Download Architecture and Biosynthesis of the Saccharomyces cerevisiae Cell

YEASTBOOK
CELL SIGNALING & DEVELOPMENT
Architecture and Biosynthesis of the Saccharomyces
cerevisiae Cell Wall
Peter Orlean1
Department of Microbiology, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801
ABSTRACT The wall gives a Saccharomyces cerevisiae cell its osmotic integrity; defines cell shape during budding growth, mating,
sporulation, and pseudohypha formation; and presents adhesive glycoproteins to other yeast cells. The wall consists of b1,3- and b1,6glucans, a small amount of chitin, and many different proteins that may bear N- and O-linked glycans and a glycolipid anchor. These
components become cross-linked in various ways to form higher-order complexes. Wall composition and degree of cross-linking vary
during growth and development and change in response to cell wall stress. This article reviews wall biogenesis in vegetative cells,
covering the structure of wall components and how they are cross-linked; the biosynthesis of N- and O-linked glycans, glycosylphosphatidylinositol membrane anchors, b1,3- and b1,6-linked glucans, and chitin; the reactions that cross-link wall components; and the
possible functions of enzymatic and nonenzymatic cell wall proteins.
TABLE OF CONTENTS
Abstract
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Introduction
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Wall Composition and Architecture
Polysaccharides
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Chitin:
b-Glucans:
Cross-links between polysaccharides:
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Cell wall mannoproteins
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GPI proteins:
Mild alkali-releasable proteins:
Disulfide-linked proteins:
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Strategies to identify CWP
Cell wall phenotypes
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Precursors and Carrier Lipids
Sugar nucleotides
Dolichol and dolichol phosphate sugars
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Dolichol phosphate synthesis:
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Continued
Copyright © 2012 by the Genetics Society of America
doi: 10.1534/genetics.112.144485
Manuscript received May 17, 2012; accepted for publication August 6, 2012
Supporting information is available online at http://www.genetics.org/lookup/suppl/doi:10.1534/genetics.112.144485/-/DC1.
1
Address for correspondence: Department of Microbiology, University of Illinois at Urbana-Champaign, B-213 Chemical and Life Sciences Laboratory, 601 South Goodwin Ave.,
Urbana, IL 61801. E-mail: [email protected]
Genetics, Vol. 192, 775–818 November 2012
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CONTENTS, continued
Dol-P-Man and Dol-P-Glc synthesis:
Biosynthesis of Wall Components Along the Secretory Pathway
N-Glycosylation
Assembly and transfer of the Dol-PP-linked precursor oligosaccharide:
Steps on the cytoplasmic face of the ER membrane:
Transmembrane translocation of Dol-PP-oligosaccharides:
Lumenal steps in Dol-PP-oligosaccharide assembly:
Oligosaccharide transfer to protein:
N-glycan processing in the ER and glycoprotein quality control:
Mannan elaboration in the Golgi:
Formation of core-type N-glycan and mannan outer chains:
Mannan side branching and mannose phosphate addition:
O-Mannosylation
Protein O-mannosyltransferases in the ER:
Extension and phosphorylation of O-linked manno-oligosaccharide chains:
Importance and functions of O-mannosyl glycans:
GPI anchoring
GPI structure and proteins that receive GPIs:
GPI structure:
Identification of GPI proteins:
Assembly of the GPI precursor and its attachment to protein in the ER:
Steps on the cytoplasmic face of ER membrane:
Lumenal steps in GPI assembly:
GPI transfer to protein:
Remodeling of protein-bound GPIs:
Sugar nucleotide transport
GDP-Man transport:
Other sugar nucleotide transport activities:
Biosynthesis of Wall Components at the Plasma Membrane
Chitin
Septum formation:
Chitin synthase biochemistry:
S. cerevisiae’s chitin synthases and auxiliary proteins:
Chitin synthase I:
Chitin synthase II and proteins impacting its localization and activity:
Chitin synthase III and proteins impacting its localization and activity:
Chitin synthesis in response to cell wall stress:
Chitin synthase III in mating and ascospore wall formation:
b1,3-Glucan
Fks family of b1,3-glucan synthases:
Roles of the Fks proteins in b1,3-glucan synthesis:
Rho1 GTPase, a regulatory subunit of b1,3-glucan synthase:
b1,6-Glucan
In vitro synthesis of b1,6-glucan
Proteins involved in b1,6-glucan assembly
ER proteins:
Homologs of the UGGT/calnexin protein quality control machinery:
Fungus-specific ER chaperones required for b1,6-glucan
synthesis:
More widely distributed secretory pathway proteins:
Kre6 and Skn1:
Kre9 and Knh1:
Plasma membrane protein Kre1:
How might b1,6-glucan be made?:
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Continued
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P. Orlean
CONTENTS, continued
Remodeling and Cross-Linking Activities at the Cell Surface
Order of incorporation of components into the cell wall
Incorporation of GPI proteins into the wall
Incorporation of PIR proteins into the wall
Cross-linkage of chitin to b1,6- and b1,3-glucan
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Cell Wall-Active and Nonenzymatic Surface Proteins and Their Functions
Known and predicted enzymes
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Chitinases:
b1,3-glucanases:
Exg1, Exg2, and Ssg/Spr1 exo-b1,3-glucanases:
Bgl2, Scw4, Scw10, and Scw11 endo-b1,3-glucanases:
Eng1/Dse4 and Eng2/Acf2 endo-b1,3-glucanases:
Gas1 family b1,3-glucanosyltransferases:
Yapsin aspartyl proteases:
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Nonenzymatic CWPs
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Structural GPI proteins:
Sps2 family:
Tip1 family:
Sed1 and Spi1:
Ccw12:
Other nonenzymatic GPI proteins:
Flocculins and agglutinins:
Non-GPI-CWP:
PIR proteins:
Scw3 (Sun4):
Srl1:
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What Is Next?
T
HE wall gives Saccharomyces cerevisiae its morphologies
during budding growth, pseudohypha formation, mating, and sporulation; it preserves the cell’s osmotic integrity;
and it provides a scaffold to present agglutinins and flocculins to other yeast cells. The wall consists of mannoproteins,
b-glucans, and a small amount of chitin, which become
cross-linked in various ways. Wall composition and organization vary during growth and development. During the
budding cycle, deposition of chitin is tightly controlled,
and expression of certain hydrolases involved in cell separation is daughter cell-specific. The wall can be weakened, and
the cell consequently stressed, by treatment with polysaccharide binding agents such as Calcofluor White (CFW), Congo
Red, sodium dodecyl sulfate (SDS), aminoglycoside antibiotics, and b-glucanase preparations or by mutational loss of
capacity to make a wall component. Such stresses commonly
activate the cell wall integrity (CWI) pathway (Levin 2011)
and result in compensatory synthesis of wall material.
Up to a quarter of the genes in S. cerevisiae have some role
in maintenance of a normal wall. From the results of a survey
of deletion strains for cell wall phenotypes, De Groot et al.
(2001) estimated that 1200 genes, not counting essential
ones, impact the wall. Most of the effects, however, are in-
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direct, and the number of genes that encode enzymes directly
involved in biosynthesis or remodeling of the wall, or nonenzymatic wall proteins, is now 180 (see Supporting Information, Table S1). This review covers these proteins, with
emphasis on the wall of vegetative cells during the budding
cycle and in response to stress. Wall synthetic activities will be
covered in the context of their cellular localization, starting
with precursors in the cytoplasm, proceeding along the secretory pathway from the endoplasmic reticulum (ER) to the
plasma membrane, and culminating with the events outside
the plasma membrane that generate covalent cross-links between wall components. Additional information about individual proteins and the phenotypes of strains lacking them is
presented in File S1, File S2, File S3, File S4, File S5, File S6,
File S7, File S8, and File S9. Earlier work on the yeast cell
wall has been reviewed by Ballou (1982), Fleet (1991),
Orlean (1997), Kapteyn et al. (1999a), Cabib et al. (2001),
Klis et al. (2002, 2006), and Lesage and Bussey (2006).
Wall Composition and Architecture
The wall accounts for 15–30% of the dry weight of a vegetative S. cerevisiae cell (Aguilar-Uscanga and François 2003;
S. cerevisiae Cell Wall
777
Yin et al. 2007). It is 110–200 nm wide, as estimated from
transmission electron micrographs and by using an atomic
force microscope to detect surface accessibility of “molecular
rulers” consisting of versions of the plasma membrane sensor
Wsc1 with different lengths (Dupres et al. 2010; Yamaguchi
et al. 2011). The wall’s major components are b1,3- and
b1,6-linked glucans, mannoproteins, and chitin, which can
be covalently joined to form higher-order complexes. The
b1,6-glucan, although quantitatively a minor component
of the wall, has a central role in cross-linking wall components (Kollar et al. 1997). Some mannoproteins have or are
predicted to have enzymatic activity as hydrolases or crosslinkers; others may have structural roles or mediate “social
activity” by serving as mating agglutinins or flocculins.
Among the latter, Flo1 and Flo11 promote formation of extensive mats of cells, or biofilms (Reynolds and Fink 2001;
Beauvais et al. 2009; Bojsen et al. 2012).
Electron micrographs of thin sections through the wall of
vegetative cells reveal two layers. The outer one is electrondense, has a brush-like surface (Osumi et al. 1998) (Baba
et al. 1989; Osumi et al. 1998); Kapteyn et al. 1999a; Hagen
et al. 2004; Yamaguchi et al. 2011), and can be removed
by proteolysis (Kopecka et al. 1974; Zlotnik et al. 1984); it
therefore consists mostly of mannoproteins. The inner layer,
more electron transparent, is microfibrillar and b-glucanasedigestible, indicating that its major components are glucans.
The relative thicknesses of the two layers and their apparent
organization can be altered in cell wall mutants.
Relative amounts and localization of individual wall
components vary depending on cell cycle or developmental
stage, growth phase, nutritional conditions, and wall stresses
imposed by hypo-osmolarity, mutational loss of wall biosynthetic activities or wall proteins, or drug treatment.
Variations in wall composition and organization impact the
extent to which the wall is a barrier to export of soluble,
secreted proteins to the medium. Some proteins can be
retained by the wall outside the plasma membrane in the
periplasmic space; in the case of Suc2, this is due to the ability
of the protein to form large multimers (Orlean 1987). The
barrier function of the wall is dependent on growth phase and
cultural conditions, with the walls of growing cells being
more porous (De Nobel and Barnett 1991). Native glycoproteins such as Cts1, as well as many heterologously expressed
soluble glycoproteins with masses up to 400 kDa, can pass
through the wall of logarithmically growing cells to the medium, whereas walls of stationary-phase cells are less porous
(De Nobel et al. 1990; Kuranda and Robbins 1991). The
relatively high porosity of walls of logarithmic-phase cells
could reflect a lower degree of cross-linking, but the dissolution of septal material that occurs when dividing cells separate could also release wall proteins to the medium (see
Order of incorporation of components into the cell wall). Perspectives on wall organization are provided by Kapteyn et al.
(1999a), Klis et al. (2002, 2006), Latgé (2007), Pitarch et al.
(2008), and Gonzalez et al. (2010a). The major wall components and strategies for isolating them are as follows.
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P. Orlean
Polysaccharides
Wall polysaccharides are typically separated into three
fractions defined on the basis of their solubility in alkali
and acid (Fleet 1991). These fractions contain differing relative amounts of b1,3- and b1,6-linked glucans and mannan
(Magnelli et al. 2002) and also differ in whether and to what
extent the glucans are cross-linked to chitin, which determines their solubility in alkali. Determination of Man-to-Glc
ratios in total acid hydrolysates of walls has been useful in
assessing the impact of mutations on wall composition (Ram
et al. 1994; Dallies et al. 1998). Digestion of isolated walls
and wall fractions with linkage-specific glycosidases has
been used to quantify wall components and determine the
fine structure of b1,6-glucan (Boone et al. 1990; Magnelli
et al. 2002; Aimanianda et al. 2009), as well as to generate
oligosaccharides for structural analysis and characterization
of linkages between polymers (Kollar et al. 1995, 1997).
Chitin: This polymer of b1,4-linked GlcNAc contributes only
1–2% of the dry weight of the wall of unstressed wild-type
cells. Chitin is normally deposited in a ring in the neck between a mother cell and its emerging bud, in the primary
division septum, and in the lateral walls of newly separated
daughter cells. Chitin can be visualized in situ by staining
with CFW, which reveals that most of it is present in division
septa and bud scars. Chitin in lateral walls and in division
septa can also be detected by immunoelectron microscopy
(Shaw et al. 1991). Chitin levels are typically determined
after extraction of walls with acid and alkali or hot SDS,
followed by acid or enzymatic hydrolysis and quantification
of GlcNAc (Kang et al. 1984; Orlean et al. 1985; Dallies et al.
1998; Magnelli et al. 2002). The average length of chitin in
b-glucanase-digested septa is 110 GlcNAc residues (Kang
et al. 1984). However, chitin occurs in three different and
polydisperse forms in the wall: in addition to free chitin,
some is bound to b1,3-glucan and present mainly in the
neck between mother and daughter cell, whereas a lesser
amount, found in lateral walls, is bound to b1,6-glucan,
which is in turn linked to mannan and b1,3-glucan (Cabib
and Duran 2005; Cabib 2009). Chitin levels increase in response to mating pheromones (Schekman and Brawley
1979; Orlean et al. 1985; see Sugar nucleotides) and delocalized chitin in lateral walls can increase to as much as 20%
of the wall in S. cerevisiae mutants mounting the cell wall
stress response (Kapteyn et al. 1997, 1999a; Popolo et al.
1997; Dallies et al. 1998; Ram et al. 1998; Osmond et al.
1999; Valdivieso et al. 2000; Magnelli et al. 2002; see Chitin
synthesis in response to cell wall stress).
b-Glucans: b-linked glucans compose 30–60% of the dry
weight of the wall and can be separated into three fractions
that contain both b1,3 and b1,6 linkages. The major fraction, which makes up 35% of the dry weight of the wall, is
an acid- and alkali-insoluble b1,3-glucan with a degree of
polymerization of 1500 and b1,3-linked glucan side chains
initiated at branching b1,6-linked glucoses that represent
Figure 1 Wall components and cross-links between them. (A) Reducing end of chitin linked to
a side-branching b1,3-Glc on b1,6-glucan. (B) Reducing end of chitin linked to a nonreducing end
of b1,3-glucan. (C) Reducing end of b1,3-glucan
chain linked to a side-branching b1,6-Glc on b1,3glucan. (D) Reducing end of GPI glycan (possibly
the a1,4-Man) to internal Glc in b1,6-glucan (linkage to nonreducing end of b1,6-glucan is also possible). (E) Ester linkages between b1,3-Glc and
g-carboxyl groups of glutamates in PIR protein internal repeats. (F) Disulfide link between CWP.
Chemical treatments used to release CWP are
indicated.
3% of the whole polymer (Fleet 1991). The nonreducing
ends of b1,3-glucan chains in this fraction can be linked to
chitin, rendering the b-glucan insoluble (Kollar et al. 1995;
see below). A second b-glucan fraction, representing 20%
of the dry weight of the wall, is similar in size and composition to the alkali-insoluble b1,3-glucan, but soluble in alkali because it is not cross-linked to chitin (Hartland et al.
1994). A third fraction, making up 5% of the dry weight of
the wall, can be released from alkali-insoluble glucan by
extraction with acid or digestion with endo-b1,3-glucanase
(Manners et al. 1973; Boone et al. 1990). This fraction is
a b1,6-glucan with a degree of polymerization of 140, in
which 14% of the b1,6-linked residues bear a side-branching
b1,3 Glc (Manners et al. 1973). A procedure involving serial
digestion with purified hydrolases has also been used to
separate and quantify b1,3- and b1,6-glucan, mannan, and
chitin (Magnelli et al. 2002). The b1,6-glucan was released
from the high-molecular-weight material remaining after
treatment of walls with a mixture of b1,3-glucanase and chitinase by digestion with recombinant endo-b1,6-glucanase.
The b1,6-glucan was therefore recovered as a mixture of
oligosaccharides whose major component was Glcb1,6Glc,
and which also contained Glcb1,6Glcb1,6Glc and smaller
amounts of Glcb1,3Glcb1,6Glc and Glcb1,6Glcb1,6Glc with
a b1,3-Glc branching from its middle Glc (Magnelli et al.
2002). The degree of b1,3 branching inferred from the oligosaccharide profile was similar to that reported by Manners
et al. (1973). This b1,6-glucan analysis would also include
the b1,6-glucan present in the alkali-soluble cell wall fraction, which is not included in procedures involving alkali
extraction. In another approach, b1,6-glucan was isolated
following extraction of intact cells with hot SDS and mercaptoethanol, treatment with hot alkali under reducing conditions, and b1,3-glucanase digestion of the alkali-insoluble
material (Aimanianda et al. 2009). The b1,3-glucanase releasable material was a b1,6-glucan of 190–200 glucoses
with, on average, a b1,3-Glc or a b1,3-Glcb1,3-Glc side
branch on every fifth b1,6-linked glucose (Aimanianda
et al. 2009).
Cross-links between polysaccharides: Three types of linkages between wall polysaccharides have been described
(Figure 1). The first is a b1,4-linkage between the reducing
end of a chitin chain and the nonreducing end of a b1,3linked glucan (Kollar et al. 1995), and up to half of the chitin
chains in the wall may be linked to b-glucan in this way.
Because there is about one chitin-b-glucan linkage per 8000
hexoses, these rare cross-links have a major impact on the
solubility of b-glucan (Kollar et al. 1995). The second linkage
is between the reducing end of chitin and the nonreducing
end of a b1,3-Glc that branches off b1,6-glucan (Kollar et al.
1997; see Remodeling and Cross-Linking Activities at the Cell
Surface). The configuration of this linkage is either b1,2- or
b1,4-. The two types of chitin-b-glucan linkage are found in
different parts of the wall. In the third linkage, the reducing
ends of b1,6-glucan chains can be attached to b1,3-glucan,
but the configuration is unknown (Kollar et al. 1997).
Cell wall mannoproteins
Yeast cell wall proteins can bear asparagine- (N-)linked
glycans, O-linked manno-oligosaccharides, and often a glycosylphosphatidylinositol (GPI) as well. The N-linked glycans can be extended with an outer chain of 50 or more
a1,6-linked Man that is extensively decorated with short
a1,2-Man side branches terminated in a1,3-Man. Phosphodiester-linked mannoses can also be attached to a1,2-linked
residues. Many glycoproteins also bear O-mannosyl glycans,
which are often present in Ser/Thr-rich stretches.
Proteins relevant to the wall can be placed into one of
three groups. The first contains those with the potential to
participate in wall construction as hydrolases or transglycosidases. The second contains nonenzymatic agglutinins, flocculins, or b1,3-glucan cross-connectors (Klis et al.
2006, 2010; Dranginis et al. 2007; Goossens and Willaert
S. cerevisiae Cell Wall
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2010). Most, if not all the proteins in these two groups are
glycosylated. Proteins that are covalently attached to cell wall
glycan are referred to as CWP (Yin et al. 2005) and fall into
the subgroups below. The third group consists of single-pass
plasma membrane proteins with short C-terminal cytoplasmic
domains and long Ser/Thr-rich extracellular regions. These
include Wsc1, Wsc2, and Wsc3, which also have N-terminal
cysteine-rich domains, as well as Mtl1 and Mid2. These are
mechanosensors that detect cell wall stress and activate the
CWI pathway (Rodicio and Heinisch 2010; Levin 2011). CWP
and cell wall-active enzymes are discussed in Cell Wall-Active
and Nonenzymatic Surface Proteins and Their Functions.
GPI proteins: These receive a GPI that initially anchors them
in the outer face of the plasma membrane, but many then
become cross-linked to b1,6-glucan via a remnant of the GPI
(Gonzalez et al. 2009). Results to date suggest that the GPI
is cleaved between its GlcN residue and Man, whereupon
the mannose’s reducing end is glycosidically linked to a nonreducing end of b1,6-glucan or to a Glc in a b1,6-Glc chain
(Kollar et al. 1997; Fujii et al. 1999). The b1,6-glucan to
which the GPI-CWP is attached is in turn linked to b1,3glucan and chitin (Kapteyn et al. 1996; Van der Vaart
et al. 1996; Kollar et al. 1997; Fujii et al. 1999; Figure 1).
Some wall-bound GPI proteins may retain enzymatic activity, whereas others may have a structural role (Yin et al.
2005). GPI-CWP are released by treatment with hydrogen
fluoride (HF)/pyridine, which cleaves the phosphodiester
of the GPI that links Man and the phosphoethanolamine
(Etn-P) moiety that is linked to protein (Yin et al. 2005).
Proteins released in this way have a C-terminal GPI signalanchor sequence, and this, and signals for wall anchorage of
GPI-CWP, are discussed in Lumenal steps in GPI assembly and
in Incorporation of GPI proteins into the cell wall. At least one
GPI-CWP, Cwp1, can additionally be linked to the wall via
an alkali-labile linkage (Kapteyn et al. 2001).
Mild alkali-releasable proteins: These include four proteins
with internal repeats (PIR proteins), which have multiple
copies of the internal repeat sequence SQ[I/V][S/T/G]DGQ
[I/V]Q[A][S/T/A] (Toh-E et al. 1993) [simplified to DGQ
[hydrophobic amino acid]Q by Klis et al. (2010)] and are
released by mild alkali or b1,3-glucanase (Mrša et al. 1997).
PIR proteins have no GPI attachment sequence and are not
linked to b1,6-glucan; rather, they are ester-linked to b1,3glucan via side chains of amino acids in the repeat sequences
(Ecker et al. 2006; see Incorporation of PIR proteins into the
cell wall). Because PIR proteins can form several linkages to
b1,3-glucan, they could interconnect glucans. Single PIR
repeats are also present in certain GPI-CWP (see Incorporation of PIR proteins into the cell wall), and additional proteins
lacking PIR sequences can be also extracted with alkali or
b1,3-glucanase (Yin et al. 2005; see Cell Wall-Active and
Nonenzymatic Surface Proteins and Their Functions).
Disulfide-linked proteins: Various proteins can be released
from the walls of living cells with sulfhydryl reagents,
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P. Orlean
indicating that they are directly attached via disulfides or
retained behind a network of disulfide-linked proteins
(Orlean et al. 1986; Cappellaro et al. 1998; Moukadiri
et al. 1999; Moukadiri and Zueco 2001; Insenser et al.
2010). Disulfide-linked mannoproteins create a barrier that
protects wall polysaccharides from externally added glycosylhydrolases, making mercaptoethanol and protease pretreatment necessary for spheroplasting with lytic enzymes
(Zlotnik et al. 1984). Furthermore, the ability of the cysteine-rich domain of Wsc1 to form disulfide cross-links is important for this mechanosensor in forming clusters and in
functioning in CWI signaling (Heinisch et al. 2010; Dupres
et al. 2011).
Strategies to identify CWP
Biochemistry and bioinformatics have been used to identify
CWP. Because proteins can be associated with the wall in
different ways, different treatments are necessary to release
them. Separation and identification of individual CWP can
be complicated by their heavy and heterogeneous glycosylation. CWP can be released from the wall by treatment with
b1,3- and b1,6-glucanases (Van der Vaart et al. 1995; Mrša
et al. 1997; Shimoi et al. 1998). In one approach, labeling of
intact cells with a membrane-impermeable biotinylation reagent, followed successively by SDS and mercaptoethanol
extraction and then mild alkali or b1,3-glucanase treatment,
led to identification of nine “soluble cell wall” (Scw) and 11
“covalently linked cell wall” (Ccw) proteins (Mrša et al.
1997). In another approach, isolated walls, extracted with
SDS, mercaptoethanol, NaCl, and EDTA, were then treated
with HF/pyridine or mild alkali, and the CWP released were
identified by mass spectrometry. Additional CWP were identified following proteolytic digestion of walls, the two procedures yielding 19 CWP, including GPI and PIR proteins
and alkali-releasable proteins without PIR sequences (Yin
et al. 2005, 2007). These studies led to the estimate that
a dividing haploid cell contains 2 · 106 covalently attached CWPs and the suggestion that CWP form a densely
packed surface layer (Yin et al. 2007). A strategy that also
permitted identification of noncovalently associated surface
proteins used treatment of intact cells with dithiothreitol
followed by two-dimensional electrophoretic separation, or
direct proteolytic digestion and isolation of peptides, and
then mass spectrometric protein fingerprinting (Insenser
et al. 2010). The 99 proteins so identified included CWP
and glycosylhydrolases, as well as proteins associated with
intracellular functions. The presence in the wall of proteins
considered cytosolic raises the possibility that they reach the
wall via a nonconventional export pathway (Nombela et al.
2006; Insenser et al. 2010). However, mercaptoethanol can
make the plasma membrane permeable to cytosolic proteins
(Klis et al. 2007).
Bioinformatics has been used identify proteins likely to
receive a GPI anchor; hence, members of the major class of
CWP. In silico surveys for GPI attachment sequences reveal
that the S. cerevisiae proteome contains 60–70 potential GPI
proteins, which often contain Ser/Thr-rich stretches (Caro
et al. 1997; Hamada et al. 1998a; De Groot et al. 2003;
Eisenhaber et al. 2004).
Cell wall phenotypes
Cell wall phenotypes that are typically scored are sensitivity
to hypo-osmotic stress, which can be tested on half-strength
yeast extract peptone medium (Valdivia and Schekman
2003); sensitivity or resistance to CFW and Congo Red; sensitivity to aminoglycosides, b1,3-glucan synthase inhibitors,
caffeine, SDS, and K1 killer toxin; and sensitivity to b1,3glucanase preparations (Ram et al. 1994; Hampsey 1997;
Lussier et al. 1997b; De Groot et al. 2001).
Precursors and Carrier Lipids
Sugar nucleotides
Glycosyltransferases involved in wall biogenesis use UDP-Glc,
UDP-GlcNAc, and GDP-Man or dolichol phosphate (Dol-P)
Man or Dol-P-Glc as donors. UDP-Glc is formed from UTP and
Glc-1-P by the essential UDPGlc pyrophosphorylase Ugp1
(Daran et al. 1995). Impairment of UDP-Glc synthesis ultimately impacts formation of cell wall b-glucans, although
cells with no more than 5% of the activities of the phosphoglucomutases and Ugp1 that generate UDP-Glc are unaffected in growth and viability (Daran et al. 1997). GDPMan is formed from Fru-6-P by the successive actions of
phosphomannose isomerase (Pmi40), phosphomannomutase
(Sec53), and GDP-Man pyrophosphorylase (Psa1/Srb1/Vig9),
which are all encoded by essential genes, and loss of any of
these enzyme activities leads to severe glycosylation and secretion defects (Hashimoto et al. 1997; Orlean 1997; Yoda
et al. 2000). Elevated expression of GDP-Man pyrophosphorylase, which presumably increases GDP-Man levels, corrects
the N-glycosylation defects in alg1 and alg2 mutants and the
mannosylation and GPI synthetic defects in dpm1 cells (Janik
et al. 2003). GDP-Man transport into the Golgi lumen is discussed in Sugar nucleotide transport.
The pathway for UDP-GlcNAc formation (Milewski et al.
2006) involves conversion of Fru-6-P to GlcN-6-P by glutamine:Fru-6-P amidotransferase Gfa1 (Watzele and Tanner
1989), N-acetylation of GlcN-6-P by Gna1 (Mio et al.
1999), conversion of GlcNAc-6-P to GlcNAc-1-P by the
GlcNAc phosphate mutase Agm1/Pcm1 (Hofmann et al.
1994), and formation of UDP-GlcNAc by the pyrophosphorylase Uap1/Qri1 (Mio et al. 1998). Deficiencies in these
enzymes lead to formation of short chains of undivided cells,
swelling, and eventual lysis, a phenomenon known as glucosamineless death (Ballou et al. 1977; Mio et al. 1998,
1999). Glucosamine supply is highly regulated and impacts
chitin levels, which increase in response to mating pheromones and cell wall stress (File S1).
dolichol (Schenk et al. 2001a; Grabinska and Palamarczyk
2002) starts with extension of trans farnesyl-PP by successive addition of cis-isoprene units by the homologous cisprenyltransferases Rer2 and Srt1 (Sato et al. 1999; Schenk
et al. 2001b). Rer2 is dominant and makes dolichols with
10–14 isoprene units, whereas dolichols made by Srt1 in
cells lacking Rer2 contain 19–22 isoprenes. rer2D strains
have severe defects in growth and in N- and O-glycosylation
(Sato et al. 1999). The next two steps are likely the removal
of the two phosphates from dehydrodolichyl diphosphate by
unknown enzymes. The a-isoprene unit of the polyprenol is
then reduced, and Dfg10 is responsible for much of this
activity (Cantagrel et al. 2010; File S1). Dolichol is likely
next phosphorylated by the CTP-dependent Dol kinase
Sec59 (Heller et al. 1992).
Dol-PP generated on the lumenal side of the ER membrane after transfer of the N-linked oligosaccharide to
protein is dephosphorylated to Dol-P and Pi on that side of
the membrane by the phosphatase Cwh8/Cax4 (Van Berkel
et al. 1999; Fernandez et al. 2001). CWH8-disruptants have
an N-glycosylation defect and a growth defect that is partially suppressed by high-level expression of RER2, SEC59,
and the lipid phosphatase gene LPP1. Cwh8 likely has a role in
recycling of Dol-PP for use in new rounds of N-glycosylation
on the cytoplasmic face of the ER membrane.
Dol-P-Man and Dol-P-Glc synthesis: Dol-P-Man and Dol-PGlc are the donors in the lumenal glycosyltransfers that
occur in protein O-mannosylation and the assembly pathways for the Dol-PP-linked precursor in N-glycosylation and
the GPI anchor precursor glycolipid. Dol-P-Man is formed
upon transfer of Man from GDP-Man to Dol-P by the Dol-PMan synthase Dpm1 (Orlean et al. 1988; Orlean 1990).
Temperature-sensitive dpm1 mutants have cell wall defects,
consistent with a general block of glycosylation and GPI
anchoring, and these phenotypes are suppressed by highlevel expression of RER2, which presumably elevates Dol-P
levels (Orlowski et al. 2007).
Dol-P-Glc is formed from UDP-Glc and Dol-P. Deletion of
the synthase gene, ALG5, is not lethal, and the disruptants
show no obvious growth defects (Te Heesen et al. 1994).
Because Dol-P-Man and Dol-P-Glc are used in lumenal reactions, and because spontaneous transmembrane translocation of these glycolipids is not favored energetically, their
translocation may be protein-mediated. Assays for Dol-PMan flipping have been reported (Haselbeck and Tanner
1982; Sanyal and Menon 2010), but a protein involved
has yet to be identified. One possibility is that the Dol-PMan and Dol-P-Glc-utilizing transferases are their own flippases (Burda and Aebi 1999).
Dolichol and dolichol phosphate sugars
Biosynthesis of Wall Components Along the
Secretory Pathway
Dolichol phosphate synthesis: Yeast dolichols contain 14–
18 isoprene units (Jung and Tanner 1973). Biosynthesis of
Cell surface proteins can be modified with N-glycans,
O-linked manno-oligosaccharides, and a GPI anchor as they
S. cerevisiae Cell Wall
781
Figure 2 Assembly of the Dol-PP-linked precursor oligosaccharide in N-glycosylation, its transfer to protein, and subsequent glycan processing. Residues
added at the cytoplasmic face of the ER membrane originate from sugar nucleotides, whereas Dol-P sugars generated at the cytoplasmic face of the
membrane are the donors in lumenal transfers. Symbols are adaptations of those used by the Consortium of Glycobiology Editors in Essentials in
Glycobiology (Varki and Sharon 2009).
transit the secretory pathway. Initial attachment of these
structures occurs in the ER lumen, and the glycans are
modified in the Golgi before the glycoproteins are deposited
in the plasma membrane or secreted from the cell, whereupon many become cross-linked to wall polysaccharides.
N-Glycosylation
N-glycosylation involves preassembly of a branched 14sugar oligosaccharide on the carrier Dol-PP in the ER
membrane and then transfer of the oligosaccharide to
selected asparagines in the ER lumen (Burda and Aebi
1999; Helenius and Aebi 2004; Lehle et al. 2006; Larkin
and Imperiali 2011). The first 7 sugars are transferred
from sugar nucleotides on the cytosolic side of the ER membrane, and the remainder from Dol-P on the lumenal side
(Figure 2).
Assembly and transfer of the Dol-PP-linked precursor
oligosaccharide: Steps on the cytoplasmic face of the ER
membrane: These steps are (i) transfer of GlcNAc-1-P from
UDP-GlcNAc to Dol-P by Alg7, the target of the N-glycosylation
inhibitor tunicamycin (Barnes et al. 1984), (ii) transfer of b1,4-
782
P. Orlean
GlcNAc from UDP-GlcNAc by heterodimeric Alg13/Alg14
(Bickel et al. 2005; Chantret et al. 2005; Gao et al. 2005),
(iii) transfer of a b1,4-linked Man by Alg1 (Couto et al.
1984), (iv) successive transfer of an a1,3 and an a1,6
Man by Alg2 (O’Reilly et al. 2006; Kämpf et al. 2009), and
(v) transfer of two a1,2-linked Man by Alg11 (Cipollo et al.
2001; O’Reilly et al. 2006; Absmanner et al. 2010). These
proteins act in higher-order complexes (Gao et al. 2004;
Noffz et al. 2009; File S2).
Transmembrane translocation of Dol-PP-oligosaccharides:
Dol-PP-GlcNAc2Man5 formed on the cytoplasmic face of the
ER membrane is somehow translocated into the lumen
(Burda and Aebi 1999; Helenius and Aebi 2002), and Rft1
is a candidate for the flippase (Helenius et al. 2002). Strains
deficient in Rft1 accumulate Dol-PP-GlcNAc2Man5, but retain
Alg3 Man-T activity and are unaffected in O-mannosylation
or in GPI assembly, ruling out deficiences in Dol-P-Man supply to the lumen. Furthermore, high level expression of
RFT1 partially suppresses the growth defect of alg11D and
leads to increased levels of lumenal Dol-PP-GlcNAc2Man6-7
and an increase in glycosylation of the reporter carboxypeptidase Y, consistent with enhanced flipping of the suboptimal
substrate Dol-PP-GlcNAc2Man3 (Helenius et al. 2002). However, although the above evidence is consistent with Rft1
being the flippase, depletion of Rft1 did not lead to loss of
flipping activity measured in vitro (Frank et al. 2008; Rush
et al. 2009; File S2).
Lumenal steps in Dol-PP-oligosaccharide assembly: Dol-PPGlcNAc2Man5 is extended by four Man and three Glc on the
lumenal side of the ER membrane using Dol-P-Man and DolP-Glc as donors. Alg3 adds the sixth, a1,3-Man to the a1,6
Man of Dol-PP-GlcNAc2Man5 (Aebi et al. 1996; Sharma et al.
2001), Alg9 then transfers an a1,2-linked Man to the Man
added by Alg3 (Burda et al. 1999; Cipollo and Trimble
2002), and Alg12 next adds the eighth, a1,6-Man to the
Man added by Alg9 (Burda et al. 1999). Alg9 acts again to
add the ninth Man, a1,2-linked Man to the Man added by
Alg12 (Frank and Aebi 2005). Two a1,3-linked Glc are successively added by Alg6 and Alg8 to extend the arm of the
heptasaccharide ending in the a1,2-linked Man transferred
by Alg11, and finally, Alg10 adds an a1,2-Glc (Stagljar et al.
1994; Reiss et al. 1996; Burda and Aebi 1998). The six DolP-sugar-utilizing transferases are members of a family of
multispanning membrane proteins that includes Man-T involved in GPI biosynthesis (Oriol et al. 2002).
Oligosaccharide transfer to protein: GlcNAc2Man9Glc3 is
transferred from Dol-PP to asparagines by the oligosaccharyltransferase complex (OST) (Knauer and Lehle 1999a; Yan
and Lennarz 2005a; Kelleher and Gilmore 2006; Lehle et al.
2006; Weerapana and Imperiali 2006; Lennarz 2007; Larkin
and Imperiali 2011). Acceptor asparagines occur in the
sequon Asn-X-Ser/Thr, where X can be any amino acid except
Pro. Mass spectrometric analyses of wall-derived peptides
revealed that 85% of sequons were completely occupied, with
preferential usage Asn-X-Thr over Asn-X-Ser sites (Schulz
and Aebi 2009). Analyses of protein-linked N-glycans in
mutants defective in the elaboration of the Dol-PP-linked
precursor indicate that structures smaller than GlcNAc2Man9Glc3 can be transferred in vivo.
Yeast OST consists of Stt3, Ost1, Ost2, Wbp1, Swp1,
Ost4, Ost5, and either of the paralogues Ost3 or Ost6. The
first five are encoded by essential genes. Two OST complexes can be formed, containing either Ost3 or Ost6
(Schwarz et al. 2005; Spirig et al. 2005; Yan and Lennarz
2005b). The Ost3-containing complex is about four times as
abundant as the Ost6-containing one (Spirig et al. 2005).
Genetic interaction studies and coimmunoprecipitation and
chemical cross-linking experiments suggest the existence of
three OST subcomplexes: (i) Swp1-Wbp1-Ost2, (ii) Stt3Ost4-Ost3, and (iii) Ost1-Ost5 (Karaoglu et al. 1997; Reiss
et al. 1997; Spirig et al. 1997; Knauer and Lehle 1999b; Kim
et al. 2003; Li et al. 2003; Kelleher and Gilmore 2006; File
S2). OST complexes themselves may function as dimers
(Chavan et al. 2006).
Stt3 is the catalytic subunit of OST. It can be chemically
cross-linked to peptides derivatized with photoactivatable
groups (Yan and Lennarz 2002; Nilsson et al. 2003), and
its bacterial and protist homologs transfer glycans to protein
substrates (Wacker et al. 2002; Kelleher and Gilmore 2006;
Kelleher et al. 2007; Nasab et al. 2008; Hese et al. 2009).
Ost3 and Ost6 have a lumenal thioreductase fold with
a CXXC motif common to proteins involved in disulfide bond
shuffling during oxidative protein folding (Kelleher and
Gilmore 2006; Schulz et al. 2009), and the proteins likely
form transient disulfide bonds with nascent proteins and
promote efficient glycosylation of Asn-X-Ser/Thr sites by
delaying oxidative protein folding (Schulz and Aebi 2009;
Schulz et al. 2009). The Swp1p, Wbp1p, and Ost2p subcomplex may confer the preference of OST for GlcNAc2Man9Glc3
(Pathak et al. 1995; Kelleher and Gilmore 2006), Ost4 is
required for recruitment of Ost3 and Ost6 to OST and also
interacts with Stt3 (Karaoglu et al. 1997; Spirig et al. 1997;
Knauer and Lehle 1999b; Kim et al. 2000, 2003; Spirig et al.
2005), and Ost1 may funnel nascent polypeptides to Stt3
(Lennarz 2007). OST may be subject to regulation by the
CWI pathway via an interaction between Pkc1 or components of the PKC pathway with Stt3 (Park and Lennarz
2000; Chavan et al. 2003a; File S2).
N-glycan processing in the ER and glycoprotein quality
control: Protein-linked GlcNAc2Man9Glc3 is processed to
glycans that are recognized by mechanisms that monitor
correct protein folding and permit export from the ER or
ensure degradation if the protein misfolds (Herscovics
1999; Aebi et al. 2010). Processing proceeds by removal of
the a1,2-linked Glc by glucosidase I, Gls1/Cwh41 (Romero
et al. 1997), and then of the two a1,3-linked Glc by soluble
glucosidase II, a heterodimer of catalytic Gls2/Rot2 and
Gtb1 (Trombetta et al. 1996; Wilkinson et al. 2006; Quinn
et al. 2009; Figure 2). ER mannosidase I, Mns1, removes
an a1,2 Man to generate GlcNAc2Man8 (Jakob et al. 1998;
Herscovics 1999), and, if correctly folded, proteins bearing
this glycan are exported from the ER. Un- or misfolded proteins are bound by protein disulfide isomerase Pdi1, some of
which is in complex with Mns1 homolog Htm1, which trims
the glycan to a GlcNAc2Man7 (Clerc et al. 2009; Gauss et al.
2011; File S2). Misfolded proteins with GlcNAc2Man7 are
targeted to the cytosol for destruction by the ER-associated
protein degradation (ERAD) system (Helenius and Aebi
2004). They are bound by the lectin Yos9 (Buschhorn et al.
2004; Bhamidipati et al. 2005; Kim et al. 2005; Szathmary
et al. 2005) and in turn directed to the HRD-ubiquitin ligase
complex of Hrd1 and Hrd3 for retrotranslocation to the cytoplasm (Bays et al. 2001; Deak and Wolf 2001; Gauss et al.
2006), where they are deglycosylated by peptide N-glycanase
Png1 (Suzuki et al. 2000; Hirayama et al. 2010).
In mammals and Schizosaccharomyces pombe, following
glucosidase II action, UDP-Glc:glycoprotein glucosyltransferase (UGGT) adds back an a1,3-Glc, allowing the monoglucosylated N-glycans to interact with the lumenal lectin
domains of calnexin or calreticulin (Parodi 1999; Caramelo
and Parodi 2007; Aebi et al. 2010). This interaction retains
partially folded or misfolded proteins in the ER and buys
them time to fold properly and be deglucosylated. Properly
S. cerevisiae Cell Wall
783
Figure 3 Formation of mannan outer chains and core-type N-glycans in the Golgi. Protein-bound Man8-GlcNAc2 structures are first acted on by the
Och1 a1,6-Man-T in the cis-Golgi. The initiating a1,6-Man is then elongated with 10 a1,6-linked Man by mannan polymerase (M-Pol)-I, and this chain
is then extended with up to 50 a1,6-linked Man by M-Pol-II. Kre2/Mnt1, Ktr1, Ktr2, Ktr3, and Yur1 collectively add a1,2-linked mannoses. Core-type
glycans are formed when an a1,2-linked Man is added to the Och1-derived a1,6-Man. Symbols are as used in Figure 2.
folded proteins are no longer recognized by UGGT and
exported to the Golgi, whereas persistent misfolders are removed by ERAD. In S. cerevisiae, however, this quality control mechanism does not operate because UGGT activity is
not detectable, and although the S. cerevisiae ER protein
Kre5 is a sequence homolog of S. pombe UGGT, expression
of the S. pombe UGGT cannot rescue the growth defect of
kre5 mutants. However, kre5, as well as glucosidase I and II
mutants and mutants in the calnexin homolog Cne1, are defective in b1,6-glucan synthesis, indicating roles for S. cerevisiae
homologs of players in the UGGT/calnexin quality control
system in b1,6-glucan synthesis (Jiang et al. 1996; Shahinian
et al. 1998; Simons et al. 1998; see b1,6-Glucan).
Mannan elaboration in the Golgi: N-linked glycans on
proteins are extended with a Man10-14 core-type structure or
with mannan outer chains containing up to 150–200 Man.
Both structures can be modified with mannose phosphate
(Figure 3) (Ballou 1990; Orlean 1997; Jigami 2008). The
mannoses all originate from GDP-Man and are transferred
by members of several families of redundant Golgi Man-T.
Formation of core-type N-glycan and mannan outer chains:
Formation of core structures and mannan is initiated in
the cis-Golgi by Och1, which transfers an a1,6-Man to the
a1,3-Man of the N-glycan that had been added by Alg2
(Nakayama et al. 1997). OCH1 deletion is lethal in some
strain backgrounds, and och1D strains have severe growth
defects, highlighting the importance of mannan.
Synthesis of the poly-a1,6-mannan backbone is carried
out in the cis-Golgi by two protein complexes: Man-Pol I,
see containing homologs Mnn9 and Van1, and Man-Pol II,
784
P. Orlean
containing Mnn9, Anp1, Hoc1, and related Mnn10 and
Mnn11 (Hashimoto and Yoda 1997; Jungmann and Munro
1998; Jungmann et al. 1999; File S2). M-Pol I acts first, with
its Mnn9 subunit adding the first a1,6-Man to the Och1derived Man, upon which 10–15 a1,6-Man are added in
Van1-requiring reactions (Stolz and Munro 2002; Rodionov
et al. 2009). This a1,6 backbone is further elongated with
40–60 a1,6-Man by M-Pol II, whose Mnn10 and Mnn11
subunits are responsible for the majority of the a1,6-ManT activity (Jungmann et al. 1999). Hoc1’s role is unclear.
Core-type N-glycans are formed when an a1,2-Man is added
to the Och1-derived Man, blocking elongation of an a1,6 mannan chain. The protein(s) involved have not been identified,
but presumably either they, or M-Pol I, can tell from the context
of an N-glycan which type of structure it is to bear (Lewis and
Ballou 1991; Stolz and Munro 2002; Rodionov et al. 2009).
Core-type structures are completed when that a1,2-Man, as
well as the two other terminal a1,2-Man on the Man8GlcNAc2
structure, receives a1,3 mannoses from Mnn1.
Mannan side branching and mannose phosphate addition:
Branching of the a1,6 mannan backbone is initiated by the
Mnn2 a1,2-Man-T, and Mnn5 adds a second a1,2-Man
(Rayner and Munro 1998). Mnn2 and Mnn5 make up one
of two Mnn1 subfamilies (Lussier et al. 1999). Five members
of the Ktr1 protein subfamily, Kre2/Mnt1, Yur1, Ktr1, Ktr2,
and Ktr3, also contribute to N-linked outer chain synthesis,
acting collectively in the addition of the second and subsequent a1,2-mannoses to mannan side branches (Lussier
et al. 1996, 1997a, 1999).
Core-type glycans and mannan can be modified with Man-P
on a1,2-linked mannoses. Mnn6/Ktr6, a Ktr1 subfamily
member, is mostly responsible for transferring Man-1-P
from GDP-Man, generating GMP (Wang et al. 1997; Jigami
and Odani 1999; File S2). Mnn4 is also involved in Man-P
addition but does not resemble glycosyltransferases and
may be regulatory (Odani et al. 1996). Levels of mannan
phosphorylation are highest in the late log and stationary
phases, when MNN4 expression is elevated (Odani et al.
1997). Terminal a1,2 mannoses and Man-1-Ps can be capped with a1,3-Man, added by Mnn1 (Ballou 1990; Yip et al.
1994).
O-Mannosylation
Many yeast proteins are modified on extracytoplasmic Ser or
Thr residues with linear manno-oligosaccharides. The first
Man is attached in a-linkage in the lumen of the ER, and up
to four further Man are added by Man-T that act mostly in
the Golgi.
Protein O-mannosyltransferases in the ER: The first Man is
transferred from Dol-P-Man (Strahl-Bolsinger et al. 1999;
Lehle et al. 2006; Lommel and Strahl 2009). Consistent with
the requirement for Dol-P-Man, O-mannosylation of the model
protein Cts1 is blocked in a dpm1-Ts mutant (Orlean 1990).
