the wall
Possibly, increased usage of the alkali-sensitive linkage between GPI-CWPs and
L1,3-glucan
More chitin in the lateral walls
[10,73,116,130,131]
Less L1,6-glucan
Defective N- and O-glycosylation
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[65,84]
[2,132^135]
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GPI-CWPs, Pir-CWPs, as well as disul¢de-linked cell
wall proteins all have been successfully used to target fusion proteins to the cell wall for covalent attachment
[27,83,85,138^145].
Finally, there is clear evidence for limited release of cell
wall proteins into the medium [146], possibly as a result of
remodeling of the cell wall during isotropic growth or
when a new bud is formed.
2.8. Predictive value of the cell wall model for
ascomycotinous yeasts
There is strong evidence that the cell wall of Candida
albicans is similarly organized as the wall of S. cerevisiae
[11]. The model presented here seems also valid for other
Candida species. For example, in Candida glabrata an authentic GPI-CWP involved in adhesion to human epithelial cells has been identi¢ed that, after expression in
S. cerevisiae cells, permits the latter to e⁄ciently adhere
to epithelial cells [147]. Pir-CWPs have been identi¢ed in
C. albicans [94,148], Kluyveromyces lactis [91], and in Zygosaccharomyces rouxii [91]. The cell wall model presented
here is only partially valid for the ¢ssion yeast Schizosaccharomyces pombe. First, unlike S. cerevisiae, the wall of
S. pombe contains a considerable amount of K1,3-glucan.
Second, Southern analysis did not reveal PIR-like genes in
S. pombe [91]. For a more detailed discussion of the cell
wall of mycelial fungi from the Ascomycotina, see
[42,149].
2.9. Identi¢cation of cell wall mutants
De Groot and co-workers have shown that cell wall
construction in S. cerevisiae is a tightly regulated process
and that about 1200 genes directly or indirectly a¡ect formation of the cell wall [150]. Using a hierarchical screening approach, the gene products can be divided in various
classes such as proteins involved in the synthesis of cell
wall macromolecules, proteins involved in remodeling or
interconnecting cell wall polymers and proteins involved in
the regulation of cell wall construction. Primary screens
for the detection of mutants with an impaired cell wall
are often based on increased sensitivity towards compounds that are known to interfere ^ either directly or
indirectly ^ with normal cell wall construction, such as
L1,3-glucanase [151,152], Calco£uor white [150^152] and
the related compound Congo red [150,153], SDS, and caffeine [150]. These compounds have in common that their
deleterious e¡ect on growth can be alleviated by osmotically stabilizing the medium [23,150,154,155]. Increased
sensitivity to sonication [150,156] and hypotonic shock
[157,158] has also been used to detect mutants with a
cell wall defect. 2-Deoxyglucose, which is incorporated in
the cell wall and causes cell lysis in growing areas, may
presumably also be e¡ectively used to identify cell wall
mutants [159,160]. Alternatively, the culture ¢ltrate may
be screened for increased amounts of cell wall proteins
or for incomplete CWP^polysaccharide complexes indicative of defective cell wall assembly [84,150]. In general, cell
wall perturbants appear to be more e¡ective at 37‡C than
at 30‡C, possibly because of the combined e¡ect of increased turgor pressure on the wall and of defective cell
wall construction due to thermal inactivation of cell wall
assembly enzymes at 37‡C (see Section 3.3).
Hypoglycosylation mutants represent a special category
of cell wall-related mutants. They are often more resistant
to orthovanadate and hypersensitive to hygromycin, which
facilitates their identi¢cation [13]. Defects in O-glycosylation can be easily detected using the heavily O-glycosylated endochitinase Cts1 as a marker protein [99], whereas
invertase is a useful marker for defects in N-glycosylation
[161]. For a more detailed analysis of O- and N-glycosylation, FACE (£uorophore-assisted carbohydrate electrophoresis) is useful [162].
Mutants with an endocytotic block show a rare cell
wall-related phenotype, namely, the mother cells of such
mutants form multiple cell wall layers [163].
3. Cell wall dynamics
An estimated 1200 genes of S. cerevisiae a¡ect the composition and organization of the cell wall [150]. Changes in
the newly formed cell wall occur depending on the phase
of the cell cycle, nutrient availability, and environmental
conditions such as pH, temperature, and the availability of
oxygen. Evidence is accumulating that all major signaling
pathways such as the mitogen-activated protein (MAP)
kinase pathways and the cAMP-dependent protein kinase
A (PKA) pathway directly or indirectly may a¡ect cell
wall construction. The Pkc1-controlled MAP kinase pathway (the Slt2/Mpk1 MAP kinase pathway) is directly involved in signaling cell wall damage. The expression of cell
wall proteins is particularly tightly controlled. For example, transcript levels of Cwp1, an abundant GPI-CWP, are
cell cycle-controlled, but they are also up-regulated during
late sporulation, under hypertonic conditions, and in case
of cell wall stress [132,133]. They are down-regulated during growth at low oxygen concentrations, low temperatures and during nitrogen depletion. This points to an
intricate network of signaling mechanisms controlling the
level of Cwp1. Regulation of the GPI-CWP Muc1/Flo11,
which is involved in pseudohyphal and invasive growth
and in bio¢lm formation (Table 2), is equally complex
[164^166]. These observations show that cell wall construction is a tightly regulated process.
3.1. Cell cycle-related changes in the cell wall
Pulse labeling of cells with FITC-concanavalin A, which
speci¢cally binds to mannoproteins, is an elegant means to
monitor cell surface growth during the cell cycle [167,168].
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This approach has revealed that cells growing in rich medium switch from polarized growth to isotropic growth
and vice versa during the cell cycle. Cells with a small
bud show mainly tip growth, but during the G2 ^M phase
they switch to isotropic growth [163,169]. As a result, they
acquire their characteristic oval shape. The formation of a
septum between mother and daughter cell directs cell wall
formation temporarily to the neck region. Released daughters tend to be smaller than mother cells [170]. They grow
mainly isotropically until they have reached a critical size
before a new budding cycle is initiated.
Immuno£uorescence labeling and GFP tagging studies
have shown that the expression of several cell wall proteins
is cell cycle regulated [27,58,100,171^173]. This has been
con¢rmed by transcript analysis, indicating that the expression of over 50% of all cell wall proteins is cell cycle
regulated [117,174]. For example, several cell wall proteins
belong to the SIC1 expression cluster of cell cycle-regulated genes. This cluster is named for its prototype gene
SIC1, which encodes an inhibitor of the Cdc28^Clb protein kinase complex. It comprises at least 27 genes that are
maximally expressed in late M and/or early G1 [117]. Using YFP (yellow £uorescent protein) reporter constructs
controlled by the promoter from the genes in the SIC1
cluster, Colman and co-workers have identi¢ed 10 genes
in this group that show daughter-speci¢c expression [100].
These include the endochitinase Cts1 [99], the putative
glucanases Scw11, Dse2 and Dse4 [57,100], and the GPICWP Pry3. The GPI-CWP Egt2, the loss of which causes
a delay in cell separation [112], also shows daughter-speci¢c expression, but only in cultures reaching saturation
[100]. In view of their enzymatic activities, these proteins
seem to be involved in cell separation by degradation of
the septum from the daughter-proximal side. This also
explains why the birth scar is much less obvious than
the bud scar. Consistent with this, chitinase has been speci¢cally localized in the cell wall at the daughter side of the
bud neck [100]. Daughter-speci¢c expression of this subset
of SIC1 genes involves preferential localization of the
transcription factor Ace2 in the nucleus of the daughter
cell, possibly as a complex with Cbk1 and Mob2 [100].
The Pir-CWP-encoding genes PIR1, PIR, and PIR3 also
belong to the SIC1 expression cluster and show a similar
expression pattern [117]. Because the encoded proteins follow the secretory pathway, they will be incorporated in the
cell wall during early G1 , which is a period of isotropic
growth. This raises the question whether they might have
a function related to this mode of growth. PIR4/CIS3, on
the other hand, is maximally expressed at the G2 phase
[117]. In agreement with this, Pir4/Cis3 is predominantly
found in the wall of growing buds [27]. The GPI-CWP
Crh1 shows a particularly interesting expression pattern.
It is deposited at the site of the future bud and, later, in
the neck region of large-budded cells, consistent with
peaking of transcript levels of CRH1 both in late G1
and in M/G1 [58].
247
Normally, the time of expression during the cell cycle
seems to correlate with where at the cell surface a cell wall
protein is incorporated. The GPI-CWP Cwp1, however,
behaves di¡erently [173]. Similar to the GPI-CWP Cwp2,
its transcript levels are maximal at G2 , but whereas Cwp2
is incorporated in the walls of the growing bud, Cwp1 is
incorporated later and targeted speci¢cally to the birth
scar of the daughter cell. For further discussion of these
and related topics, the reader is referred to [137,166,175].
3.2. Stationary phase
When cells enter the stationary phase, they form di¡erent walls. For example, they become thicker, more resistant to L1,3-glucanase digestion [22,23,119], and less permeable to macromolecules [22]. The GPI-CWP Sed1
becomes the most abundant cell wall protein [119], and
the transcript level of SPI1, which encodes a related
GPI-CWP [83], also strongly increases [120]. The level of
mannosyl phosphorylation of cell wall proteins increases
at the late-exponential and stationary phase of cell growth
[176]. Also, the number of disul¢de bridges in the wall
increases 6^7-fold [22]. Force-deformation studies with
single yeast cells entering stationary phase indicate that
they strengthen their walls by simply increasing wall thickness, but not by increased cross-linking [4]. Interestingly,
stationary phase cells have a considerably higher turgor
pressure, compared to exponentially growing cells [29],
which is probably caused by the strong increase in the
level of trehalose [177^179]. This is consistent with the
presence of a thicker cell wall.
Evidence is emerging that the Ras/cAMP-dependent signal transduction pathway modulates cell wall integrity.
Deletion of PDE2, which encodes the high a⁄nity
cAMP phosphodiesterase, results in up-regulation of this
pathway and in increased sensitivity to hypotonic shock,
indicating a loss of cell wall strength [158]. Conversely,
overexpression of PDE2, which results in down-regulation
of the pathway, suppresses the sorbitol dependence of a
mutant strain with fragile walls and thus seems to
strengthen the wall [158]. Similarly, overexpression of
PDE2 leads to earlier mannosyl phosphorylation of cell
wall proteins [176]. Entrance of the stationary phase is
accompanied by low PKA activity [180]. As PKA negatively regulates expression of genes with stress response
elements (STREs) in their promoter region [162], STREcontrolled genes, which include several cell wall-related
genes, will become more active, thus contributing to the
changes in cell wall properties observed in stationary
phase.