There are six protein O-mannosyltransferases (PMTs) in yeast.
Prototypical Pmt1 is an ER protein with seven membranespanning domains with conserved residues important for
catalysis and for interactions with acceptor peptides located
in the first lumenal loop (Strahl-Bolsinger and Scheinost
1999; Girrbach et al. 2000; Lommel et al. 2011).
Pmts function as hetero- or homodimers, and the pairs that
are formed are determined by membership of a subunit in
one of three Pmt subfamilies. Pmt1 family members Pmt1 and
Pmt5 can form heterodimers with members of the Pmt2 family (which also contains Pmt3 and Pmt6), for example, Pmt1Pmt2 and Pmt5-Pmt3 dimers, which are the most prevalent
complexes (Girrbach and Strahl 2003). Pmt4, the lone representative of the third family, forms homodimers.
Analyses of O-mannosylation of individual proteins in pmtD
strains reveal that the different Pmt complexes have specificity
for different protein substrates (File S3). Substrates for Pmt4
need to be attached to the membrane by a transmembrane
domain or a GPI anchor and have an adjacent, lumenal Ser/
Thr-rich domain, whereas Pmt1/Pmt2 substrates can be soluble or membrane-associated (Hutzler et al. 2007).
Because PMTs modify Ser and Thr, N-linked glycosylation
sites are also potential targets, and this is the case with Cwp5.
This protein contains a single sequon, NAT, that is normally
O-mannosylated by Pmt4, but which receives an N-linked
glycan in pmt4D cells (Ecker et al. 2003). O-mannosylation,
therefore, normally precedes the action of OST on Cwp5 and
may control N-glycosylation of this protein, and perhaps
others as well.
Extension and phosphorylation of O-linked mannooligosaccharide chains: The Ser- or Thr-linked Man is extended with up to four a-linked Man by GDP-Man-dependent
Man-T of the Ktr1 and Mnn1 families (Lussier et al. 1999;
Figure 4; File S3). Transfer of the first two a1,2-Man is
carried out by the Ktr1 subfamily members Ktr1, Ktr3, and
Kre2 and extension of the trisaccharide chain with one
or two a1,3-linked Man by Mnn1 family members Mnn1,
Mnt2, and Mnt3 (Lussier et al. 1997a; Romero et al. 1999).
The second a1,2-Man of an O-linked glycan can be modified
with Man-1-P by Mnn6 with the involvement of the regulator Mnn4 (Nakayama et al. 1998).
Importance and functions of O-mannosyl glycans: No
individual PMT deletion is lethal, but strains lacking certain
combinations of three Pmts, such as pmt1D pmt2D pmt4D or
pmt2D pmt3D pmt4D, are inviable, even with osmotic support, indicating that yeast must carry out some minimum
level of O-mannosylation to be viable or that one or more
essential proteins need to be O-mannosylated (Gentzsch and
Tanner 1996; Lommel et al. 2004). Moreover, strains lacking
other combinations of Pmts, such as the pmt2D pmt3D and
pmt2D pmt4D double nulls or the pmt1D pmt2D pmt3D triple null, are osmotically fragile, indicating impaired wall
assembly (Gentzsch and Tanner 1996). Analyses of pmt
mutants show that O-mannosylation can be important for
function of individual O-mannosylated proteins (File S3).
The phenotypes of pmt mutants are mimicked by treatment
with the rhodanine-3-acetic acid derivative OGT1458, which
inhibits PMT activity in vitro (Orchard et al. 2004; Arroyo et al.
2011). OGT1458 was used to analyze genome-wide transcriptional changes in response to inhibition of O-mannosylation.
Consistent with the importance of O-mannosylation in wall
construction and protein stability, consequences of defective
O-mannosylation were activation of the CWI pathway and
the unfolded protein response (Arroyo et al. 2011). Furthermore, certain genes involved in N-linked mannan outer chain
assembly were upregulated. This, together with the finding
that PMT gene transcription is elevated when N-glycosylation
is inhibited by tunicamycin (Travers et al. 2000), suggests
that the N- and O-linked glycans of cell wall mannoproteins
can compensate for one another to some extent (Arroyo
et al. 2011).
GPI anchoring
GPI structure and proteins that receive GPIs: GPI structure:
S. cerevisiae GPI anchors have the core structure protein CONH2-CH2-CH2-PO4-6-Mana1,2Mana1,6Mana1,4GlcNa1,6myoinositol phospholipid. In addition, the third, a1,2-Man,
bears a fourth a1,2-Man that is added during precursor assembly, and this Man may receive another a1,2- or a1,3linked Man in the Golgi (Fankhauser et al. 1993). The
a1,4- and a1,6-linked Man are also modified with Etn-P at
their 29- and 69-OHs, respectively, and the 2-OH of inositol is
transiently modified with palmitate (Orlean and Menon
2007; Pittet and Conzelmann 2007) (Figure 5). The lipid
moiety, initially diacylglycerol, is remodeled to a diacylglycerol with C26, acyl chains, or, in many cases, to a ceramide
(Conzelmann et al. 1992; Fankhauser et al. 1993).
S. cerevisiae Cell Wall
785
Figure 4 Biosynthesis of O-linked glycans. (A) Addition of
a-Man by protein O-mannosyltransferases in the ER lumen. Pmt4 homodimers act on membrane proteins or
GPI proteins. Representative Pmt heterodimers are shown.
(B) Extension of O-linked manno-oligosaccharides in the
Golgi. Ktr1 family members have a collective role in adding
a1,2-linked mannoses, and Mnt1 family members add
a1,3-linked mannoses. The dominant Man-T active at
each step are shown in boldface type. Man-P may be
added to saccharides with two a1,2-linked Man.
Identification of GPI proteins: Biochemical demonstrations
of a GPI on a yeast protein are rare, and the criterion of
release of a protein by treatment with Ptd-Ins-specific
phospholipase C (PI-PLC) is unreliable because although
protein-bound GPIs are mostly sensitive to PI-PLC, this
treatment does not always render the protein aqueous
soluble in the commonly used Triton X-114 fractionation
procedure (Conzelmann et al. 1990). Many GPI proteins
become covalently linked to wall polysaccharide, and release from walls by treatment with HF/pyridine is a clue
that the protein had received a GPI (see GPI proteins; Yin
et al. 2005). The presence of a GPI is usually inferred from
the results of in silico analyses of a protein’s sequence.
Features of a likely GPI protein are a hydrophobic
N-terminal secretion signal and a C-terminal GPI signalanchor sequence that includes the amino acid residue, v, to
which the GPI will be amide-linked. Amino acids N-terminal
to v are designated v(2), and those C-terminal, are designated v(+). Proceeding from the C-terminal amino acid of
the unprocessed protein, the signal anchor signal consists of
(i) a variable stretch of hydrophobic amino acids capable
of spanning the membrane; (ii) a spacer region of moderately polar amino acids in positions v+3 to v+9 or more;
(iii) the v+2 residue, restricted mostly to G, A, or S; (iv) the
v residue itself, generally G, A, S, N, D, or C; and (v)
a stretch of some 10 amino acids that may form a flexible
linker region and whose relative polarity may influence
plasma membrane or wall localization of the protein
(Nuoffer et al. 1991, 1993; see Incorporation of GPI proteins
into the cell wall). Some C-terminal sequences may contain
alternative candidates for the v and v+2 amino acids. Ev-
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P. Orlean
idence that a predicted GPI attachment sequence is functional can be obtained by fusing the sequence to the C
terminus of a reporter protein and testing whether the reporter becomes expressed at the plasma membrane or in the
wall (Hamada et al. 1998a).
Assembly of the GPI precursor and its attachment to
protein in the ER: At least 21 proteins are involved in GPI
precursor synthesis and attachment to protein (Figure 5).
Eighteen are encoded by essential genes, and mutants lacking any of the other noncatalytic proteins or GPI sidebranching enzymes have severe growth defects. Additional
information about GPI synthetic proteins and phenotypes
associated with deficiencies in them is given in File S4.
Steps on the cytoplasmic face of ER membrane: GPI assembly
starts with transfer of GlcNAc from UDP-GlcNAc to PI. A
complex of at least six proteins (GPI-GnT) is involved, of
which Gpi3 is catalytic because it can be labeled with a photoactivatable UDP-GlcNAc analog (Kostova et al. 2000). GlcNAc
transfer occurs at the cytoplasmic face of the ER membrane
(Vidugiriene and Menon 1993; Watanabe et al. 1996; Tiede
et al. 2000). Essential Gpi2, Gpi15, and Gpi19 (Leidich et al.
1995; Yan et al. 2001; Newman et al. 2005), and nonessential
Gpi1 and Eri1 (Leidich and Orlean 1996; Sobering et al.
2004), are also required for GlcNAc-PI synthesis. ERI1 and
GPI1 null mutants are temperature-sensitive. The mammalian
orthologs of these proteins form a complex (Watanabe et al.
1998; Tiede et al. 2000; Eisenhaber et al. 2003; Murakami
et al. 2005), and the yeast proteins likely also do, for Eri1 and
Gpi19 associate with Gpi2 (Sobering et al. 2004; Newman
et al. 2005). Roles of the noncatalytic subunits are unclear.
Figure 5 Biosynthesis of the GPI precursor and its transfer to protein in the ER membrane. GlcNAc addition to PI and de-N-acetylation of GlcNAc-PI to
GlcN-PI occur at the cytoplasmic face of the ER membrane, and further additions to the GPI occur on the lumenal side of the ER membrane. Gpi18 and
Mcd4 need not act in a defined order. Man3- and Man4-GPIs either bearing Etn-P on Man-2 but not Man-1 or without any Etn-Ps (not shown) have also
been detected in radiolabeling experiments with certain late-stage GPI assembly mutants.
Ras2, in its GTP-bound form, can also join GPI-GnT
(Sobering et al. 2004). Membranes from ras2D cells have
8- to 10-fold higher in vitro GPI-GnT activity than wild-type
membranes, whereas membranes from cells expressing
constitutively active Ras2-Val19 have almost undetectable
activity. These findings indicate that Ras2-GTP is a negative
regulator of GPI-GnT, and, depending on the degree to
which the GTPase is activated, this could permit about
a 200-fold range of GlcNAc-PI synthetic activity.
Once formed, GlcNAc-PI is de-N-acetylated at the cytoplasmic face of the ER membrane by Gpi12 (Vidugiriene and
Menon 1993; Watanabe et al. 1999). GlcN-PI is the precursor
likely to be translocated to the lumenal side of the ER membrane. Its flipping has been reconstituted in rat liver microsomes, but the protein involved is unknown (Vishwakarma
and Menon 2005).
Lumenal steps in GPI assembly: The inositol ring in GlcN-PI
is next acylated on its 2-OH, making the glycolipid resistant
to cleavage by PI-specific phospholipase C. The reaction uses
acyl CoA as donor (Costello and Orlean 1992), and the acyl
chain transferred in vivo is likely palmitate. Gwt1, the acyltransferase, was identified in a screen for resistance to 1-[4butylbenzyl] isoquinoline, which inhibits surface expression
of GPI proteins (Tsukahara et al. 2003; Umemura et al.
2003). Disruption of GWT1 is lethal or leads to slow growth
and temperature sensitivity, depending on the strain background (Tsukahara et al. 2003). The inositol acyl chain may
prevent GPIs from being translocated back to the cytoplasmic side of the ER membrane (Sagane et al. 2011), be important for later steps in GPI assembly or transfer to protein,
or block the action of PI-specific phospholipases.
GlcN-(acyl)PI is next extended with four Man by GPIMan-T I-IV, and the first three Man are concurrently
modified with Etn-P by Etn-P-T I, II, and III. Dol-P-Man
donates the mannoses because the dpm1 mutant accumu-
lates GlcN-(acyl)PI (Orlean 1990). The first, a1,4-linked
Man (Man-1, Figure 5) is added by Gpi14 (Maeda et al.
2001), and two additional proteins are involved at this step.
One, Arv1, was originally implicated in ceramide and sterol
metabolism. ARV1 disruptants are impaired in ER-to-Golgi
transport of GPI proteins and accumulate GlcN-(acyl)PI
in vitro (but not in vivo), although they are not defective in
in vitro GPI-Man-T-I or Dpm1 activity or in N-glycosylation,
and it was proposed that Arv1 has a role in delivering GlcN(acyl)PI to Gpi14 (Kajiwara et al. 2008). The second protein,
Pbn1, was implicated at the GPI-Man-T-I step because
expression of both GPI14 and PBN1 is necessary to complement mammalian cell lines defective in Pbn1’s mammalian
homolog Pig-X, and co-expression of PIG-X and the gene for
Gpi14’s mammalian homolog, PIG-M, partially rescues the
lethality of gpi14D (Ashida et al. 2005; Kim et al. 2007).
Furthermore, Pbn1 depletion leads to accumulation of some
of the ER form of the GPI protein Gas1, a phenotype of GPI
precursor assembly mutants (Subramanian et al. 2006;
File S4).
Addition of a1,6-linked Man-2 requires catalytic Gpi18
(Fabre et al. 2005; Kang et al. 2005) and Pga1 (Sato et al.
2007), which form a complex (Sato et al. 2007). Gpi18deficient cells accumulate both a Man1-GPI with Etn-P esterified to its Man and an unmodified Man1-GPI, suggesting that
GPI-Man-T-II can use either as acceptor (Fabre et al. 2005;
Scarcelli et al. 2012).
Gpi10 and Smp3 successively add a1,2-linked Man-3 and
Man-4 (Canivenc-Gansel et al. 1998; Sütterlin et al. 1998;
Grimme et al. 2001). Smp3-dependent addition of Man-4 is
essential because addition of this residue precedes addition
of the Etn-P that subsequently becomes linked to protein
(Grimme et al. 2001).
As the GPI glycan is extended, Etn-P moieties are added
to the 2-OH of Man-1 and to the 6-OH of Man-2 and Man-3
S. cerevisiae Cell Wall
787
(Orlean 2009). The Etn-Ps likely originate from Ptd-Etn
(Menon and Stevens 1992; Imhof et al. 2000; File S4).
The Etn-P-T-I, II, and III transferases are Mcd4, Gpi7, and
Gpi13, respectively, which are 830- to 1100-amino-acid proteins predicted to have 10–14 transmembrane domains and
a large lumenal loop containing sequences characteristic
of the alkaline phosphatase superfamily that are important
for function (Benachour et al. 1999; Gaynor et al. 1999;
Galperin and Jedrzejas 2001; File S4). GPI-Etn-P-T-II and
III also require small, hydrophobic Gpi11 for activity. mcd4
mutants accumulate unmodified Man 1 and Man 2 -GPI
(Wiedman et al. 2007; Scarcelli et al. 2012), suggesting that
both structures can serve as Etn-P acceptors. From this, and
because Gpi18-depleted cells accumulate Etn-P-modified
Man1-GPI (Fabre et al. 2005), it seems that both Mcd4
and Gpi18 can use Man1-GPI as acceptor and then modify
the GPI that the other has acted on (Figure 5; File S4). Etn-P
transfer to Man-1 and GPI-dependent processing of Gas1 are
inhibited by the terpenoid lactone YW3548 (Sütterlin et al.
1997, 1998). The Etn-P on Man-1 may enhance the ability of
Gpi10 to add Man-3, promote export of GPI proteins from
the ER, and be necessary for remodeling of the lipid moiety
to ceramide (Zhu et al. 2006).
Gpi7 is the catalytic subunit of GPI-Etn-P-T-II, and GPI7
nulls, which are viable but temperature-sensitive, accumulate
a Man4-GPI with Etn-P on Man-1 and Man-3 (Benachour
et al. 1999). Essential Gpi11 was implicated at this step
because Gpi11-deficient cells have similar GPI precursor accumulation profiles to gpi7D (Taron et al. 2000). The Etn-P
on Man-2 enhances transfer of GPIs to protein, ER-to-Golgi
transport of GPI proteins, GPI lipid remodeling to ceramide,
transfer of GPI proteins to the wall, and targeting of certain
GPI-anchored proteins in daughter cells (Benachour et al.
1999; Toh-E and Oguchi 1999; Richard et al. 2002; Fujita
et al. 2004).
Gpi13 is the catalytic subunit of GPI-Etn-P-T-III. The major GPI accumulated upon Gpi13 depletion is a Man4-GPI
with a single Etn-P on Man-1 (Flury et al. 2000; Taron et al.
2000). Gpi11 is likely involved in the GPI-Etn-P-T-III reaction because a gpi11-Ts mutant also accumulates a Man4GPI with its Etn-P on Man-1 (K. Willis and P. Orlean, unpublished results), and human Gpi11 interacts with and
stabilizes human Gpi13 (Hong et al. 2000).
GPI transfer to protein: Man4-GPIs bearing three Etn-Ps
are transferred to proteins with a C-terminal GPI signalanchor sequence in a transamidation reaction in which the
amino group of the Etn-P on Man-3 acts as nucleophile. Five
essential membrane proteins are involved: Gaa1, Gab1,
Gpi8, Gpi16, and Gpi17 (Hamburger et al. 1995; Benghezal
et al. 1996; Fraering et al. 2001; Ohishi et al. 2000, 2001;
Hong et al. 2003; Grimme et al. 2004). Gpi18 is catalytic
because it resembles cysteine proteases and mutation of
predicted active site residues eliminates its function (Meyer
et al. 2000). The five transamidase subunits form a complex
itself consisting of two subcomplexes: one containing Gaa1,
Gpi8, and Gpi16, and the other, Gab1 and Gpi17 (Fraering
788
P. Orlean
et al. 2001; Grimme et al. 2004; Zhu et al. 2005). Roles for
the noncatalytic subunits include recognition of the peptide
and glycolipid substrates (Signorell and Menon 2009), and,
in the case of Gab1 and Gpi8, possible interactions with the
actin cytoskeleton (Grimme et al. 2004; File S4)
Remodeling of protein-bound GPIs: Following GPI transfer
to protein, both the anchor’s lipid and glycan remodeled
(Figure 6; Fujita and Kinoshita 2010). The earliest event,
which occurs in the ER, is removal of the inositol acyl moiety
by lipase-related Bst1 (Tanaka et al. 2004; Fujita et al.
2006a). Next, the sn-2 acyl chain of the diacylglycerol is
removed by the ER membrane protein Per1 to generate
a lyso-GPI (Fujita et al. 2006b), whereupon a C26:0 acyl
chain is transferred to the sn-2 position by Gup1 in the ER
membrane (Bosson et al. 2006). Modifications of the GPI
lipid by Bst1, Per1, and Gup1 are necessary for efficient
transport of GPI proteins from the ER to the Golgi (File S4).
Many GPIs are next remodeled by replacement of their
diacylglycerol with ceramide by Cwh43 (Martin-Yken et al.
2001; Ghugtyal et al. 2007; Umemura et al. 2007). Ceramide
remodeling requires prior action of Bst1, and, because per1D
and gup1D strains show defects in remodeling, the exchange
reaction likely takes place after the first three lipid modification
steps. The mechanism could involve a phospholipase-like reaction that replaces diphosphatidic acid with ceramide phosphate or diacylglycerol with ceramide (Ghugtyal et al. 2007;
Fujita and Kinoshita 2010). Ceramide remodeling is not
obligatory because certain GPI proteins, such as Gas1, reach
the plasma membrane with a diacylglycerol-based anchor
(Fankhauser et al. 1993). Moreover, ceramide remodeling
does not seem to be required for incorporation of GPI proteins into the wall (Ghugtyal et al. 2007).
Further GPI processing events may be the removal of the
Etn-P moieties from Man-2 and Man-1. This is inferred from
the fact that mammalian PGAP5, which removes the sidebranching Etn-P from Man-2 (Fujita et al. 2009), has two
homologs in yeast: ER-localized Ted1 and Cdc1. Export of
Gas1 is retarded in ted1D cells, and genetic interactions
connect TED1 and CDC1 with processing and export of
GPI proteins (Haass et al. 2007). Because Etn-P side chains
are important for ceramide remodeling, they are likely removed after Cwh43 has acted.
Finally, a fifth, a1,2- or a1,3-linked Man can be added to
Man-4 of protein-bound GPIs (Fankhauser et al. 1993).
This modification is made to 20–30% of GPI proteins and
occurs in the Golgi, but none of the many Golgi Man-T seems
to be involved (Sipos et al. 1995; Pittet and Conzelmann
2007). On reaching the plasma membrane, the GPIs on
many proteins become cross-linked to b1,6-glucan (see Incorporation of GPI proteins into the cell wall), and these GPICWP play structural or enzymatic roles in the wall (see Cell
Wall-Active and Nonenzymatic Surface Proteins and Their
Functions).
No individual GPI protein is essential in unstressed
wild-type cells, so the lethality of mutations blocking GPI
Figure 6 Remodeling of protein-bound GPIs. The inositol palmitoyl group and the sn-2 acyl chain are removed by Bst1 and Per1, respectively, and Gup1
transfers a C26:0 acyl chain to the sn-2 position. Cwh43 can replace diphosphatidic acid with ceramide phosphate (shown here) or diacylglycerol with
ceramide. Etn-P on Man-1 and Man-2 may be removed by Ted1 and Cdc1. Steps through Etn-P removal occur in the ER. An a1,2- or an a1,3-linked
Man is added to Man-4 in the Golgi by as yet unknown Man-T. At the plasma membrane, the GPI can be cleaved, possibly between GlcN and Man, and
the reducing end of the GPI remnant transferred to b1,6-glucan. Symbols are as used in Figure 1 and Figure 5.
anchoring may be due to the collective effects of retarding
ER exit and plasma membrane or wall anchorage of multiple
proteins. Consistent with this, temperature-sensitive GPI
anchoring mutants grown at semipermissive temperature
have aberrant morphologies and shed wall proteins into the
medium (Leidich and Orlean 1996; Vossen et al. 1997).
Sugar nucleotide transport
GDP-Man transport: Cytoplasmically generated GDP-Man
used by Golgi Man-T is transported into the Golgi lumen by
Vrg4/Vig4. GMP, generated from GDP formed in Man-T
reactions by GDPase activity, serves as antiporter. Vrg4/
Vig4 is essential, and vrg4 mutants are defective in mannosylation of N- and O-linked glycans and mannosyl inositolphosphoceramides (Dean et al. 1997; Abe et al. 1999).
Two homologous Golgi proteins, Gda1 and Ynd1, have
GDP-hydrolyzing activity. Gda1 has the highest activity toward GDP (Abeijon et al. 1989), and, consistent with GMP’s
role as antiporter, rates of in vitro GDP-Man import into
Golgi vesicles from gdaD cells are fivefold lower than those
of vesicles from wild-type cells (Berninsone et al. 1994).
Ynd1 is a broader specificity apyrase (Gao et al. 1999) that
has a partially overlapping function with Gda1, and both
Ynd1 and Gda1 are necessary for full elongation of N- and
O-linked glycans (Gao et al. 1999; File S5).
Other sugar nucleotide transport activities: Transport
activities for UDP-Glc, UDP-GlcNAc, and UDP-Gal also occur
in S. cerevisiae (Roy et al. 1998, 2000; Castro et al. 1999), and
there are eight more candidate transporters (Dean et al.
1997; Esther et al. 2008) whose functions are unclear. UDPGlc transport activity is present in the ER (Castro et al. 1999),
and one possible need for it might be for a glucosylation reaction at an early stage of b1,6-glucan assembly (see b1,
6-Glucan). Yea4 is an ER-localized UDP-GlcNAc transporter
whose deletion impacts chitin synthesis (Roy et al. 2000; File
S6). Hut1 is a candidate UDP-Gal transporter (Kainuma et al.
2001), although galactose has not been detected on S. cerevisiae glycans. Both Hut1 and Yea4 may have broader specificity and transport UDP-Glc (Esther et al. 2008).
Biosynthesis of Wall Components at the Plasma
Membrane
Chitin
S. cerevisiae has three chitin synthase activities—CS I, CS II,
and CS III—which require the catalytic proteins Chs1, Chs2,
and Chs3, respectively. The Chs proteins are active in the
plasma membrane although they originate from the rough
ER. The pathways for trafficking and activation of Chs2 and
Chs3 involve different sets of auxiliary proteins that ensure
the correct spatial and temporal localization of chitin synthesis during septation.
Septum formation: Factors determining the site at which
a bud will be formed, and the proteins that recruit and
organize the participants in septum formation, including
septins and an actin–myosin contractile ring, are reviewed
by Cabib et al. (2001), Cabib (2004), Roncero and Sanchez
(2010), and Bi and Park (2012). Two chitin-containing
structures are made during bud emergence and septum formation (Figure 7). The first is a ring deposited in the wall
around the base of the emerging bud. This chitin is formed
by Chs3 (Shaw et al. 1991), and, after cell separation,
remains on the mother cell as a component of the bud scar.
Upon completion of mitosis, the primary septum is formed
by centripetal synthesis of chitin by Chs2 in the neck region
between mother cell and bud (Shaw et al. 1991). Upon
closure, the septum separates the plasma membranes of
S. cerevisiae Cell Wall
789
the two cells, accomplishing cytokinesis. In budding wildtype cells, the primary septum is thickened on both sides by
deposition of a secondary septum that normally contains
chitin, b1,3-glucan, b1,6-glucan, and covalently cross-linked
mannoprotein (Rolli et al. 2009), resulting in a three-layered
structure (Shaw et al. 1991).
Chs2 and Chs3 have important roles in septation and
cytokinesis although in the absence of Chs2 or Chs3, or indeed of all three chitin synthases, cytokinesis can still take
place. In chs2D mutants, the primary septum is missing, and
a thick, amorphous septum is formed that contains chitin
made by Chs3 (Shaw et al. 1991; Cabib and Schmidt
2003). chs3D mutants form a three-layered septum, but
the neck region between mother cell and bud is elongated
(Shaw et al. 1991). chs2D chs3D and chs1D chs2D chs3D
strains grow very slowly on osmotically supported medium
(Sanz et al. 2004; Schmidt 2004; File S6). The triple
mutants, however, acquired a suppressor mutation that
eliminated the need for osmotic support and conferred
a growth rate as fast as that of a chs2D mutant although
over a third of suppressed and unsuppressed cells in a culture were dead (Schmidt 2004).
For mother and daughter cells to separate, septal
material must be degraded, a process that results from
secretion of chitinase Cts1 (Kuranda and Robbins 1991),
endo-b1,3-glucanases Eng1/Dse4 and Scw11 (Cappellaro
et al. 1998; Colman-Lerner et al. 2001; Baladron et al.
2002; see Known and predicted enzymes), and possibly additional activities from the daughter cell’s side of the septum.
Daughter cell-specific expression of these enzymes is under
the control of the transcription factor Ace2 (Colman-Lerner
et al. 2001).
Chitin synthase biochemistry: Chs1, Chs2, and Chs3 use
UDP-GlcNAc as donor and are members of GT family 2 of
processive inverting glycosyltransferases, which includes
hyaluronate and cellulose synthases. Yeast’s chitin synthases
are predicted to have three to five transmembrane helices
toward their C termini, and Chs3 likely has two more transmembrane domains nearer its N terminus (Jimenez et al.
2010; Merzendorfer 2011). Amino acid residues important
for catalysis lie in a large cytoplasmic domain containing
the signature sequences QXXEY, EDRXL, and QXRRW
(Nagahashi et al. 1995; Saxena et al. 1995; Cos et al. 1998;
Yabe et al. 1998; Ruiz-Herrera et al. 2002; Merzendorfer
2011). An additional motif, (S/T)WG(X)T(R/K), predicted
to be extracellularly oriented (Merzendorfer 2011), lies near
the protein’s C terminus (Cos et al. 1998; Merzendorfer 2011).
The molecular mechanism of chitin synthesis is not yet
clear. By analogy with bacterial NodC, which synthesizes
chito-oligosaccharides, and with nonfungal chitin synthases,
chain extension would be at the nonreducing end (Kamst
et al. 1999; Imai et al. 2003). This topic, and the issue of
how the synthases overcome the steric challenge that each
sugar in a b1,4-linked polymer is rotated by 180 relative
to its neighbor, are discussed further in File S6.
790
P. Orlean
Figure 7 Roles of chitin synthases II and III in chitin deposition during
budding growth. (A) Chitin synthase III synthesizes a chitin ring (blue)
around the base of the emerging bud. (B) The plasma membrane invaginates and chitin synthase II synthesizes the primary septum (red). No
chitin is made in the lateral walls of the bud. (C) Secondary septa (green)
are laid down on the mother- and daughter-cell sides of the primary
septum, and chitin synthase III starts synthesizing lateral wall chitin in
the bud (blue). (D) After cell separation, the bud scar (which is formed
from the chitin ring made by Chs3), most of the primary septum made by
Chs2, as well as secondary septal material deposited on the mother cell
side, remain on the mother cell. The birth scar on the daughter cell
contains residual chitin from the primary septum as well as secondary
septal material. (E and F) Chitinase digestion of the primary septum from
the daughter-cell side facilitates cell separation, and lateral wall chitin
synthesis continues as the daughter cell grows. Figure is adapted from
Cabib and Duran (2005).
Chitin made in vitro by CS I or CS III contains, on average, 115–170 GlcNAc residues (Kang et al. 1984; Orlean
1987). Chitin synthases presumably make chitin chains with
a range of lengths, and the range would be predicted to shift
to shorter chains as UDP-GlcNAc concentration drops below
Km, resulting in lowered rates of chain extension. Indeed,
purified Chs1 and membranes from cells overexpressing
Chs2 make chito-oligosaccharides at low substrate concentrations (Kang et al. 1984; Yabe et al. 1998). Chitin made
in vivo is polydisperse (Cabib and Duran 2005), and increased chitin chain lengths are seen in fks1D and gas1D
mutants and CFW-treated cells, which mount the chitin
stress response, whereas shorter chains were made in
a strain expressing a Chs4 variant with lower in vitro CS
III activity (Grabinska et al. 2007). However, GlcN treatment, which stimulates chitin synthesis in vivo (Bulik et al.
2003; see Sugar nucleotides), had little effect on polymer
chain length (Grabinska et al. 2007).
S. cerevisiae’s three chitin synthases are all stimulated up
to a few fold in vitro by high concentrations of GlcNAc
(Sburlati and Cabib 1986; Orlean 1987). Possible explanations are that GlcNAc serves as a primer or allosteric activator in the chitin synthase reaction (see File S6).
S. cerevisiae’s chitin synthases and auxiliary proteins:
Chitin synthase I: Most, if not all, Chs1 activity is detectable
in vitro only after pretreatment of membranes or extensively
purified Chs1 with trypsin (Duran and Cabib 1978; Kang
et al. 1984; Orlean 1987). Proteolytically activated Chs1
has the highest in vitro activity of the chitin synthases
assayed in membranes from wild-type cells (Sburlati and
Cabib 1986; Orlean 1987), although Chs1 does not contribute measurably to chitin synthesis in vivo, even in the absence of Chs2 and Chs3 (Shaw et al. 1991). Although trypsin
activation may mimic the effect of an endogenous activating
protease, neither such an activator, nor an active, processed
form of Chs1, have been identified.
Levels of protease-elicited Chs1 activity are the same in
membranes from logarithmically growing and stationaryphase cells (Orlean 1987), and levels of Chs1 show little
change during the cell division cycle (Ziman et al. 1996).
CHS1 transcription and in vitro CS I activity increase in response to mating factors, but elevated in vitro activity is
detectable only after trypsin activation (Schekman and
Brawley 1979; Orlean 1987; Appeltauer and Achstetter
1989). However, Chs1 does not contribute to pheromoneinduced chitin synthesis (Orlean 1987).
chs1D cultures contain the occasional lysed bud, a phenotype
more pronounced in acidic medium but partially alleviated
when Cts1 chitinase is also deleted (Cabib et al. 1989). Two
explanations, which are not mutually exclusive, are that Chs1
may repair wall damage due to overdigestion of chitin by Cts1
or that Chs1 participates in septum synthesis and makes chitin
during growth in acidic medium (Cabib et al. 1989; Bulawa
1993). Chs1 promotes wall association of at least one protein
because small amounts of the GPI protein Gas1 are released
into the medium from chs1D cells (Rolli et al. 2009).
Although the contribution of Chs1 to chitin synthesis is
small, a wider role for the protein emerged from an analysis
of the networks of genes that interact synthetically with
CHS1 and CHS3 (Lesage et al. 2005). Most of the 57 genes
in the CHS1 interaction network fell into two sets. One set
contained genes that, when mutated, impact cell integrity or
that themselves interact with genes involved in b1,3-glucan
synthesis, indicating a role for Chs1 in buffering the wall
against changes impacting its robustness. The other set contained genes involved in budding and in endocytic protein
recycling, which in turn may impact Chs2 function, suggesting that Chs1 also buffers against deficiencies in Chs2. The
CHS1-interacting genes were mostly distinct from the genes
in the network that impacts Chs3 function, and, moreover,
mutations in CHS1 itself or in the genes in the CHS1 interaction set do not trigger the Chs3-dependent chitin stress
response. Chs1 and Chs3 therefore have distinct functions
and one does not buffer against defects in the other (Lesage
et al. 2005).
Chitin synthase II and proteins impacting its localization
and activity: Chs2 makes no more than 5% of the chitin in
budding cells. Activity of endogenous Chs2 is detectable
only in membranes from growing cells and can be stimulated by treatment with trypsin (Sburlati and Cabib 1986)
although, in some studies, membrane preparations as well
as partially purified Chs2 have significant in vitro activity
without prior trypsin treatment, raising the possibility that
full-size Chs2 makes chitin (Uchida et al. 1996; Oh et al.
2012). A soluble fraction from growing yeast cells, which
stimulates Chs2 activity two- to fourfold but which must itself
be pretreated with trypsin, has been described (MartínezRucobo et al. 2009). An endogenously activated, processed
form of Chs2 has not been identified (File S6).
Levels of CHS2 expression and localization of the protein
are coordinated with synthesis of the primary septum (Figure 8). CHS2 message levels peak just prior to primary septum formation at the G2/M phase (Pammer et al. 1992; Cho
et al. 1998; Spellman et al. 1998), and levels of Chs2 and CS
II activity then peak as the primary septum is made (Pammer
et al. 1992; Choi et al. 1994a; Chuang and Schekman 1996).
Upon completion of cytokinesis, levels of Chs2 and its message drop, indicating that both turn over rapidly.
Temporal and spatial localization of Chs2 is impacted at
at least two stages by protein kinases. Chs2 is synthesized in
the ER during metaphase, but its release from the ER is
coordinated with exit of the cell from mitosis and triggered
upon inactivation of mitotic kinase by Sic1 (Zhang et al.
2006). The mitotic kinase Cdk1 likely acts directly on
Chs2, which contains four CDK1 phosphorylation sites near
its N terminus, because mutation of the target Ser residues
to Glu leads to retention of Chs2 in the ER, whereas changing the serines to Ala leads to constitutive release of the
mutant Chs2 even in the presence of high Cdk1 activity
(Teh et al. 2009). Timed release of Chs2 from the ER after
chromosome separation and exit of the cells from mitosis is
triggered by dephosphorylation of the Cdk1 sites by the
Cdc14 phosphatase, the terminal component of the mitotic
exit network (MEN) cascade (Chin et al. 2012).
Exit of Chs2 from the ER and its delivery to the plasma
membrane at the mother cell–bud junction is effected by
COPII vesicles (Chuang and Schekman 1996; VerPlank
and Li 2005; Zhang et al. 2006). Localization of Chs2 at
the bud neck, correct formation of the primary septum,
and removal of Chs2 at the end of cytokinesis depend on
phosphorylation of Chs2 by the mitotic exit kinase Dbf2, also
a component of MEN (Oh et al. 2012). Inn1 and Cyk3,
whose localization to the division site is also regulated by
MEN, are also involved in activation of Chs2 for primary
septum formation (Nishihama et al. 2009; Meitinger et al.
2010; Oh et al. 2012). Overexpression of CYK3 leads to increased deposition of chitin at the division site in chs1D
chs3D cells, where Chs2 is the sole chitin synthase
(Oh et al. 2012). Cyk3 has a transglutaminase-like domain
(Nishihama et al. 2009), but the nature of Cyk3’s effect on
Chs2 is unclear and Inn1’s role in Chs2 activation is unknown.
S. cerevisiae Cell Wall
791
Additional phosphorylation sites are present in Chs2’s N-terminal domain (Martínez-Rucobo et al. 2009), but their roles
are unclear.
Chs2 resides at the site of primary septum formation for
only 7–8 min (Roh et al. 2002a; Zhang et al. 2006). The
protein is degraded upon endocytosis and delivery to the
vacuole (Chuang and Schekman 1996; Schmidt et al.
2002; VerPlank and Li 2005), and optimal endocytic turnover
of Chs2 requires components of the endosomal sorting complexes required for transport (ESCRT) pathway (McMurray
et al. 2011).
Chitin synthase III and proteins impacting its localization
and activity: Chitin synthase III is responsible for the
synthesis of .90% of the chitin in unstressed vegetative
cells, for the additional chitin made in the chitin stress response and in response to mating pheromones, and for the
synthesis of the chitin that is de-N-acetylated to chitosan
during ascospore wall formation. Cells deficient in CS III
activity are resistant to CFW (Roncero et al. 1988). Chs3 is
the transferase, but its function depends on its regulated
transport from the ER to the plasma membrane, its removal
from the plasma membrane and sequestration in intracellular vesicles called chitosomes, and its remobilization from
chitosomes to the plasma membrane. A number of proteins
are required for regulated Chs3 trafficking and for enzyme
activity (Bulawa 1993; Trilla et al. 1999; Roncero 2002).
CS III [referred to as chitin synthase II by Orlean (1987)]
is the major, if not only, activity detected in membrane fractions from logarithmically growing wild-type cells without
prior treatment with trypsin and is trypsin sensitive (Orlean
1987). CS III activity determined in this way is presumably
either due to constitutively active Chs3 or to an endogenously
activated form of the protein. A pool of trypsin-activatable
CS III was detected in detergent-treated membranes from
chs1D chs2D cells or from cells lacking Chs4, an activator
of CS III (Choi et al. 1994b; Trilla et al. 1997). The latter
finding, together with the observation that overexpression of
Chs4 lowers the extent to which trypsin activates CS III, suggested that trypsin treatment might mimic Chs4-dependent
processing of Chs3. However, because no endogenously processed forms of Chs3 have been detected (Santos and
Snyder 1997; Cos et al. 1998), and because Chs4 does not
resemble any protease, the apparent zymogenicity of Chs3
in chs4D may be an artifact (Reyes et al. 2007).
Levels of CHS3 mRNA and Chs3 vary little during the
budding cycle (Choi et al. 1994a; Chuang and Schekman
1996; Cos et al. 1998), indicating that CS III is regulated at
the post-translational level. Sequences involved include the
C-terminal extracellular region containing the motif (S/T)
WG(X)T(R/K), which is required for in vitro CS III activity
and chitin synthesis in vivo (Cos et al. 1998).
A number of proteins interact with Chs3 as it transits the
secretory pathway (Figure 9). Chs3 is palmitoylated in the
ER by Pfa4 (Lam et al. 2006; Montoro et al. 2011). pfa4D
mutants are CFW-resistant and accumulate Chs3 in the ER,
indicating a role for palmitoylation in Chs3 export (Lam et al.
792
P. Orlean
Figure 8 Trafficking and regulation of Chs2. Cell cycle-regulated expression of CHS2 peaks at the G2-M phase transition, and Chs2 is synthesized
at the ER. Phosphorylation of Chs2 by Cdk1 retains Chs2 in the ER. Upon
chromosomal separation, Cdc14-dependent dephosphorylation of Chs2
allows release of the protein from the ER and its transit to the mother
cell–bud junction. Inn1 and Cyk3, localized at the division site, are involved in Chs2 activation. After primary septum formation is complete,
Chs2 is endocytosed and degraded. Localization, function, and subsequent removal of Chs2 when the primary septum is complete depend
on phosphorylation by Dbf2. Figure is adapted from Lesage and Bussey
(2006).
2006). Chs3 has two palmitoylation sites in a cytoplasmic domain N-terminal to the proposed catalytic residues (Meissner
et al. 2010). Exit of Chs3 from the ER also requires Chs7, an
ER chaperone with six or seven transmembrane domains
(Trilla et al. 1999) that interacts with Chs3. The effects of
CHS7 deletion on chitin levels and CSIII activity are almost
as severe as those of CHS3 deletion (Trilla et al. 1999). Chs3
aggregates in the ER in chs7D cells (Lam et al. 2006), and
Chs7 is a limiting factor in export of Chs3 because simultaneous overexpression of CHS3 and CHS7 leads to elevated
CSIII activity, whereas overexpression of CHS3 alone does not.
Neither Pfa4 nor Chs7 is required for exit of Chs1 and Chs2
from the ER (Trilla et al. 1999; Lam et al. 2006). ER-membrane
proteins Rcr1 and Yea4 also impact Chs3-dependent chitin
synthesis in ways that are unclear (File S6).
Transport of new Chs3 from the trans-Golgi to the plasma
membrane, as well as Chs3 cycling from chitosomes to the
plasma membrane, requires the peripheral Golgi membrane
proteins Chs5 and Chs6 (Santos and Snyder 1997; Santos
Figure 9 Overview of Chs3 trafficking. Chs3, synthesized in the ER,
requires palmitoylation by Pfa4 and association with Chs7 to exit the
ER. In the trans-Golgi, Chs3 association with exomer components Chs5
and Chs6 facilitates incorporation of Chs3 into secretory vesicles for delivery to the plasma membrane at the site of chitin ring formation. Localization and activation of Chs3 depends on association with Chs4, whose
association with the septin ring is mediated in turn via an interaction with
Bni4. In cells with medium-sized buds, Chs3 is retrieved from the plasma
membrane and sequestered in chitosomes in an endocytic process
depending on End4 and later recruited back to the neck region in
a Chs6-dependent manner. During the cell wall stress response, Rho1
and Pkc1 trigger mobilization of Chs3 from chitosomes to the plasma
membrane for synthesis of extra chitin in the lateral wall. Figure is adapted from Lesage and Bussey (2006).
et al. 1997; Ziman et al. 1998). Chs6 and its homologs Bch1,
Bch2, and Bud7, referred to as Chs5-Arf1-binding proteins,
join with Chs5 to form exomer complexes that transiently
bind Chs3 to promote its incorporation into secretory
vesicles (Sanchatjate and Schekman 2006; Trautwein et al.
2006; Wang et al. 2006). Although Chs5 and Chs6 act in
a complex, the two have different impacts on Chs3 activity
and transport. chs5D and chs6D mutants make 25 and 10%
of wild-type amounts of chitin, respectively, but whereas
chs5D membranes lack in vitro CS III activity, this activity
is normal in chs6D membranes (Bulawa et al. 1993; Santos
et al. 1997). This may be because, in chs5D cells, Chs3 accumulates in late Golgi vesicles (Santos and Snyder 1997),
whereas, in chs6D mutants, it collects in chitosomes, where
it may encounter a chitosomal activator (Ziman et al. 1998).
Exomer has a role in the transport of the chitin-b1,3-glucan
cross-linker Crh2 to the cell surface. Cotransport of Chs3
and Crh2 would ensure colocalization of these proteins for
efficient cross-linking of chitin to b1,3-glucan.
At the plasma membrane, Chs4 (Csd4/Skt5) interacts
with Chs3 (DeMarini et al. 1997; Ono et al. 2000; Meissner
et al. 2010) and has two roles apparently specific to Chs3.
chs4D mutants lack in vitro CS III activity and make very
little chitin (Bulawa 1993; Trilla et al. 1997). Overexpression of CHS4, but not CHS3, raises in vitro CS III activity
(Bulawa 1993; Trilla et al. 1997; Ono et al. 2000) as well as
levels of Chs3 in the plasma membrane (Reyes et al. 2007),
suggesting that Chs4 is an activator of CS III. Stimulation of
CS III by Chs4 requires a region of Chs4 to bind Chs3 because the ability of truncated forms of Chs4 to elicit CS III
activity correlates with the ability of Chs4 fragments to interact with Chs3 in a two-hybrid analysis (Ono et al. 2000;
Meissner et al. 2010). Chs4 has a C-terminal farnesylation
site (Bulawa et al. 1993; Trilla et al. 1997; Grabinska et al.
2007) whose roles are discussed in File S6.
Chs4 not only activates Chs3, but also mediates Chs3
localization on the mother cell’s plasma membrane at the
site of formation of the chitin ring prior to bud emergence.
There, it interacts with the scaffold protein Bni4, which
in turn associates with the septins (DeMarini et al. 1997;
Kozubowski et al. 2003; Sanz et al. 2004). Absence of Bni4
leads to mislocalized deposition of chitin (DeMarini et al.
1997; Kozubowski et al. 2003; Sanz et al. 2004), and Chs4
is absent from the base of buds in small-budded cells (Sanz
et al. 2004). In contrast to chs4D mutants, chitin synthesis
and CS III activity are not dramatically affected in bni4D cells,
suggesting that Bni4 is not required for CS III activity per se
(Sanz et al. 2004).
Chs3 and Chs4 are associated with the plasma membrane
just before formation of the chitin ring at the site of bud
emergence and reside there in a ring at the base of the
bud in many cells with small buds, then become scarcely
detectable in cells with medium-sized buds, only to reappear, in a Bni4-independent manner, at both sides of the
neck in cells with large buds prior to cytokinesis (Chuang
and Schekman 1996; DeMarini et al. 1997; Santos and
Snyder 1997; Kozubowski et al. 2003; Sanz et al. 2004).
In between, Chs3 is retrieved from the membrane to chitosomes in an endocytic process dependent on End4/Sla2
(Chuang and Schekman 1996; Ziman et al. 1996, 1998),
but is recruited back to the plasma membrane in a Chs6dependent manner (Ziman et al. 1998; Wang et al. 2006).
Chs3’s itinerary is consistent with the overall order of events
in yeast cytokinesis.
Chitin synthesis in response to cell wall stress: Cells with
mutations affecting the formation of b-glucan, mannan,
O-linked glycans, and GPI anchors respond by depositing
additional chitin—as much as 10 times more than in wildtype cells—in their lateral walls in apparent compensation
for compromised cell integrity (Gentzsch and Tanner 1996;
Kapteyn et al. 1997, 1999a; Popolo et al. 1997; Dallies et al.
S. cerevisiae Cell Wall
793
1998; Osmond et al. 1999; Garcia-Rodriguez et al. 2000;
Valdivieso et al. 2000; Carotti et al. 2002; Lagorce et al.
2002; Magnelli et al. 2002; Sobering et al. 2004; Lesage
et al. 2005). This chitin stress response, which is accompanied by increased precursor supply (see Precursors and Carrier Lipids), requires Chs3 and is dependent on Chs4, -5, -6,
and -7 in gas1D cells (Valdivieso et al. 2000; Carotti et al.
2002). The response does not involve upregulation of the
CHS genes, but, rather, an altered distribution of Chs3,
which was seen in the plasma membrane of buds of gas1D
and fks1D cells, and Chs4 was also delocalized (GarciaRodriguez et al. 2000; Valdivieso et al. 2000; Carotti et al.
2002; Valdivia and Schekman 2003). Interestingly, gas1D
suppressed the lysed bud phenotype of chs1D, suggesting
that the chitin stress response also repaired weakened bud
walls (Valdivieso et al. 2000). The Chs3 making the stress
response originates from chitosomes, and its translocation to
the plasma membrane is regulated by Rho1 and Pkc1, which
act early in the CWI pathway that triggers the chitin stress
response (Valdivia and Schekman 2003).
Chitin synthase III in mating and ascospore wall formation: Chitin synthase III is responsible for the extra chitin
made in response to mating pheromones and for formation
of the chitosan of ascospore walls. MATa cells treated with
a-factor show a three- to fourfold increase in chitin, which is
laid down diffusely in the shmoo (Schekman and Brawley
1979). Chs3 is necessary because no extra chitin is made in
pheromone-treated chs3D cells, and the response is either
abolished or much smaller in chs5D, chs6D, and chs4D cells,
indicating that the machinery for trafficking and activation
of Chs3 is required (Orlean 1987; Roncero et al. 1988;
Bulawa 1993; Santos and Snyder 1997; Bulik et al. 2003).
Consistent with its role in chitin deposition, Chs3 is localized
at the periphery of the mating projection, and it remains
there because it is not subject to endocytic turnover as it is
in budding cells (Santos and Snyder 1997; Sacristan et al.
2012). Although the extra chitin synthesis in response to
a-factor is presumably driven by the increased amount
of UDP-GlcNAc made during the pheromone response
(Orlean et al. 1985; Bulik et al. 2003), the mechanism behind pheromone-stimulated chitin synthesis by Chs3 is unclear.
Levels of Chs3 increase sixfold upon a-factor treatment (Cos
et al. 1998), but neither CHS3 transcription nor levels of
chitin synthase III activity are elevated (Orlean 1987; Choi
et al. 1994a). Factors that might limit total Chs3 activity
might include prevention of the mobilization of the protein
to the plasma membrane in shmoos or interference with
interactions between Chs3 and regulatory proteins (Choi
et al. 1994a).
The chitosan of the ascospore wall is initially synthesized
as chitin by Chs3 (Pammer et al. 1992) and is then de-Nacetylated by chitin deacetylases Cda1 and Cda2, of which Cda2
has the dominant role (Mishra et al. 1997; Christodoulidou
et al. 1999). From the sporulation defects in mutants in proteins
involved in Chs3 trafficking in vegetative cells, Chs6 and Chs7,
794
P. Orlean
but not Chs5, have as-yet-undefined roles in ascospore maturation (Santos et al. 1997; Trilla et al. 1999). A Chs4 homolog,
Shc1, has a regulatory role in chitosan synthesis (Sanz et al.
2002; File S6). Ascospore wall structure and assembly are
reviewed by Neiman (2011).
b 1,3-Glucan
De novo b1,3-glucan synthetic activity is associated with
members of the Fks family, of which Fks1 and Fks2 require
the soluble Rho1 GTPase as a regulatory subunit. In vitro
activity is membrane-associated, uses UDP-Glc as donor, is
stimulated by GTP via Rho1, and yields a product with a
chain length of 60–80 glucoses (Shematek et al. 1980; Kang
and Cabib 1986; Drgonová et al. 1996; Mazur and Baginsky
1996; Qadota et al. 1996). In vitro b1,3-glucan synthase
activity is inhibited by acylated cyclic hexapeptides of the
echinocandin group and by papulocandins, acylated derivatives of b1,4-galactosylglucose (Debono and Gordee 1994;
Georgopapadakou and Tkacz 1995).
Fks family of b1,3-glucan synthases: Fks1 (Cwh53/Etg1/
Gsc1/Pbr1), Fks2, and Fks3 are in GT family 48, which also
contains proteins implicated in callose sythesis in plants
(Verma and Hong 2001). Fks1 has an N-terminal cytoplasmic domain that is followed by 6 transmembrane helices,
a large cytoplasmic domain, and then 10 transmembrane
helices (Inoue et al. 1995; Mazur et al. 1995; Qadota et al.
1996; Dijkgraaf et al. 2002; Okada et al. 2010). Three functional domains, mutations in which separately affect in vivo
and in vitro b1,3 glucan synthetic activity, as well as cell
polarity and endocytosis, have been distinguished (Okada
et al. 2010; File S7). The phenotypes of fks1 mutants may
in part reflect the involvement of the protein in processes
other than b1,3-glucan biosynthesis. For example, mutations in both FKS1 and FKS2 result in lowered b1,6-glucan
synthesis (Dijkgraaf et al. 2002). Fks1 is localized to the
plasma membrane at sites of polarized growth and cell wall
remodeling throughout the cell cycle, and this localization
coincides with that of actin patches (Qadota et al. 1996;
Dijkgraaf et al. 2002; Utsugi et al. 2002). Fks1 transits the
secretory pathway because it accumulates intracellularly in
vesicular transport mutants and its activity is sensitive to
phytosphingosine levels in the ER (El-Sherbeini and Clemas
1995; Abe et al. 2001; File S7).
Roles of the Fks proteins in b1,3-glucan synthesis: The Fks
proteins show a degree of specialization. Deletion of FKS1
leads to slow growth, a 75% reduction in b1,3-glucan, and
low in vitro b-1,3-glucan synthase activity, whereas the
in vitro activity of fks2D membranes is nearly that of wildtype membranes and the disruptants have no defect in vegetative growth (Inoue et al. 1995; Mazur et al. 1995). Although this suggests that Fks1 is the major contributor to
b1,3-glucan synthesis in budding cells, fks1D fks2D null
mutants are inviable, indicating that Fks1 and Fks2 have
overlapping functions (Inoue et al. 1995; Mazur et al.
1995). Consistent with this, overexpression of either FKS1
or FKS2 can partially correct the defects caused by deleting
the other of the two genes (Mazur et al. 1995; Dijkgraaf
et al. 2002), and, furthermore, the two proteins colocalize
in sites of polarized growth in budding cells, although Fks1
is the most abundant (Dijkgraaf et al. 2002).
FKS1 and FKS2 show different expression patterns. FKS1
is expressed during budding growth and transcript levels
peak in the late G1 and early S phases (Mazur et al. 1995;
Ram et al. 1995; Lesage and Bussey 2006). FKS2 mRNA, in
contrast, cannot be detected in budding cultures grown in
glucose, but appears when glucose becomes depleted; when
cells are grown on acetate, glycerol, or galactose; when cells
are treated with a-factor or Ca2+; in fks1 mutants and
mutants defective in the synthesis of other wall polymers;
and when cells are stressed by shift to high temperature
(Mazur et al. 1995; Zhao et al. 1998; Lesage and Bussey
2006). Induction of FKS2 is mediated via the PKC CWI
and calcineurin pathways (Mazur et al. 1995; Ram et al.
1995; Zhao et al. 1998; Lagorce et al. 2003).
Fks2 is important in sporulation because fks2D fks2D diploids have a severe defect in this process (Mazur et al. 1995;
Huang et al. 2005; Ishihara et al. 2007). Homozygous fks3D
fks3D diploids also form abnormal spores, indicating a role
for Fks3 in ascopore wall formation, although Fks3’s role in
sporulation does not overlap with Fks2’s. It was proposed
that Fks2 is primarily responsible for synthesis of b1,3-glucan
in the ascospore wall and that Fks3, rather than functioning
as a synthase, modulates glucan synthesis during ascospore
wall formation (Ishihara et al. 2007; File S7).
After their export through the plasma membrane, b1,3glucan chains can be cross-linked to chitin by Crh1 and Crh2
(Cabib 2009), and the polymer can be extended through the
action of Gas1 family b1,3-glucanosyltransferases (Mouyna
et al. 2000), and side-branching b1,6-linked glucoses as well
as PIR proteins may be attached (Ecker et al. 2006; see Incorporation of PIR proteins into the cell wall and Exg1, Exg2,
and Ssg/Spr1 exo-b1,3-glucanases).
Deficiencies in Fks1 are compensated for by Chs3-dependent
chitin synthesis (Garcia-Rodriguez et al. 2000; Valdivieso
et al. 2000; Carotti et al. 2002), and fks1D shows synthetic
interactions with chs3D, chs4D, chs5D, chs6D, and chs7D
(Osmond et al. 1999; Lesage et al. 2004), but correct synthesis of other wall constituents is also necessary when
b1,3-glucan synthesis is compromised (Lesage et al.
2004). Analyses of the genome-wide responses to FKS1
deletion revealed upregulation of a “cell wall compensatory cluster” of 79 coregulated genes whose products include a range of proteins involved in wall synthesis and
remodeling (Terashima et al. 2000; Lagorce et al. 2003).
An overlapping set of genes, whose products function in
the biosynthesis of chitin, b1,6-glucan, and mannan, as
well as in the function of the secretory pathway and in
maintenance of cell polarity, was identified in an analysis
of the synthetic genetic interactions of fks1D (Lesage et al.
2004). This study showed that FKS2 made interactions
only with FKS1 and that FKS3 made no interactions, consistent with differential expression of FKS2 and FKS3 (Lesage
et al. 2004).
Rho1 GTPase, a regulatory subunit of b1,3-glucan synthase:
The essential Rho1 GTPase, which activates Pkc1 in the CWI
pathway and is required for cell cycle progression and polarization of growth (Drgonová et al. 1999; Levin 2011), has
a distinct role as a regulatory subunit of b1,3-glucan synthetic complexes containing Fks proteins. Evidence for this is
that (i) Fks1 and Rho1 colocalize and coimmunoprecipitate,
(ii) membranes from a temperature-sensitive rho1 mutant
have a thermolabile b1,3-glucan synthase activity that can
be corrected by adding back purified Rho1, (iii) membranes
from cells expressing a consitutively active rho1 allele have
GTP-independent b1,3-glucan synthase activity, and (iv) inactivation of Rho1 by ADP ribosylation eliminates the
in vitro b1,3-glucan synthase activity of membranes from
fks1D and fks2D strains (Drgonová et al. 1996; Mazur and
Baginsky 1996; Qadota et al. 1996). Moreover, there are
rho1 mutations that affect regulation of b1,3-glucan synthesis, but not other Rho1 functions, and the amino acids affected are different from those whose mutation causes cell
cycle and polarization defects (Saka et al. 2001; Roh et al.
2002b). The amino acid changes in the b1,3-glucan synthesisspecific rho1 mutants might impact binding to Fks proteins,
but the interacting domains on the regulatory and catalytic
subunits have not been defined. The Rho1-Fks interaction at
the cytoplasmic face of the plasma membrane, as well as
activation of b1,3-glucan synthesis, requires Rho1 to be geranylgeranylated at its C terminus (Inoue et al. 1999).
b1,6-Glucan
Mutations in genes with products localized along the secretory
pathway impact formation of b1,6-glucan (Shahinian and
Bussey 2000; Lesage and Bussey 2006), but the biochemistry
of b1,6-glucan synthesis is unclear. In vitro synthesis of b1,
6-glucan is hard to detect, and no fungal enzyme has yet been
shown to catalyze formation of a b1,6-glucosidic linkage using
UDP-Glc as donor, although the linkage can be generated by
the Bgl2 protein in a transglycosylation reaction (Goldman et al.
1995). Synthesis of b1,6-glucan is normal in alg5D mutants,
indicating that Dol-P-Glc is not involved in formation of this
polymer (Shahinian et al. 1998; Aimanianda et al. 2009).
In vitro synthesis of b1,6-glucan
Because b1,6-glucan is a linear polymer with side branches
on average every fifth Glc (see b-glucans), it could be generated by a processive, UDP-Glc-dependent b1,6-glucan synthase and then branched or by assembly of shorter repeat
units, whose glucoses originate from UDP-Glc. Detection of
UDP-Glc-dependent formation of b1,6-glucan is complicated
by the fact that UDP-Glc is also the donor in the synthesis of
b1,3-glucan, glycogen, and glucolipids.
S. cerevisiae Cell Wall
795
Two assays of the formation of b1,6-glucan using UDPGlc as donor have been described. In the first, formation of
b1,6-glucanase-sensitive polymer by membranes was detected by dot-blot assay using an anti-b1,6 glucan antibody
(Vink et al. 2004). The reaction was distinguished from
b1,3-glucan synthase because membranes from kre5 mutants, which make little b1,6-glucan but have normal
b1,3-glucan synthetic capability, made little b1,6-glucan
in vitro but had nearly wild-type b1,3-glucan synthase activities. Comparisons of the activities of wild-type and b1,6glucan synthesis-defective strains revealed that levels of
b1,6-glucan formed de novo correlated with the reduction
in b1,6-glucan synthesis in vivo. It was proposed that the
dot-blot assay measured b1,6-glucan chain extension and
that higher rates of Glc transfer reflected the presence of
more acceptor (Vink et al. 2004). The reaction was stimulated by GTP and higher b1,6-glucan synthetic activity was
detected in membranes from cells overexpressing Rho1
GTPase, suggesting that b1,6-glucan synthase, like b1,3-glucan
synthase, is Rho1-dependent (Vink et al. 2004).
In the second approach, formation of b1,6-glucan was
measured in cells permeablized by osmotic shock and incubated with radiolabeled UDP-Glc (Aimanianda et al. 2009).
The insoluble, radiolabeled b1,6-glucan formed was chemically identical to the branched b1,6-linked glucan isolated
from cell walls, and radioactivity was distributed throughout
the in situ product, indicating that de novo polymerization of
b1,6-glucan had occurred (Aimanianda et al. 2009). Consistent with their severe in vivo defects in b1,6-glucan synthesis, permeabilized kre5 and kre9 mutants showed no in situ
b1,6-glucan synthetic activity, but made b1,3-glucan. The
b1,6-glucan synthetic activity in permeabilized cells was
not stimulated by GTP. However, because b1,3-glucan synthesis mutants make less b1,6-glucan, and vice versa, formation of the two polymers may be coordinated in another
way (Dijkgraaf et al. 2002; Aimanianda et al. 2009; see The
Fks family of b1,3-glucan synthases).
Proteins involved in b1,6-glucan assembly
Mutants defective in b1,6-glucan synthesis were identified
in screens for resistance to K1 killer toxin, which uses b1,6glucan as its receptor (Hutchins and Bussey 1983), and in
screens for CFW sensitivity (Ram et al. 1994; Lussier et al.
1997b; Orlean 1997; Shahinian and Bussey 2000; Pagé
et al. 2003). In these mutants, levels of alkali-insoluble b1,
6-glucan were lowered to different extents, and the proportions of b1,6- and b1,3-linked Glc residues in the alkaliinsoluble glucan fraction were often altered. The finding that
the proteins implicated in b1,6-glucan assembly were localized in the ER, Golgi, or plasma membrane, together with
demonstrations of epistasis relationships and genetic interactions, led to the notion of a secretory pathway-based pathway
for b1,6-glucan elaboration (Boone et al. 1990; reviewed by
Orlean 1997 and Shahinian and Bussey 2000). However,
b1,6-glucan is not detectable intracellularly (Montijn et al.
1999), and the roles of most of the proteins so far implicated
796
P. Orlean
are indirect. Proteins affecting the formation of b1,6-glucan
will be discussed in the order of their location along the
secretory pathway.
ER proteins: Homologs of the UGGT/calnexin protein quality
control machinery: Four homologs of proteins involved in the
UGGT/calnexin protein quality control system (see N-glycan
processing in the ER and glycoprotein quality control) are required for formation of normal amounts of b1,6-glucan
(Jiang et al. 1996; Abeijon and Chen 1998; Shahinian
et al. 1998; Simons et al. 1998). These are diverged UGGT
homologs Kre5, Gls1/Cwh41, Gls2/Rot2, and Cne1, of
which Kre5 has the most important role because kre5
mutants make no more than 5% of normal amounts of b1,
6-glucan (Meaden et al. 1990; Montijn et al. 1999; Levinson
et al. 2002; Aimanianda et al. 2009). The contributions of
the glucosidases and calnexin are likely indirect ones in
maintaining normal levels of unknown components of the
b1,6-glucan assembly machinery in the secretory pathway
(Shahinian et al. 1998; Lesage and Bussey 2006). The essential function of Kre5 is other than as a UGGT in protein
quality control because kre5D remained lethal in an alg8D
gls2D background in which all N-glycans stayed monoglucosylated, thereby bypassing the need for UGGT activity
(Shahinian et al. 1998). Kre5 could be a glucosyltransferase
with a specialized role in quality control of b1,6-glucan
assembly proteins (Levinson et al. 2002; Herrero et al.
2004; Lesage and Bussey 2006), or it could glucosylate
the GPI glycan of future GPI-CWPs to generate a signal
or attachment point for subsequent transfer to b1,6-glucan
(Shahinian and Bussey 2000). S. cerevisiae has the necessary ER UDP-Glc transport activity to supply the donor
(Castro et al. 1999).
N-glycosylation is important for wild-type levels of b1,6glucan to be made. For example, stt3 mutants have a severe
defect in b1,6-glucan synthesis and are synthetically lethal
with kre5 and kre9 (Chavan et al. 2003b). This may reflect
a requirement for N-glycosylation of one or more b1,6-glucan synthetic proteins or for an N-glycan to serve as acceptor
for initiation of a b1,6-glucan chain (Lesage and Bussey
2006). Interestingly, mutations such as och1 and mnn9, which
affect synthesis of the a1,6-mannan backbone, and mnn2,
which blocks addition of the first, a1,2 side-branching Man,
show elevated levels of b1,6-glucan (Magnelli et al. 2002;
Pagé et al. 2003), suggesting that a balance is normally maintained between these two polymers.
Fungus-specific ER chaperones required for b1,6-glucan
synthesis: Mutations in genes encoding the ER-localized,
fungus-specific membrane proteins Rot1, Big1, and Keg1 all
cause a b1,6-glucan synthetic defect. ROT1 and BIG1 null
mutants grow only with osmotic support, and even then
very slowly, and in this and in the severity of their b1,6glucan synthetic defect—a 95% reduction—they resemble
kre5D strains (Bickle et al. 1998; Azuma et al. 2002; Machi
et al. 2004). Levels of b1,3-glucan and chitin are elevated in
rot1D and big1D. The b1,6-glucan defect in keg1D cells is
similar to that in kre6D—about a 50% reduction (Nakamata
et al. 2007).
Rot1, Big1, and Keg1 are small proteins that show no
similarity to one another or to carbohydrate-active enzymes
(Lesage and Bussey 2006). They seem to function as ER
chaperones with varying degrees of importance for the stability of proteins involved in b1,6-glucan synthesis and may in
some cases cooperate. Observations supporting this notion
and indicating a relationship to Kre5 are discussed in File S8.
More widely distributed secretory pathway proteins: Kre6
and Skn1: Kre6 and Skn1 are homologous type 2 membrane
proteins in GH family 16 of b-1,6/b-1,3-glucanases (Henrissat
and Davies 1997; Montijn et al. 1999). kre6D cells make half
normal amounts of b1,6-glucan, whereas skn1D cells make
b1,6-glucan normally and have no growth defect. Expressed
at high copy, Skn1 restores almost normal levels of b1,6glucan to kre6D cells, and kreD skn1D double mutants are
inviable or very slow growing, depending on the strain background, and make no more than 10% of normal amounts of
b1,6-glucan (Roemer et al. 1993). From this, Kre6 and Skn1
seem to be functional homologs, with Kre6 normally having
the dominant role in b1,6-glucan synthesis. As hydrolases or
transglycosylases, Kre6 and Skn1 could act on a structure
that serves as a precursor or acceptor in elaboration of b1,6glucan or on a glycoprotein involved in b1,6-glucan synthesis (Lesage and Bussey 2006), but enzyme activity has yet to
be demonstrated for these proteins. Much of Kre6 is ERlocalized, where it interacts with Keg1, but the protein is
also detectable in the Golgi, in secretory vesicles, and at
the plasma membrane at sites of polarized growth (Li
et al. 2002; Nakamata et al. 2007; Kurita et al. 2011; File
S8). Localization of Skn1 has not been explored in detail.
Skn1 also has a role in the formation of mannosyl diinositolphosphoryl ceramide [M(IP)2C], because skn1D, but not
kre6D strains, is defective in M(IP)2C (Thevissen et al. 2005;
File S8).
Kre6 has been implicated in the mode of action of a pyridobenzimidazole derivative identified in a screen for inhibitors of cell wall incorporation of a reporter GPI-CWP
(Kitamura et al. 2009). Because cells treated with this compound showed lowered incorporation of radiolabeled Glc
into a b1,6-glucan fraction, and because a resistant mutant
had an amino acid substitution in Kre6, it was proposed that
the compound is an inhibitor of b1,6-glucan synthesis and
that Kre6 is its likely target (Kitamura et al. 2009).
Kre9 and Knh1: Kre9 and Knh1 are 30 kDa, soluble, fungus-specific, O-mannosylated proteins that are secreted into
the medium when overproduced (Brown and Bussey 1993;
Dijkgraaf et al. 1996). The two are functional homologs,
with Kre9 having the dominant role. kre9 nulls are slow
growing and show an 80% reduction in b1,6-glucan, and
the residual b1,6-glucan in them has about half the molecular mass as the wild-type polymer and is altered in its proportion of b1,6 and b1,6 linkages (Brown and Bussey 1993).
The size and structure of b1,6-glucan made in knh1D cells is
normal. Overexpression of KNH1 corrects the growth and
b1,6-glucan defects of kre9D, but the kre9 skn1 combination
is synthetically lethal (Dijkgraaf et al. 1996). kre9D, but not
knh1D, is also synthetically lethal with kre1D (see below)
and kre6D, but not with skn1D. KRE9’s genetic interactions
indicate that its product has a pleiotropic impact on b1,6glucan formation, although its effects must be exerted after
Kre5’s because the kre5D kre9D double null has the same
phenotype as kre5D (Dijkgraaf et al. 1996; Shahinian and
Bussey 2000). Neither Kre9 nor Knh1 shows similarity to
proteins of known function. If they are not enzymes, Kre9
and Knh1 may serve to anchor b1,6-glucan in the cell wall
(Lesage and Bussey 2006), but this must be reconciled with
the finding that kre9 mutants have no UDP-Glc-dependent
b1,6-glucan synthetic activity (Aimanianda et al. 2009).
Plasma membrane protein Kre1: Kre1, a GPI protein, functions at the plasma membrane or in the wall. kre1D cells
make 40% of wild-type levels of b1,6-glucan, but this glucan
is smaller and its b1,3 side branches are not extended
(Boone et al. 1990; Roemer and Bussey 1995). GPI attachment is necessary for Kre1’s function and cell surface localization (Breinig et al. 2004). The hydrophilic portion of Kre1
shows no similarity to known enzymes. Kre1 has a structural
role and becomes cross-linked to other wall proteins (Breinig
et al. 2004), and it also serves as a receptor for K1 killer
toxin (File S8).
How might b1,6-glucan be made?: Obstacles to identifying
the b1,6-glucan synthase gene might be an inherent difficulty
in obtaining hypomorphic alleles of an essential synthase
gene or the existence of multiple redundant synthase genes
whose individual mutation gives no phenotype (Lesage and
Bussey (2006). Furthermore, there are no precedents in other
organisms that could be exploited in bioinformatics-based
approaches to b1,6-glucan synthesis. Although b1,6-glucan
is widely distributed in the Fungi (Lesage and Bussey
2006), it is very rare elsewhere. The bacterium Actinobacillus
suis makes a lipopolysaccharide containing a b1,6-glucan homopolymer (Monteiro et al. 2000), but the proteins involved
in its formation are unknown. Some bacteria have GT2 family
transferases that make polymers of b1,6-linked GlcNAc
(Gerke et al. 1998; Itoh et al. 2008), but these enzymes resemble the S. cerevisiae Chs proteins. If b1,6-glucan is indeed
formed directly from UDP-Glc, the b1,6-glucosyltransferase
would represent a new GT family. Further possibilities are
that a known yeast GT may also form b1,6-glucosidic linkages
using UDP-Glc as donor or that b1,6-glucan is generated
solely by transglycosylation.
Remodeling and Cross-Linking Activities at the Cell
Surface
Order of incorporation of components into the cell wall
CWPs delivered by the secretory pathway meet up with chitin
and b-glucans at the outer face of the plasma membrane and
S. cerevisiae Cell Wall
797
undergo cross-linking reactions that incorporate them into
the wall. The order in which wall components are assembled
has been inferred from analyses of the material formed
when spheroplasts regenerate their walls and from the wall
compositions of mutants unable to make a particular component (Kreger and Kopecká 1976; Roh et al. 2002b). The
starting component is b1,3-glucan, which is necessary for
incorporation of both b1,6-glucan and mannoproteins. Because b1,6-glucan was still attached to b1,3-glucan when
GPI anchoring was inhibited (Roh et al. 2002b), and because
incorporation of GPI-CWPs is lowered in b1,6-glucan synthesis mutants (Lu et al. 1995; Kapteyn et al. 1997), GPICWPs are likely incorporated after b1,6-glucan. Because chitin became detectable in the walls of daughter cells only
after cytokinesis (Shaw et al. 1991), it was concluded that
chitin is the last component to be incorporated into the wall
(Roh et al. 2002b). The sequence b1,3-glucan/b1,6glucan/mannoprotein must be able to accommodate
changes in expression or assembly of individual components
dictated by the cell cycle, cell wall stress, mating, or sporulation, as well as remodeling of individual polysaccharides.
For example, a compensatory incorporation of PIR protein
directly attached to b1,3-glucan is seen in b1,6-glucan synthesis mutants (Kapteyn et al. 1999b).
The model for the order of incorporation of wall
components needs to be reconciled with the model for
a bilayered wall, during whose formation CWPs are propelled to the cell surface, leaving polysaccharides nearer the
plasma membrane. Furthermore, surface CWP may not be
retained at the surface of wild-type cells. Thus, wild-type
diploids expressing a Sag1-GFP fusion released a significant
basal level of that glycoprotein into the medium (Gonzales
et al. 2010). CWP may therefore routinely be shed during
vegetative growth, perhaps upon digestion of the wall between mother cell and bud, along with secreted proteins
such as chitinase and invertase (Kuranda and Robbins
1991). Cross-linking and remodeling reactions will be described next, and hydrolases of known or unknown functions, as well as nonenzymatic wall proteins are discussed
in Cell Wall-Active and Nonenzymatic Surface Proteins and
Their Functions.
Incorporation of GPI proteins into the wall
The v(2) region of a GPI protein (see Identification of GPI
proteins) influences whether the protein will be retained in
the plasma membrane in lipid-anchored form or whether it
can be transferred into the wall (Caro et al. 1997; Hamada
et al. 1998a, 1999; De Sampaïo et al. 1999; Frieman and
Cormack 2004). If this region includes two basic amino
acids, the protein will be mostly retained in the plasma
membrane (Caro et al. 1997; Frieman and Cormack 2003),
but if basic residues are absent or replaced with hydrophobic
ones, the predominant location is the wall (Hamada et al.
1998b, 1999; Frieman and Cormack 2003). However, having
two basic amino acids in the v(2) region does not guarantee
membrane localization because the additional presence of
798
P. Orlean
a longer stretch of amino acids rich in Ser and Thr will override
the dibasic motif and shift the protein to the wall (Frieman
and Cormack 2004). Furthermore, not all wall-anchored GPI
proteins have the amino acids suggested to promote incorporation into the wall (De Groot et al. 2003). In general, GPI
proteins are partitioned between the membrane and wall
to varying extents, and none may be restricted to only one
location (Gonzales et al. 2009).
The nature of a GPI-protein’s mode of cell surface attachment can be critical. Ecm33, which is required for growth at
elevated temperature (see Sps2 family), occurs mainly as
a plasma membrane-anchored GPI protein, and this localization is required for in vivo function. Replacement of v(2)
amino acids v-1-13 of Ecm33 with the corresponding amino
acid sequences from wall-localized proteins resulted in increased cross-linking of Ecm33 to the wall, but also in loss of
the protein’s ability to support growth at high temperature
(Terashima et al. 2003).
The lipid-to-wall transfer reaction could be a one-step
transglycosylation in which the GPI glycan is cleaved and its
reducing end transferred to b1,6-glucan, or it could involve
separate GPI cleavage and transglycosylation steps. Candidates for cross-linkers are Dfg5 and Dcw1, an essential, redundant pair of homologous GPI proteins that resemble an
a1,6-endomannanase and are in GH Family 76 (Kitagaki
et al. 2002). Single dfg5D and dcw1D mutants are viable,
although dcw1D is hypersensitive to Zymolyase, but the
combination of dfg5D and dcw1D is lethal (Kitagaki et al.
2002). Depletion of Dfg5 or Dcw1 by repressing their expression in the double-null background led to cell enlargement, delocalized chitin deposition, and secretion of a GPICWP protein into the growth medium (Kitagaki et al. 2002).
dcw1D was also recovered in a screen for impaired crosslinking of GPI proteins (Gonzalez et al. 2010). The defects
caused by loss of Dfg5 and Dcw1, together with the proteins’
resemblance to an a-endomannanase, are consistent with
their having a role in GPI cleavage and/or transglycosylation. Homozygous DFG5 nulls are defective in filamentous
growth (Mosch and Fink 1997).
GPI-CWP can be used to display heterologous proteins on
the yeast cell surface (Schreuder et al. 1993; Van der Vaart
et al. 1997; Gai and Wittrup 2007; Shibasaki et al. 2009). In
one such system, heterologous proteins are fused to the
Aga2 subunit of the a-mating agglutinin (see Flocculins
and agglutinins), which is disulfide-linked to its partner,
the GPI-CWP Aga1 (Boder and Wittrup 1997).
Incorporation of PIR proteins into the wall
The internal repeat (PIR) sequences of PIR proteins are
required for the alkali-labile linkage that joins these proteins
to b1,3-glucan (see Wall Composition and Architecture). Deletion of all PIR sequences from Pir1 and Pir4 leads to release of these proteins from the cells (Castillo et al. 2003;
Sumita et al. 2005), indicating that the repeats are necessary
for wall association. The more repeats, the stronger the
binding: deletion of increasing numbers of Pir1’s repeats
led to release of increasing amounts of Pir1 into the medium
(Sumita et al. 2005).
Studies of Pir4/Ccw5, which has one PIR sequence and
needs it for cell wall anchorage, revealed that the alkalilabile linkage was an ester between the g-carboxyl group
of glutamate and the hydroxyl groups of b1,3-glucan. The
linkage was generated in a transglutaminase reaction with
Q74 in the PIR sequence SQIGDGQ74[V/I]QAT[T/S] (Ecker
et al. 2006). In addition to substitutions of Q74, individual
mutations of Q69, D72, and Q76 also resulted in loss of wall
anchorage of the protein, indicating that these residues have
roles in the reaction. No transglutaminase has yet been identified, but Ecker et al. (2006) point out that, because the free
energy of hydrolysis of the amide is high enough to drive
formation of the ester linkage, the PIR proteins could catalyze their own attachment to b1,3-glucan.
The glucan attachment sequence of PIR repeats is also
found in the GPI-CWP Tip1, Tir1, Cwp1, and Cwp2 (Van der
Vaart et al. 1995). In the case of Cwp1, the PIR repeat may
be used as an additional wall anchorage point because the
protein is attached to the wall by both an alkali-labile and
a GPI-dependent linkage (Kapteyn et al. 2001). Like GPICWP, PIR proteins can be used as carriers to direct surface
expression of heterologous proteins fused to them (Andrés
et al. 2005; Shimma et al. 2006).
Cross-linkage of chitin to b1,6- and b1,3-glucan
Related Crh1 and Crh2 generate cross-linkages between the
reducing ends of chitin chains and both the nonreducing end
of b1,3-glucose side branches on b1,6-glucan and the nonreducing ends of b1,3-glucan chains (Cabib et al. 2007; Cabib
2009), and their homolog Crr1 likely does so during ascospore wall assembly. These proteins are in GH family 16, and
Crh2 and Crr1 also have a chitin-binding module (RodriguezPena et al. 2000; Cabib et al. 2008). Crh1 and Crh2 are GPI
proteins (Caro et al. 1997; Hamada et al. 1998a) whose localization matches that of Chs3. Crh1-GFP fusions are detectable at the site of bud emergence and later in the neck region
between mother cell and bud, and Crh2-GFP is seen in the
neck region throughout the budding cycle, as well as in the
lateral wall (Rodriguez-Pena et al. 2000, 2002). Single crh1
and crh2 null mutants show Calcofluor White and Congo Red
sensitivity, phenotypes enhanced in the double null, suggesting that Crh1 and Crh2 have a common wall-related function.
Single crh mutants have a higher ratio of alkali soluble- to
alkali-insoluble glucan, and this ratio is higher still in crh1D
crh2D, indicating a role for Crh1 and Crh2 in linking b-glucan
and chitin (Rodriguez-Pena et al. 2000).
Evidence that Crh1 and Crh2 are transglycosylases came
from elegant studies by Cabib and coworkers, who used fluorescent, sulforhodamine-conjugated b1,3-gluco-oligosaccharides
as acceptors and showed that they became cross-linked to
chitin in bud scars and the lateral walls of live cells (Cabib
et al. 2008). Fluorescent labeling was very weak in crh1D
crh2D cells or in cells lacking Chs3, which makes the chitin
normally bound to b1,3- and b1,6-glucan (Cabib and Duran
2005). The entire process of chitin polymerization and crosslinking could be reconstituted in detergent permeabilized,
protease-treated cells. Cross-linking of fluorescent b1,3gluco-oligosaccharides depended on the addition of UDPGlcNAc (Cabib et al. 2008). Interestingly, the nascent chitin
was generated in situ by Chs1, which is highly active in
permeabilized cells, rather than by Chs3.
CRR1 shows sporulation-specific expression. Crr1-GFP
fusions are localized on the surface of ascospores, and homozygous crr1D diploids have ascospore wall abnormalities,
with irregular deposition of the outer dityrosine and chitosan layers over the inner b-glucan layer (Gómez-Esquer
et al. 2004). Ascopores from Crr1-deficient diploids show
increased sensitivity to heat shock and lytic enzymes, and
these defects are exacerbated when the chitin deacetylases
Cda1 and Cda2 are also absent. These findings suggest
a role for Crr1 in generating cross-links between the b-glucan
and chitosan or chitin during ascospore wall maturation
(Gómez-Esquer et al. 2004).
Cell Wall-Active and Nonenzymatic Surface Proteins
and Their Functions
Secreted, membrane, or wall proteins with known or conjectured roles in wall biogenesis, adhesion, and nutrition are
surveyed here. The primary division is according to whether
proteins have or are likely to have enzymatic activity or
whether they are nonenzymatic, structural proteins. Both
groups contain GPI proteins. Cell wall proteins have been
reviewed by Klis et al. (2002, 2006), De Groot et al. (2005),
Lesage and Bussey (2006), and Gonzalez et al. (2009), glycosylhydrolases by Adams (2004), agglutinins by Dranginis
et al. (2007), and flocculins by Goossens and Willaert
(2010). Additional information about these proteins is presented in File S9.
Known and predicted enzymes
Chitinases: S. cerevisiae has two chitinases, Cts1 and Cts2.
Cts1, an endochitinase, has an N-terminal catalytic domain,
followed by a heavily O-mannosylated Ser/Thr-rich region,
and lastly, a C-terminal chitin-binding domain (Kuranda
and Robbins 1991). Cts1 is periplasmic, but much of it is
secreted into the medium of cells grown in rich medium
(Correa et al. 1982; Kuranda and Robbins 1991). Cts1 has
a key role in cell separation because cts1D strains form aggregates of cells that remain joined at their chitin-containing
septa, a phenotype mimicked when cells are treated with
the chitinase inhibitor dimethyallosamidin (Kuranda and
Robbins 1991). Cts1’s chitin-binding domain contributes to
the enzyme’s localization in the septal region because Cts1
truncations lacking it only partially complement the cts1D
separation defect (Kuranda and Robbins 1991). Cts2 may
have a role in sporulation (Dünkler et al. 2008).
b1,3-glucanases: Exg1, Exg2, and Ssg/Spr1 exo-b1,3-glucanases:
Exg1 is disulfide-linked (Cappellaro et al. 1998) whereas
S. cerevisiae Cell Wall
799
Exg2 is a surface-anchored GPI protein (Larriba et al. 1995;
Caro et al. 1997). Single- or double-null mutants in EXG1
and EXG2 have no overt defects, although exg1D cells have
slightly elevated levels of b1,6-glucan, and EXG1 overexpressers lower amounts of that polymer, suggesting roles
for Exg1 and Exg2 in b-glucan remodeling (Jiang et al.
1995; Lesage and Bussey 2006). Ssg1/Spr1 is a sporulationspecific protein (File S9).
Bgl2, Scw4, Scw10, and Scw11 endo-b1,3-glucanases:
These are GPI-less secretory proteins. Bgl2 has endo-b1,3glucanase activity in vitro (Mrša et al. 1993), but it can also
create a b1,6 linkage between the reducing end that it generates by cleaving a b1,3-gluco-oligosaccharide and the nonreducing end of another b1,3-glucan chain (Goldman et al.
1995), and so could function as a b1,3-glucan branching
enzyme. No enzymatic activity has been shown for Scw4,
Scw10, or Scw11, although mutation of predicted catalytic
residues in Scw10 abolished in vivo function (Sestak et al.
2004). Scw4, Scw10, and Bgl2 are wall-associated via disulfides (Cappellaro et al. 1998), but some Scw4 and Scw10
can also be linked to b1,3-glucan (Yin et al. 2005).
These proteins have roles in maintaining normal walls.
bgl2D, scw4D, and scw10D strains grow like wild-type cells,
but show CFW sensitivity and slightly increased chitin levels
(Klebl and Tanner 1989; Cappellaro et al. 1998; Kalebina
et al. 2003; Sestak et al. 2004), and bgl2D walls have elevated
levels of alkali-soluble glucan (Sestak et al. 2004). Strains
lacking both Scw4 and Scw10 are CFW-hypersensitive and
morphologically abnormal, have doubled chitin content and
increased alkali-soluble glucan, and show alterations in
b1,3-glucan structure and in cross-linking of mannoproteins
to the wall (Cappellaro et al. 1998; Sestak et al. 2004).
The growth and morphological defects of scw4D scw10D
are exacerbated by deletion of CHS3 or FKS2 (Sestak et al.
2004). From the phenotypes of strains expressing different relative amounts of Bgl2 and Scw10, it was proposed
that levels of Bgl2 and Scw10 need to be balanced
to ensure wall stability (Klebl and Tanner 1989; Sestak
et al. 2004; File S9). Cells lacking Scw11 have a separation
defect, and, consistent with this, Scw11 is a daughter cellspecific protein (Cappellaro et al. 1998; Colman-Lerner
et al. 2001).
Eng1/Dse4 and Eng2/Acf2 endo-b1,3-glucanases: These
related proteins have endo-b1,3-glucanase activity in vitro,
but different localizations. Eng1 is a GPI protein (Baladron
et al. 2002; De Groot et al. 2003), whereas Eng2 is likely
intracellular. Mutants lacking one or both proteins make
normal walls, but eng1D cells have a separation defect, consistent with Eng1’s localization to the daughter side of the
septum (Colman-Lerner et al. 2001; Baladron et al. 2002).
ENG2 expression increases during sporulation, although
eng2D diploids are not defective in that process (Baladron
et al. 2002). Surprisingly, loss of multiple exo- and endob1,3-glucanases is not catastrophic because cells lacking
Exg1, Exg2, Eng1, Eng2, and Bgl2 grow well and show only
the eng1D separation defect (Cabib et al. 2008).
800
P. Orlean
Gas1 family b1,3-glucanosyltransferases: This family has
five members, all of which have GPI attachment sites
(Fankhauser et al. 1993; Caro et al. 1997; Popolo and Vai
1999; De Groot et al. 2003). Gas1, Gas3, and Gas5 can also
be covalently linked to the wall (De Sampaïo et al. 1999; Yin
et al. 2005). Gas proteins have b1,3-glucanosyltransfer activity: they cleave b1,3-glucosidic linkages within b1,3-glucan
chains and then transfer the newly generated reducing end
of the cleaved glycan to the nonreducing end of another
b1,3-glucan molecule, thereby extending the recipient
b1,3-glucan chain (Mouyna et al. 2000; Carotti et al.
2004; Ragni et al. 2007b; Mazan et al. 2011; File S9).
Gas1 has a major role in vegetative wall biogenesis.
gas1D mutants are CFW-hypersensitive (Ram et al. 1994)
and have less b1,3-glucan but more chitin and mannan in
their walls (Ram et al. 1995; Popolo et al. 1997; Valdivieso
et al. 2000). gas1D cells release b1,3-glucan to the medium
(Ram et al. 1998), and Gas1’s b1,3-glucan elongase activity
may therefore be necessary for incorporation of b1,3-glucan
into the wall. In addition, analyses of the synthetic interaction network of gas1D revealed that survival in the absence
of Gas1 requires correct assembly of b1,6-glucan (Tomishige
et al. 2003; Lesage et al. 2004). Gas1 is detectable in the
lateral wall, in the chitin ring in small-budded cells, and
near the primary septum and remains in the bud scar after
cell separation, and its localization is dependent on the presence of its GPI-attachment sequence (Rolli et al. 2009).
Gas3 and Gas5 likely have wall-related functions in vegetative cells (File S9). GAS2 and GAS4 are expressed only
during sporulation, and diploids lacking both Gas2 and Gas4
have a severe sporulation defect. The inner glucan layer of
the wall of double homozygous gas2 gas4 null spores was
disorganized and detached from chitosan, suggesting that
the b1,3-glucanosyltransferase activity of Gas2 and Gas4
generates b1,3-glucan chains that associate optimally with
chitosan (Ragni et al. 2007a).
Yapsin aspartyl proteases: GPI-anchored aspartyl proteases
of the yapsin family have roles in the turnover of CWPs and
wall-localized enzymes. Yps1, Yps2/Mkc7, Yps3, and Yps6
are mostly plasma membrane-associated, whereas Yps7 is
predicted to be wall anchored (Krysan et al. 2005; GagnonArsenault et al. 2006). Yapsins cleave their substrates
C-terminally to Lys or Arg or pairs of these residues (Olsen
et al. 1998; Komano et al. 1999) and themselves undergo
proteolytic processing to generate active enzyme (File S9).
Individual yapsin null mutants are sensitive to various
wall-disrupting agents, and loss of multiple YPS genes leads
to osmotically remedial, temperature-sensitive lysis defects,
findings that indicate that the yapsins are involved in wall
maintenance (Krysan et al. 2005). Walls from yps1D yps2D
and yps1D yps2D yps3D yps6D yps7D null mutants showed
lowered b1,3- and b1,6-glucan and elevated chitin levels,
whereas mannan levels were unchanged, with the wall
alterations being most pronounced in the quintuple mutant
(Krysan et al. 2005). The b-glucan defects were due to
decreased incorporation of these polymers into the wall,
because synthesis of the two b-glucans was normal in the
deletion strains. These findings suggest that yapsins act on
wall hydrolases and transglycosidases, thereby regulating
activity of the latter, and hence, incorporation of glucans
into the wall (Krysan et al. 2005). Support for this came
from identification of Gas1, Pir4, and Msb2 as Yps1 substrates (Gagnon-Arsenault et al. 2008; Vadaie et al. 2008).
In addition to degrading or shedding proteins during wall
remodeling, yapsins also have roles in mediating release of
aberrantly folded or overexpressed GPI proteins that induce
ER stress (Miller et al. 2010).
Nonenzymatic CWPs
Structural GPI proteins: There are three families of GPICWP and several individual GPI-CWP that do not resemble
known enzymes. Strains lacking one or more of these GPICWP have wall defects, and expression of some of these
proteins can vary with cell cycle stage or be induced during
mating or sporulation in response to cell wall stress or when
oxygen levels are low. In general, GPI-CWPs have a collective
role in maintaining cell wall stability (Lesage and Bussey
2006; Ragni et al. 2007c).
Sps2 family: This group comprises Ecm33, Pst1, Sps2, and
Sps22 (Caro et al. 1997). Of these, Ecm33 has an important
role in vegetative walls. ecm33D cells are temperature-sensitive
and sensitive to various wall-perturbing agents, have a disorganized wall with a thin or absent mannoprotein layer,
and shed b1,6-glucan-linked mannoproteins and Pir2 that
is possibly linked to b1,3-glucan more than normal in the
medium (Lussier et al. 1997b; Pardo et al. 2004). pst1D cells
have no obvious phenotype, but ecm33D pst1D double nulls
show exacerbated sensitivity to various wall stresses.
Ecm33’s and Pst1’s functions partially overlap, but the proteins are not fully redundant because overexpression of
PST1 only weakly suppresses the ecm33D defects (Pardo
et al. 2004). Sps2 and Sps22 are a redundant pair required
for normal ascospore wall formation. Diploids lacking them
form spores with abnormal b-glucan, chitosan, and dityrosine layers (Coluccio et al. 2004). Sps2 and Sps22 likely act
at a similar stage in ascospore wall formation as Gas2, Gas4,
and Crr1 in the formation of the b-glucan layer.
Tip1 family: Tip1, Cwp1, Cwp2, Tir1, Tir2, Tir3, Tir4, and
Dan1/Ccw13 are mostly small, Ser- and Ala-rich GPI-CWP
that show differential expression during the cell cycle and
during aerobic or anaerobic growth and can be localized
differently on the cell surface. Cwp2 also contains a PIR repeat and so could be linked to b1,3-glucan (Klis et al. 2010).
Cwp1, Cwp2, Tip1, and Tir1 have roles in the vegetative
wall. Deletion of their genes individually leads to CFW hypersensitivity (Van der Vaart et al. 1995), and cwp1D cwp2D
double mutants show increased permeability to DNA-binding
agents relative to the single nulls (Zhang et al. 2008). In
addition, the walls of cwp2D and cwp1D cwp2D cells are
thinner than those of the wild type (Van der Vaart et al.
1995; Zhang et al. 2008). Localization of these proteins
correlates with their expression. Tip1 is expressed in G1
and found in mother cells only, whereas Cwp1, Cwp2, and
Tir1 are expressed during the S-to-G2 transition, Cwp2 being
found in small-to-medium-sized buds (Caro et al. 1998;
Smits et al. 2006). Localization of Cwp2 and Tip1 is determined by the timing of their expression in the cell cycle
(Smits et al. 2006; File S9). Tip1 and Tip2 are also heatand cold-shock-inducible, and Tir1 and Tir4 are induced by
cold shock (Kowalski et al. 1995; Abramova et al. 2001).