3.3. Activation of the Slt2/Mpk1 MAP kinase pathway by
cell wall stress
A GFP-tagged form of Pkc1, which drives the Slt2
MAP kinase pathway, is rapidly relocalized in cells chal-
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lenged with the cell wall-degrading enzyme Zymolyase.
Whereas in unchallenged cells, Pkc1 localizes in areas of
polarized growth, it moves out within minutes to multiple
spots outside this area [181]. Possibly, redistribution of
Pkc1 under cell wall stress conditions is involved in targeting the secretory vesicles to sites of cell wall damage. Similarly, the L1,3-glucan synthase complex is redistributed in
case of cell wall stress [182]. Both observations suggest
that the cell is capable of locally repairing the cell wall
in case of cell wall damage. The Slt2 MAP kinase pathway
itself is also activated by cell wall stress. For example
Zymolyase, which degrades the L1,3-glucan network, and
cell wall perturbing compounds such as Calco£uor white
or Congo red activate the Slt2 MAP kinase pathway
[23,183]. Ca¡eine, vanadate and SDS, which probably
indirectly weaken the cell wall, also activate the Slt2
MAP kinase pathway [155,184]. In addition, mutant cells
with a weakened cell wall such as fks1v and gas1v show
constitutive activation of the Slt2 MAP kinase pathway
[23].
When cells are transferred to a medium of lower osmotic strength, resulting in increased water uptake and
increased turgor pressure [185], an almost immediate activation of Slt2 is observed. Interestingly, the Slt2 MAP
kinase pathway is also activated by high growth temperatures (37^39‡C) [155,186]. Compared to the rapid activation of the Slt2 MAP kinase pathway in cells challenged
with hypotonic stress, which is maximal within 1 min and
then slowly tapers o¡ to the original state within 30 min,
this temperature stress response is delayed, becoming maximal after 15^30 min. This raises the question whether
activation of the Slt2 MAP kinase pathway may be an
indirect e¡ect of heat stress. The following observation is
pertinent to this question. When cells are transferred to
high growth temperatures (37^40‡C), the cytosolic concentration of trehalose increases rapidly within minutes and
continues to rise for up to 90 min to an estimated level of
over 0.5 M [187^191]. In addition, the intracellular glycerol level increases [192]. This raises the possibility that
these responses, and the concomitant rise in turgor pressure, are largely or entirely responsible for activating the
Slt2 MAP kinase pathway during heat stress. These observations can also explain why mutations in a member of
the Slt2 MAP kinase pathway result in cell lysis only at
high growth temperatures, but not at lower temperatures.
Conceivably, thermal inactivation of proteins involved in
cell wall construction may also result in cell wall weakening, thus aggravating the e¡ect of increased turgor pressure. Finally, in cells treated with K-pheromone not only
the Fus3 MAP kinase pathway is activated but also the
Slt2 MAP kinase pathway. This is again a late response
coinciding with the appearance of the mating structure
[183,193^195]. It seems likely that the extensive remodeling
of the cell wall required to form a mating structure leads
to local softening of the wall, thus explaining the activation of the cell wall salvage pathway in the presence of
pheromone. At the base of the mating structure chitin is
deposited [183,196]. Conceivably, the cell forms a zone of
CWP-GPI^L1,6-glucan^chitin complexes in this area.
It is not precisely known how the cell detects cell wall
stress. The observations discussed in the previous section
can be most easily explained by assuming that the cell
actually responds to acute plasma membrane stretch.
This is also consistent with the observation by Kamada
and co-workers [186] who found that chlorpromazine,
which is supposed to cause stretching of the plasma membrane, also activates the Slt2 MAP kinase pathway. Membrane stretch is accompanied by opening of stretch-activated membrane channels [197] and a rapid increase in the
cytosolic levels of free Ca2þ [198]. However, it is not
known whether these responses are related to the cell
wall salvage pathway. It is conceivable that an additional
pathway may be activated using the Ca2þ /calmodulinregulated protein phosphatase calcineurin, which co-regulates the transcript level of FKS2/GSC2 and thus in£uences the synthesis of L1,3-glucan (see Section 3.5) [199].
Several putative sensor proteins for cell wall stress have
been identi¢ed that function upstream from the Slt2 MAP
kinase pathway, such as the type I transmembrane proteins Slg1, Wsc2/3/4, Mid2, and Mtl1 [183,195,200]. They
are located in the plasma membrane and uniformly distributed over the cell surface [195,200] and act through
Rom2, a guanine nucleotide exchange factor for Rho1
[201]. Of these, Mid2 seems to play a major role in sensing
cell wall stress and in the coupled activation of the MAP
kinase Slt2 [23,155]. Mid2 is a single span transmembrane
protein with its C-terminus in the cytosol and its N-terminus outside the plasma membrane. An important feature
of the part of the protein outside the plasma membrane is
the presence of an extended serine- and threonine-rich
region, which is essential for its function [183,201]. This
is consistent with the observation that O-glycosylation of
Mid2 by the protein mannosyl transferase Pmt2 is essential for its signaling function [201]. Conceivably, Mid2,
and also the other cell wall integrity sensors, which have
a similar organization, are linked to the L1,3-glucan network through one or more of its O-linked side chains, in a
Pir-CWP-like fashion.
Recently, it has been reported that mutations in SLG1/
WSC1 and the other WSC genes result in increased sensitivity towards hydrogen peroxide and ethanol [202]. Intriguingly, these treatments are known to induce the intracellular accumulation of trehalose [187], and, thus, cause
the turgor pressure to rise. This suggests that wsc mutants
are hypersensitive to these stress treatments, because they
cannot adapt to the accompanying increase in turgor pressure.
Generally, cells with a defective cell wall as a result of
either a genetic defect or treatment with a cell wall perturbing compound can be rescued by increasing the osmotic value of the medium, for example by adding 10%
(w/v) sorbitol [154,155]. This sounds paradoxical, because
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increasing the osmotic strength of the medium is expected
to activate the Hog1 MAP kinase pathway. As a result,
the intracellular glycerol level is expected to increase, thus
counteracting the e¡ect of increasing the osmotic strength
of the medium. This raises the question whether the Hog1
MAP kinase pathway is suppressed in case of cell wall
stress. For further discussion of the cell wall salvage pathway, the reader is referred to [175,203^207].
3.4. Ultrastructural changes in the wall in response to cell
wall defects
Recent studies using genomic transcript analysis have
shown that constitutive activation of the Slt2 MAP kinase
pathway results in the up-regulation of mainly cell wallrelated genes [113,114]. These include genes that code for
biosynthetic enzymes such as Gfa1 and Chs3, which are
involved in chitin synthesis, and Gsc2/Fks2, which is involved in the synthesis of L1,3-glucan, cell wall proteins
such as Cwp1, Crh1, and the Pir-CWPs, and the MAP
kinase Slt2 itself [113,114]. A simple way to constitutively
activate the Slt2 MAP kinase pathway is based on using
cell wall mutants with a reduced amount of L1,3-glucan or
L1,6-glucan [23]. In these mutants various changes in cell
wall composition and organization have been detected
(Table 3) [65,73,116,130,131]. Although their cell wall
organization di¡ers considerably, it seems unlikely that
the cell is able to sense speci¢c cell wall defects and respond accordingly. Presumably, by activating the Slt2
MAP kinase pathway, cell wall stress leads in all cases
to the activation of multiple cell wall reinforcing reactions,
which ^ depending on the cell wall defect involved ^ may
result in various changes in cell wall organization. Generally, the chitin level in the wall of cells confronted with cell
wall stress is strongly elevated [10,65]. This is consistent
with the observation that many mutants with a cell wall
defect are synthetic lethal with chs3v and hypersensitive to
nikkomycin, an inhibitor of chitin synthesis [55,135]. Chitin deposition in response to cell wall stress seems to be
independent of Chs6 [208], which under non-stress conditions is required for proper Chs3 targeting [209].
3.5. Control of GSC2/FKS2
FKS1 and GSC2/FKS2 encode alternative subunits of
the L1,3-glucan synthase complex. FKS1 is preferentially
expressed under optimal growth conditions and is cell
cycle-regulated [210,211]. Regulation of GSC2/FKS2 expression is independently controlled by various inputs
[199]. For example, growth at 39‡C leads to increased expression of FKS2 through parallel inputs from the Slt2
MAP kinase pathway and from calcineurin, acting on different parts of the 5P upstream region of GSC2/FKS2
[199]. GSC2/FKS2 is also independently up-regulated
upon entry into the stationary phase and at low glucose
levels [199]. As discussed above, transcript levels of GSC2/
249
FKS2 also rise in case of cell wall stress in an Slt2-dependent fashion [23,113].
3.6. The Hog1 MAP kinase pathway
The Hog1 MAP kinase pathway is activated in response
to hyperosmotic stress. This seems to be accompanied by
transient usage of an altered set of cell wall proteins. The
transcript levels of the GPI-CWP-encoding genes CWP1,
SED1, and SPI1, and of SPS100 and YGP1, which encode
putative cell wall proteins (see Section 2.5), increase,
whereas the transcript levels of the GPI-CWP-encoding
genes CWP2 and EGT2 decrease [212^214]. Cells quickly
adapt to hypertonic stress, resulting in down-regulation of
many of the up-regulated genes.
There is clear evidence that the Hog1 MAP kinase signaling pathway is also required for cell wall construction
under non-stress conditions. First, both deletion of PBS2,
which codes for the MAP kinase kinase of this pathway,
and of HOG1, results in increased sensitivity of intact cells
to L1,3-glucanase, whereas multicopy expression of PBS2
results in lower sensitivity to L1,3-glucanase [192,215]. In
addition, L1,6-glucan levels are slightly increased in pbs2v
cells and slightly lowered in cells that overexpress PBS2
[216]. Third, in hog1v cells the Golgi mannosyl transferase
Mnn1 is mislocalized [217]. Fourth, as discussed in Section
3.8, cells kept at low pH become highly resistant to L1,3glucanase in a Hog1-dependent fashion [115]. Finally,
pbs2v and hog1v are more resistant to the cell wall perturbant Calco£uor white than wild-type cells, again indicating that the mutant cell wall di¡ers from the wild-type
wall [66]. How the Hog1 MAP kinase pathway a¡ects cell
wall construction still remains unclear.