CWP1 and CWP2 are downregulated upon shift to anaerobic conditions, whereas Tip1, Tir1, Tir2, Tir3, Tir4, Dan1/
Ccw13, and Dan4 are induced (Abramova et al. 2001). Of
these, the Dan proteins are strongly repressed by oxygen.
Strains lacking Tir1, Tir3, or Tir4 do not grow under anaerobic conditions. Shift to anaerobiosis therefore leads to
remodeling of the wall (Abramova et al. 2001), although
it is not clear how the anaerobically induced CWPs permit
anaerobic growth.
Sed1 and Spi1: These are two related, Ser/Thr-rich GPICWP whose expression is induced by nutrient limitation and
stress. Sed1 is releasable from walls by treatment with
b-glucanases or proteases (Shimoi et al. 1998). Association
of Sed1 with the wall is dependent on Kre6 (Bowen and
Wheals 2004), consistent with anchorage involving b1,6glucan. SED1 expression is induced in the stationary phase,
a time when the wall becomes thicker and more resistant to
lytic enzymes (De Nobel et al. 1990). Consistent with a protective role in stationary-phase walls, sed1D cells become
more sensitive to Zymolyase in that growth phase (Shimoi
et al. 1998). Elevated Sed1 expression is also part of the
compensatory response made by cells lacking multiple
GPI-CWP (Hagen et al. 2004; File S9). Expression of SPI1
is induced by weak organic acids, and Sps1 is a major contributor to the b1,3-glucanase resistance that arises in response to this stress (Simoes et al. 2003). Low external pH
also leads to formation of new alkali-labile linkages between
GPI-CWPs and b1,3-glucan (Kapteyn et al. 2001).
Ccw12: Ccw12 is a small, heavily glycosylated, Ser/Thrrich GPI-CWP with two C-terminal repeats of an amino acid
sequence critical for its function (File S9). Ccw12 is releasable by b1,3-glucanases (Mrša et al. 1999), but also has the
potential to make disulfide cross-links because it has a threeCys motif found in several S. cerevisiae flocculins (see Flocculins and agglutinins) and in wall proteins of other yeasts
(Klis et al. 2010). Ccw12 is likely abundant because its gene
has a very high codon adaptation index (Klis et al. 2010).
Ccw12 has a major role in the wall, because cells lacking it
are hypersensitive to CFW and other wall stressing agents
and rounder than wild type, with thick, disorganized walls,
lysis-prone buds, and elevated levels of chitin (Mrša et al.
1999; Hagen et al. 2004; Ragni et al. 2007c; Shankarnarayan
et al. 2008). Man-to-Glc ratios in ccw12D cells are unchanged,
but levels of alkali-soluble relative to alkali-insoluble glucan
are higher, indicating altered organization and cross-linkage
of wall components (Ragni et al. 2007c). Ccw12 localizes to
sites of active wall synthesis, including the future bud site,
S. cerevisiae Cell Wall
801
the septum, the lateral walls of enlarging daughter cells, as
well as the tips of mating projections, but then turns over,
suggesting that it may stabilize walls as daughter cells and
that mating projections are being formed (Ragni et al.
2011). Loss of Ccw12 alone activates the CWI pathway-mediated chitin stress response (Ragni et al. 2007c, 2011; see
Chitin synthesis in response to cell wall stress), but deletion of
additional GPI-CWP genes forces cells over a threshold that
leads to triggering of a new compensatory response to loss
of multiple GPI-CWP that depends on Sed1 and the nonGPI-CWP Srl1 (see File S7).
Other nonenzymatic GPI proteins: Ccw14 (Ssr1/Icwp) is
a b1,3-glucanase-extractable, Ser-rich GPI-CWP that has been
localized to the inner cell wall (Moukadiri et al. 1997; Mrša
et al. 1999; File S9). The protein has an eight-Cys-containing
CFEM domain found in various fungal surface proteins
(Kulkarni et al. 2003; De Groot et al. 2005) and, hence, a potential for disulfide formation. CCW14/SSR1 null mutants
have no obvious growth defects, but show increased sensitivity to CFW, Congo Red, and Zymolyase. Overexpression of
CCW14/SSR1 also leads to increased CFW and Congo Red
sensitivity, although not Zymolyase sensitivity, suggesting that
levels of Ccw14/Ssr1 relative to one or more other wall components need to be balanced (Moukadiri et al. 1997).
Dse2 and Egt2, which are unrelated to one another, are
daughter cell-specific proteins with roles in cell separation.
In haploids, Dse2 is concentrated in regions connecting
mother and daughter cells (Colman-Lerner et al. 2001; Doolin
et al. 2001), and Egt2 is localized to the septum (Fujita
et al. 2004). Of these two GPI proteins (Hamada et al.
1998a; Terashima et al. 2002; De Groot et al. 2003), Egt2’s
localization also depends on Gpi7 (Fujita et al. 2004). dse2D
haploids show no defects, but homozygous DSE2 nulls show
unipolar budding and form chains of cells. egt2D cells have
separation defects similar to those of eng1D cells, a phenotype exacerbated in the double null, indicating that the two
proteins act in parallel pathways involved in cell separation
(Kovacech et al. 1996; Baladron et al. 2002).
The related Ser/Thr-rich GPI proteins Fit1, Fit2, and Fit3
(Hamada et al. 1999) have a nutritional role. Their expression is induced by iron limitation, and the proteins normally
retain iron bound to ferrichrome because Zymolyase treatment of FIT-deletion mutants releases less iron from cells
(Protchenko et al. 2001). Fit1 localizes to the wall, where
it, Fit2, and Fit3 concentrate siderophore iron and facilitate
subsequent uptake of the metal, highlighting a role of the
wall in nutrient acquisition (Protchenko et al. 2001).
Flocculins and agglutinins: GPI-CWP involved in cell–cell
adhesion are the related Flo1, Flo5, Flo9, Flo10, and Flo11/
Muc1 flocculins, the Aga1 and Fig2 pair, and the a-agglutinin
Sag1 (Roy et al. 1991; Cappellaro et al. 1994; Chen et al.
1995; Caro et al. 1997; Erdman et al. 1998; Guo et al. 2000;
Shen et al. 2001; Dranginis et al. 2007; Van Mulders et al.
2009; Goossens and Willaert 2010).
Flo1, Flo5, Flo9, and Flo10 are modular proteins composed
of 1100–1500 amino acids. Major features are N-terminal
802
P. Orlean
PA14 domains that bind a-mannosides and mediate adhesion to adjacent cells, a central Ser/Thr-rich domain that is
organized in repeat sequences and heavily glycosylated,
and two or three conserved three-Cys repeats toward their
C termini (Verstrepen and Klis 2006; Goossens and Willaert
2010; Klis et al. 2010; Veelders et al. 2010; Goossens et al.
2011). In addition, the Ser/Thr-rich domains of Flo1 and
Flo11/Muc1 have short sequences enriched in Ile, Thr, and
Val that are predicted to form intramolecular b-sheetlike interactions or amyloids, and both a soluble, GPI-less
portion of Flo11/Muc1 and a Flo1-derived form fibrillar
b-aggregates in vitro (Ramsook et al. 2010). Amyloid formation correlates with flocculation in vivo, for cells expressing
Flo1 and Flo11/Muc1 that had been induced to flocculate in
the presence of Ca2+ stained more brightly with an amyloidbinding dye, and amyloid formation may be part of the
mechanism by which these proteins promote cell aggregation (Ramsook et al. 2010). FLO1, FLO5, FLO9, and FLO10
are not expressed in laboratory strains such as S288C because of a mutation in the transcriptional activator Flo8.
However, activation of individual FLO genes confers the ability to flocculate (Guo et al. 2000; Van Mulders et al. 2009).
Flo11/Muc1, a diverged Flo protein (Lambrechts et al. 1996;
Lo and Dranginis 1996), is not involved in flocculation,
but is required for pseudohypha formation by diploids, invasion of agar by haploids, and biofilm development (Lo
and Dranginis 1998; Guo et al. 2000; Reynolds and Fink
2001; Dranginis et al. 2007; Bojsen et al. 2012).
Related Aga1 and Fig2 function in mating and localize to
the mating projection (Erdman et al. 1998; Guo et al. 2000;
Jue and Lipke 2002). Aga1 is a component of a-agglutinin
that displays the Aga2 subunit, which is disulfide linked
to it, and which confers binding specificity to a-agglutinin
Sag1 (Orlean et al. 1986; Roy et al. 1991; Cappellaro et al.
1994; Shen et al. 2001). Fig2, which like Aga1 is expressed
in both mating types, is required for formation of mating
projections and maintenance of wall integrity during mating
(Erdman et al. 1998; Guo et al. 2000; Zhang et al. 2002;
File S9).
Sag1 has a long Ser/Thr-rich region in its C-terminal half
that may hold up the N-terminal, Aga2-binding portion
of the protein at the cell surface. Sag1’s N-terminal region
contains three sequential domains that resemble variable
immunoglobulin-like folds (Chen et al. 1995; Shen et al.
2001), the most C-terminal of which contains amino acids necessary for Aga2 binding (Wojciechowicz et al. 1993; Cappellaro
et al. 1994; De Nobel et al. 1996).
Non-GPI-CWP: PIR proteins: Expression and localization of
Pir1 (Ccw6), Pir2 (Ccw7/Hsp150), Pir3 (Ccw8), and Pir4
(Ccw5/Cis3) is regulated during cell cycle progression and
in response to stress. PIR1, PIR2, and PIR3 show peaks of
expression in early G1, whereas PIR4 expression is highest in
G2 (Spellman et al. 1998). PIR2 is also induced by heat
shock, treatment with CFW or Zymolyase, and nitrogen limitation (Russo et al. 1993; Toh-e et al. 1993; Yun et al. 1997;
Boorsma et al. 2004). Consistent with their upregulation
upon wall stress, all four PIR genes show elevated expression in an mpk1 mutant that constitutively activates the protein kinase C-dependent CWI pathway, an effect eliminated
in mutants lacking the PKC pathway’s target transcription
factor, Rlm1 (Jung and Levin 1999).
PIR proteins localize to different parts of the surface of
budding cells (Sumita et al. 2005; File S9). Pir1 and Pir2 are
found at bud scars of both haploids and diploids, Pir1 being
localized inside the chitin ring. Some Pir1 and Pir2 and most
Pir3 are also present in lateral walls (Yun et al. 1997). Pir4
has been reported be uniformly distributed at the cell surface or restricted to growing buds (Moukadiri et al. 1999;
Sumita et al. 2005).
Strains lacking individual PIR proteins have subtle
growth defects, but as more PIR genes are deleted, disruptants show a progressive increase in sensitivity to CFW,
Congo Red, and heat shock, and cells become larger and
irregularly shaped (Toh-e et al. 1993; Mrša and Tanner
1999). pir1D pir2D pir3D pir4D mutants show a loss of viability that is suppressed in osmotically supported medium
(Teparic et al. 2004). These findings suggest a collective role
for PIR proteins in maintenance of a normal wall. How these
proteins contribute is unclear, because the carbohydrate composition of the quadruple PIR disruptant’s wall is unaltered,
and the relative amounts of alkali-soluble and -insoluble glucan and chitin show modest changes (Teparic et al. 2004;
Mazan et al. 2008). PIR proteins, however, impact permeability of the wall because the pir1D pir2D pir3D mutant is hypersensitive to membrane-active tobacco osmotin, whereas
overexpression of PIR1, PIR2, or PIR3 confers osmotin resistance on walled cells but not spheroplasts (Yun et al. 1997).
The effects of PIR protein levels on wall permeability are
consistent with the role of these proteins in cross-linking
b1,3-glucans (see Mild alkali-releasable proteins).
Scw3 (Sun4): Haploids lacking this soluble cell wall
protein (Cappellaro et al. 1998) are larger than wild-type
cells and have a separation defect and thickened septa
(Mouassite et al. 2000). Scw3/Sun4 is a member of the
SUN family of proteins, of which Sim1 and Uth1 are also
released from cell walls by dithiothreitol treatment (Velours
et al. 2002). Uth1 and Scw3/Sun4 additionally localize to
mitochondria (Velours et al. 2002), but the significance of
this distribution of the SUN proteins is unclear. The biochemical function of the SUN proteins is unknown as they
show no similarity to known enzymes (File S9).
Srl1: This small Ser/Thr-rich protein is involved in the
compensatory response to loss of multiple GPI-CWP (Hagen
et al. 2004; File S9). It rescues the lysis defects of strains
defective in the function of the “regulation of Ace2 and polarized morphogenesis” (RAM) signaling network when overexpressed (Kurischko et al. 2005), and some of it is tightly
associated with the wall and released by b1,3-glucanase
(Terashima et al. 2002). Slr1 localizes to the periphery of
small buds (Shepard et al. 2003). srl1D mutants have no
obvious morphological defects and show modest Calcofluor
White sensitivity at 22, but are hypersensitive to this agent
at 37 (Kurischko et al. 2005). Mutants defective in RAM
function are also suppressed by overexpression of Sim1 (see
above) and Ccw12 (Kurischko et al. 2005), and slr1D and
ccw12D show a strong genetic interaction in the RAMdefective background. The srl1D ccw12D strain is CFW hypersensitive at both 22 and 37, and at 22, but not at 37,
resembles mating pheromone-treated wild-type cells (Kurischko
et al. 2005). Srl1 and Ccw12 have been proposed to have
parallel functions in activation of a CWI pathway that operates when RAM signaling is defective (Kurischko et al.
2005).
What Is Next?
The biosynthesis of most individual yeast wall components
is now understood in much detail and involves conserved
pathways such as N-glycosylation and GPI anchoring and
enzymes represented in other organisms, such as chitin and
b1,3-glucan synthases. In contrast, b1,6-glucan formation
and cross-linkage to GPI proteins and cross-linking of chitin
to b-glucans are clearly restricted to certain yeasts and filamentous fungi, and the enzymes implicated in the latter
processes, as well as certain CWP, are signatures of fungal
cell walls, whose evolution of has been reviewed by RuizHerrera and Ortiz-Castellanos (2010) and Xie and Lipke
(2010).
Much of the work necessary to take both the conserved
and the yeast-specific aspects of wall biogenesis to the next
level must be biochemical and analytical. These efforts will
involve charting new biochemical territory, such as determining how b1,6-glucan is made and defining the functions of wall-active proteins such as Ccw12, Ecm33, Kre1,
and Kre9, which have key roles in wall biogenesis, but show
no resemblance to proteins of known function and may not
be enzymes. Other biochemical challenges are the mechanism and activation of chitin and b1,3-glucan synthases, the
mechanism and conjectured flippase activities of the multispanning glycosyltransferases of the dolichol, O-mannosylation, and
GPI pathways, the functions of the Etn-P side branches on
GPIs, and the biochemical activities of predicted enzymes
such as the Dcw1/Dfg5 pair, Kre5, Kre6, Scw4, and Scw10.
The latter efforts require application of high-resolution techniques to analyze the fine structure and linkages of cell wall
glycans (Magnelli et al. 2002; Aimanianda et al. 2009),
which should highlight the reactions for which biochemists
need to develop assays and screen mutants.
With the identification of so many proteins involved in cell
wall biogenesis, and with ever-improving knowledge of wall
composition, we can look forward to deepening our understanding of the complexities of yeast cell wall biogenesis.
Acknowledgments
I thank my students for their many contributions to GPI and
wall biosynthesis. I also acknowledge the contributions of
S. cerevisiae Cell Wall
803
the late Yoshifumi Jigami to our field. I am grateful to three
reviewers for their helpful comments. Work in my laboratory
has been supported by grant GM-46220 from the National
Institutes of Health and by a Burroughs Wellcome Scholar
Award in Molecular Pathogenic Mycology.
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Communicating editor: J. Thorner
GENETICS
Supporting Information
http://www.genetics.org/lookup/suppl/doi:10.1534/genetics.112.144485/-/DC1
Architecture and Biosynthesis of the Saccharomyces
cerevisiae Cell Wall
Peter Orlean
Copyright © 2012 by the Genetics Society of America
DOI: 10.1534/genetics.112.144485
File S1 Precursors and Carrier Lipids This Supporting File contains additional information related to Precursors and Carrier Lipids. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Sugar nucleotides Regulation of glucosamine supply and chitin levels. Glucosamine supply is highly regulated and impacts chitin levels, which increase in response to mating pheromones and cell wall stress. Expression of GFA1 and AGM1 is upregulated upon treatment of MATa cells with α-­‐factor (Watzele and Tanner, 1989; Hoffman et al. 1994), and is accompanied by an increase in chitin deposition (Schekman and Brawley, 1979; Orlean et al. 1985). The cell wall stress-­‐induced increase in chitin synthesis (Popolo et al. 1997; Dallies et al. 1998; Kapteyn et al. 1999; see Wall Composition and Architecture) is also accompanied by elevated GFA1 expression (Terashima et al. 2000; Lagorce et al. 2002; Bulik et al. 2003). Elevation of glucosamine levels by other means also elicits increased chitin synthesis, for chitin levels are correlated with levels of expression of GFA1 itself (Lagorce et al. 2002; Bulik et al. 2003), and exogenous glucosamine also leads to increased chitin synthesis (Bulik et al. 2003). However, Bulik et al. (2003) found that chitin formation was not proportional to UDP-­‐GlcNAc concentration. These observations led to the conclusion that chitin synthesis is proportional to Gfa1 activity but that additional factors, for example a glucosamine metabolite or Gfa1 itself, must modulate chitin levels (Bulik et al. 2003). It is also formally possible that additional chitin is in a soluble or intracellular form and not detected in cell wall analyses. Dolichol and dolichol phosphate sugars Dolichol phosphate synthesis: Rer2 and Srt1. Biosynthesis of dolichol starts with the extension of trans farnesyl-­‐PP by successive addition of cis-­‐
isoprene units by the homologous cis-­‐prenyltransferases Rer2 and Srt1 (Sato et al. 1999; Schenk et al. 2001b). Rer2 is the dominant activity and makes dolichols with 10-­‐14 isoprene units, whereas dolichols made by Srt1 in cells lacking Rer2 contain 19-­‐22 isoprenes, like mammals. rer2Δ strains have severe defects in growth and in N-­‐ and O-­‐glycosylation, and SRT1 is a high-­‐
copy suppressor of rer2 mutants (Sato et al. 1999). The rer2Δ srt1Δ double null is inviable (Sato et al. 1999). Rer2 and Srt1 both behave as peripheral membrane proteins (Sato et al. 2001; Schenk et al. 2001b), but Rer2 is localized to the ER membrane, whereas Srt1 is detected in “lipid particles” (Sato et al. 2001). P. Orlean 1 SI Dfg10. Dfg10 has a steroid 5α reductase domain, and is responsible for much of the activity that reduces the α-­‐
isoprene unit of polyprenol activity. Both dfg10-­‐100 transposon insertion mutants and dfg10Δ strains underglycosylate carboxypeptidase Y to the same extent, and dolichol levels are decreased by 70% in dfg10-­‐100 cells, with a corresponding increase in unsaturated polyprenol (Cantagrel et al. 2010). The biosynthetic origin of the residual dolichol is not known. Membrane organization of Sec59 dolichol kinase. Sec59 is a multispanning membrane protein whose CTP-­‐binding site is oriented towards the cytoplasm (Shridas and Waechter, 2006). Dolichol chain length specificity of yeast glycosyltransferases and flippases. The enzymes that act after Rer2 and Srt1 can use shorter chain dolichols. Thus, the growth and glycosylation defects of rer2Δ cells can be complemented by expression of the E. coli cis-­‐isoprenyltransferase, which generates C55 isoprenoids, or of the Giardia homologue, which makes C55-­‐60 (Rush et al. 2010; Grabinska et al. 2010). The native glycosyltransferases and flippases must therefore also be able to use shorter chain dolichols as substrates. Dol-­‐P-­‐Man and Dol-­‐P-­‐Glc synthesis: Relationship between Dpm1 and Alg5. Alg5 and Dpm1 are most similar in their N-­‐terminal halves, which contain their GT-­‐A superfamily domain, but diverge in their C-­‐terminal halves. Both are likely to catalyze their reactions at the cytoplasmic face of the ER membrane. Literature Cited Grabinska, K. A., Cui, J., Chatterjee, A., Guan, Z., Raetz, C. R., et al., 2010 Molecular characterization of the cis-­‐prenyltransferase of Giardia lamblia. Glycobiology 20: 824-­‐832. Rush, J. S., Matveev, S., Guan, Z., Raetz, C. R. H., Waechter, C. J. 2010 Expression of functional bacterial undecaprenyl pyrophosphate synthase in the yeast rer2Δ mutant and CHO cells. Glycobiology 20: 1585-­‐1593. Sato, M., Fujisaki, S., Sato, K., Nishimura, Y., Nakano, A., 2001 Yeast Saccharomyces cerevisiae has two cis-­‐prenyltransferases with different properties and localizations. Implication for their distinct physiological roles in dolichol synthesis. Genes Cells 6: 495-­‐506. 2 SI P. Orlean Shridas, P., Waechter, C. J., 2006 Human dolichol kinase, a polytopic endoplasmic reticulum membrane protein with a cytoplasmically oriented CTP-­‐binding site. J. Biol. Chem. 281: 31696-­‐316704. P. Orlean 3 SI File S2 N-­‐glycosylation This Supporting File contains additional information related to Biosynthesis of Wall Components Along the Secretory Pathway, N-­‐glycosylation. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Assembly and transfer of the Dol-­‐PP-­‐linked precursor oligosaccharide: Steps on the cytoplasmic face of the ER membrane: Alg7. The Alg7 GlcNAc-­‐1-­‐P transferase, which carries out the first step in the assembly of the Dol-­‐PP-­‐linked precursor is highly conserved among eukaryotes and has homologues in Bacteria, for example MraY, which catalyzes transfers N-­‐
acetylmuramic acid-­‐pentapeptide from UDP to undecaprenol phosphate in peptidoglycan biosynthesis (Price and Momany, 2005). GlcNAc-­‐1-­‐P transferases such as Alg7 and MraY have multiple transmembrane domains and amino acid residues important for catalysis by members of this protein family lie in cytoplasmic loops (Dan and Lehrman; Price and Momany, 2005). Alg13/Alg14. These proteins function as a heterodimer to transfer the second, β1,4-­‐GlcNAc-­‐linked GlcNAc to Dol-­‐PP-­‐
GlcNAc (Bickel et al. 2005; Chantret et al. 2005; Gao et al. 2005). Soluble Alg13, assigned to GT Family 1, is the catalytic subunit and associates with membrane-­‐spanning Alg14 at the cytosolic face of the ER membranes (Averbeck et al. 2007; Gao et al. 2008). Alg13 and 14 are homologous to C and N-­‐terminal domains, respectively, of the bacterial MurG polypeptide, which adds N-­‐acetylmuramic acid to undecaprenol-­‐PP-­‐GlcNAc in peptidoglycan synthesis (Chantret et al. 2005). Alg1. This β1,4-­‐Man-­‐T, assigned to GT Family 33, transfers the first mannose from GDP-­‐Man to Dol-­‐PP-­‐GlcNAc2 (Couto et al. 1984). Alg2. This protein is a member of GT Family 4. Remarkably, Alg2 has both GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2Man α1,3-­‐Man-­‐T and GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2Man2 α1,6-­‐Man-­‐T activity and successively adds an α1,3-­‐Man and an α1,6 Man to the Dol-­‐PP-­‐
linked precursor (O'Reilly et al. 2006; Kämpf et al. 2009). Alg11. Alg11, also a member of GT Family 4, adds the next two α1,2-­‐linked mannoses (Cipollo et al. 2001; O'Reilly et al. 2006; Absmanner et al. 2010). alg11D mutants are viable though growth-­‐defective, and accumulate Dol-­‐PP-­‐GlcNAc2Man3, as well as some Dol-­‐PP-­‐GlcNAc2Man6-­‐7 (Cipollo et al. 2001; Helenius et al. 2002). The latter are aberrant glycan structures formed when Dol-­‐PP-­‐GlcNAc2Man3 is translocated to the lumen and acted on by lumenal Man-­‐T. 4 SI P. Orlean Heterologous expression and membrane topology of Alg1, Alg2, and Alg11. Alg1, Alg2, and Alg11 are catalytically active when expressed in E. coli (Couto et al. 1984; O'Reilly et al. 2006). The catalytic region of Alg1 is predicted to be cytoplasmic, and experimentally derived models for the membrane topology of Alg2 and Alg11 also place catalytic domains at the cytoplasmic side of the ER membrane (Kämpf et al. 2006; Absmanner et al. 2009), although not all predicted hydrophobic helices in Alg2 and Alg11 span the ER membrane, rather, they lie in its cytoplasmic face. Complex formation by early-­‐acting Alg proteins. There is evidence from analyses by coimmunoprecipitation and size exclusion chromatographic analyses for higher order organization of the proteins involved in the cytoplasmic steps of the yeast dolichol pathway. Alg7, 13, and 14 associate in a hexamer (Noffz et al. 2009). Alg1 forms separate complexes containing either Alg2 and Alg11, although the latter two do not interact with one another (Gao et al. 2004). Formation of these multienzyme complexes may in turn facilitate channeling of Dol-­‐PP-­‐linked intermediates to successive membrane-­‐associated transferases. Transmembrane translocation of Dol-­‐PP-­‐oligosaccharides: After Dol-­‐PP-­‐GlcNAc2Man5 is generated on the cytoplasmic face of the ER membrane, it is somehow translocated to the lumenal side of the membrane where subsequent sugars are transferred from Dol-­‐P-­‐sugars (Burda and Aebi, 1999; Helenius & Aebi, 2002). The presumed Dol-­‐PP-­‐oligosaccharide flippase likely prefers the heptasaccharide as substrate, but the presence of shorter oligosaccharides on proteins in both the alg2-­‐Ts and alg11Δ mutants (Jackson et al. 1989; Cippolo et al. 2001) indicates that truncated oligosaccharides can be translocated as well. The Rft1 protein is a candidate for the protein Dol-­‐PP-­‐GlcNAc2Man5 flippase (Helenius et al. 2002). Strains deficient in Rft1 accumulate Dol-­‐PP-­‐GlcNAc2Man5, but are unaffected in O-­‐mannosylation or in GPI anchor assembly, ruling out a deficiency in Dol-­‐P-­‐Man supply to the ER lumen. Because the few N-­‐glycans chains that were still transferred to the reporter protein carboxypeptidase Y in Rft1-­‐depleted cells were endoglycosidase H sensitive, the activity of Alg3, which adds the α1,3-­‐Man required for substrate recognition by endoglycosidase H, was unaffected. Moreover, high level expression of RFT1 partially suppresses the growth defect of alg11Δ and leads to increased levels of lumenal Dol-­‐PP-­‐GlcNAc2Man6-­‐7 and an increase in carboxypeptidase Y glycosylation, consistent with the notion of enhanced flipping of the suboptimal flippase substrate Dol-­‐PP-­‐
GlcNAc2Man3 (Helenius et al. 2002). However, although the above findings are consistent with Rft1 being the flippase itself, this role could not be demonstrated in biochemical assays for flippase activity, for sealed microsomal vesicles or proteoliposomes depleted of Rft1 retained flippase activity, and in fractionation experiments, flippase activity could be separated from Rft1 (Franck et al. 2008; Rush et al. 2009). P. Orlean 5 SI Lumenal steps in Dol-­‐PP-­‐oligosaccharide assembly: Alg3. This α1,3-­‐Man-­‐T is a member of GT Family 58, and transfers the precursor’s sixth, α1,3-­‐Man from Dol-­‐P-­‐Man, making the glycan sensitive to endoglycosidase H (Aebi et al. 1996; Sharma et al. 2001). Alg3’s Dol-­‐P-­‐Man:Dol-­‐PP-­‐GlcNAc2Man5 Man-­‐T activity can be selectively immunoprecipitated from detergent extracts of membranes (Sharma et al. 2001), providing strong evidence that Alg3 and its yeast homologues in the dolichol and GPI assembly pathways are indeed glycosyltransferases. Alg9 and Alg12. Alg9, a member of GT Family 22, transfers the seventh, α1,2-­‐linked Man to the α1,3-­‐Man added by Alg3 (Burda et al. 1999; Cipollo and Trimble, 2000). Alg12, also a GT22 Family member, next adds the eighth, α1,6-­‐Man to the α1,2-­‐linked Man just added by Alg9 (Burda et al. 1999), whereupon Alg9 acts again to add the ninth Man, in α1,2 linkage, to the α1,6-­‐Man added by Alg12 (Frank and Aebi 2005). The second activity of Alg9 was uncovered in in vitro assays in which alg9Δ and alg12Δ membranes were tested for their ability to elongate acceptor Dol-­‐PP-­‐GlcNAc2Man7 isolated from alg12Δ cells. These experiments established that Alg12 requires prior addition of the seventh Man by Alg9, even though Alg12 does not transfer its Man to that residue, and that the Alg12 reaction precedes Alg9’s second α1,2 mannosyltransfer (Frank and Aebi 2005). Alg6, Alg8, and Alg10. Alg6 and Alg8, members of GT Family 57, act successively to transfer two α1,3-­‐linked glucoses to extend the second α1,2-­‐Man added by Alg11, and lastly, Alg10, assigned to GT Family 59, completes the 14-­‐sugar Dol-­‐PP-­‐
linked oligosaccharide by adding a third, α1,2-­‐Glc (Reiss et al., 1996; Stagljar et al., 1994; Burda and Aebi, 1998). Shared transmembrane topology of Dol-­‐P-­‐sugar-­‐utilizing transferases. The six Dol-­‐P-­‐sugar-­‐utilizing transferases are members of a larger protein family that includes the Dol-­‐P-­‐Man-­‐utilizing Man-­‐T involved in GPI anchor biosynthesis (Oriol et al. 2002). The results of in silico analyses of the sequences of these proteins suggested they have a common membrane topology and 12 transmembrane segments, and a membrane organization recalling that of membrane transporters, which is consistent with the idea that each protein translocates its own Dol-­‐P-­‐linked sugar substrate (Burda and Aebi, 1999; Helenius and Aebi, 2002). It also plausible that these transferases operate in multienzyme complexes to facilitate substrate channeling. Oligosaccharide transfer to protein: Truncated oligosaccharides can be transferred to protein. The results of analyses of the N-­‐linked glycans present on protein in mutants defective in the assembly of the Dol-­‐PP-­‐linked precursor oligosaccharide indicate that a range of structures smaller than GlcNAc2Man9Glc3 can be transferred in vivo. However, full-­‐size Dol-­‐PP-­‐GlcNAc2Man9Glc3 is the preferred OST substrate in vitro, and the observation that mutants that make smaller precursor oligosaccharides have a synthetic phenotype 6 SI P. Orlean with OST mutants indicates the preference exists in vivo as well (Knauer and Lehle, 1999; Zufferey et al. 1995; Reiss et al. 1997; Karaoglu et al. 2001). This preference does not reflect differences between the binding affinities of Dol-­‐PP-­‐GlcNAc2Man9Glc3 and smaller oligosaccharides at the OST active site, rather, it has been proposed that OST has an allosteric site that binds GlcNAc2Man9Glc3 as well as smaller oligosaccharides, in turn activating the catalytic site for GlcNAc2Man9Glc3 and acceptor peptide binding. Binding of a truncated oligosaccharide at the allosteric site, however, enhances GlcNAc2Man9Glc3 binding more strongly, and so ensures preferential utilization of the full-­‐size precursor (Karaoglu et al., 2001; Kelleher and Gilmore, 2006). Purification and protein-­‐protein interactions of OST. Complete heterooctomeric OST complexes have been affinity purified (Karaoglu et al. 1997; Spirig et al. 1997; Karaoglu et al. 2001; Chavan et al. 2006), and the subunits appear to be present in stoichiometric amounts (Karaoglu et al. 1997). The OST complexes themselves may themselves function as dimers (Chavan et al. 2006). The results of genetic interaction studies and coimmunoprecipitation-­‐ and chemical cross-­‐linking experiments suggest the existence of three sub-­‐complexes i) Swp1-­‐Wbp1-­‐Ost2, ii) Stt3-­‐Ost4-­‐Ost3, and iii) Ost1-­‐Ost5 (Spirig et al. 1997; Karaoglu et al. 1997; Reiss et al. 1997; Li et al. 2003; Kim et al. 2003; reviewed by Knauer and Lehle, 1999; Kelleher and Gilmore, 2006). It has been noted, however, that treatment of OST with non-­‐ionic detergents does not yield these three subcomplexes (Kelleher and Gilmore, 2006). Furthermore, additional interactions between OST subunits have been detected using chemical cross-­‐linking approaches and membrane protein two-­‐hybrid analyses (Yan et al. 2003, 2005). OST also interacts with the Sec61 translocon complex and large ribosomal subunit (Chavan et al. 2005; Harada et al. 2009), suggesting that the complex is poised to act on nascent, freshly translocated proteins. However, protein O-­‐mannosyltransferases can compete for the hydroxyamino acids in a freshly translocated sequon (Ecker et al. 2003; see O-­‐mannosylation). Stt3 is the catalytic subunit of OST. There is strong evidence that Stt3, which has a soluble, lumenal domain towards its C-­‐terminus preceded by 11 transmembrane domains (Kim et al. 2005), is the catalytic subunit of OST. First, it can be crosslinked to peptides derivatized with a photoactivatable group and containing an N-­‐X-­‐T glycosylation site, or to nascent polypeptide chains containing the sequon-­‐mimicking, cryptic glycosylation site Q-­‐X-­‐T and a photoactivable side chain (Yan and Lennarz, 2002; Nilson et al. 2003). Second, Stt3 homologues are present in all eukarya, as well as in certain Bacteria and many Archaea, in which diverse types of glycan are transferred to protein (Kelleher and Gilmore, 2006; Kelleher et al. 2007). The Stt3 homologue from Campylobacter jejuni, PglB, was shown to be required for transfer of that bacterium’s characteristic glycan to Asn in a substrate peptide when the C. jejuni pgl gene cluster was heterologously expressed in E. coli (Wicker et al. 2002). Third, Stt3 homologues from the protist Leishmania major, whose proteome contains no other OST subunits, complement the S. P. Orlean 7 SI cerevisiae stt3Δ mutants as well as null mutations in the genes for the essential OST subunits Ost1, Ost2, Swp1, and Wbp1, indicating that the protist Stt3 functions autonomously as an OST (Nasab et al. 2008; Hese et al. 2009). Stt3 has been assigned to GT Family 66. Ost3 and Ost6: role of a thioredoxin domain. The other OST subunits for which catalytic activity has been demonstrated are the paralogues Ost3 and Ost6. ost3Δ ost6Δ double mutants have a more severe glycosylation defect than the single nulls (Knauer and Lehle, 1999b). The two proteins confer a degree of acceptor preference to the OST complexes that contain them (Schulz and Aebi, 2009) because they each have peptide binding grooves lined by amino acids whose side chains are complementary in hydrophobicity and charge to different substrate peptides (Jamaluddin et al. 2011). Ost3 and Ost6 are predicted to have four transmembrane domains at their C-­‐termini and an N-­‐terminal domain containing a thioredoxin fold with the CXXC motif common to proteins involved in disulfide bond shuffling during oxidative protein folding (Kelleher and Gilmore, 2006; Schulz et al. 2009). This domain most likely lies in the lumen (Kelleher and Gilmore, 2006). Mutations of the cysteines in the CXXC motifs of Ost3 and Ost6 lead to site-­‐specific underglycosylation, indicating the importance of the thioreductase motif. This was confirmed by the demonstration that the thioredoxin domain of Ost6, expressed in E. coli, had oxidoreductase activity towards a peptide substrate (Schulz et al. 2009). These findings led to a model in which Ost3/Ost6 form transient disulfide bonds with nascent proteins and promote efficient glycosylation of more Asn-­‐X-­‐Ser/Thr sites by delaying oxidative protein folding (Schulz et al. 2009). Structural analyses of the thioredoxin domain of Ost6 showed that the peptide binding groove is present only when the CXXC motif is oxidized (Jamaluddin et al. 2011). Recruitment of Ost3 or Ost6 to OST requires Ost4, a hydrophobic 36 amino protein (Kim et al. 2000, 2003; Spirig et al. 2005). Ost4 also interacts with Stt3 (Karaoglu et al. 1997; Spirig et al. 1997; Knauer and Lehle, 1999; Kim et al. 2003). ost4Δ strains are temperature-­‐sensitive and severely underglycosylate protein (Chi et al. 1996). Possible roles for other OST subunits. A sub-­‐complex of Swp1p, Wbp1p, and Ost2p, has been suggested to confer the preference for GlcNAc2Man9Glc3, possibly by providing the allosteric site (Kelleher and Gilmore, 2006). Evidence for a role of complex subunits other than Stt3 was obtained with Trypanosoma cruzi Stt3, which transfers GlcNAc2Man7-­‐9 to protein in vitro as efficiently as it does glucosylated oligosaccharides. When expressed in S. cerevisiae in place of native Stt3, trypanosomal Stt3 now preferentially transferred GlcNAc2Man9Glc3 to protein in vitro and in vivo (Castro et al. 2006). Similarly, when Leishmania Stt3 is expressed in the context of the other S. cerevisiae OST subunits, the Leishmania protein acquires a preference for transferring glucosylated oligosaccharides, rather than the non-­‐glucosylated oligosaccharides that it transfers in the protist itself (Hese et al. 2009). Wbp1 may be involved in recognition of Dol-­‐PP-­‐GlcNAc2Man9Glc3, because alkylation of a key cysteine 8 SI P. Orlean residue in this subunit inactivates OST, whereas inactivation is prevented by prior incubation with Dol-­‐PP-­‐GlcNAc2 (Pathak et al. 1995). The protein’s single transmembrane domain contains sequences important for incorporation into the OST complex, possibly by making interactions with Ost2 and Swp1 (Li et al. 2003). Other than their membership in proposed OST subcomplexes and interactions with other OST subunits, little is known about the function of Swp1, Ost1, Ost2, and Ost5, although it has been suggested that Ost1 has a role in funneling nascent polypeptides to Stt3 (Lennarz, 2007). Regulation of OST by the CWI pathway. Oligosaccharyltransferase may be regulated by the PKC-­‐dependent CWI pathway or by Pkc1 itself, a notion that arose from the identification of STT3 in a screen for mutants sensitive to the PKC inhibitor staurosporine and to elevated temperature (Yoshida et al. 1995). Although this suggested that adequate levels of N-­‐
glycosylation are needed for cells to overcome defects in CWI signaling, staurosporine sensitivity proved not to be a general consequence of deficient N-­‐glycosylation, because only a subset of stt3 alleles were sensitive to the drug, and mutants in most other OST subunits, with the exception of Ost4, were resistant (Chavan et al. 2003; Levin, 2005). A more direct link between Stt3 and the Pkc1-­‐dependent signaling emerged from the findings that STT3 mutations that lead to staurosporine sensitivity are located in N-­‐terminal, predicted cytosolic domains of Stt3, and that pkc1Δ mutants have half of wild type OST activity in vitro (Chavan et al. 2003; Park and Lennarz, 2000). This led to the suggestion that CWI pathway regulates OST via an interaction between Pkc1 or components of the PKC pathway with the N-­‐terminal domain of Stt3, and perhaps Stt3-­‐interacting Ost4 as well (Chavan et al. 2003). N-­‐glycan processing in the ER and glycoprotein quality control: Glucosidase II. This is a heterodimer of catalytic Gls2/Rot2 and Gtb1, the latter of which is necessary for, and influences the rate of, Glc trimming (Trombetta et al. 1996; Wilkinson et al., 2006; Quinn et al. 2009). Glycoprotein recognition by Pdi1 and the Pdi1-­‐Htm1 complex. Unfolded or misfolded proteins are bound by protein disulfide isomerase Pdi1, a subset of which is in complex with Mns1 homolog Htm1. A stochastic model has been proposed in which both Pdi1 and the Pdi1-­‐Htm1 complex recognize un-­‐ or misfolded proteins, but persistently misfolded proteins stand an increased chance of encountering Pdi1-­‐Htm1 whose Htm1 component trims a Man from N-­‐linked glycans, yielding a GlcNAc2Man7 structure bearing a terminal α1,6 Man (Clerc et al. 2009; Gauss et al. 2011).
Mannan elaboration in the Golgi: Formation of core type N-­‐glycan and mannan outer chains: P. Orlean 9 SI Elucidation of the pathway for formation of mannan outer chains. Two groups of proteins, the Mnn9/Anp1/Van1 trio, and the Mnn10 and Mnn11 pair, had been implicated in formation of the poly-­‐α1,6-­‐linked mannan backbone, but because strains deficient in these proteins retained mannosyltransferase activity and still made mannan containing α1,6 linkages, these proteins were considered more likely to affect mannan formation indirectly (reviewed by Orlean, 1997; Dean, 1999). Two key sets of findings led to clarification of mannan biosynthesis. First, co-­‐immunoprecipitation and colocalization experiments established that Mnn9, Anp1, and Van1 occurred in two different protein complexes in the cis-­‐Golgi, one containing Mnn9 and Van1 (subsequently named M-­‐Pol I), the other, Mnn9, Anp1, Hoc1 (homologous to Och1), and the related Mnn10 and Mnn11 proteins (M-­‐Pol II) (Hashimoto and Yoda, 1997; Jungmann and Munro, 1998; Jungmann et al. 1999). Second, both immunoprecipitated protein complexes had α1,6 mannosyltransferase activity, indicating that one or more of the Mnn9/Anp1/ Van1 group was an α1,6 mannosyltransferase (Jungmann and Munro, 1998; Jungmann et al. 1999). Consistent with their being glycosyltransferases, all five proteins have the GT-­‐A fold protein topology and a “DXD motif” common to enzymes that have sugar nucleotides as donors and use the aspartyl carboxylates to coordinate divalent cations and the ribose of the donor (Wiggins and Munro, 1998; Lairson et al. 2008). The contributions of the individual subunits to α1,6 mannan synthesis by each complex, and the roles of the two complexes in mannan formation, were explored in deletion mutants and in point mutants abolishing catalytic activity but otherwise preserving complex stability. The sizes of the mannans and the residual in vitro activities of the M-­‐Pol complexes in these mutants led to the current model for mannan synthesis (Jungmann et al. 1999; Munro, 2001; Figure 3 in main text). In it, M-­‐Pol I, a heterodimer, acts first to extend the Och1-­‐derived Man with further α1,6-­‐linked mannoses. Analyses of mutants in the DXD motifs of Mnn9 and Van1 indicated that Mnn9 likely adds the first α1,6-­‐liked Man, which is extended with 10-­‐15 α1,6 mannoses in Van1-­‐requiring reactions (Stolz and Munro, 2002; Rodionov et al. 2009). This α1,6 backbone is then elongated with 40-­‐60 α1,6 Man by M-­‐Pol II. Assays of M-­‐Pol ll from strains lacking Mnn10 or Mnn11 indicated that these proteins are responsible for the majority of the α1,6 mannosyltransferase activity in that complex (Jungmann et al., 1999). The contribution of Hoc1, a homologue of the Och1 α1,6-­‐Man-­‐T is not clear, for HOC1 deletion neither alters M-­‐Pol II activity nor impacts mannan size. Localization of Och1 and Man-­‐Pol complexes. The localization dynamics of Mnn9-­‐containing M-­‐Pol complexes and Och1 seem inconsistent with the order in which they act in mannan assembly, with Mnn9 showing a steady state localization in the cis-­‐Golgi and continuously cycling between that compartment and the ER, but with Och1 cycling between the ER and cis-­‐ and trans-­‐Golgi (Harris and Waters, 1996; Todorow et al. 2000; Karhinen and Makarow, 2004). It has been suggested that 10 SI P. Orlean substrate specificity, rather than transferase localization, determines their order in which the enzymes act (Okamoto et al. 2008). The size of N-­‐linked mannan can be impacted by deficiencies in proteins required for localization of Golgi mannosyltransferases. For example, deletion of VPS74, also identified as MNN3, eliminates a protein that interacts with the cytoplasmic tails of certain transferases normally resident in the cis and medial Golgi compartments. The resulting mislocalization of several mannosyltransferases would explain the underglycosylation phenotype of mnn3 mutants (Schmitz et al. 2008; Corbacho et al. 2010). Mutations in SEC20, which encodes a protein involved in Golgi to ER retrograde transport, also result in diminished Golgi mannosyltransferase activity, even though this glycosylation defect is not correlated with the secretory pathway defect (Schleip et al. 2001). The reason for this is not clear. Mannan side branching and mannose phosphate addition: Roles of the Ktr1 Man-­‐T family members in mannan side branching. Five members of the Ktr1 family of Type II membrane proteins, Kre2/Mnt1, Yur1, Ktr1, Ktr2, Ktr3, also contribute to N-­‐linked outer chain synthesis, as judged by the impact of null mutations on the mobility of reporter proteins (Lussier et al. 1996; 1997a; 1999). Of these proteins, Kre2/Mnt1, Ktr1, Ktr2, and Yur1 have been shown to have α1,2 Man-­‐T activity. These Ktr1 family members, perhaps along with uncharacterized homologues Ktr4, Ktr5, and Ktr7 (Lussier et al. 1999) have a collective role in adding the second, and perhaps subsequent α1,2-­‐mannoses to mannan side branches. Members of the Ktr1 family have been assigned to GT Family 15. Addition and function of mannose phosphate. Both core type N-­‐glycans and mannan can be modified with mannose phosphate on α1,2-­‐linked mannoses in the context of an oligosaccharide containing at least one α1,2-­‐linked mannobiose structure. Mannose phosphates confer a negative charge, an attribute exploited early on to isolate mannan synthesis mutants on the basis of their inability to bind the cationic dye Alcian Blue (Ballou, 1982; 1990). Mnn6/Ktr6, a member of the Ktr1 family, is the major activity responsible for transferring Man-­‐1-­‐P from GDP-­‐Man to both mannan outer chains and, in vitro, to core N-­‐
glycans, generating GMP. However, because deletion of MNN6 did not eliminate in vivo mannose phosphorylation in och1Δ strains that make only core type N-­‐glycans, additional, as yet unidentified, core phosphorylating proteins must exist (Wang et al. 1997; Jigami and Odani, 1999). The Mnn4 protein is also involved in Man-­‐P addition, but its role differs from Mnn6’s in that deletion of Mnn4 reduces Man-­‐P on core-­‐type glycans (Odani et al. 1996). Mnn4 does not resemble glycosyltransferases, but does have a LicD domain found in nucleotidyltransferases and phosphotransferases involved in lipopolysaccharide synthesis. The mnn4Δ mutation is dominant, and Mnn4 has been proposed to have a positive regulatory role (Jigami and Odani, 1999). Levels of mannan phosphorylation are highest in the late log and stationary phases, when MNN4 expression is elevated (Odani et al. 1997). Transcriptional regulation may involve the RSC chromatin remodeling complex because strains lacking Rcs14, a P. Orlean 11 SI subunit of that complex, show drastically reduced Alcian Blue binding and down-­‐regulated expression of MNN4 and MNN6 (Conde et al. 2007).