3.7. Responses to anaerobic conditions
When oxygen levels decrease, the cell seems to switch
from one set of GPI-CWPs to another [121]. Northern
analysis reveals that the transcript level of CWP1 and
CWP2, which encode two abundant GPI-CWPs, decrease,
whereas the transcript levels of DAN1/CCW13, DAN2,
DAN3, and TIR1, TIR2, TIR3, and TIR4, and TIP1 are
strongly up-regulated. Similarly, cells growing in stagnant
cultures, which results in hypoxic growth conditions, show
a strong increase in the transcript level of TIR1 and the
level of Tir1 in the wall [123]. Interestingly, tir1v, tir2v,
and tir3v cells arrest in the unbudded phase when transferred to anaerobic conditions, suggesting that Tir1, Tir2,
and Tir3 are required for growth during G1 [121]. Genome-wide transcript analysis of chemostat-cultured cells
grown under anaerobic conditions are largely consistent
with these data and also reveal up-regulation of MUC1/
FLO11, a GPI-CWP involved in pseudohyphal growth,
and strong down-regulation of SPS100 [218]. It has further been shown that nearly the entire family of PAU
genes (see Section 2.5) is up-regulated under anaerobic
FEMSRE 738 1-8-02
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F.M. Klis et al. / FEMS Microbiology Reviews 26 (2002) 239^256
conditions [124,218]. Biochemical data of the molecular
organization of cell walls from cells grown under anaerobic conditions are scarce. For information about the regulation of CWP-encoding genes, induced under anaerobic
conditions, the reader is referred to [124,219^221].
3.8. Responses to extreme environmental pH values
In its natural environment, for example in rotting fruits,
S. cerevisiae is often confronted with low pH values.
Growth at low pH leads to various adaptations in cell
wall composition and organization. First, the cells become
highly resistant to the recombinant L1,3-glucanase Quantazyme [115]. As this is not due to an increase in chitin
content of the wall, this suggests that the mannoprotein
layer has become less permeable to Quantazyme and, presumably, to other fungal cell wall-degrading enzymes normally found in plant tissues as well. Second, the cells use
the alkali-sensitive linkage between cell wall proteins and
L1,3-glucan more extensively, resulting in more e⁄cient
incorporation of Pir-CWPs and an increase in doublelinked GPI-CWPs [115]. Genomic transcript analysis
shows that the transcript levels of the GPI-CWP-encoding
genes CWP1, HOR7, and SPI1 are increased [115]. Also
the transcript level of the putative cell wall protein-encoding gene YGP1 (Section 2.5) is increased. A similar genomic transcript analysis, comparing pH 4 and 8, identi¢ed only two alkali-inducible GPI-CWP-encoding genes,
namely, FIT2, and FIT3 [222], believed to be involved in
iron uptake [127]. Interestingly, the increase in Quantazyme resistance at low pH is dependent on Hog1, but
not on Slt2 [115], supporting the notion that also the
Hog1 MAP kinase pathway in£uences cell wall organization (see also Section 3.6).
in yeasts and other fungi is still in its infancy. Above all,
biochemical and also crystallographic studies of these enzymes are urgently needed to complement the vast amount
of molecular genetic information currently available.
The cell wall of yeast has long been viewed as a relatively static structure with limited changes in composition
and structure depending on environmental changes. It is
now clear that cell wall construction just like in bacteria is
a very dynamic process and that the cell tends to continually adapt its newly formed wall to changing conditions
both in terms of cell wall organization and with respect to
the cell wall proteins presented at the cell surface. How all
this is regulated is still largely unclear. Additional questions are how the cell behaves when it is confronted with
multiple stress conditions simultaneously. For example,
yeast cells growing on rotting grapes are confronted with
a rather acidic, extremely glucose-rich environment in
which also fungal cell wall-degrading enzymes abound.
Are perhaps some signaling pathways epistatic with respect to each other? For example, one may imagine that
the cAMP PKA pathway, which when activated leads to a
decrease in cell wall strength, is down-regulated in case of
cell wall stress. Better understanding of these regulatory
processes may help to further and more e⁄ciently exploit
this organism, and other fungi, for the bene¢t of mankind.
Acknowledgements
We would like to acknowledge the ¢nancial support
from the Netherlands Technology Foundation (STW)
and from EUROFAN II. We also thank Dr. Han de
Winde for critical reading of the manuscript.
References
4. Perspectives
The molecular organization of the cell wall of S. cerevisiae, growing in rich medium, has now largely been elucidated. This, however, is only part of the answer. In view
of the dynamic nature of the cell wall, a study of cell wall
organization using chemostat-cultured cells limited for
carbon, nitrogen and phosphate may reveal the limits of
the variability in cell wall composition of yeast. In addition, a study of cell wall construction in pseudohyphal
cells, in which polarized growth predominates and isotropic growth is limited, seems worthwhile.
The construction of the fungal cell wall is an attractive
target for the development of new antifungal compounds.
The fact that the molecular organization of the cell wall is
now largely elucidated makes it possible to have a closer
look at more speci¢c targets such as cell wall polymer
synthases, cell wall assembly enzymes and enzymes involved in remodeling of cell wall polymers. With some
exceptions, however, detailed research of these enzymes
[1] Orlean, P. (1997) Biogenesis of yeast wall and surface components.
In: The Molecular and Cellular Biology of the Yeast Saccharomyces.
Cell Cycle and Biology (Pringle, J.R., Broach, J.R. and Jones, E.W.,
Eds.), pp. 229^362. Cold Spring Harbor Laboratory Press, Cold
Spring Harbor, NY.
[2] Kapteyn, J.C., Van Den Ende, H. and Klis, F.M. (1999) The contribution of cell wall proteins to the organization of the yeast cell
wall. Biochim. Biophys. Acta 1426, 373^383.
[3] Lipke, P.N. and Ovalle, R. (1998) Cell wall architecture in yeast: new
structure and new challenges. J. Bacteriol. 180, 3735^3740.
[4] Smith, A.E., Zhang, Z., Thomas, C.R., Moxham, K.E. and Middelberg, A.P. (2000) The mechanical properties of Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. USA 97, 9871^9874.
[5] Cappellaro, C., Baldermann, C., Rachel, R. and Tanner, W. (1994)
Mating type-speci¢c cell-cell recognition of Saccharomyces cerevisiae :
cell wall attachment and active sites of a- and alpha-agglutinin.
EMBO J. 13, 4737^4744.
[6] Osumi, M. (1998) The ultrastructure of yeast: cell wall structure and
formation. Micron 29, 207^233.
[7] Marquardt, H. (1962) Der Feinbau von Hefezellen im Elektronenmikroskop II. Mitt.: Saccharomyces cerevisiae-Sta«mme. Z. Naturforsch. 17, 689^695.
FEMSRE 738 1-8-02
F.M. Klis et al. / FEMS Microbiology Reviews 26 (2002) 239^256
[8] Fleet, G.H. (1991) Cell walls. In: The Yeasts, 2nd edn., Vol. 4 (Rose
A.H. and Harrison, J.S., Eds.), pp. 199^277. Academic Press, New
York.
[9] Hartland, R.P., Vermeulen, C.A., Klis, F.M., Sietsma, J.H. and Wessels, J.G. (1994) The linkage of (1-3)-beta-glucan to chitin during cell
wall assembly in Saccharomyces cerevisiae. Yeast 10, 1591^1599.
[10] Dallies, N., Francois, J. and Paquet, V. (1998) A new method for
quantitative determination of polysaccharides in the yeast cell wall.
Application to the cell wall defective mutants of Saccharomyces cerevisiae. Yeast 14, 1297^1306.
[11] Klis, F.M., De Groot, P. and Hellingwerf, K. (2001) Molecular organization of the cell wall of Candida albicans. Med. Mycol. 39 (Suppl.
1), 1^8.
[12] Montijn, R.C., Vink, E., Muller, W.H., Verkleij, A.J., Van Den
Ende, H., Henrissat, B. and Klis, F.M. (1999) Localization of synthesis of beta1,6-glucan in Saccharomyces cerevisiae. J. Bacteriol. 181,
7414^7420.
[13] Dean, N. (1999) Asparagine-linked glycosylation in the yeast Golgi.
Biochim. Biophys. Acta 1426, 309^322.
[14] Manners, D.J., Masson, A.J., Patterson, J.C., Bjorndal, H. and Lindberg, B. (1973) The structure of a beta-(1^6)-D-glucan from yeast cell
walls. Biochem. J. 135, 31^36.
[15] Boone, C., Sommer, S.S., Hensel, A. and Bussey, H. (1990) Yeast
KRE genes provide evidence for a pathway of cell wall beta-glucan
assembly. J. Cell Biol. 110, 1833^1843.
[16] Manners, D.J., Masson, A.J. and Patterson, J.C. (1973) The structure
of a beta-(1 leads to 3)-D-glucan from yeast cell walls. Biochem. J.
135, 19^30.
[17] Holan, Z., Pokorny, V., Beran, K., Gemperle, A., Tuzar, Z. and
Baldrian, J. (1981) The glucan-chitin complex in Saccharomyces cerevisiae. V. Precise location of chitin and glucan in bud scar and their
physico-chemical characterization. Arch. Microbiol. 130, 312^318.
[18] Baba, M. and Osumi, M. (1987) Transmission and scanning electron
microscopic examination of intracellular organelles in freeze-substituted Kloeckera and Saccharomyces cerevisiae yeast cells. J. Electron
Microsc. Tech. 5, 249^261.
[19] Lipke, P.N. and Kurjan, J. (1992) Sexual agglutination in budding
yeasts: structure, function, and regulation of adhesion glycoproteins.
Microbiol. Rev. 56, 180^194.
[20] Teunissen, A.W. and Steensma, H.Y. (1995) Review: the dominant
£occulation genes of Saccharomyces cerevisiae constitute a new subtelomeric gene family. Yeast 11, 1001^1013.
[21] Reynolds, T.B. and Fink, G.R. (2001) Bakers’ yeast, a model for
fungal bio¢lm formation. Science 291, 878^881.
[22] De Nobel, J.G., Klis, F.M., Priem, J., Munnik, T. and van den Ende,
H. (1990) The glucanase-soluble mannoproteins limit cell wall porosity in Saccharomyces cerevisiae. Yeast 6, 491^499.
[23] De Nobel, H., Ruiz, C., Martin, H., Morris, W., Brul, S., Molina, M.
and Klis, F.M. (2000) Cell wall perturbation in yeast results in dual
phosphorylation of the Slt2/Mpk1 MAP kinase and in an Slt2-mediated increase in FKS2-lacZ expression, glucanase resistance and thermotolerance. Microbiology 146, 2121^2132.
[24] Zlotnik, H., Fernandez, M.P., Bowers, B. and Cabib, E. (1984) Saccharomyces cerevisiae mannoproteins form an external cell wall layer
that determines wall porosity. J. Bacteriol. 159, 1018^1026.
[25] Jigami, Y. and Odani, T. (1999) Mannosylphosphate transfer to yeast
mannan. Biochim. Biophys. Acta 1426, 335^345.
[26] Orlean, P., Ammer, A., Watzele, M. and Tanner, W. (1986) Synthesis
of an O-glycosylated cell surface protein induced in yeast by alphafactor. Proc. Natl. Acad. Sci. USA 83, 6263^6266.
[27] Moukadiri, I., Jaafar, L. and Zueco, J. (1999) Identi¢cation of two
mannoproteins released from cell walls of a Saccharomyces cerevisiae
mnn1 mnn9 double mutant by reducing agents. J. Bacteriol. 181,
4741^4745.
[28] Morris, G.J., Winters, L., Coulson, G.E. and Clarke, K.J. (1986)
E¡ect on osmotic stress on the ultrastructure and viability of the
yeast Saccharomyces cerevisiae. J. Gen. Microbiol. 129, 2023^2034.
251
[29] Martinez de Maranon, I., Marechal, P.A. and Gervais, P. (1996)
Passive response of Saccharomyces cerevisiae to osmotic shifts : cell
volume variations depending on the physiological state. Biochem.
Biophys. Res. Commun. 227, 519^523.
[30] Scherrer, R., Louden, L. and Gerhardt, P. (1974) Porosity of the
yeast cell wall and membrane. J. Bacteriol. 118, 534^540.
[31] De Nobel, J.G. and Barnett, J.A. (1991) Passage of molecules
through yeast cell walls : a brief essay-review. Yeast 7, 313^323.
[32] De Nobel, J.G., Klis, F.M., Munnik, T., Priem, J. and van den Ende,
H. (1990) An assay of relative cell wall porosity in Saccharomyces
cerevisiae, Kluyveromyces lactis and Schizosaccharomyces pombe.
Yeast 6, 483^490.
[33] Young, S.H. and Jacobs, R.R. (1998) Sodium hydroxide-induced
conformational change in schizophyllan detected by the £uorescence
dye, aniline blue. Carbohydr. Res. 310, 91^99.
[34] Beauvais, A., Bruneau, J.M., Mol, P.C., Buitrago, M.J., Legrand, R.
and Latge, J.P. (2001) Glucan synthase complex of Aspergillus fumigatus. J. Bacteriol. 183, 2273^2279.
[35] Rees, D.A., Morris, E.R., Thom, D. and Madden, J.K. (1982) Shapes
and interactions of carbohydrate chains. In: The Polysaccharides,
Vol. I (Aspinall, G.O., Ed.), pp. 196^290. Academic Press, New
York.
[36] Krainer, E., Stark, R.E., Naider, F., Alagramam, K. and Becker,
J.M. (1994) Direct observation of cell wall glucans in whole cells of
Saccharomyces cerevisiae by magic-angle spinning 13 C-NMR. Biopolymers 34, 1627^1635.
[37] Kreger, D.R. and Kopecka, M. (1975) On the nature and formation
of the ¢brillar nets produced by protoplasts of Saccharomyces cerevisiae in liquid media : an electronmicroscopic, X-ray di¡raction and
chemical study. J. Gen. Microbiol. 92, 207^220.
[38] Mu«ller, A., Ensley, H., Pretus, H., McNamee, R., Jones, E.,
McLaughlin, E., Chandley, W., Browder, W., Lowman, D. and Williams, D. (1997) The application of various protic acids in the extraction of (1C3)-beta-D-glucan from Saccharomyces cerevisiae. Carbohydr. Res. 299, 203^208.
[39] Cabib, E., Drgonova, J. and Drgon, T. (1998) Role of small G proteins in yeast cell polarization and wall biosynthesis. Annu. Rev.
Biochem. 67, 307^333.
[40] Whistler, R.L. (1973) Solubility of polysaccharides and their behavior
in solution. Adv. Chem. Ser. 117, 242^255.
[41] De Nobel, H., Dijkers, C., Hooijberg, E. and Klis, F.M. (1989)
Increased cell wall porosity in Saccharomyces cerevisiae after treatment with dithiothreitol or EDTA. J. Gen. Microbiol. 135, 2077^
2084.
[42] De Nobel, H., Sietsma, J.H., van den Ende, H. and Klis, F.M. (2001)
Molecular organization and construction of the fungal cell wall. In:
The Mycota VIII. Biology of the Fungal Cell (Howard, R.J. and
Gow, N.A.R., Eds.). Springer Verlag, Berlin.
[43] De Nobel, H., van Den Ende, H. and Klis, F.M. (2000) Cell wall
maintenance in fungi. Trends Microbiol. 8, 344^345.
[44] Furukawa, K., Tagaya, M., Tanizawa, K. and Fukui, T. (1993) Role
of the conserved Lys-X-Gly-Gly sequence at the ADP-glucose-binding site in Escherichia coli glycogen synthase. J. Biol. Chem. 268,
23837^23842.
[45] Hochstenbach, F., Klis, F.M., van den Ende, H., van Donselaar, E.,
Peters, P.J. and Klausner, R.D. (1998) Identi¢cation of a putative
alpha-glucan synthase essential for cell wall construction and morphogenesis in ¢ssion yeast. Proc. Natl. Acad. Sci. USA 95, 9161^
9166.
[46] Moor, H. and Mu«hlethaler, K. (1963) Fine structure in frozen-etched
yeast cells. J. Cell Biol. 17, 609^628.
[47] Cui, X., Shin, H., Song, C., Laosinchai, W., Amano, Y. and Brown
Jr., R.M. (2001) A putative plant homolog of the yeast beta-1,3glucan synthase subunit FKS1 from cotton (Gossypium hirsutum L.)
¢bers. Planta 213, 223^230.
[48] Batra, K.K., Nordin, J.H. and Kirkwood, S. (1969) Biosynthesis of
the L-D-glucan of Sclerotium rolfsii SACC. Direction of chain prop-
FEMSRE 738 1-8-02
252
[49]
[50]
[51]
[52]
[53]
[54]
[55]
[56]
[57]
[58]
[59]
[60]
[61]
[62]
[63]
[64]
[65]
[66]
[67]
F.M. Klis et al. / FEMS Microbiology Reviews 26 (2002) 239^256
agation and the insertion of the branch residues. Carbohydr. Res. 9,
221^229.
Koyama, M., Helbert, W., Imai, T., Sugiyama, J. and Henrissat, B.
(1997) Parallel-up structure evidences the molecular directionality
during biosynthesis of bacterial cellulose. Proc. Natl. Acad. Sci.
USA 94, 9091^9095.
Sugiyama, J., Boisset, C., Hashimoto, M. and Watanabe, T. (1999)
Molecular directionality of beta-chitin biosynthesis. J. Mol. Biol. 286,
247^255.
Scheller, H.V., Doong, R.L., Ridley, B.L. and Mohnen, D. (1999)
Pectin biosynthesis: a solubilized K1,4-galacturonosyltransferase
from tobacco catalyzes the transfer of galacturonic acid from UDPgalacturonic acid onto the non-reducing end of homogalacturonan.
Planta 207, 512^517.
Goldman, R.C., Sullivan, P.A., Zakula, D. and Capobianco, J.O.
(1995) Kinetics of beta-1,3 glucan interaction at the donor and acceptor sites of the fungal glucosyltransferase encoded by the BGL2
gene. Eur. J. Biochem. 227, 372^378.
Yu, L., Goldman, R., Sullivan, P., Walker, G.F. and Fesik, S.W.
(1993) Heteronuclear NMR studies of 13C-labeled yeast cell wall
beta-glucan oligosaccharides. J. Biomol. NMR 3, 429^441.
Shematek, E.M., Braatz, J.A. and Cabib, E. (1980) Biosynthesis of
the yeast cell wall. II. Preparation and properties of L-(1C3)glucan
synthetase. J. Biol. Chem. 255, 888^894.
Popolo, L. and Vai, M. (1999) The Gas1 glycoprotein, a putative wall
polymer cross-linker. Biochim. Biophys. Acta 1426, 385^400.
Mouyna, I., Fontaine, T., Vai, M., Monod, M., Fonzi, W.A., Diaquin, M., Popolo, L., Hartland, R.P. and Latge, J.P. (2000) Glycosylphosphatidylinositol-anchored glucanosyltransferases play an active role in the biosynthesis of the fungal cell wall. J. Biol. Chem.
275, 14882^14889.
Cappellaro, C., Mrsa, V. and Tanner, W. (1998) New potential cell
wall glucanases of Saccharomyces cerevisiae and their involvement in
mating. J. Bacteriol. 180, 5030^5037.
Rodriguez-Pena, J.M., Cid, V.J., Arroyo, J. and Nombela, C. (2000)
A novel family of cell wall-related proteins regulated di¡erently during the yeast life cycle. Mol. Cell. Biol. 20, 3245^3255.
Mouassite, M., Camougrand, N., Schwob, E., Demaison, G., Laclau,
M. and Guerin, M. (2000) The ‘SUN’ family: yeast SUN4/SCW3 is
involved in cell septation. Yeast 16, 905^919.
Molano, J., Bowers, B. and Cabib, E. (1980) Distribution of chitin in
the yeast cell wall. An ultrastructural and chemical study. J. Cell Biol.
85, 199^212.
Shaw, J.A., Mol, P.C., Bowers, B., Silverman, S.J., Valdivieso, M.H.,
Duran, A. and Cabib, E. (1991) The function of chitin synthases 2
and 3 in the Saccharomyces cerevisiae cell cycle. J. Cell Biol. 114,
111^123.
Cabib, E., Roh, D.H., Schmidt, M., Crotti, L.B. and Varma, A.
(2001) The yeast cell wall and septum as paradigms of cell growth
and morphogenesis. J. Biol. Chem. 276, 19679^19682.
Wood, P.J. (1980) Speci¢city in the interaction of direct dyes with
polysaccharides. Carbohydr. Res. 85, 271^287.
Pringle, J.R. (1991) Staining of bud scars and other cell wall chitin
with calco£uor. Methods Enzymol. 194, 732^735.
Kapteyn, J.C., Van Egmond, P., Sievi, E., Van Den Ende, H., Makarow, M. and Klis, F.M. (1999) The contribution of the O-glycosylated protein Pir2p/Hsp150 to the construction of the yeast cell wall
in wild-type cells and beta 1,6-glucan-de¢cient mutants. Mol. Microbiol. 31, 1835^1844.
Garcia-Rodriguez, L.J., Duran, A. and Roncero, C. (2000) Calco£uor antifungal action depends on chitin and a functional high-osmolarity glycerol response (HOG) pathway: evidence for a physiological role of the Saccharomyces cerevisiae HOG pathway under
noninducing conditions. J. Bacteriol. 182, 2428^2437.
Valdivieso, M.H., Ferrario, L., Vai, M., Duran, A. and Popolo, L.
(2000) Chitin synthesis in a gas1 mutant of Saccharomyces cerevisiae.
J. Bacteriol. 182, 4752^4757.
[68] Cabib, E., Bowers, B. and Roberts, R.L. (1983) Vectorial synthesis of
a polysaccharide by isolated plasma membranes. Proc. Natl. Acad.