A Golgi GlcNAc-­‐T. S. cerevisiae also has the capacity to add GlcNAc to the non-­‐reducing end of N-­‐linked glycans. Heterologously expressed lysozyme received a GlcNAc2Man8-­‐12 glycan additionally bearing a GlcNAc residue, and the responsible GlcNAc transferase proved to be Gnt1, whose localization mostly coincides with that of Mnn1 in the medial Golgi (Yoko-­‐o et al. 2003). GNT1 disruptants have no discernible phenotype, and Gnt1 may rarely act on native yeast glycans; its activity would require that UDP-­‐GlcNAc be transported into the Golgi lumen (Yoko-­‐o et al. 2003). Literature Cited Averbeck, N., Keppler-­‐Ross, S., Dean, N., 2007 Membrane topology of the Alg14 endoplasmic reticulum UDP-­‐GlcNAc transferase subunit. J. Biol. Chem. 282: 29081-­‐29088. Castro, O., Movsichoff, F., Parodi, A. J., 2006 Preferential transfer of the complete glycan is determined by the oligosaccharyltransferase complex and not by the catalytic subunit. Proc. Natl. Acad. Sci. USA. 103: 14756-­‐14760. Chavan, M., Yan, A., Lennarz, W. J. 2005 Subunits of the translocon interact with components of the oligosaccharyl transferase complex. J. Biol. Chem. 280: 22917–22924. Chi, J. H., Roos, J., Dean, N., 1996 The OST4 gene of Saccharomyces cerevisiae encodes an unusually small protein required for normal levels of oligosaccharyltransferase activity. J. Biol. Chem. 271: 3132–3140. Conde, R., Cueva, R., Larriba, G., 2007 Rsc14-­‐controlled expression of MNN6, MNN4 and MNN1 regulates mannosylphosphorylation of Saccharomyces cerevisiae cell wall mannoproteins. FEMS Yeast Res. 7: 1248-­‐1255. Corbacho, I., Olivero, I., Hernández, M., 2010 Identification of the MNN3 gene of Saccharomyces cerevisiae. Glycobiology 20: 1336-­‐1340. 12 SI P. Orlean Dan, N., Lehrman, M. A., 1997 Oligomerization of hamster UDP-­‐GlcNAc:dolichol-­‐P GlcNAc-­‐1-­‐P transferase, an enzyme with multiple transmembrane spans. J. Biol. Chem. 272: 14214-­‐14219. Dean, N. 1999 Asparagine-­‐linked glycosylation in the yeast Golgi. Biochim. Biophys. Acta 1426: 309–322. Gao, X. D., Moriyama, S., Miura, N., Dean, N., Nishimura, S., 2008 Interaction between the C termini of Alg13 and Alg14 mediates formation of the active UDP-­‐N-­‐acetylglucosamine transferase complex. J. Biol. Chem. 283: 32534-­‐32541. Harada, Y., Li, H., Li, H., Lennarz, W. J., 2009 Oligosaccharyltransferase directly binds to ribosome at a location near the translocon-­‐binding site. Proc. Natl. Acad. Sci. USA 106: 6945-­‐6949. Harris, S. L., Waters, M. G., 1996 Localization of a yeast early Golgi mannosyltransferase, Och1p, involves retrograde transport. J. Cell Biol. 132: 985-­‐998. Jackson, B. J., Warren, C. D., Bugge, B., Robbins, P. W., 1989 Synthesis of lipid-­‐linked oligosaccharides in Saccharomyces cerevisiae: Man2GlcNAc2 and Man1GlcNAc2 are transferred from dolichol to protein in vivo. Arch. Biochem. Biophys. 272: 203-­‐
209. Jamaluddin, M. F., Bailey, U. M., Tan, N. Y., Stark, A. P., Schulz, B. L., 2011 Polypeptide binding specificities of Saccharomyces cerevisiae oligosaccharyltransferase accessory proteins Ost3p and Ost6p. Protein Sci. 20: 849-­‐555. Karaoglu, D., Kelleher, D. J., Gilmore, R., 2001 Allosteric regulation provides a molecular mechanism for preferential utilization of the fully assembled dolichol-­‐linked oligosaccharide by the yeast oligosaccharyltransferase. Biochemistry: 40: 12193–12206. Karhinen, L., Makarow, M., 2004 Activity of recycling Golgi mannosyltransferases in the yeast endoplasmic reticulum. J. Cell Sci. 117: 351-­‐358. P. Orlean 13 SI Kim, H., von Heijne, G., Nilsson, I., 2005 Membrane topology of the STT3 subunit of the oligosaccharyl transferase complex. J. Biol. Chem. 280: 20261-­‐20267. Lairson, L. L., Henrissat, B., Davies, G. J., Withers, S. G., 2008 Glycosyltransferases: structures, functions, and mechanisms. Annu. Rev. Biochem. 77: 521-­‐555. Munro, S., 2001 What can yeast tell us about N-­‐linked glycosylation in the Golgi apparatus? FEBS Lett. 498: 223-­‐227. Okamoto, M., Yoko-­‐o, T., Miyakawa T., Jigami, Y., 2008 The cytoplasmic region of α-­‐1,6-­‐mannosyltransferase Mnn9p is crucial for retrograde transport from the Golgi apparatus to the endoplasmic reticulum in Saccharomyces cerevisiae. Eukaryot. Cell 7: 310-­‐318. Price, N. P., Momany, F. A., 2005. Modeling bacterial UDP-­‐HexNAc: polyprenol-­‐P HexNAc-­‐1-­‐P transferases. Glycobiology 15: 29R-­‐42R. Schleip, I., Heiss, E., Lehle, L., 2001 The yeast SEC20 gene is required for N-­‐ and O-­‐glycosylation in the Golgi. Evidence that impaired glycosylation does not correlate with the secretory defect. J. Biol. Chem. 276: 28751-­‐28758. Schmitz, K. R., Liu, J. X., Li, S. L., Setty T. G., Wood, C. S., et al., 2008 Golgi localization of glycosyltransferases requires a Vps74p oligomer. Dev. Cell 14: 523-­‐534. Todorow, Z., Spang, A., Carmack, E., Yates, J., Schekman, R., 2000 Active recycling of yeast Golgi mannosyltransferase complexes through the endoplasmic reticulum. Proc. Natl. Acad. Sci. USA. 97: 13643-­‐13548. Wiggins, C. A., Munro, S., 1998 Activity of the yeast MNN1 α-­‐1,3-­‐mannosyltransferase requires a motif conserved in many other families of glycosyltransferases. Proc. Natl. Acad. Sci. USA. 95: 7945-­‐7950. 14 SI P. Orlean Yan, A., Ahmed, E., Yan, Q., Lennarz, W. J., 2003 New findings on interactions among the yeast oligosaccharyl transferase subunits using a chemical cross-­‐linker. J. Biol. Chem. 278: 33078–33087. Yan, A., Wu. E., Lennarz, W. J., 2005 Studies of yeast oligosaccharyl transferase subunits using the split-­‐ubiquitin system: topological features and in vivo interactions. Proc. Natl. Acad. Sci. USA 102: 7121–7126. Yoko-­‐o, T., Wiggins, C. A., Stolz, J., Peak-­‐Chew, S. Y., Munro, S., 2003 An N-­‐acetylglucosaminyltransferase of the Golgi apparatus of the yeast Saccharomyces cerevisiae that can modify N-­‐linked glycans. Glycobiology 13: 581-­‐589. Yoshida, S., Ohya, Y., Nakano, A., Anraku, Y., 1995. STT3, a novel essential gene related to the PKC1/STT1 protein kinase pathway, is involved in protein glycosylation in yeast. Gene 164: 167-­‐172. Zufferey, R., Knauer, R., Burda, P., Stagljar, I., te Heesen, S., et al., 1995 STT3, a highly conserved protein required for yeast oligosaccharyl transferase activity in vivo. EMBO J. 14: 4949-­‐4960. P. Orlean 15 SI File S3 O-­‐Mannosylation This Supporting File contains additional information related to Biosynthesis of Wall Components Along the Secretory Pathway, O-­‐mannosylation. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Protein O-­‐mannosyltransferases in the ER: Substrate proteins for different Pmt complexes. Analyses of glycosylation of individual proteins in pmtΔ strains showed that Pmt1/Pmt2 complexes are primarily involved in O-­‐mannosylation of Aga2, Bar1, Cts1, Kre9, and Pir2, whereas homodimeric Pmt4 modifies Axl2, Fus1, Gas1, Kex2 (Gentzsch and Tanner 1997; Ecker et al. 2003; Proszynski et al. 2004; Sanders et al. 1999). However, some proteins, including Mid2, the WSC proteins, and Ccw5, are modified by both complexes, although the Pmt1/Pmt2 and Pmt4/Pmt4 dimers modify different domains of these target proteins (Ecker et al. 2003; Lommel et al. 2004). Mutations in substrate proteins can cause them to be O-­‐mannosylated by a different PMT, and PMTs can also have a role in quality control of protein folding in the ER (see N-­‐glycan processing in the ER and glycoprotein quality control). Thus, wild type Gas1 is normally O-­‐mannosylated by Pmt4, whereas Gas1
G291R
, a model misfolded protein, is hypermannosylated by Pmt1-­‐
Pmt2 as well as targeted to the HRD-­‐ubiquitin ligase complex for degradation by the ERAD system (Hirayama et al. 2008; Goder and Melero, 2011). The latter, chaperone-­‐like function of Pmt1-­‐Pmt2 may be distinct from Pmt1-­‐Pmt2’s O-­‐mannosyltransferase activity (Goder and Melero, 2011). Extension and phosphorylation of O-­‐linked manno-­‐oligosaccharide chains: Extension with α-­‐linked mannoses. The Ser-­‐ or Thr-­‐linked Man is extended with up to four α-­‐linked Man that are added by GDP-­‐Man-­‐dependent Man-­‐T of the Ktr1 and Mnn1 families (Lussier et al. 1999; Figure 4 in main text). The contributions of these proteins was deduced from the sizes of the O-­‐linked chains that accumulated in strains in which Man-­‐T genes had been deleted singly or in different combinations. Transfer of the first two α1,2-­‐Man is carried out by Ktr1 sub-­‐family members Ktr1, Ktr3, and Kre2, which have overlapping roles in the process, although Kre2 has the dominant role in addition of the second, α1,2-­‐Man (Lussier et al. 1997a). The major O-­‐linked glycan made in the ktr1Δ ktr3Δ kre2Δ triple mutant consists of a single Man (Lussier et al. 1997a). Ktr1, Ktr3, and Kre2 are also involved in making α1,2-­‐branches to mannan outer chains (see Mannan elaboration in the Golgi). 16 SI P. Orlean Extension of the trisaccharide chain with one or two α1,3-­‐linked Man is the shared responsibility of Mnn1 family members Mnn1, Mnt2, and Mnt3, with Mnn1 having the major role in adding the fourth Man but Mnt2 and Mnt3 dominating when the fifth is added (Romero et al. 1999). Mnn1 also transfers Man to N-­‐linked outer chains. The α1,2 Man-­‐T have been localized to the medial Golgi, and the Mnn1 α1,3 Man-­‐T to the medial and trans-­‐Golgi (Graham et al. 1994). Because protein-­‐
bound O-­‐mannosyl glycans pulse-­‐labeled in mutants defective in ER to Golgi transport such as sec12, sec18, and sec20 contain two, sometimes more mannoses, GDP-­‐Man-­‐dependent O-­‐glycan extension can occur at the level of the ER (Haselbeck and Tanner, 1983; Zueco et al. 1986; D'Alessio et al. 2005). The process is independent of nucleotide sugar diphosphatases (see Sugar nucleotide transport; D'Alessio et al. 2005), but presumably mediated in the ER by Man-­‐T en route to the Golgi. Importance and function of O-­‐mannosyl glycans: Importance of O-­‐mannosylation for function of specific proteins. Analyses of single and conditionally lethal double pmt mutants show that O-­‐mannosylation can be important for function of individual O-­‐mannosylated proteins. For example, pmt4Δ haploids show a unipolar, rather than the normal axial budding pattern, which is due to defective O-­‐mannosylation and resulting instability and mislocalization of Axl2, which normally marks the axial budding site (Sanders et al. 1999). Pmt4-­‐initiated O-­‐mannosylation is also necessary for cell surface delivery of Fus1, because the unglycosylated protein accumulates in the late Golgi (Proszynski et al. 2004). Defects in Pmt4-­‐dependent O-­‐glycosylation of Msb2 (as well as N-­‐glycosyation) of osmosensor Msb2 lead to activation of the filamentous growth signaling pathway (Yang et al. 2009). In this case, underglycosylation may unmask a domain that normally is exposed and makes interactions when the signaling pathway is activated legitimately. O-­‐
mannosylation of Wsc1, Wsc2, and Mid2 is necessary for these Type I membrane proteins to fulfill their functions as sensors that activate the CWI pathway. Underglycosylation of the CWI pathway-­‐triggering mechanosensor Wsc1 in a pmt4Δ mutant eliminates the stiffness of this rod-­‐like glycoprotein and abolishes its “nanospring” properties, impairing Wsc1’s function as a mechanosensor (Dupres et al. 2009). Further, in pmt2Δ pmt4Δ mutants, which, like CWI pathway mutants, require osmotic stabilization, deficient O-­‐mannosylation results in incorrect proteolytic processing and instability of the sensors (Philip and Levin, 2001; Lommel et al. 2004). Literature Cited D'Alessio, C., Caramelo, J. J., Parodi, A. J., 2005 Absence of nucleoside diphosphatase activities in the yeast secretory pathway does not abolish nucleotide sugar-­‐dependent protein glycosylation. J. Biol. Chem. 280: 40417-­‐40427. P. Orlean 17 SI Dupres, V., Alsteens, D., Wilk, S., Hansen, B., Heinisch, J. J., Dufrêne, Y. F. 2009 The yeast Wsc1 cell surface sensor behaves like a nanospring in vivo. Nat. Chem. Biol. 5: 857-­‐862. Gentzsch, M., Tanner, W., 1997 Protein-­‐O-­‐glycosylation in yeast: protein-­‐specific mannosyltransferases. Glycobiology 7: 481-­‐
486. Goder, V., Melero, A., 2011 Protein O-­‐mannosyltransferases participate in ER protein quality control. J. Cell Sci. 124: 144-­‐153. Graham, T. R., Seeger, M., Payne, G. S., MacKay, V. L., Emr, S. D., 1994 Clathrin-­‐dependent localization of α1,3 mannosyltransferase to the Golgi complex of Saccharomyces cerevisiae. J. Cell Biol. 127: 667-­‐678. Haselbeck, A., Tanner, W., 1983 O-­‐glycosylation in Saccharomyces cerevisiae is initiated at the endoplasmic reticulum. FEBS Lett. 158: 335-­‐338. Hirayama, H., Fujita, M., Yoko-­‐o, T., Jigami, Y., 2008 O-­‐mannosylation is required for degradation of the endoplasmic reticulum-­‐
associated degradation substrate Gas1*p via the ubiquitin/proteasome pathway in Saccharomyces cerevisiae. J. Biochem. 143: 555-­‐567. Philip, B., 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. Proszynski, T. J., Simons, K., Bagnat, M., 2004 O-­‐Glycosylation as a sorting determinant for cell surface delivery in yeast. Mol. Biol. Cell 15: 1533-­‐1543. 18 SI P. Orlean Sanders, S. L., Gentzsch, M., Tanner, W., Herskowitz, I., 1999 O-­‐glycosylation of Axl2/Bud10p by Pmt4p is required for its stability, localization, and function in daughter cells. J. Cell Biol. 145: 1177-­‐1188. Yang, H. Y., Tatebayashi, K., Yamamoto, K., Saito, H., 2009 Glycosylation defects activate filamentous growth Kss1 MAPK and inhibit osmoregulatory Hog1 MAPK. EMBO J. 28: 1380-­‐1389. Zueco, J., Mormeneo, S., Sentandreu, R., 1986 Temporal aspects of the O-­‐glycosylation of Saccharomyces cerevisiae mannoproteins. Biochim. Biophys. Acta 884: 93-­‐100. P. Orlean 19 SI File S4 GPI anchoring This Supporting File contains additional information related to Biosynthesis of Wall Components Along the Secretory Pathway, GPI anchoring. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Assembly of the GPI precursor and its attachment to protein in the ER: Steps on the cytoplasmic face of ER membrane: Gpi3. Gpi3 is a member of GT Family 4 and has an EX7E motif conserved in a range of glycosyltransferases (Coutinho et al. 2003). Mutational analyses indicate that the glutamates are be important for function of Gpi3 and certain EX7E motif glycosyltransferases, although the comparative importance of the two glutamates varies between different transferases (Kostova et al. 2003). However, in the case of Alg2, the EX7E motif is not important for protein function (Kämpf et al. 2009). Formation of GlcNAc-­‐PI by GPI-­‐GnT. The acyl chains of the PI species that receive are the same length as those in other membrane phospholipids (Sipos et al. 1997). Evidence that GlcNAc transfer occurs at the cytoplasmic face of the ER membrane is that i) the catalytic domain of Gpi3’s human orthologue faces the cytoplasm (Watanabe et al. 1996; Tiede et al. 2000), and ii) GlcNAc-­‐PI can be labeled with membrane topological probes on the cytoplasmic side of the mammalian ER membrane (Vidugiriene and Menon, 1993). Significance of Ras2 regulation of GPI-­‐GnT. A clue to the significance of Ras2 regulation of GPI-­‐GnT came from the observation that conditional mutants in GPI-­‐GnT subunits show the phenotype of hyperactive Ras mutants, filamentous growth and invasion of agar. This led to the suggestion that Ras2-­‐mediated modulation of GPI synthesis may be involved in the cell wall and morphogenetic changes that occur in the dimorphic transition to filamentous growth (Sobering et al. 2003; 2004). Location of GlcNAc-­‐PI de-­‐N-­‐acetylation. The de-­‐acetylase reaction likely occurs at the cytoplasmic face of the ER membrane, because the bulk of Gpi12’s mammalian orthologue is cytoplasmic, and because newly synthesized GlcN-­‐PI is accessible on the cytoplasmic face of intact ER vesicles (Vidugiriene and Menon, 1993). Transmembrane translocation of GlcN-­‐PI. GlcN-­‐PI is the precursor species most likely to be translocated to the lumenal side of the ER membrane. Flipping of GlcN-­‐PI as well as GlcNAc-­‐PI has been reconstituted in rat liver microsomes, but the protein involved has not been identified, and the possibility has been raised that GlcN-­‐PI translocation may be mediated by a generic ER phospholipid flippase (Vishwakarma and Menon, 2006). 20 SI P. Orlean Lumenal steps in GPI assembly: Inositol acylation. The acyl chain transferred to GlcN-­‐(acyl)PI in vivo is likely palmitate, although a range of different acyl chains can be transferred from their corresponding CoA derivatives in vitro (Costello and Orlean, 1992; Franzot and Doering, 1999). Because mutants blocked in formation of all mannosylated GPIs accumulated inositol-­‐acylated GlcN-­‐PI (Orlean, 1990; Costello and Orlean, 1992), and because mannosylated GPI intermediates lacking an inositol acyl chain have not been reported, it is likely that inositol acylation precedes mannosylation in vivo. Gwt1, the acyltransferase, is likely to be catalytic because its affinity-­‐purified mammalian orthologue transfers palmitate from palmitoyl CoA to a dioctanoyl analogue of GlcN-­‐PI (Murakami et al. 2003). The protein has 13 transmembrane domains (Murakami et al. 2003; Sagane et al. 2011), and amino acid residues critical for function all face the lumen, indicating acyl transfer is a lumenal event (Sagane et al. 2011), although it is not yet known how acyl CoAs enter the ER lumen. Despite Gwt1’s multispanning topology, the possibility that this inositol acyltransferase is also a GlcN-­‐PI transporter is unlikely, because non-­‐acylated, mannosylated GPIs can be formed in cell lines deficient in Gwt1’s mammalian orthologue (Murakami et al. 2003). GPI Man-­‐T-­‐I. The α1,4-­‐Man-­‐T Gpi14 shows greatest similarity to Alg3, is predicted to have 12 transmembrane segments (Oriol et al. 2002), and is assigned to GT Family 50. Two additional proteins, Arv1 and Pbn1, are involved in the GPI-­‐
Man-­‐T-­‐I step along with Gpi14. arv1Δ cells grow at 30°C but not at 37°C, and are delayed in ER to Golgi transport of GPI-­‐
anchored proteins, and accumulate GlcN-­‐(acyl)PI in vitro (though not in vivo) (Kajiwara et al. 2008). Further, their temperature sensitivity is suppressed by overexpression the genes for most of the subunits of GPI-­‐GnT, suggesting a functional link between ARV1 and GPI assembly (Kajiwara et al. 2008). However, arv1Δ cells were not defective in Dol-­‐P-­‐Man synthase activity or in N-­‐
glycosylation, nor were mild detergent-­‐treated arv1Δ membranes defective in GPI-­‐Man-­‐T-­‐I activity, suggesting that Arv1 is not a Dol-­‐P-­‐Man flippase or directly involved in mannosyltransfer, and leading to the proposal that Arv1 is involved in delivering GlcN-­‐(acyl)PI to GPI-­‐Man-­‐T-­‐I (Kajiwara et al. 2008). Essential Pbn1 has been implicated at the GPI-­‐Man-­‐T-­‐I step in yeast because expression of both GPI14 and PBN1 is necessary to complement mammalian cell lines defective in Pbn1’s mammalian homologue Pig-­‐X, and likewise, co-­‐expression of PIG-­‐X and the gene for Gpi14’s mammalian homologue, PIG-­‐M, partially rescues the lethality of gpi14Δ (Ashida et al. 2005; Kim et al. 2007). Repression of PBN1 expression leads to accumulation of some of the ER form of the GPI protein Gas1, a phenotype seen in GPI precursor assembly mutants (Subramanian et al. 2006). However, it has not been reported whether pbn1 mutants accumulate the predicted GPI intermediate GlcN-­‐(acyl)PI. Because Pbn1 is also involved in processing a number of non-­‐GPI proteins that pass though the ER to the vacuole, the vacuolar membrane, and the plasma membrane, it must have additional functions in the ER (Subramanian et al. 2006). P. Orlean 21 SI GPI Man-­‐T-­‐II. Unlike the other Dol-­‐P-­‐Man-­‐utilizing transferases of the GPI assembly and dolichol pathways, the α1,6-­‐
Man-­‐T Gpi18 is predicted to have 8 transmembrane domains (Fabre et al. 2005; Kang et al. 2005). This protein and its orthologues have been assigned to GT Family 76. GPI Man-­‐T-­‐III and IV. These two α1,2-­‐Man-­‐T, together with their homologues in the dolichol pathway, Alg9 and Alg12, are predicted to have 12 transmembrane domains and are assigned to GT Family 22 (Oriol et al. 2002). Overexpression of GPI10 does not rescue the lethal smp3Δ null mutation, and vice versa, indicating that the two α1,2-­‐Man-­‐T have very strict acceptor specificities (Grimme et al. 2001). Phosphoethanolamine addition: origin of Etn-­‐P from Ptd-­‐Etn. There is good evidence that the Etn-­‐Ps, at least those on Man-­‐1 and Man3, originate from Ptd-­‐Etn. Yeast mutants unable to make CDP-­‐Etn or CDP-­‐Cho from exogenously supplied Etn, 3
but still capable of making Ptd-­‐Etn by decarboxylation of Ptd-­‐Ser, do not incorporate [ H]Etn into protein-­‐bound GPIs or into a 3
Man2-­‐GPI precursor that otherwise receives Etn-­‐P on Man-­‐1. However, radioactivity supplied as [ H]Ser is incorporated into the 3
Man2-­‐GPI after formation and decarboxylation of Ptd-­‐[ H]Ser (Menon and Stevens, 1992; Imhoff et al. 2000). The importance of Ptd-­‐Ser decarboxylation for GPI anchoring is underscored by the finding that the combination of a conditional gpi13 mutation, defective in the EtnP-­‐T-­‐III, with psd1Δ and psd2Δ, nulls in the two Ptd-­‐Ser decarboxylase genes, are inviable (Toh-­‐e and Oguchi, 2002). Direct transfer of Etn-­‐P from Ptd-­‐Etn to a GPI remains to be demonstrated in vitro. Phosphoethanolamine addition: importance of the alkaline phosphatase domain of Mcd4, Gpi7, and Gpi13. These three proteins all have a large lumenal loop of some 400 amino acids that contains sequences characteristic of the alkaline phosphatase superfamily (Gaynor et al. 1999; Benachour et al. 1999, Galperin and Jedrzejas, 2001), consistent with 227
involvement in formation or cleavage of a phosphodiester. This domain is important for function, because the G
E substitution that results in temperature-­‐sensitive growth and a conditional block in GPI precursor assembly in the mcd4-­‐174 mutant (Gaynor et al. 1999) lies in one of the two metal-­‐binding sites in alkaline phosphatase family members (Galperin and Jedrzejas, 2001). The metal is commonly zinc, and in vitro Etn-­‐P addition from an endogenous donor is zinc dependent (Sevlever 2+
et al. 2001) and Zn suppresses the temperature sensitivity of a gpi13 allele. Phosphoethanolamine addition: Man2-­‐GPI may be Mcd4’s preferred substrate. Three sets of findings suggest that Mcd4 may act preferentially on Man2-­‐GPI: i) treatment of wild type cells with the terpenoid lactone YW3548, which inhibits addition of Etn-­‐P to Man-­‐1, leads to accumulation of Man2-­‐GPI (Sütterlin et al. 1997, 1998), ii) Man2-­‐GPI is the most abundant of the accumulating GPIs in mcd4-­‐174, and iii) Man2-­‐GPI is the largest GPI formed in vitro by mcd4 membranes (Zhu et al. 2006). 22 SI P. Orlean Phosphoethanolamine addition: importance of the Etn-­‐P added to Man-­‐1 by Mcd4 and additional possible functions for Mcd4. The finding that mcd4 mutants accumulate unmodified Man2-­‐GPI suggests that the presence of Etn-­‐P on Man-­‐1 is important for GPI-­‐Man-­‐T-­‐III to add the third Man. The requirement, though, is not absolute because mcd4Δ cells can be partially rescued by overexpression of Gpi10 (Wiedman et al. 2007). In addition to enhancing the efficiency of mannosylation by Gpi10, the Etn-­‐P moiety on Man-­‐1 may be important for additional reasons. mcd4Δ cells expressing human or trypanosomal Gpi10 orthologues, Man-­‐T known to mannosylate Man2-­‐GPIs lacking Etn-­‐P on Man-­‐1 efficiently, still grow slowly (Zhu et al. 2006; Wiedman et al. 2007). Further, mcd4Δ cells expressing trypanosomal Gpi10 are retarded in export of GPI-­‐proteins from the ER, unable to remodel their GPI lipid moiety to ceramide, and are defective in selection of axial budding sites (Zhu et al. 2006). How the presence of Etn-­‐P on Man-­‐1 influences these processes is not yet known. Mutations in MCD4 also impact cellular processes that are not directly connected with GPI biosynthesis. Cells expressing the Mcd4-­‐P
301
L variant, but not G
227
E, are defective in the transport of Ptd-­‐Ser to the Golgi and vacuole for decarboxylation, but unaffected in GPI anchoring suggesting an additional role for Mcd4 in transport dependent Ptd-­‐Ser metabolism (Storey et al. 2001). Further, yeast overexpressing Mcd4 (as well as Gpi7 and Gpi13) release ATP into the medium, and Golgi vesicles from the Mcd4 overexpressers were enriched in that protein and showed elevated levels of ATP uptake (Zhong et al. 2003). It was suggested that Mcd4 normally mediates symport of ATP and Ptd-­‐Etn into the ER lumen, and that overexpression of the protein leads ATP to accumulate in secretory vesicles, which eventually fuse with the plasma membrane (Zhong et al. 2003). Phosphoethanolamine addition to Man-­‐2 and its possible functions. GPI-­‐Etn-­‐P-­‐II consists of catalytic Gpi7 and non-­‐
catalytic Gpi11. Both gpi7Δ and temperature-­‐sensitive gpi11Δ disruptants complemented by the human Gpi11 orthologue PIG-­‐
F accumulate a Man4-­‐GPI bearing Etn-­‐P on Man-­‐1 and Man-­‐3 but missing one on Man-­‐2 (Benachour et al. 1999; Taron et al. 2000). Because loss of GPI-­‐Etn-­‐P function leads to accumulation of a Man4-­‐GPI with Etn-­‐Ps on Man-­‐1 and Man-­‐3, GPI-­‐Etn-­‐P-­‐II may normally add Etn-­‐P to Man-­‐2 after GPI-­‐Etn-­‐P-­‐T-­‐III has modified Man-­‐3. However, because Man3-­‐ and Man4-­‐GPIs with a single Etn-­‐P on Man-­‐2 accumulate in the smp3 mutants and in temperature-­‐sensitive gpi11Δ strains complemented by the human Gpi11 orthologue (Taron et al. 2000; Grimme et al. 2001), GPI-­‐Etn-­‐P-­‐II has the capacity to act on Etn-­‐P-­‐free GPIs. Diverse phenotypes of gpi7Δ cells indicate that the Etn-­‐P moiety on Man-­‐2 is important for a number of reasons. First, the combination of gpi7Δ with the GPI transamidase mutation gpi8 leads to a synthetic growth defect, indicating that an Etn-­‐P on Man-­‐2 enhances transfer of GPIs to protein (Benachour et al. 1999). Second, gpi7Δ cells have defects in ER to Golgi transport of GPI-­‐proteins and GPI lipid remodeling to ceramide (Benachour et al. 1999). Third, GPI7 deletion leads to cell wall defects and P. Orlean 23 SI shedding of GPI-­‐proteins, indicating defective transfer of such proteins into the wall (Toh-­‐e and Oguchi, 1999; Richard et al., 2002). Lastly, gpi7Δ cells show a cell separation defect that results from mistargeting of Egt2, a GPI protein expressed in daughter cells and implicated in degradation of the septum (Fujita et al. 2004). These phenotypes suggest that the Etn-­‐P group on Man-­‐2 is recognized by GPI transamidase, the intracellular transport machinery, GPI lipid remodeling enzymes, and cell wall crosslinkers. An inability to remove Etn-­‐P from Man-­‐2 also leads to phenotypes (see Remodeling of protein bound GPIs). Phosphoethanolamine addition to Man-­‐3 by Gpi13 and the role of Gpi11. Gpi13 is the catalytic subunit of GPI-­‐Etn-­‐P-­‐T-­‐
III, and, as expected from the fact that it adds the Etn-­‐P that participates in the GPI transamidase reaction, GPI13 is essential. The major GPI accumulated by yeast strains depleted of Gpi13 is a Man4-­‐GPI with a single Etn-­‐P on Man-­‐1 (Flury et al. 2000; Taron et al. 2000). Gpi11 is likely involved in the GPI-­‐Etn-­‐P-­‐T-­‐III reaction as well, because a recently isolated gpi11-­‐Ts mutant also accumulates a Man4-­‐GPI with its Etn-­‐P on Man-­‐1 (K. Willis and P. Orlean, unpublished results), and human Gpi11 interacts with and stabilizes human Gpi13 (Hong et al. 2000). Human Gpi11 (Pig-­‐F) also interacts with human Gpi7 (Shishioh et al. 2005). The lipid accumulation phenotypes observed in various types of gpi11 mutants may prove to be explainable in terms of differential abilities of wild type Gpi11, mutant Gpi11, and human Gpi11 to interact with Gpi7, Gpi13, and possibly even Mcd4, and permit varying extents of Etn-­‐P modification. Because GPIs with the same chromatographic mobilities may be isoforms modified with Etn-­‐P at different positions, and because accumulating GPIs may be mixtures of isoforms, detailed structural analyses should give a clearer picture of the role of Gpi11 in Etn-­‐P modification. GPI transfer to protein: Depletion of Gab1 and Gpi8 leads to actin bar formation. Additional functions for Gab and Gpi18 are suggested by the finding that depletion of Gab1 or Gpi8 from yeast, but not of Gaa1, Gpi16, or Gpi17, leads to accumulation of bar-­‐like structures of actin that associate with the perinuclear ER and are decorated with cofilin (Grimme et al. 2004). This phenotype, which is not a general result of defective GPI anchoring, might reflect disruption of some functional interaction between resident ER membrane proteins and the actin cytoskeleton and consequent collapse of the ER around the nucleus (Grimme et al. 2004). Remodeling of protein-­‐bound GPIs: Roles of Bst1, Per1, and Gup1 in ER exit and transport of GPI proteins. Modifications of the GPI lipid by Bst1, Per1, and Gup1 are necessary for efficient transport of GPI proteins from the ER to the Golgi. Loss of Bst1 function leads to retarded transport of GPI-­‐proteins from the ER to the Golgi (Vashist et al. 2001), and delayed ER degradation of misfolded GPI proteins, suggesting that inositol deacylation generates sorting signals for ER exit of GPI proteins and for recognition by a quality control 24 SI P. Orlean mechanism for GPI-­‐proteins (Fujita et al. 2006; Fujita and Jigami, 2008). per1Δ and gup1Δ cells also show significantly delayed ER to Golgi transport of GPI-­‐proteins (Bosson et al. 2006; Fujita et al. 2006b). Lipid remodeling events generate a GPI able to associate with and be concentrated in membrane microdomains at ER exit sites prior to their export from the ER (Castillon et al. 2009). At these sites, the p24 complex of membrane proteins then serves as an adapter between GPI-­‐proteins and the COP II machinery to promote incorporation of GPI proteins into COP II vesicles specialized for transport of GPI-­‐proteins from the ER. Remodeled GPIs may bind p24 with higher affinity, therefore promoting export of the proteins bearing them (Castillon et al. 2011). In the Golgi, GPI-­‐proteins with remodeled anchors are released and proceed onwards along the secretory pathway. However, p24 complexes, which cycle between the ER and Golgi, again monitor the remodeling status of GPIs and exert a quality control function in the Golgi by sensing and retrieving proteins with unmodified GPIs to the ER, where they may encounter the resident ER remodeling enzymes (Castillon et al. 2011). Remodeling of the GPI lipid moiety to ceramide by Cwh43. Cwh43, which replaces the diacylglycerol moiety of GPIs with ceramide, is a large protein with 19 predicted transmembrane domains (Martin-­‐Yken et al. 2001; Ghugtyal et al. 2007; Umemura et al. 2007). cwh43Δ cells accumulate GPI-­‐proteins whose lipids are diacylglycerols with a very long acyl chain similar to the lipid generated after action of Bst1, Per1, and Gup1. Because ceramide remodeling requires prior action of Bst1, and per1Δ and gup1Δ strains show severe defects in remodeling, the exchange reaction seems to take place after the first three lipid modification steps. The mechanism is so far unknown, but could involve a phospholipase-­‐like reaction that replaces diphosphatidic acid with ceramide phosphate or diacylglycerol with ceramide (Ghugtyal et al. 2007; Fujita and Kinoshita, 2010). However, alternatives to such a linear remodeling pathway, in which Cwh43 acts instead on the Bst1 or Per1 products, have been discussed (Umemura et al. 2007). The C-­‐terminal domain of Cwh43 contains a motif that may be involved in recognition of inositol phosphate (Umemura et al. 2007). Because mcd4 and gpi7, mutants defective in addition of Etn-­‐P to Man-­‐1 and Man-­‐2, are affected in ceramide remodeling, Cwh43 may also recognize Etn-­‐P side-­‐branches. Cwh43 appears to act in the ER, where it remodels GPIs with a ceramide consisting of phytosphingosine bearing a C26 acyl chain, as well as in the Golgi, where the ceramide it introduces contains phytosphingosine with a hydroxy-­‐C26 acyl group (Reggiori et al. 1997). Removal of Etn-­‐P moieties from Man-­‐1 and Man-­‐2. The ER-­‐localized Ted1 and Cdc1 proteins are homologous to mammalian PGAP5, which removes EtN-­‐P moieties from Man-­‐2 (Fujita et al. 2009), and genetic interactions connect these two proteins processing and export of GPI-­‐proteins. Export of Gas1 is retarded in ted1Δ cells, and ted1Δ’s buffering genetic interactions with emp24Δ and erv5Δ, mutants deficient in two components of the p24 complex involved in maturation and trafficking of GPI proteins, indicate a functional relationship between the three proteins (Haass et al. 2007). Further, cdc1 P. Orlean 25 SI mutations are suppressed by per1/cos16 and gup1 mutations (Paidhungat and Garrett, 1998; Losev et al. 2008). Ted1 and Cdc1 contain a lumenal metallophosphoesterase domain (Haass et al. 2007; Losev et al. 2008), and, consistent with this, cdc1’s 2+
temperature-­‐sensitivity is suppressed by Mn , the cation required by PGAP5 (Fujita et al. 2009). These findings are in turn consistent with Ted1 and Cdc1 being GPI-­‐Etn-­‐P phosphodiesterases, but this possibility awaits biochemical confirmation. Literature Cited Castillon, G. A., Aguilera-­‐Romero, A., Manzano-­‐Lopez, J., Epstein, S., Kajiwara, K., et al., 2011 The yeast p24 complex regulates GPI-­‐anchored protein transport and quality control by monitoring anchor remodeling. Mol. Biol. Cell. 22: 2924-­‐2936. Castillon, G. A., Watanabe, R., Taylor, M., Schwabe, T. M., Riezman, H., 2009 Concentration of GPI-­‐anchored proteins upon ER exit in yeast. Traffic 10: 186–200. Coutinho, P. M., Deleury, E., Davies, G. J., Henrissat, B., 2003 An evolving hierarchical family classification for glycosyltransferases. J. Mol. Biol. 328: 307-­‐317 Franzot, S. P, Doering, T. L. 1999 Inositol acylation of glycosylphosphatidylinositols in the pathogenic fungus Cryptococcus neoformans and the model yeast Saccharomyces cerevisiae. Biochem. J. 340: 25-­‐32. Fujita, M., Jigami, Y., 2008 Lipid remodeling of GPI-­‐anchored proteins and its function. Biochim. Biophys. Acta 1780: 410-­‐420. Kostova, Z., Yan, B. C., Vainauskas, S., Schwartz, R., Menon, A. K., et al. 2003 Comparative importance in vivo of conserved glutamates in the EX7E-­‐motif retaining glycosyltransferase Gpi3p, the UDP-­‐GlcNAc-­‐binding subunit of the first enzyme in glycosylphosphatidylinositol assembly. Eur. J. Biochem. 270: 4507-­‐4514. Losev, E., Papanikou, E., Rossanese, O. W., Glick, B. S., 2008 Cdc1p is an endoplasmic reticulum-­‐localized putative lipid 2+
phosphatase that affects Golgi inheritance and actin polarization by activating Ca signaling. Mol. Cell. Biol. 28: 3336–3343. 26 SI P. Orlean Murakami, Y., Siripanyapinyo, U., Hong, Y., Kang, J. Y., Ishihara, S., Nakakuma, H., et al., 2003 PIG-­‐W is critical for inositol acylation but not for flipping of glycosylphosphatidylinositol-­‐anchor. Mol. Biol. Cell 14: 4285-­‐4295. 2+
Paidhungat, M., Garrett, S., 1998 Cdc1 and the vacuole coordinately regulate Mn homeostasis in the yeast Saccharomyces cerevisiae. Genetics 148: 1787–1798. Reggiori, F., Canivenc-­‐Gansel, E., Conzelmann, A., 1997 Lipid remodeling leads to the introduction and exchange of defined ceramides on GPI proteins in the ER and Golgi of Saccharomyces cerevisiae. EMBO J. 16: 3506-­‐3518. Sevlever, D., Mann, K. J., Medof, M. E., 2001, Differential effect of 1,10-­‐phenanthroline on mammalian, yeast, and parasite glycosylphosphatidylinositol anchor synthesis. Biochem. Biophys. Res. Commun. 288: 1112-­‐1118. Shishioh, N., Hong, Y., Ohishi, K., Ashida, H., Maeda, Y., et al., 2005 GPI7 is the second partner of PIG-­‐F and involved in modification of glycosylphosphatidylinositol. J. Biol. Chem. 280: 9728-­‐9734. Sipos, G., Reggiori, F., Vionnet, C., Conzelmann, A., 1997 Alternative lipid remodelling pathways for glycosylphosphatidylinositol membrane anchors in Saccharomyces cerevisiae. EMBO J. 16: 3494-­‐3505. Sobering, A. K., Romeo, M. J., Vay, H. A., Levin, D. E., 2003 A novel Ras inhibitor, Eri1, engages yeast Ras at the endoplasmic reticulum. Mol. Cell. Biol. 23: 4983-­‐49890. Storey, M. K., Wu, W. I., Voelker, D. R., 2001 A genetic screen for ethanolamine auxotrophs in Saccharomyces cerevisiae identifies a novel mutation on Mcd4p, a protein implicated in glycosylphosphatidylinositol anchor synthesis. Biochim. Biophys. Acta. 1532: 234-­‐247. Toh-­‐e, A., Oguchi, T., 2002 Genetic characterization of genes encoding enzymes catalyzing addition of phospho-­‐ethanolamine to the glycosylphosphatidylinositol anchor in Saccharomyces cerevisiae. Genes Genet. Syst. 77: 309-­‐322. P. Orlean 27 SI Vashist, S., Kim, W., Belden, W. J., Spear, E. D., Barlowe, C., et al., 2001 Distinct retrieval and retention mechanisms are required for the quality control of endoplasmic reticulum protein folding. J. Cell Biol. 155: 355-­‐368. Zhong, X., Malhotra, R., Guidotti, G., 2003 ATP uptake in the Golgi and extracellular release require Mcd4 protein and the +