Sci. USA 80, 3318^3321.
[69] Saxena, I.M. and Brown Jr., R.M. (1997) Identi¢cation of cellulose
synthase(s) in higher plants: sequence analysis of processive L-glycosyltransferases with the common motif ‘D, D, D35Q(R,Q)XRW’.
Cellulose 4, 33^49.
[70] Kollar, R., Petrakova, E., Ashwell, G., Robbins, P.W. and Cabib, E.
(1995) Architecture of the yeast cell wall. The linkage between chitin
and beta(1C3)-glucan. J. Biol. Chem. 270, 1170^1178.
[71] Kollar, R., Reinhold, B.B., Petrakova, E., Yeh, H.J., Ashwell, G.,
Drgonova, J., Kapteyn, J.C., Klis, F.M. and Cabib, E. (1997) Architecture of the yeast cell wall. Beta(1C6)-glucan interconnects mannoprotein, beta(1C)3-glucan, and chitin. J. Biol. Chem. 272, 17762^
17775.
[72] Inoue, S.B., Takewaki, N., Takasuka, T., Mio, T., Adachi, M., Fujii,
Y., Miyamoto, C., Arisawa, M., Furuichi, Y. and Watanabe, T.
(1995) Characterization and gene cloning of 1,3-beta-D-glucan synthase from Saccharomyces cerevisiae. Eur. J. Biochem. 231, 845^
854.
[73] Kapteyn, J.C., Ram, A.F., Groos, E.M., Kollar, R., Montijn, R.C.,
Van Den Ende, H., Llobell, A., Cabib, E. and Klis, F.M. (1997)
Altered extent of cross-linking of beta1,6-glucosylated mannoproteins
to chitin in Saccharomyces cerevisiae mutants with reduced cell wall
beta1,3-glucan content. J. Bacteriol. 179, 6279^6284.
[74] Shahinian, S. and Bussey, H. (2000) Beta-1,6-glucan synthesis in Saccharomyces cerevisiae. Mol. Microbiol. 35, 477^489.
[75] Mrsa, V., Seidl, T., Gentzsch, M. and Tanner, W. (1997) Speci¢c
labelling of cell wall proteins by biotinylation. Identi¢cation of four
covalently linked O-mannosylated proteins of Saccharomyces cerevisiae. Yeast 13, 1145^1154.
[76] Jentoft, N. (1990) Why are proteins O-glycosylated ? Trends Biochem.
Sci. 15, 291^294.
[77] Chen, M.H., Shen, Z.M., Bobin, S., Kahn, P.C. and Lipke, P.N.
(1995) Structure of Saccharomyces cerevisiae alpha-agglutinin. Evidence for a yeast cell wall protein with multiple immunoglobulinlike domains with atypical disul¢des. J. Biol. Chem. 270, 26168^
26177.
[78] Ibeas, J.I., Lee, H., Damsz, B., Prasad, D.T., Pardo, J.M., Hasegawa,
P.M., Bressan, R.A. and Narasimhan, M.L. (2000) Fungal cell wall
phosphomannans facilitate the toxic activity of a plant PR-5 protein.
Plant J. 23, 375^383.
[79] Caro, L.H., Tettelin, H., Vossen, J.H., Ram, A.F., van den Ende, H.
and Klis, F.M. (1997) In silicio identi¢cation of glycosyl-phosphatidylinositol-anchored plasma-membrane and cell wall proteins of Saccharomyces cerevisiae. Yeast 13, 1477^1489.
[80] Kapteyn, J.C., Montijn, R.C., Vink, E., de la Cruz, J., Llobell, A.,
Douwes, J.E., Shimoi, H., Lipke, P.N. and Klis, F.M. (1996) Retention of Saccharomyces cerevisiae cell wall proteins through a phosphodiester-linked beta-1,3-/beta-1,6-glucan heteropolymer. Glycobiology 6, 337^345.
[81] Fujii, T., Shimoi, H. and Iimura, Y. (1999) Structure of the glucanbinding sugar chain of Tip1p, a cell wall protein of Saccharomyces
cerevisiae. Biochim. Biophys. Acta 1427, 133^144.
[82] Hamada, K., Terashima, H., Arisawa, M. and Kitada, K. (1998)
Amino acid sequence requirement for e⁄cient incorporation
of glycosylphosphatidylinositol-associated proteins into the cell
wall of Saccharomyces cerevisiae. J. Biol. Chem. 273, 26946^
26953.
[83] Hamada, K., Terashima, H., Arisawa, M., Yabuki, N. and Kitada,
K. (1999) Amino acid residues in the omega-minus region participate
in cellular localization of yeast glycosylphosphatidylinositol-attached
proteins. J. Bacteriol. 181, 3886^3889.
[84] Lu, C.F., Montijn, R.C., Brown, J.L., Klis, F., Kurjan, J., Bussey, H.
and Lipke, P.N. (1995) Glycosyl phosphatidylinositol-dependent
cross-linking of alpha-agglutinin and beta 1,6-glucan in the Saccharomyces cerevisiae cell wall. J. Cell Biol. 128, 333^340.
FEMSRE 738 1-8-02
F.M. Klis et al. / FEMS Microbiology Reviews 26 (2002) 239^256
[85] Schreuder, M.P., Brekelmans, S., van den Ende, H. and Klis, F.M.
(1993) Targeting of a heterologous protein to the cell wall of Saccharomyces cerevisiae. Yeast 9, 399^409.
[86] Imhof, I., Canivenc-Gansel, E., Meyer, U. and Conzelmann, A.
(2000) Phosphatidylethanolamine is the donor of the phosphorylethanolamine linked to the alpha1,4-linked mannose of yeast GPI
structures. Glycobiology 10, 1271^1275.
[87] Taron, C.H., Wiedman, J.M., Grimme, S.J. and Orlean, P. (2000)
Glycosylphosphatidylinositol biosynthesis defects in Gpi11p- and
Gpi13p-de¢cient yeast suggest a branched pathway and implicate
gpi13p in phosphoethanolamine transfer to the third mannose.
Mol. Biol. Cell 11, 1611^1630.
[88] Gaynor, E.C., Mondesert, G., Grimme, S.J., Reed, S.I., Orlean, P.
and Emr, S.D. (1999) MCD4 encodes a conserved endoplasmic reticulum membrane protein essential for glycosylphosphatidylinositol
anchor synthesis in yeast. Mol. Biol. Cell 10, 627^648.
[89] Mu«ller, G., Schubert, K., Fiedler, F. and Bandlow, W. (1992) The
cAMP-binding ectoprotein from Saccharomyces cerevisiae is membrane-anchored by glycosyl-phosphatidylinositol. J. Biol. Chem.
267, 25337^25346.
[90] Kainuma, M., Chiba, Y., Takeuchi, M. and Jigami, Y. (2001) Overexpression of HUT1 gene stimulates in vivo galactosylation by enhancing UDP-galactose transport activity in Saccharomyces cerevisiae. Yeast 18, 533^541.
[91] Toh-e, A., Yasunaga, S., Nisogi, H., Tanaka, K., Oguchi, T. and
Matsui, Y. (1993) Three yeast genes, PIR1, PIR2 and PIR3, containing internal tandem repeats, are related to each other, and PIR1
and PIR2 are required for tolerance to heat shock. Yeast 9, 481^
494.
[92] Mrsa, V. and Tanner, W. (1999) Role of NaOH-extractable cell wall
proteins Ccw5p, Ccw6p, Ccw7p and Ccw8p (members of the Pir
protein family) in stability of the Saccharomyces cerevisiae cell
wall. Yeast 15, 813^820.
[93] Yun, D.J., Zhao, Y., Pardo, J.M., Narasimhan, M.L., Damsz, B.,
Lee, H., Abad, L.R., D’Urzo, M.P., Hasegawa, P.M. and Bressan,
R.A. (1997) Stress proteins on the yeast cell surface determine resistance to osmotin, a plant antifungal protein. Proc. Natl. Acad.
Sci. USA 94, 7082^7087.
[94] Kapteyn, J.C., Hoyer, L.L., Hecht, J.E., Muller, W.H., Andel, A.,
Verkleij, A.J., Makarow, M., Van Den Ende, H. and Klis, F.M.
(2000) The cell wall architecture of Candida albicans wild-type cells
and cell wall-defective mutants. Mol. Microbiol. 35, 601^611.
[95] Law, D.T. and Segall, J. (1988) The SPS100 gene of Saccharomyces
cerevisiae is activated late in the sporulation process and contributes
to spore wall maturation. Mol. Cell. Biol. 8, 912^922.
[96] Destruelle, M., Holzer, H. and Klionsky, D.J. (1994) Identi¢cation
and characterization of a novel yeast gene: the YGP1 gene product
is a highly glycosylated secreted protein that is synthesized in response to nutrient limitation. Mol. Cell. Biol. 14, 2740^2754.
[97] Watzele, M., Klis, F. and Tanner, W. (1988) Puri¢cation and characterization of the inducible a agglutinin of Saccharomyces cerevisiae. EMBO J. 7, 1483^1488.
[98] Lu, C.F., Kurjan, J. and Lipke, P.N. (1994) A pathway for cell wall
anchorage of Saccharomyces cerevisiae alpha-agglutinin. Mol. Cell.
Biol. 14, 4825^4833.
[99] Kuranda, M.J. and Robbins, P.W. (1991) Chitinase is required for
cell separation during growth of Saccharomyces cerevisiae. J. Biol.
Chem. 266, 19758^19767.
[100] Colman-Lerner, A., Chin, T.E. and Brent, R. (2001) Yeast Cbk1
and Mob2 activate daughter-speci¢c genetic programs to induce
asymmetric cell fates. Cell 107, 739^750.
[101] Klis, F.M. (1994) Review: cell all assembly in yeast. Yeast 10, 851^
869.
[102] Lopez-Ribot, J.L. and Cha⁄n, W.L. (1996) Members of the Hsp70
family of proteins in the cell wall of Saccharomyces cerevisiae.
J. Bacteriol. 178, 4724^4726.
[103] Delgado, M.L., O’Connor, J.E., Azorin, I., Renau-Piqueras, J., Gil,
[104]
[105]
[106]
[107]
[108]
[109]
[110]
[111]
[112]
[113]
[114]
[115]
[116]
[117]
[118]
[119]
[120]
253
M.L. and Gozalbo, D. (2001) The glyceraldehyde-3-phosphate dehydrogenase polypeptides encoded by the Saccharomyces cerevisiae
TDH1, TDH2 and TDH3 genes are also cell wall proteins. Microbiology 147, 411^417.
Pardo, M., Monteoliva, L., Pla, J., Sanchez, M., Gil, C. and Nombela, C. (1999) Two-dimensional analysis of proteins secreted by
Saccharomyces cerevisiae regenerating protoplasts: a novel approach
to study the cell wall. Yeast 15, 459^472.