vacuolar H -­‐ATPase. J. Biol. Chem. 278: 33436-­‐33444. 28 SI P. Orlean File S5 Sugar nucleotide transport This Supporting File contains additional information related to Biosynthesis of Wall Components Along the Secretory Pathway, Sugar nucleotide transport. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. GDP-­‐Man transport: The GDP-­‐Man transporter, Vrg4/Vig4. This protein forms homodimers (Abe et al. 1999; Gao and Dean, 2000), shows a wide distribution in the Golgi, and contains a GALNK motif involved in GDP-­‐Man binding (Gao et al. 2001). Gda1 and Ynd1. Evidence these proteins have partially overlapping functions is as follows. i) Deletion of either GDA1 or YND1 impacts mannosylation of N-­‐ and O-­‐glycans, ii) high-­‐level expression of YND1 corrects some of gda1Δ’s glycosylation defects, and iii) gda1Δ ynd1Δ double mutants have a synthetic phenotype and show growth and cell wall defects (Gao et al. 1999). However, gda1Δ ynd1Δ double mutants are viable and capable of some mannosylation of N-­‐ and O-­‐linked glycans, indicating that GDP-­‐Man can enter the Golgi in their absence, and suggesting there may be a mechanism for GDP exit independent of GDP hydrolysis (D’Alessio et al. 2005). GMP generated upon Man-­‐P transfer to glycoproteins could also be a source of antiporter, but it is not a significant one because because the glycans made gda1Δ or gda1Δ ynd1Δ strains are not affected by disruption of MNN4 or MNN6 (Jigami and Odani, 1999; D’Alessio et al. 2005). Other sugar nucleotide transport activities: Transport activities for UDP-­‐Glc, UDP-­‐GlcNAc, and UDP-­‐Gal also occur in S. cerevisiae (Roy et al. 1998; 2000 Castro et al. 1999), and there are eight further candidate transporters (Dean et al. 1997; Esther et al. 2008), a couple of which have been associated with these transport activities. Some of the transporters may have specificity for more than one sugar nucleotide. In the case of UDP-­‐Glc, transport activity was present in the ER (Castro et al. 1999), but the responsible protein for that activity has yet to identified, although broad specificity Yea4 and Hut1 (see below) may transport UDP-­‐Glc (Esther et al. 2008). One possible need for UDP-­‐Glc transport into the ER might be for a glucosylation reaction at an early stage of β1,6-­‐glucan assembly (Section VI). The Hut1 protein is a candidate for the UDP-­‐Gal transporter (Kainuma et al. 2001), but whether that is Hut1’s primary role in vivo is unclear because galactose has not been detected on S. cerevisiae glycans. Yea4 was characterized as an ER-­‐localized UDP-­‐GlcNAc transporter and its deletion impacts chitin synthesis (Roy et al. 2000; Section V). Of the other P. Orlean 29 SI transporter homologs, Hvg1 resembles Vrg4 most closely, but hvgΔ cells have neither a mannosylation nor a GDP-­‐Man transport defect (Dean et al. 1997). The roles of the other proteins in sugar nucleotide transport, if any, is unknown. One or more transporters may supply the Golgi GlcNAc-­‐T Gnt1 with its substrate (Section IV.1.c.ii). Literature Cited D'Alessio, C., Caramelo, J. J., Parodi, A. J., 2005 Absence of nucleoside diphosphatase activities in the yeast secretory pathway does not abolish nucleotide sugar-­‐dependent protein glycosylation. J. Biol. Chem. 280: 40417-­‐40427. Gao, X. D., Dean, N., 2000 Distinct protein domains of the yeast Golgi GDP-­‐mannose transporter mediate oligomer assembly and export from the endoplasmic reticulum. J. Biol. Chem. 275: 17718-­‐17727. Gao, X. D., Nishikawa, A., Dean, N., 2001 Identification of a conserved motif in the yeast Golgi GDP-­‐mannose transporter required for binding to nucleotide sugar. J. Biol. Chem. 276: 4424-­‐4432. 30 SI P. Orlean File S6 Chitin This Supporting File contains additional information and discussion related to Biosynthesis of Wall Components at the Plasma Membrane, Chitin. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Septum formation: Phenotypes of chs1Δ chs2Δ chs3Δ triple mutants. chs1Δ chs2Δ chs3Δ strains grew very slowly but acquired a suppressor mutation that conferred a growth rate as fast as that of a chs2Δ mutant, although over a third of suppressed or unsuppressed cells in a culture were dead (Schmidt, 2004). Membranes from the triple mutants had no detectable chitin synthase activity. Unsuppressed triple mutants formed chains of up to eight cells that appeared to be connected by “cytoplasmic stalks”, whereas suppressed strains formed shorter chains. Nuclear division continued in the mutant, but in some cells, nuclear segregation was unsuccessful. Ultrastructural analysis showed that in both suppressed and unsuppressed mutants, a bulky remedial septum arises upon thickening of the lateral walls in the mother cell-­‐bud neck region. The suppressor was not identified, but its effect was to allow the remedial septa to be formed more efficiently. The phenotypes of the triple chitin synthase mutants indicate that although it is possible for S. cerevisiae to grow without chitin, Chs3-­‐dependent chitin synthesis is nonetheless important for remedial septum formation in chs2Δ cells. Chitin synthase biochemistry: Directionality and mechanism of extension of β1,4-­‐linked polysaccharide chains. Although the bacterial chitin synthase homologue NodC extends chito-­‐oligosaccharides at their non-­‐reducing ends (Kamst et al. 1999), both reducing-­‐ and non-­‐reducing end extension has been reported for Chs-­‐related vertebrate Class I hyaluronate synthases (Weigel and DeAngelis, 2007), and extension by insertion of Glc at the reducing end of a glycan chain has also been proposed for a bacterial cellulose synthase (Han and Robyt, 1998). The latter mechanism was suggested to involve a lipid pyrophosphate intermediate. However, no evidence has been obtained for any lipid-­‐linked intermediate in chitin synthesis. The growing glycan chain may be extruded through the plasma membrane through a pore made up by a bundle of transmembrane helices, which occur towards the C-­‐
terminus of chitin synthases (Delmer, 1999; Guerriero et al. 2010; Merzendorfer, 2011; Carpita, 2011). Separate proteins might mediate chitin translocation, but no candidates have been identified. With non-­‐reducing end extension, a nascent chitin chain would be extruded into the cell wall reducing end first, which would be compatible with the formation of linkages between P. Orlean 31 SI chitin and non-­‐reducing ends of β-­‐glucans (see Cross-­‐linkage of chitin to β1,6-­‐ and β1,3-­‐glucan; Kollar et al. 1995, 1997; Cabib and Duran, 2005; Cabib, 2009). The stereochemical challenge in formation of β1,4-­‐linked polysaccharides. Each sugar in a β1,4-­‐linked polymer is rotated by about 180° relative to its neighbor, which presents the synthase with a steric challenge, because with successive rounds of addition of a β1,4-­‐linked GlcNAc, the new acceptor 4-­‐OH would alternate between two positions relative to incoming substrate and catalytic residues. Various ways of overcoming this, without invoking movements of the enzyme or the acceptor glycan, have been considered. The first possibility, that UDP-­‐di-­‐N-­‐acetylchitobiose is the donor, has been ruled out by the finding that yeast membranes make no chitin when supplied with synthetic UDP-­‐GlcNAc2 (Chang et al. 2003). The second possibility is that β1,4-­‐linked polysaccharide synthases have two UDP-­‐sugar binding sites that orient the monosaccharides such that neither enzyme nor polymer needs to rotate, then catalyzes two glycosyltransfers (Saxena et al. 1995; Guerriero et al. 2010; Carpita, 2011). Evidence supportive of a two active site mechanism came from the finding that a bivalent UDP-­‐GlcNAc analog consisting of two tethered uridine mimetics, envisaged to bind in both active sites, was a better inhibitor than the monomeric analog (Yaeger and Finney, 2004). The observation that the NodC protein, Chs1, and Chs2 all synthesize odd-­‐ as well as even-­‐numbered chito-­‐ooligosaccharides in vitro (Kang et al. 1984; Yabe et al. 1998; Kamst et al. 1999) is consistent with extension by addition with single GlcNAcs, but extension of GlcNAc, GlcNAc3, or GlcNAc5 by two GlcNAcs at a time would also generate odd-­‐numbered chito-­‐oligosaccharides, if these oligosaccharides are indeed used as primers. Third, it is possible that a chain is extended by a dimeric synthase whose subunits alternately add GlcNAcs, as discussed for cellulose synthase (Carpita, 2011). Consistent with this notion, a two-­‐hybrid analysis indicated that Chs3 can interact with itself (DeMarini et al. 1997). The molecular weight of purified native Chs1 was estimated to be around 570,000, approximately consistent with a tetramer, but the authors noted the result may have been due to protein aggregation (Kang et al. 1984). 14
In vitro properties of yeast chitin synthases. Chitin synthase assays typically detect the transfer of [ C]GlcNAc from 14
UDP[ C]GlcNAc to insoluble chitin that is then collected on filters, but a high-­‐throughput method that relies on product binding to immobilized wheat germ agglutinin has also been described (Lucero et al. 2002). Of the two procedures, the filtration method would not detect chito-­‐oligosaccharides (Yabe et al. 1998). CS I, CS II, and CS III activities differ in their pH optima and their responses to divalent cations (Sburlati and Cabib, 1986; Orlean, 1987; Choi and Cabib, 1994). The three chitin synthase activities have Kms for UDP-­‐GlcNAc in the range of 0.5-­‐1.3 mM (Kang et al. 1984; Sburlati and Cabib, 1986; Orlean, 1987; Uchida et al. 1996). At low substrate concentrations relative to Km (0.03-­‐0.1 mM), purified Chs1 and membranes from cells overexpressing CHS2 make chito-­‐oligosaccharides (Kang et al. 1984; Yabe et al. 1998). Whether these are bona fide chitin 32 SI P. Orlean synthase products whose formation reflects low rates of chain extension, or whether the oligosaccharides are generated by chitinase activity on longer nascent chains is not clear (Kang et al. 1984). Effects of free GlcNAc and chitin oligosaccharides on chitin synthesis. S. cerevisiae’s three chitin synthases are all stimulated up to a few fold in vitro by high concentrations of free GlcNAc (e.g. 32 mM; Sburlati and Cabib, 1986; Orlean, 1987). Neither the mechanistic basis nor the physiological relevance of this are clear, but possible explanations are that GlcNAc serves as a primer or allosteric activator in the chitin synthetic reaction. Results of a kinetic analysis of the chitin synthase activity in wild type membranes led to the proposal that GlcNAc participates along with UDP-­‐GlcNAc in a two substrate reaction with an ordered mechanism in which UDP-­‐GlcNAc binds first (Fähnrich and Ahlers, 1981). Consistent with the idea that GlcNAc serves as a primer or co-­‐substrate, the bacterial NodC chitin synthase homologue incorporates free GlcNAc at the reducing end of chito-­‐
oligosaccharide chains that are extended at their non-­‐reducing end by GlcNAc transfer from UDP-­‐GlcNAc (Kamst et al. 1999). However, were free GlcNAc to serve as a co-­‐substrate or activator of chitin synthases in vivo, there would have to be a mechanism to generate it, for example from GlcNAc-­‐1-­‐P or GlcNAc-­‐6-­‐P (see Precursors and Carrier Lipids) or by turnover of GlcNAc-­‐containing molecules. Growing chitin chains presumably serve as acceptors for further GlcNAc addition, but such a primer function has not been shown using short oligosaccharides. NodC did not use short chito-­‐oligosaccharides as GlcNAc acceptor from UDP-­‐GlcNAc (Kamst et al. 1999), nor did purified Chs1 elongate chitotetraose into insoluble chitin in the presence of UDP-­‐GlcNAc (Kang et al. 1984). However, inclusion of 1 mM GlcNAc5 and GlcNAc8 in assays of membrane preparations expressing predominantly Chs1 led to about a 1.25-­‐fold increase in incorporation of GlcNAc into chitin from UDP-­‐GlcNAc in the presence of free GlcNAc (Becker et al. 2011), suggesting a primer function for longer chito-­‐oligosaccharides. The initiation and early elongation steps in chitin synthesis clearly still need to be defined. S. cerevisiae’s chitin synthases and auxiliary proteins: Chitin synthase classes. Fungal chitin synthases can be classified into five to seven classes on the basis of amino acid sequence similarity, with S. cerevisiae Chs1, Chs2, and Chs3 being assigned to Classes I, II, and IV respectively (Roncero, 2002; Ruiz-­‐Herrera et al. 2002; Van Dellen et al. 2006; Merzendorfer, 2011). Members of the other classes are found in filamentous fungi. S. cerevisiae’s chitin synthases show most amino acid sequence divergence in their amino terminal halves, and these non-­‐
homologous regions may make interactions with proteins involved in regulation or trafficking of the individual synthases (Ford et al. 1996). Deletion analyses have shown that amino acids in Chs3’s hydrophilic C-­‐terminal region are also important for function (Cos et al. 1998). P. Orlean 33 SI Chitin synthase I: Activity of N-­‐terminally truncated Chs1. N-­‐terminally truncated forms of Chs1 lacking up to 390 amino acids show a gradual lowering of both specific activity and their ability to be activated by trypsin (Ford et al. 1996). Chitin synthase II and proteins impacting its localization and activity: Detection of Chs2’s activity. Studies of Chs2 enzymology use membranes from strains overexpressing the protein because the activity of genomically encoded Chs2 in membranes of cells grown in minimal medium is negligible (Nagahashi et al. 1995). The high amounts of in vitro activity obtained by overexpressing Chs2 indicate that levels of Chs2 activity are not tightly limited by endogenous activating or regulatory proteins, in contrast to Chs3. Effects of proteolysis on wild type and truncated forms of Chs2. Although endogenously activated, processed forms of Chs2 have not been identified, trypsin treatment of partially purified, full-­‐size and N-­‐terminally truncated Chs2 generated a range of discrete protein fragments. The smallest of these, a 35 kDa protein containing the amino acid sequences proposed to be involved in catalysis, was suggested to be sufficient for catalysis, although the instablity of this form prevented its purification to test this notion (Uchida et al. 1996). Some 220 amino terminal amino acids of Chs2 are dispensable for in vivo function (Ford et al. 1996), and moreover, Chs2 versions lacking these amino terminal amino acids have higher in vitro activity than the full-­‐length protein, and this activity is stimulated by trypsin (Uchida et al. 1996; Martínez-­‐Rucobo et al. 2009). Other truncated forms of Chs2, or forms with amino acid substitutions, also vary in their extent of activation by trypsin (Ford et al. 1996; Uchida et al. 1996). It has been noted that amino acid deletions or substitutions in Chs2 could perturb interactions with native mechanisms for activation and localization of the protein (Ford et al. 1996). Chitin synthase III and proteins impacting its localization and activity: Relationship between Pfa4 and Chs7 and their roles in Chs3 exit from the ER. Chs3 interacts with Chs7 and is palmitoylated by Pfa4. The Chs3-­‐Chs7 interaction also occurs in pfa4Δ cells, though to a slightly reduced extent, and Chs3 can still be palmitoylated, likewise to a lesser extent, in chs7Δ cells, indicating that Chs3 palmitoylation is not obligatory for Chs3 recognition by Chs7 (Lam et al. 2006). Pfa4 does not palmitoyate Chs7. It seems that Pfa4 and Chs7 act in parallel, though not wholly independently, to promote folding of Chs3 prior to the synthase’s exit from the ER. These roles of Pfa4 and Chs7 are specific to Chs3, for neither is required for exit of Chs1 and Chs2 from the ER (Trilla et al. 1999; Lam et al. 2006). Rcr1 and Yea4 in Chs3-­‐dependent chitin synthesis. These proteins have both been localized to the ER membrane. Rcr1 has a slight negative regulatory effect on Chs3-­‐dependent chitin synthesis. High copy RCR1 confers resistance to Congo Red, a dye that binds chitin (as well as β1,3-­‐glucan (Kopecká and Gabriel, 1992)), whereas rcr1Δ cells showed slightly increased 34 SI P. Orlean sensitivity to Congo Red and CFW (Imai et al. 2005). Wild type cells overexpressing RCR1 have 70% of the chitin in control cells, and rcr1Δ cells make 115% of wild type levels of chitin. However, RCR1 overexpression affects neither the amount nor localization of Chs3, Chs5, and Chs7, nor do Rcr1 and Chs7 physically interact (Imai et al. 2005). The role of Rcr1 in Chs3-­‐
dependent chitin synthesis is therefore not clear, but the protein has also been reported to act after the ER and have a role in an endosome-­‐vacuole pathway that impacts trafficking of plasma membrane nutrient transporters (Kota et al. 2007). The second ER membrane protein, Yea4, was identified through its homology to the Kluyveromyces lactis UDP-­‐GlcNAc transporter (Roy et al. 2000). Membrane vesicles from cells overexpressing Yea4 have 8-­‐fold elevated levels of UDP-­‐GlcNAc transport activity, consistent with Yea4’s function as a transporter (Roy et al. 2000). yea4Δ cells contain 65% of wild type levels of chitin, implicating Yea4 in chitin synthesis, but whether and how Yea4’s transport activity contributes to this process is unclear. Role of exomer in transport of wall related proteins other than Chs3. Exomer has roles in polarized transport of other wall related proteins to the cell surface. Thus, transport of Fus1, which promotes cell fusion during mating, requires Chs5 for transport to the shmoo tip (Santos and Snyder, 2003), along with the ChAPs Bch1 and Bus7, but not Chs6 (Barfield et al. 2009). Further, much of the GPI-­‐anchored chitin-­‐β1,3-­‐glucan cross-­‐linker Crh2 (see Cross-­‐linkage of chitin to β1,6-­‐ and β1,3-­‐glucan) fails to reach sites of polarized growth and accumulates intracellularly in chs5Δ, although another GPI-­‐protein, Cwp1, was unaffected (Rodriguez-­‐Pena et al. 2002). Co-­‐transport of Chs3 and Crh2 would ensure colocalization of these proteins for efficient cross linking of nascent chitin to β1,3-­‐glucan. Role of Chs4 farnesylation in the activation and localization of Chs3. Chs4 has a C-­‐terminal farnesylation site (Bulawa et al. 1993; Trilla et al. 1997) that is used (Grabinska et al. 2007) and the consensus of studies of the importance of the prenyl group is that the modification has roles in Chs4 function and localization. Mutants expressing a non-­‐farnesylatable Cys to Ser variant of Chs4 make one third of normal amounts of chitin, have lower in vitro CS III activity, and show CFW resistance (Grabinska et al. 2007; Meissner et al. 2010). In two of three studies, the prenylation site mutant of Chs4 was found in the cytoplasm, suggesting that lipidation is important for membrane localization of the protein (Reyes et al. 2007; Meissner et al. 2010). Chs4 reaches the plasma membrane in mutants affected in Chs3 transport, indicating it is transported there independently of Chs3 (Reyes et al. 2007), but two sets of findings raise the possibility that Chs3 interacts with Chs4 at the level of the ER. First, two-­‐hybrid analyses established that cytoplasmic domains of Chs3 and the ER-­‐localized CAAX protease Ste24 interact. Second, ste24Δ cells exhibit moderate CFW resistance, chitin content is reduced, and less Chs3 was localized at the bud neck. Vice versa, high-­‐copy expression of STE24 leads to CFW sensitivity and some increase in cellular chitin (Meissner et al. 2010). Chs4 localization, though, was not affected in ste24Δ, nor was an interaction detected between Chs4 and Ste24. It was P. Orlean 35 SI suggested that Chs3 recruits farnesylated Chs4 in the ER for processing by Ste24, and that the modification contributes to subsequent correct localization of Chs3 and activation of CS III (Meissner et al. 2010). Chitin synthase III in mating and ascospore wall formation: Regulation of Chs3 during chitosan synthesis. The Chs4 homologue Shc1, which is 43% identical to Chs4 but expressed only during sporulation, has a role in chitosan synthesis, because homozygous shc1Δ shc1Δ diploids make ascospores with very little chitosan (Sanz et al. 2002). Shc1 and Chs4 are functionally related because when Shc1 is expressed in vegetative cells, it can activate CS III, and when Chs4 is overexpressed in shc1Δ shc1Δ diploids, it partially corrects the sporulation defect (Sanz et al. 2002). However, although Shc1 serves as CS III activator in chs4Δ cells, it does so without properly localizing Chs3 to septins as Chs4 does in vegetative cells, likely because it cannot interact with Bni4 (Sanz et al. 2002). Haploid chs4Δ shc1Δ cells do not show a synthetic growth defect, indicating they are not an essential redundant pair, and indeed, analyses of the SHC1 genetic interaction network suggests Shc1 may have additional roles distinct from those of Chs4 that are not directly related to chitin synthesis (Lesage et al. 2005). Sporulation-­‐specific kinase Sps1, regulates mobilization of Chs3 as well as sporulation-­‐specific β1,3-­‐glucan synthase Fks2/Gsc2 (see β1,3-­‐glucan) to the prospore membrane (Iwamoto et al. 2005). Literature Cited Barfield, R. M., Fromme, J. C., Schekman, R., 2009 The exomer coat complex transports Fus1p to the plasma membrane via a novel plasma membrane sorting signal in yeast. Mol. Biol. Cell 20: 4985-­‐4996. Becker, H.F., Piffeteau, A., Thellend, A. 2011 Saccharomyces cerevisiae chitin biosynthesis activation by N-­‐acetylchitooses depends on size and structure of chito-­‐oligosaccharides. BMC Res. Notes. 4: 454. Carpita, N. C., 2011 Update on mechanisms of plant cell wall biosynthesis: how plants make cellulose and other (1→4)-­‐β-­‐D-­‐
glycans. Plant Physiol. 155: 171-­‐184. Chang, R., Yeager, A. R. Finney, N. S., 2003 Probing the mechanism of a fungal glycosyltransferase essential for cell wall biosynthesis. UDP-­‐chitobiose is not a substrate for chitin synthase. Org. Biomol. Chem. 1: 39-­‐41. 36 SI P. Orlean Choi, W. J., Cabib, E., 1994 The use of divalent cations and pH for the determination of specific yeast chitin synthetases. Anal. Biochem. 219: 368-­‐372. Delmer, D. P., 1999 Cellulose biosynthesis: exciting times for a difficult field of study. Annu. Rev. Plant Physiol. Plant Mol. Biol. 50: 245-­‐276. Fähnrich, M., Ahlers, J. 1981 Improved assay and mechanism of the reaction catalyzed by the chitin synthase from Saccharomyces cerevisiae. Eur. J. Biochem. 121: 113-­‐118. Ford, R. A., Shaw, J. A., Cabib, E., 1996 Yeast chitin synthases 1 and 2 consist of a non-­‐homologous and dispensable N-­‐terminal region and of a homologous moiety essential for function. Mol. Gen. Genet. 252: 420-­‐428. Imai, K., Noda, Y., Adachi, H., Yoda, K., 2005 A novel endoplasmic reticulum membrane protein Rcr1 regulates chitin deposition in the cell wall of Saccharomyces cerevisiae. J. Biol. Chem. 280: 8275-­‐828. Kopecká, M., Gabriel, M., 1992 The influence of congo red on the cell wall and (1-­‐3)-­‐β-­‐D-­‐glucan microfibril biogenesis in Saccharomyces cerevisiae. Arch Microbiol. 158: 115-­‐126. Guerriero, G., Fugelstad, J., Bulone, V. 2010 What do we really know about cellulose biosynthesis in higher plants? J. Integr. Plant Biol. 52: 161-­‐175. Iwamoto, M. A., Fairclough, S. R., Rudge, S. A., Engebrecht, J., 2005 Saccharomyces cerevisiae Sps1p regulates trafficking of enzymes required for spore wall synthesis. Eukaryot. Cell 4: 536-­‐544. Kota, J., Melin-­‐Larsson, M., Ljungdahl, P. O., Forsberg, H., 2007 Ssh4, Rcr2 and Rcr1 affect plasma membrane transporter activity in Saccharomyces cerevisiae. Genetics 175: 1681-­‐1694. P. Orlean 37 SI Lucero, H. A., Kuranda M. J., Bulik, D. A., 2002 A nonradioactive, high throughput assay for chitin synthase activity. Anal. Biochem. 305: 97-­‐105. Nan, N. S., Robyt, J. F. 1998. The mechanism of Acetobacter xylinum cellulose biosynthesis: direction of chain elongation and the role of lipid pyrophosphate intermediates in the cell membrane. Carbohydrate Res. 313: 125-­‐133. Santos, B., Snyder, M., 2003. Specific protein targeting during cell differentiation: polarized localization of Fus1p during mating depends on Chs5p in Saccharomyces cerevisiae. Eukaryot. Cell 2: 821–825. Van Dellen, K. L., Bulik, D. A., Specht, C. A., Robbins, P. W., Samuelson, J. C., 2006 Heterologous expression of an Entamoeba histolytica chitin synthase in Saccharomyces cerevisiae. Eukaryot. Cell. 5: 203-­‐206. Weigel, P. H., DeAngelis, P. L., 2007 Hyaluronan synthases: a decade-­‐plus of novel glycosyltransferases. J. Biol. Chem. 282: 36777-­‐36781. Yaeger, A.R., Finney, N. S., 2004 The first direct evaluation of the two-­‐active site mechanism for chitin synthase. J. Org. Chem. 69: 613-­‐618. 38 SI P. Orlean File S7 β 1,3-­‐glucan This Supporting File contains additional information and discussion related to Biosynthesis of Wall Components at the Plasma Membrane, β 1,3-­‐glucan. The subheadings used in the main text are retained, and new subheadings are underlined. Fks family of β 1,3-­‐glucan synthases: Identification of Fks1, Fks2, and Fks3. Fks1 (Cwh53/Etg1/Gsc1/Pbr1) was identified in screens for hypersensitivity to the calcineurin inhibitors FK506 and cyclosporin A and to CFW, for resistance to echinocandin and papulocandin, and following purification of β1,3-­‐glucan synthase activity (reviewed by Orlean, 1997 and Lesage and Bussey, 2006). Cross-­‐hybridization with FKS1 and copurification with Fks1 led to identification of Fks2/Gsc2, which is 88% identical to Fks1 (Inoue et al. 1995; Mazur et al. 1995). The S. cerevisiae proteome also contains Fks3, which is 55% identical to Fks1 and Fks2 (Dijkgraaf et al. 2002). The Fks proteins are assigned to GT Family 48, and a strong case can be made for them being processive β1,3-­‐glucan synthases themselves, although roles as glucan exporters cannot yet be excluded (Mazur et al. 1995; Dijkgraaf et al. 2002; Lesage and Bussey, 2006). Functional domains of Fks1. Fks1 is predicted to have an N-­‐terminal cytoplasmic domain of some 300 amino acids that is followed by six transmembrane helices, a second cytoplasmic domain of about 600 amino acids, then 10 transmembrane helices (Inoue et al. 1995; Mazur et al. 1995; Qadota et al. 1996; Dijkgraaf et al. 2002; Okada et al. 2010). Three functional domains have been distinguished (Okada et al. 2010). Amino acids important for β1,3 glucan synthesis in vivo are located in the first cytoplasmic domain. Mutations here have little impact on in vitro activity and do not affect the protein’s interaction with Rho1, but cells have a lowered β1,3 glucan content. Mutations in the second cytoplasmic domain that lie close to the C-­‐
terminus of the sixth helix lead to a loss of cell polarity as well as defects in endocytosis, but have little effect on in vitro and in vivo b-­‐glucan synthesis, and this part of Fks1 may interact with factors involved in cell polarity (Okada et al. 2010). Mutations in Fks1 in residues more distal to the sixth helix lead to low in vitro glucan synthase activity and large decreases in in vivo 14
incorporation of [ C]glucose into β1,3 glucan, suggesting that if Fks1 is a synthase, this part of the protein contains the catalytic site (Dijkgraaf et al. 2002; Okada et al. 2010). Fatty acid elongases and phytosphingosine and Fks1 function. The ER-­‐localized fatty acid elongase Elo2/Gns1 may impact Fks1 at the level of that organelle, because gns1 mutants, isolated on account of their resistance to a papulocandin analogue, have very low in vitro β1,3-­‐glucan synthase activity (el-­‐Sherbeini and Clemas, 1995) and accumulate P. Orlean 39 SI phytosphingosine in the ER membrane (Abe et al. 2001). Phytosphingosine inhibits β1,3 glucan synthase in vitro, leading to the idea that this sphingolipid synthetic intermediate is a negative regulator of β1,3-­‐glucan synthesis at the level of the ER (Abe et al. 2001). Roles of the Fks proteins in β 1,3-­‐glucan synthesis Roles of Fks3 and Fks3 in sporulation. Fks2 is important in sporulation because fks2Δ fks2Δ diploids have a severe defect in this process (Mazur et al. 1995; Huang et al. 2005), and form disorganized ascospore walls with lower relative amounts of hexose in their alkali-­‐insoluble fraction and a lower alkali soluble β1,3-­‐glucan content (Ishihara et al. 2007). Homozygous fks3Δ fks3Δ diploids also form abnormal spores, indicating a role for the third Fks homologue in ascopore wall formation, but showed no alteration in the distribution of hexoses between alkali soluble-­‐ and insoluble fractions (Ishihara et al. 2007). However, the walls of ascospores formed in diploids lacking both Fks2 and Fks3 were more disorganized than those of ascospores made by fks2Δ fks2Δ diploids (Ishihara et al. 2007). Expression of FKS2 or FKS1 under the control of the FKS2 promoter, but not the FKS1 promoter, corrected the sporulation defect of homozygous fks1Δ fks2Δ diploids, suggesting that the function of Fks2 in sporulating diploids resembles that of Fks1 in vegetative cells. In contrast, overexpression of FKS3 did not suppress the phenotype of fks2Δ spores, and FKS1 or FKS2 overexpression does not correct the defect in fks3Δ spores, indicating Fks3’s function in sporulation does not overlap with that of Fks2. It was proposed that Fks2 is primarily responsible for synthesis of β1,3-­‐glucan in the ascospore wall, and that Fks3, rather than functioning as a synthase, modulates glucan synthesis by interacting with glucan synthase regulators such as Rho1 (Ishihara et al. 2007). 40 SI P. Orlean File S8 β1,6-­‐Glucan This Supporting File contains additional information and discussion related to β 1,6-­‐Glucan. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Proteins involved in β 1,6-­‐glucan assembly ER proteins: Fungus-­‐specific ER chaperones required for β1,6-­‐glucan synthesis: Evidence for the chaperone function of Rot1, Big1, and Keg1 in β1,6-­‐glucan synthesis. Rot1, Big1, and Keg1, which do not resemble known carbohydrate-­‐active enzymes, seem unlikely to catalyze formation of β1,6-­‐glucan (Lesage and Bussey, 2006). Rather, they seem to function as ER chaperones with varying degrees of importance for the stability of proteins involved in β1,6-­‐glucan synthesis, and in some cases, they may cooperate. Observations supporting this notion, and indicating a relationship to Kre5, are as follows. Analyses of levels of β1,6-­‐glucan synthesis-­‐related proteins in a rot1-­‐Ts mutant indicate that Kre6 has the strongest dependence on Rot1 for stability, although Kre5 and Big1 show appreciable dependence as well (Takeuchi et al. 2008). Keg1, a protein essential for growth in osmotically supported medium, physically interacts with Kre6 in the ER membrane, and a keg1-­‐Ts mutant is suppressed at high copy by ROT1, though not BIG1; however, a physical interaction between Keg1 and Rot1 could not be detected (Nakamata et al. 2007). Because the big1Δ rot1Δ double mutant has the same growth rate as each single mutant, it was suggested that Rot1 and Big1 impact β1,6-­‐glucan synthesis in the same way, and possibly function in the same compartment or even in a complex (Machi et al. 2004). However, although rot1, big1, and kre5 mutations individually all lower β1,6-­‐glucan levels to the same extent, the kre5 big1 double mutant, but apparently not a kre5 rot1 strain (Lesage and Bussey, 2006), shows a reduced growth rate and lowered β1,6-­‐glucan content compared with each single mutant, suggesting the function of Rot1 is partly distinct from that of Kre5 (Azuma et al. 2002; Lesage and Bussey, 2006). Indeed, the non-­‐conditional rot1-­‐1 mutant shows a synthetic growth and N-­‐glycosylation defect in combination with ost3Δ (though not ost6Δ), as well as a partial defect in O-­‐mannosylation of the chitinase Cts1, indicating a wider role for Rot1 in glycosylation (Pasikowska et al. 2012). More widely distributed secretory pathway proteins: Kre6 and Skn1: P. Orlean 41 SI Localization and transport of Kre6. Recent studies indicate that much of Kre6 is ER-­‐localized, where it interacts with Keg1, but Kre6 is also detectable in secretory vesicles and at the plasma membrane at sites of polarized growth (Nakamata et al. 2007; Kurita et al. 2011). In addition to Kre6’s lumenal domain, the protein’s cytoplasmic tail is important for Kre6’s function in β1,6-­‐glucan assembly and its transport to the plasma membrane (Li et al. 2002; Kurita et al. 2011). A truncated form of Kre6 lacking its 230 N-­‐terminal amino acids failed to be localized to the plasma membrane, and did not correct the β1,6-­‐glucan synthetic defect of kre6Δ, although it appeared stable (Kurita et al. 2011). It was concluded that transport of Kre6 to the plasma membrane is necessary for the protein to fulfill its role in β1,6-­‐glucan synthesis (Kurita et al. 2002). Localization of Skn1 has not been explored in detail. Skn1 and plant defensin resistance. skn1Δ, but not kre6Δ strains, are defective in M(IP)2C synthesis and resistant to a plant defensin that interacts with this sphingolipid to exert its antifungal activity (Thevissen et al. 2005). Defensin-­‐susceptibility is unconnected with cellular β1,6-­‐glucan content because other β1,6-­‐glucan synthesis mutants are defensin-­‐sensitive (Thevissen et al. 2005). Plasma membrane protein Kre1: Kre1 as receptor for K1 killer toxin. Membrane anchored Kre1 has an additional role as receptor for K1 killer toxin. Spheroplasts of kre1Δ cells are resistant to this toxin, but expression of the C-­‐terminal 63 amino acids of Kre1 was sufficient to make spheroplasts, but not intact cells, toxin sensitive again, leading to the proposal that Kre1’s GPI-­‐modified C-­‐terminus serves as the membrane receptor for K1 toxin after initial toxin binding to β1,6-­‐glucan (Breinig et al. 2002). Literature Cited Breinig, F., Tipper D. J., Schmitt, M. J., 2002 Kre1p, the plasma membrane receptor for the yeast K1 viral toxin. Cell 108: 395-­‐
405. Pasikowska, M., Palamarczyk, G., Lehle, L. (2012) The essential endoplasmic reticulum chaperone Rot1 is required for protein N-­‐ and O-­‐glycosylation in yeast. Glycobiology 22: 939-­‐947. 42 SI P. Orlean Takeuchi, M., Kimata, Y., Kohno, K., 2008 Saccharomyces cerevisiae Rot1 is an essential molecular chaperone in the endoplasmic reticulum. Mol. Biol. Cell 19: 3514-­‐3525. P. Orlean 43 SI File S9 Cell Wall-­‐Active and Nonenzymatic Surface Proteins and Their Functions This Supporting File contains additional information and discussion related to Cell Wall-­‐Active and Nonenzymatic Surface Proteins and Their Functions. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Known and predicted enzymes Chitinases: S. cerevisiae’s two chitinases, Cts1 and Cts2, are both members of GH Family 18, but of the two, Cts1 resembles plant-­‐
type chitinases, whereas the predicted Cts2 protein is more similar to the bacterial chitinase subfamily (Hurtado-­‐Guerrero and van Aalten, 2007). Cts1 has endochitinase activity, a pH optimum of 2.5, and is more active on nascent than on preformed chitin (Correa et al. 1982). The structure of the catalytic domain, which has chitinase activity on its own, has been determined (Hurtado-­‐Guerrero and van Aalten, 2007). Little is known about Cts2, but because CTS2 complements a defect in the sporulation-­‐specific chitinase of Ashbya gossypii (Dünkler et al. 2008), Cts2 may have a role in sporulation. β 1,3-­‐glucanases: Exg1, Exg2 and Ssg/Spr1 exo-­‐β1,3-­‐glucanases: These proteins are members of GH Family 5 and were originally characterized biochemically as exo-­‐β1,3-­‐glucanases (Larriba et al. 1995). Exg1 is a soluble cell wall protein released upon treatment with dithiothreitol (Cappellaro et al. 1998), whereas Exg2 may normally be membrane-­‐ or wall-­‐anchored because it has a potential GPI attachment site (Caro et al. 1997), whose deletion results in release of the protein into the medium (Larriba et al. 1995). Single or double null mutants in EXG1 and EXG2 have no obvious defects, although exg1Δ cells have slightly elevated levels of β1,6 glucan and EXG1 overexpressers lower amounts of that polymer. This, together with the finding that the Exg proteins can act on the β1,6-­‐glucan pustulan in vitro (Nebreda et al. 1986), raises the possibility that Exg1 and Exg2 have roles in β-­‐glucan remodeling (Jiang et al. 1995; Lesage and Bussey, 2006). Ssg1/Spr1 is a sporulation-­‐specific protein. Its mRNA is expressed late in sporulation, and homozygous null diploids show a delay in the onset of ascus formation (Muthukumar et al. 1993; San Segundo et al. 1993). Bgl2, Scw4, Scw10 endo-­‐β1,3-­‐glucanases:
These proteins are members of GH Family 17. Scw4, Scw10, and Bgl2 can be extracted from the wall with dithiothreitol (Capellaro et al. 1998), suggesting wall association via disulfides. However, a population of Scw4 and Scw10 44 SI P. Orlean resists extraction by hot SDS and β-­‐mercaptoethanol, and is released instead by mild alkali or by β1,3-­‐glucanase digestion, indicating a covalent linkage to β1,3-­‐glucan (Yin et al. 2005). However, Scw4 and Scw10 lack PIR sequences. Purified Bgl2 binds both β1,3-­‐glucan and chitin (Klebl and Tanner, 1989), but whether these non-­‐covalent interactions represent an additional mode of wall association, or reflect an enzyme-­‐substrate interaction, is unexplored. Levels of Bgl2 and Scw10 need to be balanced in order to ensure cell wall stability (Sestak et al. 2004). This proposal is based on the findings that deletion of BGL2 in the scw4Δ scw10Δ background (but not of SCW11, EXG1, CRH1, or CRH2) alleviated many of the phenotypes of that double mutant, that overexpression of BGL2 is lethal in a wild type background, and that high level expression of SCW10 in bgl2Δ significantly increases the strain’s CFW sensitivity (Klebl and Tanner, 1989; Sestak et al. 2004). Bgl2 and Scw10 may also contribute to compensatory responses to mutationally induced wall stress, because BGL2 and SCW10, as well as EXT1 and CRH1, are upregulated in mnn9, kre6, mnn9, and gas1 mutants (Lagorce et al. 2003). What Bgl2 and Scw10’s precise biochemical roles are, and how they antagonize one another, are intriguing questions. Eng1/Dse4 and Eng2/Acf2 endo-­‐β1,3-­‐glucanases: These two related proteins are members of GH family 81. ENG1 expression is highest at the M to G1-­‐phase transition and shut down during sporulation. Eng1 localizes to the daughter side of the septum, consistent with a hydrolytic role during cell separation (see Septum formation; Baladron et al. 2002). Eng2 recognizes β1,3-­‐glucans of at least five residues and releases trisaccharides from the non-­‐reducing end of the substrate, but has no detectable transglycosidase activity (Martín-­‐Cuadrado et al. 2008). Gas1 family β 1,3-­‐glucanosyltransferases: Domain organization and mechanism of Gas proteins. Gas1 and its four paralogues, Gas2, Gas3, Gas, 4, and Gas5 (Popolo and Vai, 1999), are members of the GH Family 72. The catalytic domain of Gas proteins lies in their N-­‐terminal half, and in the case of Gas1 and Gas2, is followed by a cysteine-­‐rich domain that is a member of the CBM43 group of carbohydrate binding modules. The other Gas proteins lack this module but have a serine and threonine-­‐rich sequence instead, and Gas1 has both (Popolo and Vai, 1999). The biochemical activity of Gas proteins was first defined for the Aspergillus fumigatus Gas1 homologue, Gel1, but S. cerevisiae Gas1, Gas2, Gas4, and Gas5 all proved to carry out the same reaction in vitro (Mouyna et al. 2000; Carotti et al. 2004; Ragni et al. 2007b; Mazan et al. 2011). The proteins have β1,3-­‐glucanosyltransfer or “elongase” activity, which involves cleavage of a β1,3 glucosidic linkage within a β1,3-­‐glucan chain, then transfer of the newly generated reducing end of the P. Orlean 45 SI cleaved glycan to the non-­‐reducing end of another β1,3 glucan molecule, thus extending the acceptor β1,3-­‐glucan chain (Mouyna et al. 2000). The structure of a soluble form of Gas2 in complex with β1,3-­‐gluco-­‐oligosaccharides revealed the presence of two oligosaccharide binding sites and led to a base-­‐occlusion hypothesis for how transglycosylation could be favored over hydrolysis. In the hypothesized mechanism, one binding site is occupied by the donor glucan, which is hydrolyzed with formation of an enzyme-­‐oligosaccharide intermediate, whereupon the other, acceptor, site is transiently filled by the second product of the hydrolysis reaction. Occupancy of the acceptor site has the effect of occluding the catalytic base on the enzyme, preventing any incoming water molecule from being activated for nucleophilic attack on the enzyme-­‐saccharide intermediate. The gluco-­‐oligosaccharide in the acceptor site is then displaced by a longer and tighter binding acceptor glucan with concomitant formation of the new β1,3-­‐glucosidic linkage (Hurtado-­‐Guerrero et al. 2009). In the case of Gas1 and Gas2, the cysteine-­‐rich domain is necessary for catalytic activity, being required for proper folding of the catalytic domain, for substrate binding, or for both (Popolo et al. 2008). This domain, however, is not necessary for activity of Gas4 or Gas5, which lack it, and, because Gas4 and Gas5 generate profiles of oligosaccharides from β1,3-­‐gluco-­‐
oligosaccharide substrates that are different from those released by Gas1 and Gas2, it is possible that the cysteine-­‐rich domain influences cleavage site preference (Ragni et al. 2007b). Nonetheless, expression of Gas4, but not Gas2, in a gas1Δ strain fully complemented the gas1Δ growth defect in media with a pH of 6.5 or above (Ragni et al. 2007a).
Localization of Gas1. Gas1 fused to GFP but retaining its N-­‐ and C-­‐terminal signal sequences is detectable in the lateral wall, in the chitin ring in small-­‐budded cells, and near the primary septum, and remains in the bud scar after cell separation (Rolli et al. 2009). Gas1 localization to the chitin ring and bud scars was abolished in cells lacking the chitin-­‐β1,3-­‐glucan cross-­‐
linkers Crh1 and Crh2, suggesting that Gas1 anchorage to chitin was dependent on linkage of a Gas1-­‐β1,6-­‐glucan-­‐β1,3-­‐glucan complex to chitin (Rolli et al. 2009). Consistent with this, Gas1 was shed into the medium from chs3Δ cells, which are unable to make the chitin known to be cross-­‐linked to β-­‐glucan (Cabib and Duran, 2005). Because the released Gas1 was not significantly larger than Gas1 in lysates of wild type cells (Rolli et al., 2009), the β1,6-­‐glucan-­‐β1,3-­‐glucan presumed to link the protein to chitin must be quite small. Some Gas1 was also released from chs2Δ cells, suggesting that localization of Gas1 near the primary septum requires Chs2-­‐dependent chitin synthesis (Rolli et al. 2009). However, because the chitin made by Chs2 is free of cross-­‐
links (Cabib and Duran, 2005), its association with Gas1 would be indirect. Cell-­‐associated Gas1 was distributed throughout the remedial septum made in chs2Δ cells (Section V.1.a). Intriguingly, Gas1 was also shed from chs1Δ cells, though at reduced levels when the medium was buffered to lower chitinase activity. Amounts and localization of cell-­‐associated Gas1 appeared 46 SI P. Orlean unchanged, however, presumably because Chs2 and Chs3 still make chitin. Nonetheless, this observation indicates that Chs1 or its product contribute to wall association of some Gas1 (Rolli et al. 2009). Functions of Gas2, Gas3, Gas4, and Gas5. The following findings indicate that Gas5 and Gas3 have wall-­‐related functions in vegetative cells. GAS5 is expressed during vegetative growth but repressed during sporulation, and gas5Δ strains are Calcofluor White sensitive (Caro et al. 1997). Purified Gas3 is inactive (Ragni et al. 2007b), and gas3Δ strains make no genetic interactions with strains with single or double deletions in other GAS genes (Rolli et al. 2010). Moreover, Gas3 cannot substitute for Gas1, but overexpression in gas1Δ of wild type GAS3 or a gas3 mutant encoding catalytically inactive Gas3 exacerbated the gas1Δ growth defect, indicating that high levels of Gas3 are toxic (Rolli et al. 2010). Gas2 and Gas4 have overlapping functions in ascospore wall assembly. Their genes are expressed only during sporulation, and although diploids homozygous for single GAS2 or GAS4 deletions sporulate normally, diploids lacking both Gas2 and Gas4 have a severe sporulation defect (Ragni et al. 2007a). The inner glucan layer of the spore wall from by double homozygous gas2 gas4 nulls was disorganized and detached from chitosan, and dityrosine, though present, was less abundant and diffusely distributed. The absence of β1,3-­‐glucanosyltransferase activity may result in shorter β1,3-­‐glucan chains that are more loosely associated with chitosan. Gas2 and Gas4 likely need to be GPI anchored to fulfill their key roles in ascospore wall formation, which in part explains the severe sporulation defect of homozygous gpi1/gpi1 and gpi2/gpi2 diploids (Leidich and Orlean, 1996). Because such diploids lack dityrosine, additional GPI-­‐proteins must normally be involved in ascospore wall assembly. Yapsin aspartyl proteases: Yapsin processing. Yapsins are synthesized as zymogens and undergo proteolytic processing to generate a mature active enzyme. The steps include removal of a propeptide and excision of an internal segment flanked by basic amino acids that separates the enzyme’s two catalytic domains, which remain disulfide-­‐linked (Gagnon-­‐Arsenault et al. 2006, 2008). In the case of Yps1, the propeptide removal and excision steps are likely autocatalytic at an environmental pH of 3, but involve other proteases, including yapsins, at pH 6 (Gagnon-­‐Arsenault et al. 2008). Cell wall phenotypes of yapsin-­‐deficient strains. Strains lacking individual yapsin genes are sensitive to various cell wall disrupting agents, though their sensitivity profiles differ. For example, yps7Δ is the only yps null hypersensitive to CFW, but yps1Δ the only mutant sensitive to the β1,3-­‐glucan synthase inhibitor caspofungin (Krysan et al. 2005). The quintuple yps1Δ yps2Δ yps3Δ yps6Δ yps7Δ null mutant is viable, but undergoes osmotically remedial lysis at 30°C, as does the yps1Δ yps2Δ P. Orlean 47 SI yps3Δ triple deletion strain, and to a slightly lesser extent, the yps1Δ yps2Δ double null (Krysan et al. 2005). The temperature-­‐
sensitive lysis phenotype of strains lacking multiple yapsins is consistent with a role for these proteins when cell walls are stressed, and indeed, expression of YPS1, YPS2, YPS3, and YPS6 is upregulated under such conditions (Garcia et al. 2004; Krysan et al. 2005). Non-­‐enzymatic CWPs Structural GPI proteins: Sps2 family: Ecm33. Mannan outer chains produced by ecm33Δ cells are slightly smaller than normal, although O-­‐mannosylation and core-­‐type N-­‐glycans are not affected. Epitope-­‐tagged Pst1 is most abundant at the surface of buds, but Ecm33’s localization is uncertain because tagging Ecm33 abolishes its in vivo function (Pardo et al. 2004). Ecm33 occurs in both plasma membrane and wall-­‐anchored forms, but must retain its GPI anchor and plasma membrane localization for in vivo function (see Incorporation of GPI proteins into the wall; Terashima et al. 2003; Yin et al. 2005). Expression of a minimal amount of GPI-­‐
anchored Ecm33 may be necessary for growth at high temperature, because the temperature-­‐sensitivity of mcd4, gpi7, gpi13 and gpi14 mutants is suppressed by overexpression of ECM33 (Toh-­‐e & Oguchi, 2002; A. Sembrano and P. Orlean, unpublished). Tip1 family: Localization of Cwp2 and Tip1 is influenced by the timing of their expression. A swap of the promoters of CWP2 and TIP1 caused these genes’ products to exchange their cellular location, indicating that the localization of Cwp2 and Tip1, and perhaps that of other CWPs, is influenced by the timing of their expression in the cell cycle (Smits et al. 2006). Cwp1, however, is localized to the birth scar in a manner that depends on normal septum formation, but, because neither Tip1 nor Cwp2 is targeted to the birth scar when expressed behind CWP1‘s promoter, additional CWP1 sequences are required for Cwp1 localization (Smits et al. 2006). Ccw12: Structural features of Ccw12. Ccw12 has a predicted mass of 13 kDa but migrates on denaturing polyacrylamide gels with an apparent molecular weight of a least 200 kDa. Elimination of Ccw12’s three N-­‐linked sites shows that N-­‐linked glycans are mostly responsible for this apparent size increase, but these modifications are not necessary for in vivo function, because Ccw12 lacking its N-­‐linked sites complements ccw12Δ phenotypes (Ragni et al. 2007c). O-­‐mannosylation contributes some 42 kDa to the apparent size of Ccw12 (Hagen et al. 2004). The protein is not obviously related to any known enzymes, but contains 48 SI P. Orlean two repeats of the sequence TTEAPKNGTSTAAP (Mrša et al. 1999). Deletion of one or both of these does not affect cross-­‐
linkage Ccw12 to the wall, but the repeats are nonetheless critical for in vivo function because proteins lacking them do not restore the growth and cell wall defects of ccw12Δ (Ragni et al. 2007c). Four sequences similar to the Ccw12 repeat are present in Sed1 (Mrša et al. 1999; Ragni et al. 2007c). Certain Tip1 family members and Slr1 also migrate in denaturing polyacrylamide gels with much higher molecular weights than would be expected (van der Vaart et al. 1995; Terashima et al. 2002). A new mechanism for compensating loss of multiple GPI-­‐CWP uncovered in ccw12Δ . Deletion of additional genes for GPI-­‐CWP in the ccw12Δ background uncovered a mechanism for compensating for loss of multiple GPI-­‐CWPs. Rather than showing an exacerbated phenotype, the ccw12Δ ccw14Δ double null was less sensitive to CFW compared with ccw12Δ, and the ccw12Δ ccw14Δ dan1Δ mutant showed wild type levels of sensitivity to CFW and nearly normal levels of chitin. Moreover, additional deletion of CWP1 and TIP1 had no further effect on CFW sensitivity, although walls of the quintuple mutant had a thicker inner glucan layer and a thinner but more ragged outer mannoprotein layer (Hagen et al. 2004). It seems that although loss of Ccw12 alone activates the CWI pathway-­‐mediated chitin stress response (Ragni et al. 2007c, 2011; see Chitin synthesis in response to cell wall stress), deletion of additional GPI-­‐CWP genes forces cells over a threshold that leads to triggering of a new compensatory response, whereupon the chitin response becomes less important. This new response depends on Sed1 and the non-­‐GPI-­‐CWP Srl1. Not only is their expression upregulated in the ccw12Δ ccw14Δ dan1Δ cwp1Δ tip1Δ strain, but deletion of either in the ccw12Δ ccw14Δ dan1Δ background reverts the strain to the high-­‐chitin phenotype of ccw12Δ (Hagen et al. 2004). In addition, the cell wall remodeling genes SCW10 and BGL2 are upregulated and CRH2 downregulated, suggesting that the response involves alterations of the structure of the β-­‐glucan layer (Hagen et al. 2004). More generally, the phenotypes of the multiple GPI-­‐CWP mutants indicate that GPI-­‐CWPs have a collective role in maintaining cell wall stability (Lesage and Bussey, 2006; Ragni et al. 2007c). Ccw12 and Slr1 also have parallel functions in a pathway that relieves defects in a polarized morphogenesis signaling network (see Slr1). Other non-­‐enzymatic GPI-­‐proteins: Ccw14/Ssr1/Icwp as an inner cell wall protein. A monoclonal antibody that recognizes Ccw14/Ssr1 on immunoblots does not detect the protein on intact cells, whereas it does have access to the glycoprotein in tunicamycin-­‐treated cells or in mnn1 mnn9 mutants (Moukadiri et al. 1997). Assuming that the antibody would have had access to its epitope on Ccw14/Ssr1 if the protein were at the surface of wild type cells, this finding is consistent with Ccw14/Ssr1 being a protein of the inner cell wall P. Orlean 49 SI (Moukadiri et al. 1997). Flocculins and agglutinins: Roles and interactions of Aga1 and Fig2 in mating. Deletion of FIG2 in MATa cells with the W303 background, but not MATa cells, increases the agglutinability of MATα cells, suggesting a role for Fig2 in attenuating agglutination of MATa cells (Erdman et al. 1998; Jue and Lipke, 2002). Both Aga1 and Fig2 have an additional, additive role in mating in MATα strains that is unconnected with Aga2, because simultaneous deletion of AGA1 and FIG2 in certain MATα sag1Δ backgrounds leads to a severe mating defect on solid medium, whereas individually deleting the AGA1 and FIG2 in those strain backgrounds does not (Guo et al. 2000). An explanation for the expanded roles for Aga1 and Fig2 in mating came from detection of heterotypic adhesive interactions between Aga1 and Fig2, and homotypic interactions between Fig2 and Fig2, which are mediated by WPCL and CX4C domains present in both proteins (Huang et al. 2009). Non-­‐GPI-­‐CWP: PIR proteins: PIR protein localization. Fusions of Pir1 and Pir2 with red fluorescent protein are found at bud scars of both haploid and diploid cells, with Pir1 being localized inside the chitin ring. This localization of Pir1 is independent of normal chitin ring and primary septum formation because the protein is still transported to the budding site in chs2Δ and chs3Δ cells, although in the absence of the chitin ring in chs3Δ, Pir1 no longer shows a ring-­‐like distribution (Sumita et al. 2005). Some Pir1 and Pir2, and most Pir3, are also present in lateral walls, where these proteins can be detected by immunoelectron microscopy using antibody to Pir3 (Yun et al. 1997). Pir4 has been reported to show a uniform distribution at the cell surface, but in one study, this distribution was restricted to growing buds (Moukadiri et al. 1999; Sumita et al. 2005). A Kex2 processing site in PIR proteins. The four PIR proteins contain a site for processing by the Kex2 protease, but although Kex2 acts on the PIR proteins in vivo, wall localization of these proteins is unaffected in kex2Δ, so the significance of this processing event is unclear (Mrša et al. 1997). Scw3 (Sun4): SUN proteins. Members of this family of highly glycosylated proteins have a common C-­‐terminal domain of some 250 amino acids in which the spacing of four cysteines is conserved (Velours et al. 2002). The SUN proteins other than Scw3/Sun4 (Sim1, Uth1, and Nca3) have been implicated in various cellular functions unrelated to the cell wall, but SUN family members have been assumed to be glucanases because they are homologous to Candida wickerhamii BglA, an additional protein 50 SI P. Orlean identified in a screen of a cDNA expression library for proteins that reacted with an antibody to a cell-­‐bound β-­‐glucosidase (Skory and Freer, 1995). However, glycosidase activity has not been verified for BglA and the SUN proteins show no homology to any carbohydrate active enzymes, making it doubtful they are glycosidases. Literature Cited Garcia, R., Bermejo, C., Grau, C., Perez, R., Rodriguez-­‐Pena, et al., 2004 The global transcriptional response to transient cell wall damage in Saccharomyces cerevisiae and its regulation by the cell integrity signaling pathway. J. Biol. Chem. 279: 15183-­‐15195. Huang, G., Dougherty, S. D., Erdman, S. E., 2009 Conserved WCPL and CX4C domains mediate several mating adhesin interactions in Saccharomyces cerevisiae. Genetics 182: 173-­‐189. Hurtado-­‐Guerrero, R., Schüttelkopf, A. W., Mouyna, I., Ibrahim, A. F. M., Shepherd, S., et al., 2009 Molecular mechanisms of yeast cell wall glucan remodeling. J. Biol. Chem. 284: 8461-­‐8469. Hurtado-­‐Guerrero, R., van Aalten, D. M., 2007 Structure of Saccharomyces cerevisiae chitinase 1 and screening-­‐based discovery of potent inhibitors. Chem. Biol. 14: 589-­‐599. Martín-­‐Cuadrado, A. B., Fontaine, T., Esteban, P. F., del Dedo, J. E., de Medina-­‐Redondo, M., et al., 2008 Characterization of the endo-­‐β-­‐1,3-­‐glucanase activity of S. cerevisiae Eng2 and other members of the GH81 family. Fungal Genet. Biol. 45: 542-­‐553. Muthukumar, G., Suhng, S. H., Magee, P. T., Jewell, R. D., Primerano, D. A., 1993 The Saccharomyces cerevisiae SPR1 gene encodes a sporulation-­‐specific exo-­‐1,3-­‐β-­‐glucanase which contributes to ascospore thermoresistance. J. Bacteriol. 175: 386-­‐
394. Nebreda, A. R., Villa, T. G., Villanueva, J. R., del Rey, F., 1986 Cloning of genes related to exo-­‐β-­‐glucanase production in Saccharomyces cerevisiae: characterization of an exo-­‐β-­‐glucanase structural gene. Gene 47: 245-­‐529. P. Orlean 51 SI Popolo, L., Ragni, E., Carotti, C., Palomares, O., Aardema, R., et al., 2008 Disulfide bond structure and domain organization of yeast β(1,3)-­‐glucanosyltransferases involved in cell wall biogenesis. J. Biol. Chem. 283: 18553-­‐18565. Rolli, E., Ragni, E., Rodriguez-­‐Peña, J. M., Arroyo, J., Popolo, L., 2010 GAS3, a developmentally regulated gene, encodes a highly mannosylated and inactive protein of the Gas family of Saccharomyces cerevisiae. Yeast 27: 597-­‐610. San Segundo, P., Correa, J., Vazquez de Aldana, C. R., del Rey, F., 1993 SSG1, a gene encoding a sporulation-­‐specific 1,3-­‐β-­‐
glucanase in Saccharomyces cerevisiae. J. Bacteriol. 175: 3823-­‐3837. Skory, C. D., Freer, S. N., 1995 Cloning and characterization of a gene encoding a cell-­‐bound, extracellular β-­‐glucosidase in the yeast Candida wickerhamii. Appl. Environ. Microbiol. 61: 518-­‐525. 52 SI P. Orlean Table S1 Proteins involved in cell wall biogenesis in Saccharomyces cerevisiae Process or protein type Protein name Activity or Function Ugp1 UDPGlc pyrophosphorylase Pmi40 phosphomannose isomerase Sec53 phosphomannomutase 1
CAZy Family Precursor supply Psa1/Srb1/Vig9 GDP-­‐Man pyrophosphorylase Gfa1 glutamine: Fru-­‐6-­‐P amidotransferase Gna1 GlcN-­‐6-­‐P N-­‐acetylase Agm1/Pcm1 GlcNAc phosphate mutase Uap1/Qri1 UDPGlcNAc pyrophosphorylase Rer2 cis-­‐prenyltransferase (Dol10-­‐14) Srt1 cis-­‐prenyltransferase (Dol19-­‐22) Dfg10 dehydrodolichol reductase Sec59 Dol-­‐kinase Cwh8/Cax4 Dolichyl pyrophosphate phosphatase Dpm1 GDP-­‐mannose:dolichyl-­‐phosphate Man-­‐T GT2 Alg5 UDP-­‐glucose:dolichyl-­‐phosphate Glc-­‐T GT2 Yea4 UDP-­‐GlcNAc transporter GT1 Vrg4/Vig4 GDP-­‐Man transporter Gda1 GDPase Ynd1 Apyrase Alg7 UDP-­‐GlcNAc: Dol-­‐P GlcNAc-­‐1-­‐P-­‐T N-­‐glycosylation Alg13 + Alg14 UDP-­‐GlcNAc: Dol-­‐PP-­‐GlcNAc β1,4-­‐GlcNAc-­‐T P. Orlean 53 SI 54 SI Alg1 GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2 β1,4-­‐Man-­‐T Alg2 Alg11 GT33 GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2Man α1,3-­‐Man-­‐T and GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2Man2 α1,6-­‐Man-­‐T GT4 GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2Man3 α1,2-­‐Man-­‐T and GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2Man4 α1,2-­‐Man-­‐T GT4 Rft1 Candidate Dol-­‐PP-­‐oligosaccharide flippase Alg3 Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man5 α1,3-­‐Man-­‐T GT58 Alg9 Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man6 α1,2-­‐Man-­‐T and Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man8 α1,2-­‐Man-­‐T GT22 Alg12 Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man7 α1,6-­‐Man-­‐T GT22 Alg6 Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man9 α1,3-­‐Glc-­‐T GT57 Alg8 Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man9Glc α1,3-­‐Glc-­‐T GT57 Alg10 Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man9Glc2 α1,2-­‐Glc-­‐T GT59 Stt3 OST catalytic subunit GT66 Wbp1 OST subunit Swp1 OST subunit Ost1 OST subunit Ost2 OST subunit Ost3 OST subunit; cysteine oxidoreductase Ost6 OST subunit; cysteine oxidoreductase Gls1/Cwh41 ER glucosidase I (α1,2 exoglucosidase); indirectly affects β1,6-­‐glucan GH63 Gls2/Rot2 ER glucosidase II (α1,3 exoglucosidase α-­‐subunit); indirectly affects β1,6-­‐glucan GH31 Gtb1 ER glucosidase II (regulatory subunit) Mns1 ER α-­‐mannosidase I GH47 GH47 Htm1/Mnl1 ER-­‐degradation enhancing a-­‐mannosidase-­‐like protein Yos9 Lectin, recognizes α1,6-­‐Man on glucosidase II product, targets misfolded protein for ERAD Png1 Cytosolic peptide N-­‐glycanase Och1 Initiating α1,6-­‐Man-­‐T GT32 Mnn9 M-­‐Pol I α1,6-­‐Man-­‐T GT62 P. Orlean Van1 M-­‐Pol I α1,6-­‐Man-­‐T GT62 Mnn9 M-­‐Pol II α1,6-­‐Man-­‐T GT62 Anp1 M-­‐Pol II α1,6-­‐Man-­‐T GT62 Mnn10 M-­‐Pol II α1,6-­‐Man-­‐T GT34 Mnn11 M-­‐Pol II α1,6-­‐Man-­‐T GT34 Hoc1 M-­‐Pol II α1,6-­‐Man-­‐T GT32 Mnn2 α1,2-­‐Man-­‐T; Mnn1 subfamily; major role in mannan side chain branching GT71 Mnn5 α1,2-­‐Man-­‐T; Mnn1 subfamily; major role in mannan side chain branching GT71 Mnn4 Positive regulator of Man phosphorylation Mnn6/Ktr6 α-­‐Man-­‐P-­‐T; acts on N-­‐ and O-­‐glycans in Golgi GT15 Mnn1 α1,3-­‐Man-­‐T; acts on N-­‐ and O-­‐glycans in Golgi GT71 Kre2/Mnt1 α1,2-­‐Man-­‐T; acts on N-­‐ and O-­‐glycans in Golgi GT15 Ktr1 α1,2-­‐Man-­‐T; acts on N-­‐ and O-­‐glycans in Golgi GT15 Ktr2 α1,2-­‐Man-­‐T; acts on N-­‐glycans in Golgi GT15 Ktr3 α1,2-­‐Man-­‐T; acts on N-­‐ and O-­‐glycans in Golgi GT15 Yur1 α1,2-­‐Man-­‐T; acts on N-­‐glycans in Golgi GT15 Ktr4 Putative α-­‐ManT GT15 Ktr5 Putative α-­‐ManT GT15 Ktr7 Putative α-­‐ManT GT15 Gnt1 GlcNAc-­‐T GT8 Vrg4 GDP-­‐Man transporter Gda1 GDPase Ynd1 Apyrase Pmt1 Dol-­‐P-­‐Man: protein: O-­‐Man-­‐T; Pmt1 family GT39 Pmt2 Dol-­‐P-­‐Man: protein: O-­‐Man-­‐T; Pmt2 family GT39 O-­‐mannosylation P. Orlean 55 SI Pmt3 Dol-­‐P-­‐Man: protein: O-­‐Man-­‐T; Pmt2 family Pmt4 Pmt5 GT39 Dol-­‐P-­‐Man: protein: O-­‐Man-­‐T; specific for membrane proteins GT39 Dol-­‐P-­‐Man: protein: O-­‐Man-­‐T; Pmt1 family GT39 Pmt6 Dol-­‐P-­‐Man: protein: O-­‐Man-­‐T; Pmt2 family GT39 Mnt2 α1,3-­‐Man-­‐T; Mnn1 subfamily; acts on O-­‐glycans in Golgi GT71 Mnt3 α1,3-­‐Man-­‐T; Mnn1 subfamily; acts on O-­‐glycans in Golgi GT71
Gpi1 GPI-­‐Gnt subunit Gpi2 GPI-­‐Gnt subunit Gpi3 GPI-­‐Gnt subunit, UDP-­‐GlcNAc: Ptd-­‐Ins α1,6-­‐GlcNAc transferase GT4 Gpi15 GPI-­‐Gnt subunit Gpi19 GPI-­‐Gnt subunit Eri1 GPI-­‐Gnt subunit Ras2 Negative regulator of GPI-­‐Gnt Gpi12 GPI-­‐Ins-­‐deacetylase Gwt1 GPI-­‐Ins-­‐acyltransferase Gpi14 GPI-­‐ManT-­‐I: Dol-­‐P-­‐Man: GlcN-­‐Ptd-­‐(acyl)Ins α1,4-­‐Man-­‐T GT50 Pbn1 Putative subunit of GPI-­‐Man-­‐T-­‐I Arv1 Proposed to present GlcN-­‐(acyl)PI to Gpi14 Mcd4 GPI-­‐Etn-­‐P-­‐T-­‐I Gpi18 GPI-­‐ManT-­‐II: Dol-­‐P-­‐Man: Man-­‐GlcN-­‐Ptd-­‐(acyl)Ins α1,6-­‐Man-­‐T GT76 Pga1 GPI-­‐ManT-­‐II subunit Gpi10 GPI-­‐Man-­‐T-­‐III: Dol-­‐P-­‐Man: Man2-­‐GlcN-­‐Ptd-­‐(acyl)Ins α1,2-­‐Man-­‐T GT22 Smp3 GPI-­‐Man-­‐T-­‐IV: Dol-­‐P-­‐Man: Man3-­‐GlcN-­‐Ptd-­‐(acyl)Ins α1,2-­‐Man-­‐T GT22 Gpi13 GPI-­‐Etn-­‐P-­‐T-­‐III Gpi11 Subunit of GPI-­‐Etn-­‐P-­‐T-­‐II and GPI-­‐Etn-­‐P-­‐T-­‐III GPI anchoring 56 SI P. Orlean Gpi7 GPI-­‐Etn-­‐P-­‐T-­‐II Gpi8 GPI transamidase catalytic subunit Gaa1 GPI transamidase subunit Gab1 GPI transamidase subunit Gpi16 GPI transamidase subunit Gpi17 GPI transamidase subunit Bst1 GlcN-­‐(acyl)PI inositol deacylase Per1 Removes acyl chain at sn-­‐2 position of protein-­‐bound GPIs Gup1 MBOAT O-­‐acyltransferase, transfers C26 acyl chain to sn-­‐2 position of protein-­‐bound GPIs Cwh43 Replaces GPI diacylglycerol with ceramide Cdc1 Homologue of mammalian PGAP5; possible GPI-­‐Etn-­‐P phosphodiesterase Ted1 Homologue of mammalian PGAP5; possible GPI-­‐Etn-­‐P phosphodiesterase Chitin and chitosan synthesis Chs1 Chitin synthase I catalytic protein GT2 Chs2 Chitin synthase II catalytic protein GT2 Chs3 Chitin synthase catalytic subunit GT2 Cdk1 Mitotic protein kinase, phosphorylates Chs2 Cdc14 Phosphoprotein phosphatase, dephosphorylates Chs2 Dbf2 Mitotic exit kinase, phosphorylates Chs2 Inn1 Localized to mother cell-­‐bud junction with Chs2 and Cyk3, implicated in Chs2 activation Cyk3 Localized to mother cell-­‐bud junction with Chs2 and Inn1, implicated in Chs2 activation Pfa4 Protein acyltransferase, palmitoylates Chs3 Chs7 Chaperone required for ER exit of Chs3 Rcr1 ER protein, small negatve effect on Chs3-­‐dependent chitin synthesis Yea4 ER protein and UDP-­‐GlcNAc transporter, yea4Δ has 65% of wild type levels of chitin. Chs5 Exomer component, involved in Chs3 trafficking P. Orlean 57 SI Chs6 Exomer component, involved in Chs3 trafficking Chs4/Skt5 Prenylated protein that interacts with, activates, and anchors Chs3 to septin ring Bni4 Scaffold protein, tethers Chs3 and Chs4 to septins Shc1 Sporulation-­‐specific Chs4 homologue Cda1 Chitin de-­‐N-­‐acetylase Cda2 Chitin de-­‐N-­‐acetylase β-­‐1,3 glucan synthesis Fks1/Gsc1/Cwh53/ Etg1/Pbr1 Probable β1,3-­‐glucan synthase, major role in vegetative cells GT48 Fks2/Gsc2 Probable β1,3-­‐glucan synthase, stress-­‐induced, role in sporulation GT48 Fks3 Probable β1,3-­‐glucan synthase, role in sporulation GT48 Rho1 GTPase; activator of Fks1 and Fks2 Kre5 Diverged UDP-­‐Glc: glycoprotein Glc-­‐T homologue GT24 Rot1 Fungus-­‐specific ER chaperone Big1 Fungus-­‐specific ER chaperone Keg1 Fungus-­‐specific ER chaperone Kre6 Resembles β-­‐1,6/β-­‐1,3 glucanases GH16 Skn1 Sequence and functional Kre6 homologue; additional role in MIPC synthesis GH16 Kre9 Fungus-­‐specific O-­‐mannosylated protein Knh1 Kre9 homologue Kre1 GPI-­‐protein, secondary receptor for K1 killer toxin β-­‐1,6 glucan formation Glycosidases, cross-­‐linking enzymes, and proteases Cts1 Endo-­‐chitinase GH18 Cts2 Chitinase GH18 GH5 Exg1/Bgl1 58 SI Major exo-­‐β-­‐1,3-­‐glucanase of the cell wall; soluble P. Orlean Exg2 GPI-­‐anchored plasma membrane exo-­‐β1,3-­‐glucanase GH5 Ssg1/Spr1 Sporulation-­‐specific exo-­‐β-­‐1,3-­‐glucanase GH5 Bgl2 Endo-­‐β1,3-­‐glucanase; can make β1,6-­‐linked Glc side branch GH17 Scw4 Endo-­‐β1,3-­‐endoglucanase-­‐like GH17 Scw10 Endo-­‐β1,3-­‐endoglucanase-­‐like GH17 Scw11 Endo-­‐β1,3-­‐endoglucanase-­‐like GH17 Eng1/Dse4 Endo-­‐β1,3-­‐endoglucanase GH81 Eng2/Acf2 Endo-­‐β1,3-­‐endoglucanase GH81 Dcw1 GPI-­‐protein, resembles α1,6-­‐endomannanase GH76 Dfg5 GPI-­‐protein, resembles α1,6-­‐endomannanase; Dcw1 homologue GH76 Crh1 GPI-­‐protein, chitin β-­‐1,6/1,3-­‐glucanosyltransferase GH16 Crh2/Utr2 GPI-­‐protein, chitin β-­‐1,6/1,3-­‐glucanosyltransferase GH16 Crr1 GPI-­‐protein, chitin β-­‐1,6/1,3-­‐glucanosyltransferase; sporulation-­‐specific GH16 Gas1 GPI-­‐protein, β-­‐1,3-­‐glucanosyltransferase GH72 Gas2 GPI-­‐protein, β-­‐1,3-­‐glucanosyltransferase; sporulation specific GH72 Gas3 GPI-­‐protein, β-­‐1,3-­‐glucanosyltransferase GH72 Gas4 GPI-­‐protein, β-­‐1,3-­‐glucanosyltransferase; sporulation specific GH72 Gas5 GPI-­‐protein, β-­‐1,3-­‐glucanosyltransferase GH72 Yps1 GPI-­‐protein, yapsin aspartyl protease Yps2/Mkc7 GPI-­‐protein, yapsin aspartyl protease Yps3 GPI-­‐protein, yapsin aspartyl protease Yps6 GPI-­‐protein, yapsin aspartyl protease Ecm33 Sps2 family; structural/non-­‐enzymatic Pst1 Sps2 family; structural/non-­‐enzymatic Sps2 Sps2 family; structural/non-­‐enzymatic; required for ascospore wall formation GPI-­‐CWP P. Orlean 59 SI 60 SI Sps22 Sps2 family; structural/non-­‐enzymatic; required for ascospore wall formation Cwp1 Tip1 family Cwp2 Tip1 family Tip1 Tip1 family; anaerobically induced Tir1 Tip1 family; anaerobically induced Tir2 Tip1 family; anaerobically induced Tir3 Tip1 family; anaerobically induced Tir4 Tip1 family; anaerobically induced Dan1/Ccw13 Tip1 family; anaerobically induced Dan4 Tip1 family; anaerobically induced Sed1 Induced in stationary phase Spi1 Induced by stress with weak organic acids; related to Sed1 Ccw12 Major role in stabilizing walls of daughter cells walls and mating projections Ccw14/Ssr1 Inner cell wall protein Dse2 Daughter cell specific, role in cell separation Egt2 Daughter cell specific, role in cell separation Fit1 Iron binding Fit2 Iron binding Fit3 Iron binding Flo1 Flocculin Flo5 Flocculin Flo9 Flocculin Flo10 Flocculin Flo11/Muc1 Required for pseudohypha formation by diploids and agar invasion by haploids Aga1 MATa agglutinin subunit, disulfide-­‐linked to Aga2, which binds MATα agglutinin Sag1 Fig2 Aga1-­‐related adhesin P. Orlean Sag1 MATα agglutinin Non-­‐GPI-­‐CWP Pir1/Ccw6 “Protein with internal repeat”, ester-­‐linked via Glu (originally Gln in repeats) to β1,3-­‐glucan Pir2/Hsp150/Ccw7 “Protein with internal repeat”, ester-­‐linked via Glu (originally Gln in repeats) to β1,3-­‐glucan Pir3/Ccw8 “Protein with internal repeat”, ester-­‐linked via Glu (originally Gln in repeats) to β1,3-­‐glucan Pir4/Cis3/ Ccw5/Ccw11 One “internal repeat” sequence”, ester-­‐linked via Glu (originally Gln in repeats) to β1,3-­‐glucan Scw3/Sun4 Member of SUN family Srl1 Acts in parallel with Ccw12 in pathway operative when regulation of Ace2 and polarized morphogenesis are defective 1
CAZy glycosyltransferase (GT) and glycosylhydrolase (GH) families are defined in the Carbohydrate Active Enzymes database (http://www.cazy.org/) (Cantarel, B. L., Coutinho, P. M., Rancurel, C., Bernard, T., Lombard, V., et al., 2009 The Carbohydrate-­‐Active EnZymes database (CAZy): an expert resource for Glycogenomics. Nucleic Acids Res. 37: D233-­‐
238). P. Orlean 61 SI File S1 Precursors and Carrier Lipids This Supporting File contains additional information related to Precursors and Carrier Lipids. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Sugar nucleotides Regulation of glucosamine supply and chitin levels. Glucosamine supply is highly regulated and impacts chitin levels, which increase in response to mating pheromones and cell wall stress. Expression of GFA1 and AGM1 is upregulated upon treatment of MATa cells with α-­‐factor (Watzele and Tanner, 1989; Hoffman et al. 1994), and is accompanied by an increase in chitin deposition (Schekman and Brawley, 1979; Orlean et al. 1985). The cell wall stress-­‐induced increase in chitin synthesis (Popolo et al. 1997; Dallies et al. 1998; Kapteyn et al. 1999; see Wall Composition and Architecture) is also accompanied by elevated GFA1 expression (Terashima et al. 2000; Lagorce et al. 2002; Bulik et al. 2003). Elevation of glucosamine levels by other means also elicits increased chitin synthesis, for chitin levels are correlated with levels of expression of GFA1 itself (Lagorce et al. 2002; Bulik et al. 2003), and exogenous glucosamine also leads to increased chitin synthesis (Bulik et al. 2003). However, Bulik et al. (2003) found that chitin formation was not proportional to UDP-­‐GlcNAc concentration. These observations led to the conclusion that chitin synthesis is proportional to Gfa1 activity but that additional factors, for example a glucosamine metabolite or Gfa1 itself, must modulate chitin levels (Bulik et al. 2003). It is also formally possible that additional chitin is in a soluble or intracellular form and not detected in cell wall analyses. Dolichol and dolichol phosphate sugars Dolichol phosphate synthesis: Rer2 and Srt1. Biosynthesis of dolichol starts with the extension of trans farnesyl-­‐PP by successive addition of cis-­‐
isoprene units by the homologous cis-­‐prenyltransferases Rer2 and Srt1 (Sato et al. 1999; Schenk et al. 2001b). Rer2 is the dominant activity and makes dolichols with 10-­‐14 isoprene units, whereas dolichols made by Srt1 in cells lacking Rer2 contain 19-­‐22 isoprenes, like mammals. rer2Δ strains have severe defects in growth and in N-­‐ and O-­‐glycosylation, and SRT1 is a high-­‐
copy suppressor of rer2 mutants (Sato et al. 1999). The rer2Δ srt1Δ double null is inviable (Sato et al. 1999). Rer2 and Srt1 both behave as peripheral membrane proteins (Sato et al. 2001; Schenk et al. 2001b), but Rer2 is localized to the ER membrane, whereas Srt1 is detected in “lipid particles” (Sato et al. 2001). P. Orlean 1 SI Dfg10. Dfg10 has a steroid 5α reductase domain, and is responsible for much of the activity that reduces the α-­‐
isoprene unit of polyprenol activity. Both dfg10-­‐100 transposon insertion mutants and dfg10Δ strains underglycosylate carboxypeptidase Y to the same extent, and dolichol levels are decreased by 70% in dfg10-­‐100 cells, with a corresponding increase in unsaturated polyprenol (Cantagrel et al. 2010). The biosynthetic origin of the residual dolichol is not known. Membrane organization of Sec59 dolichol kinase. Sec59 is a multispanning membrane protein whose CTP-­‐binding site is oriented towards the cytoplasm (Shridas and Waechter, 2006). Dolichol chain length specificity of yeast glycosyltransferases and flippases. The enzymes that act after Rer2 and Srt1 can use shorter chain dolichols. Thus, the growth and glycosylation defects of rer2Δ cells can be complemented by expression of the E. coli cis-­‐isoprenyltransferase, which generates C55 isoprenoids, or of the Giardia homologue, which makes C55-­‐60 (Rush et al. 2010; Grabinska et al. 2010). The native glycosyltransferases and flippases must therefore also be able to use shorter chain dolichols as substrates. Dol-­‐P-­‐Man and Dol-­‐P-­‐Glc synthesis: Relationship between Dpm1 and Alg5. Alg5 and Dpm1 are most similar in their N-­‐terminal halves, which contain their GT-­‐A superfamily domain, but diverge in their C-­‐terminal halves. Both are likely to catalyze their reactions at the cytoplasmic face of the ER membrane. Literature Cited Grabinska, K. A., Cui, J., Chatterjee, A., Guan, Z., Raetz, C. R., et al., 2010 Molecular characterization of the cis-­‐prenyltransferase of Giardia lamblia. Glycobiology 20: 824-­‐832. Rush, J. S., Matveev, S., Guan, Z., Raetz, C. R. H., Waechter, C. J. 2010 Expression of functional bacterial undecaprenyl pyrophosphate synthase in the yeast rer2Δ mutant and CHO cells. Glycobiology 20: 1585-­‐1593. Sato, M., Fujisaki, S., Sato, K., Nishimura, Y., Nakano, A., 2001 Yeast Saccharomyces cerevisiae has two cis-­‐prenyltransferases with different properties and localizations. Implication for their distinct physiological roles in dolichol synthesis. Genes Cells 6: 495-­‐506. 2 SI P. Orlean Shridas, P., Waechter, C. J., 2006 Human dolichol kinase, a polytopic endoplasmic reticulum membrane protein with a cytoplasmically oriented CTP-­‐binding site. J. Biol. Chem. 281: 31696-­‐316704. P. Orlean 3 SI File S2 N-­‐glycosylation This Supporting File contains additional information related to Biosynthesis of Wall Components Along the Secretory Pathway, N-­‐glycosylation. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Assembly and transfer of the Dol-­‐PP-­‐linked precursor oligosaccharide: Steps on the cytoplasmic face of the ER membrane: Alg7. The Alg7 GlcNAc-­‐1-­‐P transferase, which carries out the first step in the assembly of the Dol-­‐PP-­‐linked precursor is highly conserved among eukaryotes and has homologues in Bacteria, for example MraY, which catalyzes transfers N-­‐
acetylmuramic acid-­‐pentapeptide from UDP to undecaprenol phosphate in peptidoglycan biosynthesis (Price and Momany, 2005). GlcNAc-­‐1-­‐P transferases such as Alg7 and MraY have multiple transmembrane domains and amino acid residues important for catalysis by members of this protein family lie in cytoplasmic loops (Dan and Lehrman; Price and Momany, 2005). Alg13/Alg14. These proteins function as a heterodimer to transfer the second, β1,4-­‐GlcNAc-­‐linked GlcNAc to Dol-­‐PP-­‐
GlcNAc (Bickel et al. 2005; Chantret et al. 2005; Gao et al. 2005). Soluble Alg13, assigned to GT Family 1, is the catalytic subunit and associates with membrane-­‐spanning Alg14 at the cytosolic face of the ER membranes (Averbeck et al. 2007; Gao et al. 2008). Alg13 and 14 are homologous to C and N-­‐terminal domains, respectively, of the bacterial MurG polypeptide, which adds N-­‐acetylmuramic acid to undecaprenol-­‐PP-­‐GlcNAc in peptidoglycan synthesis (Chantret et al. 2005). Alg1. This β1,4-­‐Man-­‐T, assigned to GT Family 33, transfers the first mannose from GDP-­‐Man to Dol-­‐PP-­‐GlcNAc2 (Couto et al. 1984). Alg2. This protein is a member of GT Family 4. Remarkably, Alg2 has both GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2Man α1,3-­‐Man-­‐T and GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2Man2 α1,6-­‐Man-­‐T activity and successively adds an α1,3-­‐Man and an α1,6 Man to the Dol-­‐PP-­‐
linked precursor (O'Reilly et al. 2006; Kämpf et al. 2009). Alg11. Alg11, also a member of GT Family 4, adds the next two α1,2-­‐linked mannoses (Cipollo et al. 2001; O'Reilly et al. 2006; Absmanner et al. 2010). alg11D mutants are viable though growth-­‐defective, and accumulate Dol-­‐PP-­‐GlcNAc2Man3, as well as some Dol-­‐PP-­‐GlcNAc2Man6-­‐7 (Cipollo et al. 2001; Helenius et al. 2002). The latter are aberrant glycan structures formed when Dol-­‐PP-­‐GlcNAc2Man3 is translocated to the lumen and acted on by lumenal Man-­‐T. 4 SI P. Orlean Heterologous expression and membrane topology of Alg1, Alg2, and Alg11. Alg1, Alg2, and Alg11 are catalytically active when expressed in E. coli (Couto et al. 1984; O'Reilly et al. 2006). The catalytic region of Alg1 is predicted to be cytoplasmic, and experimentally derived models for the membrane topology of Alg2 and Alg11 also place catalytic domains at the cytoplasmic side of the ER membrane (Kämpf et al. 2006; Absmanner et al. 2009), although not all predicted hydrophobic helices in Alg2 and Alg11 span the ER membrane, rather, they lie in its cytoplasmic face. Complex formation by early-­‐acting Alg proteins. There is evidence from analyses by coimmunoprecipitation and size exclusion chromatographic analyses for higher order organization of the proteins involved in the cytoplasmic steps of the yeast dolichol pathway. Alg7, 13, and 14 associate in a hexamer (Noffz et al. 2009). Alg1 forms separate complexes containing either Alg2 and Alg11, although the latter two do not interact with one another (Gao et al. 2004). Formation of these multienzyme complexes may in turn facilitate channeling of Dol-­‐PP-­‐linked intermediates to successive membrane-­‐associated transferases. Transmembrane translocation of Dol-­‐PP-­‐oligosaccharides: After Dol-­‐PP-­‐GlcNAc2Man5 is generated on the cytoplasmic face of the ER membrane, it is somehow translocated to the lumenal side of the membrane where subsequent sugars are transferred from Dol-­‐P-­‐sugars (Burda and Aebi, 1999; Helenius & Aebi, 2002). The presumed Dol-­‐PP-­‐oligosaccharide flippase likely prefers the heptasaccharide as substrate, but the presence of shorter oligosaccharides on proteins in both the alg2-­‐Ts and alg11Δ mutants (Jackson et al. 1989; Cippolo et al. 2001) indicates that truncated oligosaccharides can be translocated as well. The Rft1 protein is a candidate for the protein Dol-­‐PP-­‐GlcNAc2Man5 flippase (Helenius et al. 2002). Strains deficient in Rft1 accumulate Dol-­‐PP-­‐GlcNAc2Man5, but are unaffected in O-­‐mannosylation or in GPI anchor assembly, ruling out a deficiency in Dol-­‐P-­‐Man supply to the ER lumen. Because the few N-­‐glycans chains that were still transferred to the reporter protein carboxypeptidase Y in Rft1-­‐depleted cells were endoglycosidase H sensitive, the activity of Alg3, which adds the α1,3-­‐Man required for substrate recognition by endoglycosidase H, was unaffected. Moreover, high level expression of RFT1 partially suppresses the growth defect of alg11Δ and leads to increased levels of lumenal Dol-­‐PP-­‐GlcNAc2Man6-­‐7 and an increase in carboxypeptidase Y glycosylation, consistent with the notion of enhanced flipping of the suboptimal flippase substrate Dol-­‐PP-­‐
GlcNAc2Man3 (Helenius et al. 2002). However, although the above findings are consistent with Rft1 being the flippase itself, this role could not be demonstrated in biochemical assays for flippase activity, for sealed microsomal vesicles or proteoliposomes depleted of Rft1 retained flippase activity, and in fractionation experiments, flippase activity could be separated from Rft1 (Franck et al. 2008; Rush et al. 2009). P. Orlean 5 SI Lumenal steps in Dol-­‐PP-­‐oligosaccharide assembly: Alg3. This α1,3-­‐Man-­‐T is a member of GT Family 58, and transfers the precursor’s sixth, α1,3-­‐Man from Dol-­‐P-­‐Man, making the glycan sensitive to endoglycosidase H (Aebi et al. 1996; Sharma et al. 2001). Alg3’s Dol-­‐P-­‐Man:Dol-­‐PP-­‐GlcNAc2Man5 Man-­‐T activity can be selectively immunoprecipitated from detergent extracts of membranes (Sharma et al. 2001), providing strong evidence that Alg3 and its yeast homologues in the dolichol and GPI assembly pathways are indeed glycosyltransferases. Alg9 and Alg12. Alg9, a member of GT Family 22, transfers the seventh, α1,2-­‐linked Man to the α1,3-­‐Man added by Alg3 (Burda et al. 1999; Cipollo and Trimble, 2000). Alg12, also a GT22 Family member, next adds the eighth, α1,6-­‐Man to the α1,2-­‐linked Man just added by Alg9 (Burda et al. 1999), whereupon Alg9 acts again to add the ninth Man, in α1,2 linkage, to the α1,6-­‐Man added by Alg12 (Frank and Aebi 2005). The second activity of Alg9 was uncovered in in vitro assays in which alg9Δ and alg12Δ membranes were tested for their ability to elongate acceptor Dol-­‐PP-­‐GlcNAc2Man7 isolated from alg12Δ cells. These experiments established that Alg12 requires prior addition of the seventh Man by Alg9, even though Alg12 does not transfer its Man to that residue, and that the Alg12 reaction precedes Alg9’s second α1,2 mannosyltransfer (Frank and Aebi 2005). Alg6, Alg8, and Alg10. Alg6 and Alg8, members of GT Family 57, act successively to transfer two α1,3-­‐linked glucoses to extend the second α1,2-­‐Man added by Alg11, and lastly, Alg10, assigned to GT Family 59, completes the 14-­‐sugar Dol-­‐PP-­‐
linked oligosaccharide by adding a third, α1,2-­‐Glc (Reiss et al., 1996; Stagljar et al., 1994; Burda and Aebi, 1998). Shared transmembrane topology of Dol-­‐P-­‐sugar-­‐utilizing transferases. The six Dol-­‐P-­‐sugar-­‐utilizing transferases are members of a larger protein family that includes the Dol-­‐P-­‐Man-­‐utilizing Man-­‐T involved in GPI anchor biosynthesis (Oriol et al. 2002). The results of in silico analyses of the sequences of these proteins suggested they have a common membrane topology and 12 transmembrane segments, and a membrane organization recalling that of membrane transporters, which is consistent with the idea that each protein translocates its own Dol-­‐P-­‐linked sugar substrate (Burda and Aebi, 1999; Helenius and Aebi, 2002). It also plausible that these transferases operate in multienzyme complexes to facilitate substrate channeling. Oligosaccharide transfer to protein: Truncated oligosaccharides can be transferred to protein. The results of analyses of the N-­‐linked glycans present on protein in mutants defective in the assembly of the Dol-­‐PP-­‐linked precursor oligosaccharide indicate that a range of structures smaller than GlcNAc2Man9Glc3 can be transferred in vivo. However, full-­‐size Dol-­‐PP-­‐GlcNAc2Man9Glc3 is the preferred OST substrate in vitro, and the observation that mutants that make smaller precursor oligosaccharides have a synthetic phenotype 6 SI P. Orlean with OST mutants indicates the preference exists in vivo as well (Knauer and Lehle, 1999; Zufferey et al. 1995; Reiss et al. 1997; Karaoglu et al. 2001). This preference does not reflect differences between the binding affinities of Dol-­‐PP-­‐GlcNAc2Man9Glc3 and smaller oligosaccharides at the OST active site, rather, it has been proposed that OST has an allosteric site that binds GlcNAc2Man9Glc3 as well as smaller oligosaccharides, in turn activating the catalytic site for GlcNAc2Man9Glc3 and acceptor peptide binding. Binding of a truncated oligosaccharide at the allosteric site, however, enhances GlcNAc2Man9Glc3 binding more strongly, and so ensures preferential utilization of the full-­‐size precursor (Karaoglu et al., 2001; Kelleher and Gilmore, 2006). Purification and protein-­‐protein interactions of OST. Complete heterooctomeric OST complexes have been affinity purified (Karaoglu et al. 1997; Spirig et al. 1997; Karaoglu et al. 2001; Chavan et al. 2006), and the subunits appear to be present in stoichiometric amounts (Karaoglu et al. 1997). The OST complexes themselves may themselves function as dimers (Chavan et al. 2006). The results of genetic interaction studies and coimmunoprecipitation-­‐ and chemical cross-­‐linking experiments suggest the existence of three sub-­‐complexes i) Swp1-­‐Wbp1-­‐Ost2, ii) Stt3-­‐Ost4-­‐Ost3, and iii) Ost1-­‐Ost5 (Spirig et al. 1997; Karaoglu et al. 1997; Reiss et al. 1997; Li et al. 2003; Kim et al. 2003; reviewed by Knauer and Lehle, 1999; Kelleher and Gilmore, 2006). It has been noted, however, that treatment of OST with non-­‐ionic detergents does not yield these three subcomplexes (Kelleher and Gilmore, 2006). Furthermore, additional interactions between OST subunits have been detected using chemical cross-­‐linking approaches and membrane protein two-­‐hybrid analyses (Yan et al. 2003, 2005). OST also interacts with the Sec61 translocon complex and large ribosomal subunit (Chavan et al. 2005; Harada et al. 2009), suggesting that the complex is poised to act on nascent, freshly translocated proteins. However, protein O-­‐mannosyltransferases can compete for the hydroxyamino acids in a freshly translocated sequon (Ecker et al. 2003; see O-­‐mannosylation). Stt3 is the catalytic subunit of OST. There is strong evidence that Stt3, which has a soluble, lumenal domain towards its C-­‐terminus preceded by 11 transmembrane domains (Kim et al. 2005), is the catalytic subunit of OST. First, it can be crosslinked to peptides derivatized with a photoactivatable group and containing an N-­‐X-­‐T glycosylation site, or to nascent polypeptide chains containing the sequon-­‐mimicking, cryptic glycosylation site Q-­‐X-­‐T and a photoactivable side chain (Yan and Lennarz, 2002; Nilson et al. 2003). Second, Stt3 homologues are present in all eukarya, as well as in certain Bacteria and many Archaea, in which diverse types of glycan are transferred to protein (Kelleher and Gilmore, 2006; Kelleher et al. 2007). The Stt3 homologue from Campylobacter jejuni, PglB, was shown to be required for transfer of that bacterium’s characteristic glycan to Asn in a substrate peptide when the C. jejuni pgl gene cluster was heterologously expressed in E. coli (Wicker et al. 2002). Third, Stt3 homologues from the protist Leishmania major, whose proteome contains no other OST subunits, complement the S. P. Orlean 7 SI cerevisiae stt3Δ mutants as well as null mutations in the genes for the essential OST subunits Ost1, Ost2, Swp1, and Wbp1, indicating that the protist Stt3 functions autonomously as an OST (Nasab et al. 2008; Hese et al. 2009). Stt3 has been assigned to GT Family 66. Ost3 and Ost6: role of a thioredoxin domain. The other OST subunits for which catalytic activity has been demonstrated are the paralogues Ost3 and Ost6. ost3Δ ost6Δ double mutants have a more severe glycosylation defect than the single nulls (Knauer and Lehle, 1999b). The two proteins confer a degree of acceptor preference to the OST complexes that contain them (Schulz and Aebi, 2009) because they each have peptide binding grooves lined by amino acids whose side chains are complementary in hydrophobicity and charge to different substrate peptides (Jamaluddin et al. 2011). Ost3 and Ost6 are predicted to have four transmembrane domains at their C-­‐termini and an N-­‐terminal domain containing a thioredoxin fold with the CXXC motif common to proteins involved in disulfide bond shuffling during oxidative protein folding (Kelleher and Gilmore, 2006; Schulz et al. 2009). This domain most likely lies in the lumen (Kelleher and Gilmore, 2006). Mutations of the cysteines in the CXXC motifs of Ost3 and Ost6 lead to site-­‐specific underglycosylation, indicating the importance of the thioreductase motif. This was confirmed by the demonstration that the thioredoxin domain of Ost6, expressed in E. coli, had oxidoreductase activity towards a peptide substrate (Schulz et al. 2009). These findings led to a model in which Ost3/Ost6 form transient disulfide bonds with nascent proteins and promote efficient glycosylation of more Asn-­‐X-­‐Ser/Thr sites by delaying oxidative protein folding (Schulz et al. 2009). Structural analyses of the thioredoxin domain of Ost6 showed that the peptide binding groove is present only when the CXXC motif is oxidized (Jamaluddin et al. 2011). Recruitment of Ost3 or Ost6 to OST requires Ost4, a hydrophobic 36 amino protein (Kim et al. 2000, 2003; Spirig et al. 2005). Ost4 also interacts with Stt3 (Karaoglu et al. 1997; Spirig et al. 1997; Knauer and Lehle, 1999; Kim et al. 2003). ost4Δ strains are temperature-­‐sensitive and severely underglycosylate protein (Chi et al. 1996). Possible roles for other OST subunits. A sub-­‐complex of Swp1p, Wbp1p, and Ost2p, has been suggested to confer the preference for GlcNAc2Man9Glc3, possibly by providing the allosteric site (Kelleher and Gilmore, 2006). Evidence for a role of complex subunits other than Stt3 was obtained with Trypanosoma cruzi Stt3, which transfers GlcNAc2Man7-­‐9 to protein in vitro as efficiently as it does glucosylated oligosaccharides. When expressed in S. cerevisiae in place of native Stt3, trypanosomal Stt3 now preferentially transferred GlcNAc2Man9Glc3 to protein in vitro and in vivo (Castro et al. 2006). Similarly, when Leishmania Stt3 is expressed in the context of the other S. cerevisiae OST subunits, the Leishmania protein acquires a preference for transferring glucosylated oligosaccharides, rather than the non-­‐glucosylated oligosaccharides that it transfers in the protist itself (Hese et al. 2009). Wbp1 may be involved in recognition of Dol-­‐PP-­‐GlcNAc2Man9Glc3, because alkylation of a key cysteine 8 SI P. Orlean residue in this subunit inactivates OST, whereas inactivation is prevented by prior incubation with Dol-­‐PP-­‐GlcNAc2 (Pathak et al. 1995). The protein’s single transmembrane domain contains sequences important for incorporation into the OST complex, possibly by making interactions with Ost2 and Swp1 (Li et al. 2003). Other than their membership in proposed OST subcomplexes and interactions with other OST subunits, little is known about the function of Swp1, Ost1, Ost2, and Ost5, although it has been suggested that Ost1 has a role in funneling nascent polypeptides to Stt3 (Lennarz, 2007). Regulation of OST by the CWI pathway. Oligosaccharyltransferase may be regulated by the PKC-­‐dependent CWI pathway or by Pkc1 itself, a notion that arose from the identification of STT3 in a screen for mutants sensitive to the PKC inhibitor staurosporine and to elevated temperature (Yoshida et al. 1995). Although this suggested that adequate levels of N-­‐
glycosylation are needed for cells to overcome defects in CWI signaling, staurosporine sensitivity proved not to be a general consequence of deficient N-­‐glycosylation, because only a subset of stt3 alleles were sensitive to the drug, and mutants in most other OST subunits, with the exception of Ost4, were resistant (Chavan et al. 2003; Levin, 2005). A more direct link between Stt3 and the Pkc1-­‐dependent signaling emerged from the findings that STT3 mutations that lead to staurosporine sensitivity are located in N-­‐terminal, predicted cytosolic domains of Stt3, and that pkc1Δ mutants have half of wild type OST activity in vitro (Chavan et al. 2003; Park and Lennarz, 2000). This led to the suggestion that CWI pathway regulates OST via an interaction between Pkc1 or components of the PKC pathway with the N-­‐terminal domain of Stt3, and perhaps Stt3-­‐interacting Ost4 as well (Chavan et al. 2003). N-­‐glycan processing in the ER and glycoprotein quality control: Glucosidase II. This is a heterodimer of catalytic Gls2/Rot2 and Gtb1, the latter of which is necessary for, and influences the rate of, Glc trimming (Trombetta et al. 1996; Wilkinson et al., 2006; Quinn et al. 2009). Glycoprotein recognition by Pdi1 and the Pdi1-­‐Htm1 complex. Unfolded or misfolded proteins are bound by protein disulfide isomerase Pdi1, a subset of which is in complex with Mns1 homolog Htm1. A stochastic model has been proposed in which both Pdi1 and the Pdi1-­‐Htm1 complex recognize un-­‐ or misfolded proteins, but persistently misfolded proteins stand an increased chance of encountering Pdi1-­‐Htm1 whose Htm1 component trims a Man from N-­‐linked glycans, yielding a GlcNAc2Man7 structure bearing a terminal α1,6 Man (Clerc et al. 2009; Gauss et al. 2011).