Pardo, M., Ward, M., Bains, S., Molina, M., Blackstock, W., Gil,
C. and Nombela, C. (2000) A proteomic approach for the study of
Saccharomyces cerevisiae cell wall biogenesis. Electrophoresis 21,
3396^3410.
Molina, M., Gil, C., Pla, J., Arroyo, J. and Nombela, C. (2000)
Protein localisation approaches for understanding yeast cell wall
biogenesis. Microsc. Res. Tech. 51, 601^612.
Roy, A., Lu, C.F., Marykwas, D.L., Lipke, P.N. and Kurjan, J.
(1991) The AGA1 product is involved in cell surface attachment
of the Saccharomyces cerevisiae cell adhesion glycoprotein a-agglutinin. Mol. Cell. Biol. 11, 4196^4206.
Erdman, S., Lin, L., Malczynski, M. and Snyder, M. (1998) Pheromone-regulated genes required for yeast mating di¡erentiation.
J. Cell Biol. 140, 461^483.
Guo, B., Styles, C.A., Feng, Q. and Fink, G.R. (2000) A Saccharomyces gene family involved in invasive growth, cell-cell adhesion,
and mating. Proc. Natl. Acad. Sci. USA 97, 12158^12163.
Bony, M., Thines-Sempoux, D., Barre, P. and Blondin, B. (1997)
Localization and cell surface anchoring of the Saccharomyces cerevisiae £occulation protein Flo1p. J. Bacteriol. 179, 4929^4936.
Kobayashi, O., Hayashi, N., Kuroki, R. and Sone, H. (1998) Region of FLO1 proteins responsible for sugar recognition. J. Bacteriol. 180, 6503^6510.
Kovacech, B., Nasmyth, K. and Schuster, T. (1996) EGT2 gene
transcription is induced predominantly by Swi5 in early G1. Mol.
Cell. Biol. 16, 3264^3274.
Terashima, H., Yabuki, N., Arisawa, M., Hamada, K. and Kitada,
K. (2000) Up-regulation of genes encoding glycosylphosphatidylinositol (GPI)-attached proteins in response to cell wall damage caused
by disruption of FKS1 in Saccharomyces cerevisiae. Mol. Gen. Genet. 264, 64^74.
Jung, U.S. and Levin, D.E. (1999) Genome-wide analysis of gene
expression regulated by the yeast cell wall integrity signalling pathway. Mol. Microbiol. 34, 1049^1057.
Kapteyn, J.C., ter Riet, B., Vink, E., Blad, S., De Nobel, H., Van
Den Ende, H. and Klis, F.M. (2001) Low external pH induces
HOG1-dependent changes in the organization of the Saccharomyces
cerevisiae cell wall. Mol. Microbiol. 39, 469^479.
Ram, A.F., Kapteyn, J.C., Montijn, R.C., Caro, L.H., Douwes,
J.E., Baginsky, W., Mazur, P., van den Ende, H. and Klis, F.M.
(1998) Loss of the plasma membrane-bound protein Gas1p in Saccharomyces cerevisiae results in the release of beta1,3-glucan into the
medium and induces a compensation mechanism to ensure cell wall
integrity. J. Bacteriol. 180, 1418^1424.
Spellman, P.T., Sherlock, G., Zhang, M.Q., Iyer, V.R., Anders, K.,
Eisen, M.B., Brown, P.O., Botstein, D. and Futcher, B. (1998) Comprehensive identi¢cation of cell cycle-regulated genes of the yeast
Saccharomyces cerevisiae by microarray hybridization. Mol. Biol.
Cell 9, 3273^3297.
Chu, S., DeRisi, J., Eisen, M., Mulholland, J., Botstein, D., Brown,
P.O. and Herskowitz, I. (1998) The transcriptional program of sporulation in budding yeast. Science 282, 699^705.
Shimoi, H., Kitagaki, H., Ohmori, H., Iimura, Y. and Ito, K. (1998)
Sed1p is a major cell wall protein of Saccharomyces cerevisiae in the
stationary phase and is involved in lytic enzyme resistance. J. Bacteriol. 180, 3381^3387.
Puig, S. and Perez-Ortin, J.E. (2000) Stress response and expression
patterns in wine fermentations of yeast genes induced at the diauxic
shift. Yeast 16, 139^148.
FEMSRE 738 1-8-02
254
F.M. Klis et al. / FEMS Microbiology Reviews 26 (2002) 239^256
[121] Abramova, N., Sertil, O., Mehta, S. and Lowry, C.V. (2001) Reciprocal regulation of anaerobic and aerobic cell wall mannoprotein
gene expression in Saccharomyces cerevisiae. J. Bacteriol. 183,
2881^2887.
[122] Donzeau, M., Bourdineaud, J.P. and Lauquin, G.J. (1996) Regulation by low temperatures and anaerobiosis of a yeast gene specifying
a putative GPI-anchored plasma membrane protein. Mol. Microbiol. 20, 449^459.
[123] Kitagaki, H., Shimoi, H. and Itoh, K. (1997) Identi¢cation and
analysis of a static culture-speci¢c cell wall protein, Tir1p/Srp1p in
Saccharomyces cerevisiae. Eur. J. Biochem. 249, 343^349.
[124] Kwast, K.E., Lai, L.C., Menda, N., James 3rd, D.T., Aref, S. and
Burke, P.V. (2002) Genomic analyses of anaerobically induced genes
in Saccharomyces cerevisiae : functional roles of Rox1 and other
factors in mediating the anoxic response. J. Bacteriol. 184, 250^265.
[125] Kondo, K. and Inouye, M. (1991) TIP1, a cold shock-inducible gene
of Saccharomyces cerevisiae. J. Biol. Chem. 266, 17537^17544.
[126] Horsted, M.W., Dey, E.S., Holmberg, S. and Kielland-Brandt, M.C.
(1998) A novel esterase from Saccharomyces carlsbergensis, a possible function for the yeast TIP1 gene. Yeast 14, 793^803.
[127] Protchenko, O., Ferea, T., Rashford, J., Tiedeman, J., Brown, P.O.,
Botstein, D. and Philpott, C.C. (2001) Three cell wall mannoproteins facilitate the uptake of iron in Saccharomyces cerevisiae. J. Biol.
Chem. 276, 49244^49250.
[128] Mrsa, V., Ecker, M., Strahl-Bolsinger, S., Nimtz, M., Lehle, L. and
Tanner, W. (1999) Deletion of new covalently linked cell wall glycoproteins alters the electrophoretic mobility of phosphorylated wall
components of Saccharomyces cerevisiae. J. Bacteriol. 181, 3076^
3086.
[129] Montijn, R.C., van Rinsum, J., van Schagen, F.A. and Klis, F.M.
(1994) Glucomannoproteins in the cell wall of Saccharomyces cerevisiae contain a novel type of carbohydrate side chain. J. Biol.
Chem. 269, 19338^19342.
[130] Popolo, L., Gilardelli, D., Bonfante, P. and Vai, M. (1997) Increase
in chitin as an essential response to defects in assembly of cell wall
polymers in the ggp1delta mutant of Saccharomyces cerevisiae.
J. Bacteriol. 179, 463^469.
[131] Garcia-Rodriguez, L.J., Trilla, J.A., Castro, C., Valdivieso, M.H.,
Duran, A. and Roncero, C. (2000) Characterization of the chitin
biosynthesis process as a compensatory mechanism in the fks1 mutant of Saccharomyces cerevisiae. FEBS Lett. 478, 84^88.
[132] Costanzo, M.C., Hogan, J.D., Cusick, M.E., Davis, B.P., Fancher,
A.M., Hodges, P.E., Kondu, P., Lengieza, C., Lew-Smith, J.E.,
Lingner, C., Roberg-Perez, K.J., Tillberg, M., Brooks, J.E. and
Garrels, J.I. (2000) The Yeast Proteome Database (YPD) and Caenorhabditis elegans Proteome Database (WormPD) : comprehensive
resources for the organization and comparison of model organism
protein information. Nucleic Acids Res. 28, 73^76.
[133] Costanzo, M.C., Crawford, M.E., Hirschman, J.E., Kranz, J.E.,
Olsen, P., Robertson, L.S., Skrzypek, M.S., Braun, B.R., Hopkins,
K.L., Kondu, P., Lengieza, C., Lew-Smith, J.E., Tillberg, M. and
Garrels, J.I. (2001) YPD1, PombePD1, and WormPD1: model
organism volumes of the BioKnowledge1 library, an integrated
resource for protein information. Nucleic Acids Res. 29, 75^79.
[134] Gentzsch, M. and Tanner, W. (1996) The PMT gene family: protein
O-glycosylation in Saccharomyces cerevisiae is vital. EMBO J. 15,
5752^5759.
[135] Mondesert, G. and Reed, S.I. (1996) BED1, a gene encoding a
galactosyltransferase homologue, is required for polarized growth
and e⁄cient bud emergence in Saccharomyces cerevisiae. J. Cell
Biol. 132, 137^151.
[136] Bom, I.J., Klis, F.M., de Nobel, H. and Brul, S. (2001) A new
strategy for inhibition of the spoilage yeasts Saccharomyces cerevisiae and Zygosaccharomyces bailii based on combination of a membrane-active peptide with an oligosaccharide that leads to an impaired glycosylphosphatidylinositol (GPI)-dependent yeast wall
protein layer. FEMS Yeast Res. 1, 187^194.
[137] Smits, G.J., Kapteyn, J.C., van den Ende, H. and Klis, F.M. (1999)
Cell wall dynamics in yeast. Curr. Opin. Microbiol. 2, 348^352.
[138] Boder, E.T. and Wittrup, K.D. (1997) Yeast surface display for
screening combinatorial polypeptide libraries. Nat. Biotechnol. 15,
553^557.
[139] Kieke, M.C., Cho, B.K., Boder, E.T., Kranz, D.M. and Wittrup,
K.D. (1997) Isolation of anti-T cell receptor scFv mutants by yeast
surface display. Protein Eng. 10, 1303^1310.
[140] Murai, T., Ueda, M., Atomi, H., Shibasaki, Y., Kamasawa, N.,
Osumi, M., Kawaguchi, T., Arai, M. and Tanaka, A. (1997) Genetic
immobilization of cellulase on the cell surface of Saccharomyces
cerevisiae. Appl. Microbiol. Biotechnol. 48, 499^503.
[141] Murai, T., Ueda, M., Yamamura, M., Atomi, H., Shibasaki, Y.,
Kamasawa, N., Osumi, M., Amachi, T. and Tanaka, A. (1997)
Construction of a starch-utilizing yeast by cell surface engineering.
Appl. Environ. Microbiol. 63, 1362^1366.
[142] Schreuder, M.P., Mooren, A.T., Toschka, H.Y., Verrips, C.T. and
Klis, F.M. (1996) Immobilizing proteins on the surface of yeast
cells. Trends Biotechnol. 14, 115^120.
[143] Van der Vaart, J.M., te Biesebeke, R., Chapman, J.W., Toschka,
H.Y., Klis, F.M. and Verrips, C.T. (1997) Comparison of cell wall
proteins of Saccharomyces cerevisiae as anchors for cell surface expression of heterologous proteins. Appl. Environ. Microbiol. 63,
615^620.
[144] Shibasaki, Y., Kamasawa, N., Shibasaki, S., Zou, W., Murai, T.,
Ueda, M., Tanaka, A. and Osumi, M. (2000) Cytochemical evaluation of localization and secretion of a heterologous enzyme displayed on yeast cell surface. FEMS Microbiol. Lett. 192, 243^248.
[145] Washida, M., Takahashi, S., Ueda, M. and Tanaka, A. (2001)
Spacer-mediated display of active lipase on the yeast cell surface.
Appl. Microbiol. Biotechnol. 56, 681^686.
[146] Kratky, Z., Biely, P. and Bauer, S. (1975) Wall mannan of Saccharomyces cerevisiae. Metabolic stability and release into growth medium. Biochim. Biophys. Acta 404, 1^6.
[147] Cormack, B.P., Ghori, N. and Falkow, S. (1999) An adhesin of the
yeast pathogen Candida glabrata mediating adherence to human
epithelial cells. Science 285, 578^582.
[148] Kandasamy, R., Vediyappan, G. and Cha⁄n, W.L. (2000) Evidence
for the presence of Pir-like proteins in Candida albicans. FEMS
Microbiol. Lett. 186, 239^243.
[149] Bernard, M. and Latge¤, J.P. (2001) Aspergillus fumigatus cell wall :
composition and biosynthesis. Med. Mycol. 39 (Suppl. 1), 9^17.
[150] De Groot, P.W.J., Ruiz, C., Vazquez de Aldana, C.R., Duenas, E.,
Cid, V.J., Del Rey, F., Rodriquez-Pena, J.M., Pe¤rez, P., Andel, A.,
Caubin, J., Arroyo, J., Garcia, J.C., Gil, C., Molina, M., Garcia,
L.J., Nombela, C. and Klis, F.M. (2001) A genomic approach for
the identi¢cation and classi¢cation of genes involved in cell wall
formation and its regulation in Saccharomyces cerevisiae. Comp.
Funct. Genomics 2, 124^142.
[151] Ram, A.F., Wolters, A., Ten Hoopen, R. and Klis, F.M. (1994) A
new approach for isolating cell wall mutants in Saccharomyces cerevisiae by screening for hypersensitivity to calco£uor white. Yeast
10, 1019^1030.
[152] Lussier, M., White, A.M., Sheraton, J., di Paolo, T., Treadwell, J.,
Southard, S.B., Horenstein, C.I., Chen-Weiner, J., Ram, A.F., Kapteyn, J.C., Roemer, T.W., Vo, D.H., Bondoc, D.C., Hall, J., Zhong,
W.W., Sdicu, A.M., Davies, J., Klis, F.M., Robbins, P.W. and
Bussey, H. (1997) Large scale identi¢cation of genes involved in
cell surface biosynthesis and architecture in Saccharomyces cerevisiae. Genetics 147, 435^450.
[153] van der Vaart, J.M., Caro, L.H., Chapman, J.W., Klis, F.M. and
Verrips, C.T. (1995) Identi¢cation of three mannoproteins in the cell
wall of Saccharomyces cerevisiae. J. Bacteriol. 177, 3104^3110.
[154] Frost, D.J., Brandt, K.D., Cugier, D. and Goldman, R. (1995) A
whole-cell Candida albicans assay for the detection of inhibitors
towards fungal cell wall synthesis and assembly. J. Antibiot. 48,
306^310.
FEMSRE 738 1-8-02
F.M. Klis et al. / FEMS Microbiology Reviews 26 (2002) 239^256
[155] Martin, H., Rodriguez-Pachon, J.M., Ruiz, C., Nombela, C. and
Molina, M. (2000) Regulatory mechanisms for modulation of signaling through the cell integrity Slt2-mediated pathway in Saccharomyces cerevisiae. J. Biol. Chem. 275, 1511^1519.
[156] Ruiz, C., Cid, V.J., Lussier, M., Molina, M. and Nombela, C.
(1999) A large-scale sonication assay for cell wall mutant analysis
in yeast. Yeast 15, 1001^1008.
[157] Stateva, L.I., Oliver, S.G., Trueman, L.J. and Venkov, P.V. (1991)
Cloning and characterization of a gene which determines osmotic
stability in Saccharomyces cerevisiae. Mol. Cell. Biol. 11, 4235^4243.
[158] Tomlin, G.C., Hamilton, G.E., Gardner, D.C., Walmsley, R.M.,
Stateva, L.I. and Oliver, S.G. (2000) Suppression of sorbitol dependence in a strain bearing a mutation in the SRB1/PSA1/VIG9 gene
encoding GDP-mannose pyrophosphorylase by PDE2 overexpression suggests a role for the Ras/cAMP signal-transduction pathway
in the control of yeast cell-wall biogenesis. Microbiology 146, 2133^
2146.
[159] Biely, P., Kratky, Z., Kovarik, J. and Bauer, S. (1971) E¡ect of
2-deoxyglucose on cell wall formation in Saccharomyces cerevisiae
and its relation to cell growth inhibition. J. Bacteriol. 107, 121^129.
[160] Johnson, B.F. (1968) Lysis of yeast cell walls induced by 2-deoxyglucose at their sites of glucan synthesis. J. Bacteriol. 95, 1169^
1172.
[161] Ballou, L., Hitzeman, R.A., Lewis, M.S. and Ballou, C.E. (1991)
Vanadate-resistant yeast mutants are defective in protein glycosylation. Proc. Natl. Acad. Sci. USA 88, 3209^3212.
[162] Goins, T.L. and Cutler, J.E. (2000) Relative abundance of oligosaccharides in Candida species as determined by £uorophore-assisted
carbohydrate electrophoresis. J. Clin. Microbiol. 38, 2862^2869.
[163] Pruyne, D. and Bretscher, A. (2000) Polarization of cell growth in
yeast. II. The role of the cortical actin cytoskeleton. J. Cell Sci. 113,
571^585.
[164] Rupp, S., Summers, E., Lo, H.J., Madhani, H. and Fink, G. (1999)
MAP kinase and cAMP ¢lamentation signaling pathways converge
on the unusually large promoter of the yeast FLO11 gene. EMBO
J. 18, 1257^1269.
[165] Gancedo, J.M. (2001) Control of pseudohyphae formation in Saccharomyces cerevisiae. FEMS Microbiol. Rev. 25, 107^123.
[166] Rua, D., Tobe, B.T. and Kron, S.J. (2001) Cell cycle control of
yeast ¢lamentous growth. Curr. Opin. Microbiol. 4, 720^727.
[167] Tkacz, J.S. and Lampen, J.O. (1972) Wall replication in Saccharomyces species: use of £uorescein-conjugated concanavalin A to reveal the site of mannan insertion. J. Gen. Microbiol. 72, 243^247.
[168] Bidlingmaier, S., Weiss, E.L., Seidel, C., Drubin, D.G. and Snyder,
M. (2001) The Cbk1p pathway is important for polarized cell
growth and cell separation in Saccharomyces cerevisiae. Mol. Cell.
Biol. 21, 2449^2462.
[169] Pruyne, D. and Bretscher, A. (2000) Polarization of cell growth in
yeast. I. Establishment and maintenance of polarity states. J. Cell
Sci. 113, 365^375.
[170] Hartwell, L.H. and Unger, M.W. (1977) Unequal division in Saccharomyces cerevisiae and its implications for the control of cell
division. J. Cell Biol. 75, 422^435.
[171] Ram, A.F., Van den Ende, H. and Klis, F.M. (1998) Green £uorescent protein-cell wall fusion proteins are covalently incorporated
into the cell wall of Saccharomyces cerevisiae. FEMS Microbiol.
Lett. 162, 249^255.
[172] Caro, L.H., Smits, G.J., van Egmond, P., Chapman, J.W. and Klis,
F.M. (1998) Transcription of multiple cell wall protein-encoding
genes in Saccharomyces cerevisiae is di¡erentially regulated during
the cell cycle. FEMS Microbiol. Lett. 161, 345^349.
[173] Smits, G.J. (2000) The Dynamics of Cell Wall Biogenesis in Yeast.
Thesis. University of Amsterdam.
[174] Cho, R.J., Campbell, M.J., Winzeler, E.A., Steinmetz, L., Conway,
A., Wodicka, L., Wolfsberg, T.G., Gabrielian, A.E., Landsman, D.,
Lockhart, D.J. and Davis, R.W. (1998) A genome-wide transcriptional analysis of the mitotic cell cycle. Mol. Cell 2, 65^73.
255
[175] Smits, G.J., van den Ende, H. and Klis, F.M. (2001) Di¡erential
regulation of cell wall biogenesis during growth and development in
yeast. Microbiology 147, 781^794.
[176] Odani, T., Shimma, Y., Wang, X.H. and Jigami, Y. (1997) Mannosylphosphate transfer to cell wall mannan is regulated by the
transcriptional level of the MNN4 gene in Saccharomyces cerevisiae.
FEBS Lett. 420, 186^190.
[177] Lillie, S.H. and Pringle, J.R. (1980) Reserve carbohydrate metabolism in Saccharomyces cerevisiae : responses to nutrient limitation.
J. Bacteriol. 143, 1384^1394.
[178] Hottiger, T., Boller, T. and Wiemken, A. (1989) Correlation of
trehalose content and heat resistance in yeast mutants altered in
the RAS/adenylate cyclase pathway : is trehalose a thermoprotectant? FEBS Lett. 255, 431^434.
[179] Elliott, B., Haltiwanger, R.S. and Futcher, B. (1996) Synergy between trehalose and Hsp104 for thermotolerance in Saccharomyces
cerevisiae. Genetics 144, 923^933.
[180] Francois, J. and Parrou, J.L. (2001) Reserve carbohydrates metabolism in the yeast Saccharomyces cerevisiae. FEMS Microbiol. Rev.
25, 125^145.
[181] Andrews, P.D. and Stark, M.J. (2000) Dynamic, Rho1p-dependent
localization of Pkc1p to sites of polarized growth. J. Cell Sci. 113,
2685^2693.
[182] Delley, P.A. and Hall, M.N. (1999) Cell wall stress depolarizes cell
growth via hyperactivation of RHO1. J. Cell Biol. 147, 163^174.
[183] Ketela, T., Green, R. and Bussey, H. (1999) Saccharomyces cerevisiae Mid2p is a potential cell wall stress sensor and upstream activator of the PKC1-MPK1 cell integrity pathway. J. Bacteriol. 181,
3330^3340.
[184] Cosano, I.C., Martin, H., Flandez, M., Nombela, C. and Molina,
M. (2001) Pim1, a MAP kinase involved in cell wall integrity in
Pichia pastoris. Mol. Genet. Genomics 265, 604^614.
[185] Davenport, K.R., Sohaskey, M., Kamada, Y., Levin, D.E. and
Gustin, M.C. (1995) A second osmosensing signal transduction
pathway in yeast. Hypotonic shock activates the PKC1 protein kinase-regulated cell integrity pathway. J. Biol. Chem. 270, 30157^
30161.
[186] Kamada, Y., Jung, U.S., Piotrowski, J. and Levin, D.E. (1995) The
protein kinase C-activated MAP kinase pathway of Saccharomyces
cerevisiae mediates a novel aspect of the heat shock response. Genes
Dev. 9, 1559^1571.
[187] Att¢eld, P.V. (1987) Trehalose accumulates in Saccharomyces cerevisiae during exposure to agents that induce heat shock response.
FEBS Lett. 225, 259^263.
[188] Hottiger, T., Boller, T. and Wiemken, A. (1987) Rapid changes of
heat and desiccation tolerance correlated with changes of trehalose
content in Saccharomyces cerevisiae cells subjected to shifts. FEBS
Lett. 220, 113^115.
[189] Hottiger, T., De Virgilio, C., Hall, M.N., Boller, T. and Wiemken,
A. (1994) The role of trehalose synthesis for the acquisition of thermotolerance in yeast. II. Physiological concentrations of trehalose
increase the thermal stability of proteins in vitro. Eur. J. Biochem.
219, 187^193.
[190] Neves, M.-J. and FrancZois, J. (1992) On the mechanism by which a
heat shock induces trehalose accumulation in Saccharomyces cerevisiae. Biochem. J. 288, 859^864.
[191] Singer, M.A. and Lindquist, S. (1998) Thermotolerance in Saccharomyces cerevisiae : the Yin and Yang of trehalose. Trends Biotechnol. 16, 460^468.
[192] Alonso-Monge, R., Real, E., Wojda, I., Bebelman, J.P., Mager,
W.H. and Siderius, M. (2001) Hyperosmotic stress response and
regulation of cell wall integrity in Saccharomyces cerevisiae share
common functional aspects. Mol. Microbiol. 41, 717^730.
[193] Errede, B., Cade, R.M., Yashar, B.M., Kamada, Y., Levin, D.E.,
Irie, K. and Matsumoto, K. (1995) Dynamics and organization of
MAP kinase signal pathways. Mol. Reprod. Dev. 42, 477^485.
[194] Zarzov, P., Mazzoni, C. and Mann, C. (1996) The SLT2(MPK1)
FEMSRE 738 1-8-02
256
[195]
[196]
[197]
[198]
[199]
[200]
[201]
[202]
[203]
[204]
[205]
[206]
[207]
[208]
[209]
F.M. Klis et al. / FEMS Microbiology Reviews 26 (2002) 239^256
MAP kinase is activated during periods of polarized cell growth in
yeast. EMBO J. 15, 83^91.
Rajavel, M., Philip, B., Buehrer, B.M., Errede, B. and Levin, D.E.
(1999) Mid2 is a putative sensor for cell integrity signaling in Saccharomyces cerevisiae. Mol. Cell. Biol. 19, 3969^3976.
Schekman, R. and Brawley, V. (1979) Localized deposition of chitin
on the yeast cell surface in response to mating pheromone. Proc.
Natl. Acad. Sci. USA 76, 645^649.
Gustin, M.C., Zhou, X.L., Martinac, B. and Kung, C. (1988) A
mechanosensitive ion channel in the yeast plasma membrane. Science 242, 762^765.
Batiza, A.F., Schulz, T. and Masson, P.H. (1996) Yeast responds to
hypotonic shock with a calcium pulse. J. Biol. Chem. 271, 23357^
23362.
Zhao, C., Jung, U.S., Garrett-Engele, P., Roe, T., Cyert, M.S. and
Levin, D.E. (1998) Temperature-induced expression of yeast FKS2 is
under the dual control of protein kinase C and calcineurin. Mol.
Cell. Biol. 18, 1013^1022.
Verna, J., Lodder, A., Lee, K., Vagts, A. and Ballester, R. (1997) A
family of genes required for maintenance of cell wall integrity and
for the stress response in Saccharomyces cerevisiae. Proc. Natl.
Acad. Sci. USA 94, 13804^13809.
Philip, B. and Levin, D.E. (2001) Wsc1 and Mid2 are cell surface
sensors for cell wall integrity signaling that act through Rom2, a
guanine nucleotide exchange factor for Rho1. Mol. Cell. Biol. 21,
271^280.
Zu, T., Verna, J. and Ballester, R. (2001) Mutations in WSC genes
for putative stress receptors result in sensitivity to multiple stress
conditions and impairment of Rlm1-dependent gene expression in
Saccharomyces cerevisiae. Mol. Genet. Genomics 266, 142^155.
Cid, V.J., Duran, A., del Rey, F., Snyder, M.P., Nombela, C. and
Sanchez, M. (1995) Molecular basis of cell integrity and morphogenesis in Saccharomyces cerevisiae. Microbiol. Rev. 59, 345^386.
Gustin, M.C., Albertyn, J., Alexander, M. and Davenport, K.
(1998) MAP kinase pathways in the yeast Saccharomyces cerevisiae.
Microbiol. Mol. Biol. Rev. 62, 1264^1300.
Banuett, F. (1998) Signalling in the yeasts: an informational cascade
with links to the ¢lamentous fungi. Microbiol. Mol. Biol. Rev. 62,
249^274.
Popolo, L., Gualtieri, T. and Ragni, E. (2001) The yeast cell-wall
salvage pathway. Med. Mycol. 39 (Suppl. 1), 111^121.
Heinisch, J.J., Lorberg, A., Schmitz, H.P. and Jacoby, J.J. (1999)
The protein kinase C-mediated MAP kinase pathway involved in the
maintenance of cellular integrity in Saccharomyces cerevisiae. Mol.
Microbiol. 32, 671^680.
Osmond, B.C., Specht, C.A. and Robbins, P.W. (1999) Chitin synthase III: synthetic lethal mutants and ‘stress related’ chitin synthesis that bypasses the CSD3/CHS6 localization pathway. Proc.
Natl. Acad. Sci. USA 96, 11206^11210.
Ziman, M., Chuang, J.S., Tsung, M., Hamamoto, S. and Schekman,
R. (1998) Chs6p-dependent anterograde transport of Chs3p from
the chitosome to the plasma membrane in Saccharomyces cerevisiae.
Mol. Biol. Cell 9, 1565^1576.
[210] Mazur, P., Morin, N., Baginsky, W., el-Sherbeini, M., Clemas, J.A.,
Nielsen, J.B. and Foor, F. (1995) Di¡erential expression and function of two homologous subunits of yeast 1,3-beta-D-glucan synthase. Mol. Cell. Biol. 15, 5671^5681.
[211] Ram, A.F., Brekelmans, S.S., Oehlen, L.J. and Klis, F.M. (1995)
Identi¢cation of two cell cycle regulated genes a¡ecting the beta 1,3glucan content of cell walls in Saccharomyces cerevisiae. FEBS Lett.
358, 165^170.
[212] Posas, F., Chambers, J.R., Heyman, J.A., Hoe¥er, J.P., de Nadal,
E. and Arino, J. (2000) The transcriptional response of yeast to
saline stress. J. Biol. Chem. 275, 17249^17255.
[213] Rep, M., Krantz, M., Thevelein, J.M. and Hohmann, S. (2000) The
transcriptional response of Saccharomyces cerevisiae to osmotic
shock. Hot1p and Msn2p/Msn4p are required for the induction of
subsets of high osmolarity glycerol pathway-dependent genes. J. Biol.
Chem. 275, 8290^8300.
[214] Gasch, A.P., Spellman, P.T., Kao, C.M., Carmel-Harel, O., Eisen,
M.B., Storz, G., Botstein, D. and Brown, P.O. (2000) Genomic
expression programs in the response of yeast cells to environmental
changes. Mol. Biol. Cell 11, 4241^4257.
[215] Lai, M.H., Silverman, S.J., Gaughran, J.P. and Kirsch, D.R. (1997)
Multiple copies of PBS2, MHP1 or LRE1 produce glucanase resistance and other cell wall e¡ects in Saccharomyces cerevisiae. Yeast
13, 199^213.
[216] Jiang, B., Ram, A.F., Sheraton, J., Klis, F.M. and Bussey, H. (1995)
Regulation of cell wall beta-glucan assembly: PTC1 negatively affects PBS2 action in a pathway that includes modulation of EXG1
transcription. Mol. Gen. Genet. 248, 260^269.
[217] Reynolds, T.B., Hopkins, B.D., Lyons, M.R. and Graham, T.R.
(1998) The high osmolarity glycerol response (HOG) MAP kinase
pathway controls localization of a yeast golgi glycosyltransferase.
J. Cell Biol. 143, 935^946.
[218] ter Linde, J.J., Liang, H., Davis, R.W., Steensma, H.Y., van Dijken,
J.P. and Pronk, J.T. (1999) Genome-wide transcriptional analysis of
aerobic and anaerobic chemostat cultures of Saccharomyces cerevisiae. J. Bacteriol. 181, 7409^7413.
[219] Rachidi, N., Martinez, M.J., Barre, P. and Blondin, B. (2000) Saccharomyces cerevisiae PAU genes are induced by anaerobiosis. Mol.
Microbiol. 35, 1421^1430.
[220] Abramova, N.E., Cohen, B.D., Sertil, O., Kapoor, R., Davies, K.J.
and Lowry, C.V. (2001) Regulatory mechanisms controlling expression of the DAN/TIR mannoprotein genes during anaerobic remodeling of the cell wall in Saccharomyces cerevisiae. Genetics 157,
1169^1177.
[221] Cohen, B.D., Sertil, O., Abramova, N.E., Davies, K.J. and Lowry,
C.V. (2001) Induction and repression of DAN1 and the family of
anaerobic mannoprotein genes in Saccharomyces cerevisiae occurs
through a complex array of regulatory sites. Nucleic Acids Res.
29, 799^808.
[222] Lamb, T.M., Xu, W., Diamond, A. and Mitchell, A.P. (2001) Alkaline response genes of Saccharomyces cerevisiae and their relationship to the RIM101 pathway. J. Biol. Chem. 276, 1850^1856.
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