Mannan elaboration in the Golgi: Formation of core type N-­‐glycan and mannan outer chains: P. Orlean 9 SI Elucidation of the pathway for formation of mannan outer chains. Two groups of proteins, the Mnn9/Anp1/Van1 trio, and the Mnn10 and Mnn11 pair, had been implicated in formation of the poly-­‐α1,6-­‐linked mannan backbone, but because strains deficient in these proteins retained mannosyltransferase activity and still made mannan containing α1,6 linkages, these proteins were considered more likely to affect mannan formation indirectly (reviewed by Orlean, 1997; Dean, 1999). Two key sets of findings led to clarification of mannan biosynthesis. First, co-­‐immunoprecipitation and colocalization experiments established that Mnn9, Anp1, and Van1 occurred in two different protein complexes in the cis-­‐Golgi, one containing Mnn9 and Van1 (subsequently named M-­‐Pol I), the other, Mnn9, Anp1, Hoc1 (homologous to Och1), and the related Mnn10 and Mnn11 proteins (M-­‐Pol II) (Hashimoto and Yoda, 1997; Jungmann and Munro, 1998; Jungmann et al. 1999). Second, both immunoprecipitated protein complexes had α1,6 mannosyltransferase activity, indicating that one or more of the Mnn9/Anp1/ Van1 group was an α1,6 mannosyltransferase (Jungmann and Munro, 1998; Jungmann et al. 1999). Consistent with their being glycosyltransferases, all five proteins have the GT-­‐A fold protein topology and a “DXD motif” common to enzymes that have sugar nucleotides as donors and use the aspartyl carboxylates to coordinate divalent cations and the ribose of the donor (Wiggins and Munro, 1998; Lairson et al. 2008). The contributions of the individual subunits to α1,6 mannan synthesis by each complex, and the roles of the two complexes in mannan formation, were explored in deletion mutants and in point mutants abolishing catalytic activity but otherwise preserving complex stability. The sizes of the mannans and the residual in vitro activities of the M-­‐Pol complexes in these mutants led to the current model for mannan synthesis (Jungmann et al. 1999; Munro, 2001; Figure 3 in main text). In it, M-­‐Pol I, a heterodimer, acts first to extend the Och1-­‐derived Man with further α1,6-­‐linked mannoses. Analyses of mutants in the DXD motifs of Mnn9 and Van1 indicated that Mnn9 likely adds the first α1,6-­‐liked Man, which is extended with 10-­‐15 α1,6 mannoses in Van1-­‐requiring reactions (Stolz and Munro, 2002; Rodionov et al. 2009). This α1,6 backbone is then elongated with 40-­‐60 α1,6 Man by M-­‐Pol II. Assays of M-­‐Pol ll from strains lacking Mnn10 or Mnn11 indicated that these proteins are responsible for the majority of the α1,6 mannosyltransferase activity in that complex (Jungmann et al., 1999). The contribution of Hoc1, a homologue of the Och1 α1,6-­‐Man-­‐T is not clear, for HOC1 deletion neither alters M-­‐Pol II activity nor impacts mannan size. Localization of Och1 and Man-­‐Pol complexes. The localization dynamics of Mnn9-­‐containing M-­‐Pol complexes and Och1 seem inconsistent with the order in which they act in mannan assembly, with Mnn9 showing a steady state localization in the cis-­‐Golgi and continuously cycling between that compartment and the ER, but with Och1 cycling between the ER and cis-­‐ and trans-­‐Golgi (Harris and Waters, 1996; Todorow et al. 2000; Karhinen and Makarow, 2004). It has been suggested that 10 SI P. Orlean substrate specificity, rather than transferase localization, determines their order in which the enzymes act (Okamoto et al. 2008). The size of N-­‐linked mannan can be impacted by deficiencies in proteins required for localization of Golgi mannosyltransferases. For example, deletion of VPS74, also identified as MNN3, eliminates a protein that interacts with the cytoplasmic tails of certain transferases normally resident in the cis and medial Golgi compartments. The resulting mislocalization of several mannosyltransferases would explain the underglycosylation phenotype of mnn3 mutants (Schmitz et al. 2008; Corbacho et al. 2010). Mutations in SEC20, which encodes a protein involved in Golgi to ER retrograde transport, also result in diminished Golgi mannosyltransferase activity, even though this glycosylation defect is not correlated with the secretory pathway defect (Schleip et al. 2001). The reason for this is not clear. Mannan side branching and mannose phosphate addition: Roles of the Ktr1 Man-­‐T family members in mannan side branching. Five members of the Ktr1 family of Type II membrane proteins, Kre2/Mnt1, Yur1, Ktr1, Ktr2, Ktr3, also contribute to N-­‐linked outer chain synthesis, as judged by the impact of null mutations on the mobility of reporter proteins (Lussier et al. 1996; 1997a; 1999). Of these proteins, Kre2/Mnt1, Ktr1, Ktr2, and Yur1 have been shown to have α1,2 Man-­‐T activity. These Ktr1 family members, perhaps along with uncharacterized homologues Ktr4, Ktr5, and Ktr7 (Lussier et al. 1999) have a collective role in adding the second, and perhaps subsequent α1,2-­‐mannoses to mannan side branches. Members of the Ktr1 family have been assigned to GT Family 15. Addition and function of mannose phosphate. Both core type N-­‐glycans and mannan can be modified with mannose phosphate on α1,2-­‐linked mannoses in the context of an oligosaccharide containing at least one α1,2-­‐linked mannobiose structure. Mannose phosphates confer a negative charge, an attribute exploited early on to isolate mannan synthesis mutants on the basis of their inability to bind the cationic dye Alcian Blue (Ballou, 1982; 1990). Mnn6/Ktr6, a member of the Ktr1 family, is the major activity responsible for transferring Man-­‐1-­‐P from GDP-­‐Man to both mannan outer chains and, in vitro, to core N-­‐
glycans, generating GMP. However, because deletion of MNN6 did not eliminate in vivo mannose phosphorylation in och1Δ strains that make only core type N-­‐glycans, additional, as yet unidentified, core phosphorylating proteins must exist (Wang et al. 1997; Jigami and Odani, 1999). The Mnn4 protein is also involved in Man-­‐P addition, but its role differs from Mnn6’s in that deletion of Mnn4 reduces Man-­‐P on core-­‐type glycans (Odani et al. 1996). Mnn4 does not resemble glycosyltransferases, but does have a LicD domain found in nucleotidyltransferases and phosphotransferases involved in lipopolysaccharide synthesis. The mnn4Δ mutation is dominant, and Mnn4 has been proposed to have a positive regulatory role (Jigami and Odani, 1999). Levels of mannan phosphorylation are highest in the late log and stationary phases, when MNN4 expression is elevated (Odani et al. 1997). Transcriptional regulation may involve the RSC chromatin remodeling complex because strains lacking Rcs14, a P. Orlean 11 SI subunit of that complex, show drastically reduced Alcian Blue binding and down-­‐regulated expression of MNN4 and MNN6 (Conde et al. 2007).
A Golgi GlcNAc-­‐T. S. cerevisiae also has the capacity to add GlcNAc to the non-­‐reducing end of N-­‐linked glycans. Heterologously expressed lysozyme received a GlcNAc2Man8-­‐12 glycan additionally bearing a GlcNAc residue, and the responsible GlcNAc transferase proved to be Gnt1, whose localization mostly coincides with that of Mnn1 in the medial Golgi (Yoko-­‐o et al. 2003). GNT1 disruptants have no discernible phenotype, and Gnt1 may rarely act on native yeast glycans; its activity would require that UDP-­‐GlcNAc be transported into the Golgi lumen (Yoko-­‐o et al. 2003). Literature Cited Averbeck, N., Keppler-­‐Ross, S., Dean, N., 2007 Membrane topology of the Alg14 endoplasmic reticulum UDP-­‐GlcNAc transferase subunit. J. Biol. Chem. 282: 29081-­‐29088. Castro, O., Movsichoff, F., Parodi, A. J., 2006 Preferential transfer of the complete glycan is determined by the oligosaccharyltransferase complex and not by the catalytic subunit. Proc. Natl. Acad. Sci. USA. 103: 14756-­‐14760. Chavan, M., Yan, A., Lennarz, W. J. 2005 Subunits of the translocon interact with components of the oligosaccharyl transferase complex. J. Biol. Chem. 280: 22917–22924. Chi, J. H., Roos, J., Dean, N., 1996 The OST4 gene of Saccharomyces cerevisiae encodes an unusually small protein required for normal levels of oligosaccharyltransferase activity. J. Biol. Chem. 271: 3132–3140. Conde, R., Cueva, R., Larriba, G., 2007 Rsc14-­‐controlled expression of MNN6, MNN4 and MNN1 regulates mannosylphosphorylation of Saccharomyces cerevisiae cell wall mannoproteins. FEMS Yeast Res. 7: 1248-­‐1255. Corbacho, I., Olivero, I., Hernández, M., 2010 Identification of the MNN3 gene of Saccharomyces cerevisiae. Glycobiology 20: 1336-­‐1340. 12 SI P. Orlean Dan, N., Lehrman, M. A., 1997 Oligomerization of hamster UDP-­‐GlcNAc:dolichol-­‐P GlcNAc-­‐1-­‐P transferase, an enzyme with multiple transmembrane spans. J. Biol. Chem. 272: 14214-­‐14219. Dean, N. 1999 Asparagine-­‐linked glycosylation in the yeast Golgi. Biochim. Biophys. Acta 1426: 309–322. Gao, X. D., Moriyama, S., Miura, N., Dean, N., Nishimura, S., 2008 Interaction between the C termini of Alg13 and Alg14 mediates formation of the active UDP-­‐N-­‐acetylglucosamine transferase complex. J. Biol. Chem. 283: 32534-­‐32541. Harada, Y., Li, H., Li, H., Lennarz, W. J., 2009 Oligosaccharyltransferase directly binds to ribosome at a location near the translocon-­‐binding site. Proc. Natl. Acad. Sci. USA 106: 6945-­‐6949. Harris, S. L., Waters, M. G., 1996 Localization of a yeast early Golgi mannosyltransferase, Och1p, involves retrograde transport. J. Cell Biol. 132: 985-­‐998. Jackson, B. J., Warren, C. D., Bugge, B., Robbins, P. W., 1989 Synthesis of lipid-­‐linked oligosaccharides in Saccharomyces cerevisiae: Man2GlcNAc2 and Man1GlcNAc2 are transferred from dolichol to protein in vivo. Arch. Biochem. Biophys. 272: 203-­‐
209. Jamaluddin, M. F., Bailey, U. M., Tan, N. Y., Stark, A. P., Schulz, B. L., 2011 Polypeptide binding specificities of Saccharomyces cerevisiae oligosaccharyltransferase accessory proteins Ost3p and Ost6p. Protein Sci. 20: 849-­‐555. Karaoglu, D., Kelleher, D. J., Gilmore, R., 2001 Allosteric regulation provides a molecular mechanism for preferential utilization of the fully assembled dolichol-­‐linked oligosaccharide by the yeast oligosaccharyltransferase. Biochemistry: 40: 12193–12206. Karhinen, L., Makarow, M., 2004 Activity of recycling Golgi mannosyltransferases in the yeast endoplasmic reticulum. J. Cell Sci. 117: 351-­‐358. P. Orlean 13 SI Kim, H., von Heijne, G., Nilsson, I., 2005 Membrane topology of the STT3 subunit of the oligosaccharyl transferase complex. J. Biol. Chem. 280: 20261-­‐20267. Lairson, L. L., Henrissat, B., Davies, G. J., Withers, S. G., 2008 Glycosyltransferases: structures, functions, and mechanisms. Annu. Rev. Biochem. 77: 521-­‐555. Munro, S., 2001 What can yeast tell us about N-­‐linked glycosylation in the Golgi apparatus? FEBS Lett. 498: 223-­‐227. Okamoto, M., Yoko-­‐o, T., Miyakawa T., Jigami, Y., 2008 The cytoplasmic region of α-­‐1,6-­‐mannosyltransferase Mnn9p is crucial for retrograde transport from the Golgi apparatus to the endoplasmic reticulum in Saccharomyces cerevisiae. Eukaryot. Cell 7: 310-­‐318. Price, N. P., Momany, F. A., 2005. Modeling bacterial UDP-­‐HexNAc: polyprenol-­‐P HexNAc-­‐1-­‐P transferases. Glycobiology 15: 29R-­‐42R. Schleip, I., Heiss, E., Lehle, L., 2001 The yeast SEC20 gene is required for N-­‐ and O-­‐glycosylation in the Golgi. Evidence that impaired glycosylation does not correlate with the secretory defect. J. Biol. Chem. 276: 28751-­‐28758. Schmitz, K. R., Liu, J. X., Li, S. L., Setty T. G., Wood, C. S., et al., 2008 Golgi localization of glycosyltransferases requires a Vps74p oligomer. Dev. Cell 14: 523-­‐534. Todorow, Z., Spang, A., Carmack, E., Yates, J., Schekman, R., 2000 Active recycling of yeast Golgi mannosyltransferase complexes through the endoplasmic reticulum. Proc. Natl. Acad. Sci. USA. 97: 13643-­‐13548. Wiggins, C. A., Munro, S., 1998 Activity of the yeast MNN1 α-­‐1,3-­‐mannosyltransferase requires a motif conserved in many other families of glycosyltransferases. Proc. Natl. Acad. Sci. USA. 95: 7945-­‐7950. 14 SI P. Orlean Yan, A., Ahmed, E., Yan, Q., Lennarz, W. J., 2003 New findings on interactions among the yeast oligosaccharyl transferase subunits using a chemical cross-­‐linker. J. Biol. Chem. 278: 33078–33087. Yan, A., Wu. E., Lennarz, W. J., 2005 Studies of yeast oligosaccharyl transferase subunits using the split-­‐ubiquitin system: topological features and in vivo interactions. Proc. Natl. Acad. Sci. USA 102: 7121–7126. Yoko-­‐o, T., Wiggins, C. A., Stolz, J., Peak-­‐Chew, S. Y., Munro, S., 2003 An N-­‐acetylglucosaminyltransferase of the Golgi apparatus of the yeast Saccharomyces cerevisiae that can modify N-­‐linked glycans. Glycobiology 13: 581-­‐589. Yoshida, S., Ohya, Y., Nakano, A., Anraku, Y., 1995. STT3, a novel essential gene related to the PKC1/STT1 protein kinase pathway, is involved in protein glycosylation in yeast. Gene 164: 167-­‐172. Zufferey, R., Knauer, R., Burda, P., Stagljar, I., te Heesen, S., et al., 1995 STT3, a highly conserved protein required for yeast oligosaccharyl transferase activity in vivo. EMBO J. 14: 4949-­‐4960. P. Orlean 15 SI File S3 O-­‐Mannosylation This Supporting File contains additional information related to Biosynthesis of Wall Components Along the Secretory Pathway, O-­‐mannosylation. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Protein O-­‐mannosyltransferases in the ER: Substrate proteins for different Pmt complexes. Analyses of glycosylation of individual proteins in pmtΔ strains showed that Pmt1/Pmt2 complexes are primarily involved in O-­‐mannosylation of Aga2, Bar1, Cts1, Kre9, and Pir2, whereas homodimeric Pmt4 modifies Axl2, Fus1, Gas1, Kex2 (Gentzsch and Tanner 1997; Ecker et al. 2003; Proszynski et al. 2004; Sanders et al. 1999). However, some proteins, including Mid2, the WSC proteins, and Ccw5, are modified by both complexes, although the Pmt1/Pmt2 and Pmt4/Pmt4 dimers modify different domains of these target proteins (Ecker et al. 2003; Lommel et al. 2004). Mutations in substrate proteins can cause them to be O-­‐mannosylated by a different PMT, and PMTs can also have a role in quality control of protein folding in the ER (see N-­‐glycan processing in the ER and glycoprotein quality control). Thus, wild type Gas1 is normally O-­‐mannosylated by Pmt4, whereas Gas1
G291R
, a model misfolded protein, is hypermannosylated by Pmt1-­‐
Pmt2 as well as targeted to the HRD-­‐ubiquitin ligase complex for degradation by the ERAD system (Hirayama et al. 2008; Goder and Melero, 2011). The latter, chaperone-­‐like function of Pmt1-­‐Pmt2 may be distinct from Pmt1-­‐Pmt2’s O-­‐mannosyltransferase activity (Goder and Melero, 2011). Extension and phosphorylation of O-­‐linked manno-­‐oligosaccharide chains: Extension with α-­‐linked mannoses. The Ser-­‐ or Thr-­‐linked Man is extended with up to four α-­‐linked Man that are added by GDP-­‐Man-­‐dependent Man-­‐T of the Ktr1 and Mnn1 families (Lussier et al. 1999; Figure 4 in main text). The contributions of these proteins was deduced from the sizes of the O-­‐linked chains that accumulated in strains in which Man-­‐T genes had been deleted singly or in different combinations. Transfer of the first two α1,2-­‐Man is carried out by Ktr1 sub-­‐family members Ktr1, Ktr3, and Kre2, which have overlapping roles in the process, although Kre2 has the dominant role in addition of the second, α1,2-­‐Man (Lussier et al. 1997a). The major O-­‐linked glycan made in the ktr1Δ ktr3Δ kre2Δ triple mutant consists of a single Man (Lussier et al. 1997a). Ktr1, Ktr3, and Kre2 are also involved in making α1,2-­‐branches to mannan outer chains (see Mannan elaboration in the Golgi). 16 SI P. Orlean Extension of the trisaccharide chain with one or two α1,3-­‐linked Man is the shared responsibility of Mnn1 family members Mnn1, Mnt2, and Mnt3, with Mnn1 having the major role in adding the fourth Man but Mnt2 and Mnt3 dominating when the fifth is added (Romero et al. 1999). Mnn1 also transfers Man to N-­‐linked outer chains. The α1,2 Man-­‐T have been localized to the medial Golgi, and the Mnn1 α1,3 Man-­‐T to the medial and trans-­‐Golgi (Graham et al. 1994). Because protein-­‐
bound O-­‐mannosyl glycans pulse-­‐labeled in mutants defective in ER to Golgi transport such as sec12, sec18, and sec20 contain two, sometimes more mannoses, GDP-­‐Man-­‐dependent O-­‐glycan extension can occur at the level of the ER (Haselbeck and Tanner, 1983; Zueco et al. 1986; D'Alessio et al. 2005). The process is independent of nucleotide sugar diphosphatases (see Sugar nucleotide transport; D'Alessio et al. 2005), but presumably mediated in the ER by Man-­‐T en route to the Golgi. Importance and function of O-­‐mannosyl glycans: Importance of O-­‐mannosylation for function of specific proteins. Analyses of single and conditionally lethal double pmt mutants show that O-­‐mannosylation can be important for function of individual O-­‐mannosylated proteins. For example, pmt4Δ haploids show a unipolar, rather than the normal axial budding pattern, which is due to defective O-­‐mannosylation and resulting instability and mislocalization of Axl2, which normally marks the axial budding site (Sanders et al. 1999). Pmt4-­‐initiated O-­‐mannosylation is also necessary for cell surface delivery of Fus1, because the unglycosylated protein accumulates in the late Golgi (Proszynski et al. 2004). Defects in Pmt4-­‐dependent O-­‐glycosylation of Msb2 (as well as N-­‐glycosyation) of osmosensor Msb2 lead to activation of the filamentous growth signaling pathway (Yang et al. 2009). In this case, underglycosylation may unmask a domain that normally is exposed and makes interactions when the signaling pathway is activated legitimately. O-­‐
mannosylation of Wsc1, Wsc2, and Mid2 is necessary for these Type I membrane proteins to fulfill their functions as sensors that activate the CWI pathway. Underglycosylation of the CWI pathway-­‐triggering mechanosensor Wsc1 in a pmt4Δ mutant eliminates the stiffness of this rod-­‐like glycoprotein and abolishes its “nanospring” properties, impairing Wsc1’s function as a mechanosensor (Dupres et al. 2009). Further, in pmt2Δ pmt4Δ mutants, which, like CWI pathway mutants, require osmotic stabilization, deficient O-­‐mannosylation results in incorrect proteolytic processing and instability of the sensors (Philip and Levin, 2001; Lommel et al. 2004). Literature Cited D'Alessio, C., Caramelo, J. J., Parodi, A. J., 2005 Absence of nucleoside diphosphatase activities in the yeast secretory pathway does not abolish nucleotide sugar-­‐dependent protein glycosylation. J. Biol. Chem. 280: 40417-­‐40427. P. Orlean 17 SI Dupres, V., Alsteens, D., Wilk, S., Hansen, B., Heinisch, J. J., Dufrêne, Y. F. 2009 The yeast Wsc1 cell surface sensor behaves like a nanospring in vivo. Nat. Chem. Biol. 5: 857-­‐862. Gentzsch, M., Tanner, W., 1997 Protein-­‐O-­‐glycosylation in yeast: protein-­‐specific mannosyltransferases. Glycobiology 7: 481-­‐
486. Goder, V., Melero, A., 2011 Protein O-­‐mannosyltransferases participate in ER protein quality control. J. Cell Sci. 124: 144-­‐153. Graham, T. R., Seeger, M., Payne, G. S., MacKay, V. L., Emr, S. D., 1994 Clathrin-­‐dependent localization of α1,3 mannosyltransferase to the Golgi complex of Saccharomyces cerevisiae. J. Cell Biol. 127: 667-­‐678. Haselbeck, A., Tanner, W., 1983 O-­‐glycosylation in Saccharomyces cerevisiae is initiated at the endoplasmic reticulum. FEBS Lett. 158: 335-­‐338. Hirayama, H., Fujita, M., Yoko-­‐o, T., Jigami, Y., 2008 O-­‐mannosylation is required for degradation of the endoplasmic reticulum-­‐
associated degradation substrate Gas1*p via the ubiquitin/proteasome pathway in Saccharomyces cerevisiae. J. Biochem. 143: 555-­‐567. Philip, B., 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. Proszynski, T. J., Simons, K., Bagnat, M., 2004 O-­‐Glycosylation as a sorting determinant for cell surface delivery in yeast. Mol. Biol. Cell 15: 1533-­‐1543. 18 SI P. Orlean Sanders, S. L., Gentzsch, M., Tanner, W., Herskowitz, I., 1999 O-­‐glycosylation of Axl2/Bud10p by Pmt4p is required for its stability, localization, and function in daughter cells. J. Cell Biol. 145: 1177-­‐1188. Yang, H. Y., Tatebayashi, K., Yamamoto, K., Saito, H., 2009 Glycosylation defects activate filamentous growth Kss1 MAPK and inhibit osmoregulatory Hog1 MAPK. EMBO J. 28: 1380-­‐1389. Zueco, J., Mormeneo, S., Sentandreu, R., 1986 Temporal aspects of the O-­‐glycosylation of Saccharomyces cerevisiae mannoproteins. Biochim. Biophys. Acta 884: 93-­‐100. P. Orlean 19 SI File S4 GPI anchoring This Supporting File contains additional information related to Biosynthesis of Wall Components Along the Secretory Pathway, GPI anchoring. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Assembly of the GPI precursor and its attachment to protein in the ER: Steps on the cytoplasmic face of ER membrane: Gpi3. Gpi3 is a member of GT Family 4 and has an EX7E motif conserved in a range of glycosyltransferases (Coutinho et al. 2003). Mutational analyses indicate that the glutamates are be important for function of Gpi3 and certain EX7E motif glycosyltransferases, although the comparative importance of the two glutamates varies between different transferases (Kostova et al. 2003). However, in the case of Alg2, the EX7E motif is not important for protein function (Kämpf et al. 2009). Formation of GlcNAc-­‐PI by GPI-­‐GnT. The acyl chains of the PI species that receive are the same length as those in other membrane phospholipids (Sipos et al. 1997). Evidence that GlcNAc transfer occurs at the cytoplasmic face of the ER membrane is that i) the catalytic domain of Gpi3’s human orthologue faces the cytoplasm (Watanabe et al. 1996; Tiede et al. 2000), and ii) GlcNAc-­‐PI can be labeled with membrane topological probes on the cytoplasmic side of the mammalian ER membrane (Vidugiriene and Menon, 1993). Significance of Ras2 regulation of GPI-­‐GnT. A clue to the significance of Ras2 regulation of GPI-­‐GnT came from the observation that conditional mutants in GPI-­‐GnT subunits show the phenotype of hyperactive Ras mutants, filamentous growth and invasion of agar. This led to the suggestion that Ras2-­‐mediated modulation of GPI synthesis may be involved in the cell wall and morphogenetic changes that occur in the dimorphic transition to filamentous growth (Sobering et al. 2003; 2004). Location of GlcNAc-­‐PI de-­‐N-­‐acetylation. The de-­‐acetylase reaction likely occurs at the cytoplasmic face of the ER membrane, because the bulk of Gpi12’s mammalian orthologue is cytoplasmic, and because newly synthesized GlcN-­‐PI is accessible on the cytoplasmic face of intact ER vesicles (Vidugiriene and Menon, 1993). Transmembrane translocation of GlcN-­‐PI. GlcN-­‐PI is the precursor species most likely to be translocated to the lumenal side of the ER membrane. Flipping of GlcN-­‐PI as well as GlcNAc-­‐PI has been reconstituted in rat liver microsomes, but the protein involved has not been identified, and the possibility has been raised that GlcN-­‐PI translocation may be mediated by a generic ER phospholipid flippase (Vishwakarma and Menon, 2006). 20 SI P. Orlean Lumenal steps in GPI assembly: Inositol acylation. The acyl chain transferred to GlcN-­‐(acyl)PI in vivo is likely palmitate, although a range of different acyl chains can be transferred from their corresponding CoA derivatives in vitro (Costello and Orlean, 1992; Franzot and Doering, 1999). Because mutants blocked in formation of all mannosylated GPIs accumulated inositol-­‐acylated GlcN-­‐PI (Orlean, 1990; Costello and Orlean, 1992), and because mannosylated GPI intermediates lacking an inositol acyl chain have not been reported, it is likely that inositol acylation precedes mannosylation in vivo. Gwt1, the acyltransferase, is likely to be catalytic because its affinity-­‐purified mammalian orthologue transfers palmitate from palmitoyl CoA to a dioctanoyl analogue of GlcN-­‐PI (Murakami et al. 2003). The protein has 13 transmembrane domains (Murakami et al. 2003; Sagane et al. 2011), and amino acid residues critical for function all face the lumen, indicating acyl transfer is a lumenal event (Sagane et al. 2011), although it is not yet known how acyl CoAs enter the ER lumen. Despite Gwt1’s multispanning topology, the possibility that this inositol acyltransferase is also a GlcN-­‐PI transporter is unlikely, because non-­‐acylated, mannosylated GPIs can be formed in cell lines deficient in Gwt1’s mammalian orthologue (Murakami et al. 2003). GPI Man-­‐T-­‐I. The α1,4-­‐Man-­‐T Gpi14 shows greatest similarity to Alg3, is predicted to have 12 transmembrane segments (Oriol et al. 2002), and is assigned to GT Family 50. Two additional proteins, Arv1 and Pbn1, are involved in the GPI-­‐
Man-­‐T-­‐I step along with Gpi14. arv1Δ cells grow at 30°C but not at 37°C, and are delayed in ER to Golgi transport of GPI-­‐
anchored proteins, and accumulate GlcN-­‐(acyl)PI in vitro (though not in vivo) (Kajiwara et al. 2008). Further, their temperature sensitivity is suppressed by overexpression the genes for most of the subunits of GPI-­‐GnT, suggesting a functional link between ARV1 and GPI assembly (Kajiwara et al. 2008). However, arv1Δ cells were not defective in Dol-­‐P-­‐Man synthase activity or in N-­‐
glycosylation, nor were mild detergent-­‐treated arv1Δ membranes defective in GPI-­‐Man-­‐T-­‐I activity, suggesting that Arv1 is not a Dol-­‐P-­‐Man flippase or directly involved in mannosyltransfer, and leading to the proposal that Arv1 is involved in delivering GlcN-­‐(acyl)PI to GPI-­‐Man-­‐T-­‐I (Kajiwara et al. 2008). Essential Pbn1 has been implicated at the GPI-­‐Man-­‐T-­‐I step in yeast because expression of both GPI14 and PBN1 is necessary to complement mammalian cell lines defective in Pbn1’s mammalian homologue Pig-­‐X, and likewise, co-­‐expression of PIG-­‐X and the gene for Gpi14’s mammalian homologue, PIG-­‐M, partially rescues the lethality of gpi14Δ (Ashida et al. 2005; Kim et al. 2007). Repression of PBN1 expression leads to accumulation of some of the ER form of the GPI protein Gas1, a phenotype seen in GPI precursor assembly mutants (Subramanian et al. 2006). However, it has not been reported whether pbn1 mutants accumulate the predicted GPI intermediate GlcN-­‐(acyl)PI. Because Pbn1 is also involved in processing a number of non-­‐GPI proteins that pass though the ER to the vacuole, the vacuolar membrane, and the plasma membrane, it must have additional functions in the ER (Subramanian et al. 2006). P. Orlean 21 SI GPI Man-­‐T-­‐II. Unlike the other Dol-­‐P-­‐Man-­‐utilizing transferases of the GPI assembly and dolichol pathways, the α1,6-­‐
Man-­‐T Gpi18 is predicted to have 8 transmembrane domains (Fabre et al. 2005; Kang et al. 2005). This protein and its orthologues have been assigned to GT Family 76. GPI Man-­‐T-­‐III and IV. These two α1,2-­‐Man-­‐T, together with their homologues in the dolichol pathway, Alg9 and Alg12, are predicted to have 12 transmembrane domains and are assigned to GT Family 22 (Oriol et al. 2002). Overexpression of GPI10 does not rescue the lethal smp3Δ null mutation, and vice versa, indicating that the two α1,2-­‐Man-­‐T have very strict acceptor specificities (Grimme et al. 2001). Phosphoethanolamine addition: origin of Etn-­‐P from Ptd-­‐Etn. There is good evidence that the Etn-­‐Ps, at least those on Man-­‐1 and Man3, originate from Ptd-­‐Etn. Yeast mutants unable to make CDP-­‐Etn or CDP-­‐Cho from exogenously supplied Etn, 3
but still capable of making Ptd-­‐Etn by decarboxylation of Ptd-­‐Ser, do not incorporate [ H]Etn into protein-­‐bound GPIs or into a 3
Man2-­‐GPI precursor that otherwise receives Etn-­‐P on Man-­‐1. However, radioactivity supplied as [ H]Ser is incorporated into the 3
Man2-­‐GPI after formation and decarboxylation of Ptd-­‐[ H]Ser (Menon and Stevens, 1992; Imhoff et al. 2000). The importance of Ptd-­‐Ser decarboxylation for GPI anchoring is underscored by the finding that the combination of a conditional gpi13 mutation, defective in the EtnP-­‐T-­‐III, with psd1Δ and psd2Δ, nulls in the two Ptd-­‐Ser decarboxylase genes, are inviable (Toh-­‐e and Oguchi, 2002). Direct transfer of Etn-­‐P from Ptd-­‐Etn to a GPI remains to be demonstrated in vitro. Phosphoethanolamine addition: importance of the alkaline phosphatase domain of Mcd4, Gpi7, and Gpi13. These three proteins all have a large lumenal loop of some 400 amino acids that contains sequences characteristic of the alkaline phosphatase superfamily (Gaynor et al. 1999; Benachour et al. 1999, Galperin and Jedrzejas, 2001), consistent with 227
involvement in formation or cleavage of a phosphodiester. This domain is important for function, because the G
E substitution that results in temperature-­‐sensitive growth and a conditional block in GPI precursor assembly in the mcd4-­‐174 mutant (Gaynor et al. 1999) lies in one of the two metal-­‐binding sites in alkaline phosphatase family members (Galperin and Jedrzejas, 2001). The metal is commonly zinc, and in vitro Etn-­‐P addition from an endogenous donor is zinc dependent (Sevlever 2+
et al. 2001) and Zn suppresses the temperature sensitivity of a gpi13 allele. Phosphoethanolamine addition: Man2-­‐GPI may be Mcd4’s preferred substrate. Three sets of findings suggest that Mcd4 may act preferentially on Man2-­‐GPI: i) treatment of wild type cells with the terpenoid lactone YW3548, which inhibits addition of Etn-­‐P to Man-­‐1, leads to accumulation of Man2-­‐GPI (Sütterlin et al. 1997, 1998), ii) Man2-­‐GPI is the most abundant of the accumulating GPIs in mcd4-­‐174, and iii) Man2-­‐GPI is the largest GPI formed in vitro by mcd4 membranes (Zhu et al. 2006). 22 SI P. Orlean Phosphoethanolamine addition: importance of the Etn-­‐P added to Man-­‐1 by Mcd4 and additional possible functions for Mcd4. The finding that mcd4 mutants accumulate unmodified Man2-­‐GPI suggests that the presence of Etn-­‐P on Man-­‐1 is important for GPI-­‐Man-­‐T-­‐III to add the third Man. The requirement, though, is not absolute because mcd4Δ cells can be partially rescued by overexpression of Gpi10 (Wiedman et al. 2007). In addition to enhancing the efficiency of mannosylation by Gpi10, the Etn-­‐P moiety on Man-­‐1 may be important for additional reasons. mcd4Δ cells expressing human or trypanosomal Gpi10 orthologues, Man-­‐T known to mannosylate Man2-­‐GPIs lacking Etn-­‐P on Man-­‐1 efficiently, still grow slowly (Zhu et al. 2006; Wiedman et al. 2007). Further, mcd4Δ cells expressing trypanosomal Gpi10 are retarded in export of GPI-­‐proteins from the ER, unable to remodel their GPI lipid moiety to ceramide, and are defective in selection of axial budding sites (Zhu et al. 2006). How the presence of Etn-­‐P on Man-­‐1 influences these processes is not yet known. Mutations in MCD4 also impact cellular processes that are not directly connected with GPI biosynthesis. Cells expressing the Mcd4-­‐P
301
L variant, but not G
227
E, are defective in the transport of Ptd-­‐Ser to the Golgi and vacuole for decarboxylation, but unaffected in GPI anchoring suggesting an additional role for Mcd4 in transport dependent Ptd-­‐Ser metabolism (Storey et al. 2001). Further, yeast overexpressing Mcd4 (as well as Gpi7 and Gpi13) release ATP into the medium, and Golgi vesicles from the Mcd4 overexpressers were enriched in that protein and showed elevated levels of ATP uptake (Zhong et al. 2003). It was suggested that Mcd4 normally mediates symport of ATP and Ptd-­‐Etn into the ER lumen, and that overexpression of the protein leads ATP to accumulate in secretory vesicles, which eventually fuse with the plasma membrane (Zhong et al. 2003). Phosphoethanolamine addition to Man-­‐2 and its possible functions. GPI-­‐Etn-­‐P-­‐II consists of catalytic Gpi7 and non-­‐
catalytic Gpi11. Both gpi7Δ and temperature-­‐sensitive gpi11Δ disruptants complemented by the human Gpi11 orthologue PIG-­‐
F accumulate a Man4-­‐GPI bearing Etn-­‐P on Man-­‐1 and Man-­‐3 but missing one on Man-­‐2 (Benachour et al. 1999; Taron et al. 2000). Because loss of GPI-­‐Etn-­‐P function leads to accumulation of a Man4-­‐GPI with Etn-­‐Ps on Man-­‐1 and Man-­‐3, GPI-­‐Etn-­‐P-­‐II may normally add Etn-­‐P to Man-­‐2 after GPI-­‐Etn-­‐P-­‐T-­‐III has modified Man-­‐3. However, because Man3-­‐ and Man4-­‐GPIs with a single Etn-­‐P on Man-­‐2 accumulate in the smp3 mutants and in temperature-­‐sensitive gpi11Δ strains complemented by the human Gpi11 orthologue (Taron et al. 2000; Grimme et al. 2001), GPI-­‐Etn-­‐P-­‐II has the capacity to act on Etn-­‐P-­‐free GPIs. Diverse phenotypes of gpi7Δ cells indicate that the Etn-­‐P moiety on Man-­‐2 is important for a number of reasons. First, the combination of gpi7Δ with the GPI transamidase mutation gpi8 leads to a synthetic growth defect, indicating that an Etn-­‐P on Man-­‐2 enhances transfer of GPIs to protein (Benachour et al. 1999). Second, gpi7Δ cells have defects in ER to Golgi transport of GPI-­‐proteins and GPI lipid remodeling to ceramide (Benachour et al. 1999). Third, GPI7 deletion leads to cell wall defects and P. Orlean 23 SI shedding of GPI-­‐proteins, indicating defective transfer of such proteins into the wall (Toh-­‐e and Oguchi, 1999; Richard et al., 2002). Lastly, gpi7Δ cells show a cell separation defect that results from mistargeting of Egt2, a GPI protein expressed in daughter cells and implicated in degradation of the septum (Fujita et al. 2004). These phenotypes suggest that the Etn-­‐P group on Man-­‐2 is recognized by GPI transamidase, the intracellular transport machinery, GPI lipid remodeling enzymes, and cell wall crosslinkers. An inability to remove Etn-­‐P from Man-­‐2 also leads to phenotypes (see Remodeling of protein bound GPIs). Phosphoethanolamine addition to Man-­‐3 by Gpi13 and the role of Gpi11. Gpi13 is the catalytic subunit of GPI-­‐Etn-­‐P-­‐T-­‐
III, and, as expected from the fact that it adds the Etn-­‐P that participates in the GPI transamidase reaction, GPI13 is essential. The major GPI accumulated by yeast strains depleted of Gpi13 is a Man4-­‐GPI with a single Etn-­‐P on Man-­‐1 (Flury et al. 2000; Taron et al. 2000). Gpi11 is likely involved in the GPI-­‐Etn-­‐P-­‐T-­‐III reaction as well, because a recently isolated gpi11-­‐Ts mutant also accumulates a Man4-­‐GPI with its Etn-­‐P on Man-­‐1 (K. Willis and P. Orlean, unpublished results), and human Gpi11 interacts with and stabilizes human Gpi13 (Hong et al. 2000). Human Gpi11 (Pig-­‐F) also interacts with human Gpi7 (Shishioh et al. 2005). The lipid accumulation phenotypes observed in various types of gpi11 mutants may prove to be explainable in terms of differential abilities of wild type Gpi11, mutant Gpi11, and human Gpi11 to interact with Gpi7, Gpi13, and possibly even Mcd4, and permit varying extents of Etn-­‐P modification. Because GPIs with the same chromatographic mobilities may be isoforms modified with Etn-­‐P at different positions, and because accumulating GPIs may be mixtures of isoforms, detailed structural analyses should give a clearer picture of the role of Gpi11 in Etn-­‐P modification. GPI transfer to protein: Depletion of Gab1 and Gpi8 leads to actin bar formation. Additional functions for Gab and Gpi18 are suggested by the finding that depletion of Gab1 or Gpi8 from yeast, but not of Gaa1, Gpi16, or Gpi17, leads to accumulation of bar-­‐like structures of actin that associate with the perinuclear ER and are decorated with cofilin (Grimme et al. 2004). This phenotype, which is not a general result of defective GPI anchoring, might reflect disruption of some functional interaction between resident ER membrane proteins and the actin cytoskeleton and consequent collapse of the ER around the nucleus (Grimme et al. 2004). Remodeling of protein-­‐bound GPIs: Roles of Bst1, Per1, and Gup1 in ER exit and transport of GPI proteins. Modifications of the GPI lipid by Bst1, Per1, and Gup1 are necessary for efficient transport of GPI proteins from the ER to the Golgi. Loss of Bst1 function leads to retarded transport of GPI-­‐proteins from the ER to the Golgi (Vashist et al. 2001), and delayed ER degradation of misfolded GPI proteins, suggesting that inositol deacylation generates sorting signals for ER exit of GPI proteins and for recognition by a quality control 24 SI P. Orlean mechanism for GPI-­‐proteins (Fujita et al. 2006; Fujita and Jigami, 2008). per1Δ and gup1Δ cells also show significantly delayed ER to Golgi transport of GPI-­‐proteins (Bosson et al. 2006; Fujita et al. 2006b). Lipid remodeling events generate a GPI able to associate with and be concentrated in membrane microdomains at ER exit sites prior to their export from the ER (Castillon et al. 2009). At these sites, the p24 complex of membrane proteins then serves as an adapter between GPI-­‐proteins and the COP II machinery to promote incorporation of GPI proteins into COP II vesicles specialized for transport of GPI-­‐proteins from the ER. Remodeled GPIs may bind p24 with higher affinity, therefore promoting export of the proteins bearing them (Castillon et al. 2011). In the Golgi, GPI-­‐proteins with remodeled anchors are released and proceed onwards along the secretory pathway. However, p24 complexes, which cycle between the ER and Golgi, again monitor the remodeling status of GPIs and exert a quality control function in the Golgi by sensing and retrieving proteins with unmodified GPIs to the ER, where they may encounter the resident ER remodeling enzymes (Castillon et al. 2011). Remodeling of the GPI lipid moiety to ceramide by Cwh43. Cwh43, which replaces the diacylglycerol moiety of GPIs with ceramide, is a large protein with 19 predicted transmembrane domains (Martin-­‐Yken et al. 2001; Ghugtyal et al. 2007; Umemura et al. 2007). cwh43Δ cells accumulate GPI-­‐proteins whose lipids are diacylglycerols with a very long acyl chain similar to the lipid generated after action of Bst1, Per1, and Gup1. Because ceramide remodeling requires prior action of Bst1, and per1Δ and gup1Δ strains show severe defects in remodeling, the exchange reaction seems to take place after the first three lipid modification steps. The mechanism is so far unknown, but could involve a phospholipase-­‐like reaction that replaces diphosphatidic acid with ceramide phosphate or diacylglycerol with ceramide (Ghugtyal et al. 2007; Fujita and Kinoshita, 2010). However, alternatives to such a linear remodeling pathway, in which Cwh43 acts instead on the Bst1 or Per1 products, have been discussed (Umemura et al. 2007). The C-­‐terminal domain of Cwh43 contains a motif that may be involved in recognition of inositol phosphate (Umemura et al. 2007). Because mcd4 and gpi7, mutants defective in addition of Etn-­‐P to Man-­‐1 and Man-­‐2, are affected in ceramide remodeling, Cwh43 may also recognize Etn-­‐P side-­‐branches. Cwh43 appears to act in the ER, where it remodels GPIs with a ceramide consisting of phytosphingosine bearing a C26 acyl chain, as well as in the Golgi, where the ceramide it introduces contains phytosphingosine with a hydroxy-­‐C26 acyl group (Reggiori et al. 1997). Removal of Etn-­‐P moieties from Man-­‐1 and Man-­‐2. The ER-­‐localized Ted1 and Cdc1 proteins are homologous to mammalian PGAP5, which removes EtN-­‐P moieties from Man-­‐2 (Fujita et al. 2009), and genetic interactions connect these two proteins processing and export of GPI-­‐proteins. Export of Gas1 is retarded in ted1Δ cells, and ted1Δ’s buffering genetic interactions with emp24Δ and erv5Δ, mutants deficient in two components of the p24 complex involved in maturation and trafficking of GPI proteins, indicate a functional relationship between the three proteins (Haass et al. 2007). Further, cdc1 P. Orlean 25 SI mutations are suppressed by per1/cos16 and gup1 mutations (Paidhungat and Garrett, 1998; Losev et al. 2008). Ted1 and Cdc1 contain a lumenal metallophosphoesterase domain (Haass et al. 2007; Losev et al. 2008), and, consistent with this, cdc1’s 2+
temperature-­‐sensitivity is suppressed by Mn , the cation required by PGAP5 (Fujita et al. 2009). These findings are in turn consistent with Ted1 and Cdc1 being GPI-­‐Etn-­‐P phosphodiesterases, but this possibility awaits biochemical confirmation. Literature Cited Castillon, G. A., Aguilera-­‐Romero, A., Manzano-­‐Lopez, J., Epstein, S., Kajiwara, K., et al., 2011 The yeast p24 complex regulates GPI-­‐anchored protein transport and quality control by monitoring anchor remodeling. Mol. Biol. Cell. 22: 2924-­‐2936. Castillon, G. A., Watanabe, R., Taylor, M., Schwabe, T. M., Riezman, H., 2009 Concentration of GPI-­‐anchored proteins upon ER exit in yeast. Traffic 10: 186–200. Coutinho, P. M., Deleury, E., Davies, G. J., Henrissat, B., 2003 An evolving hierarchical family classification for glycosyltransferases. J. Mol. Biol. 328: 307-­‐317 Franzot, S. P, Doering, T. L. 1999 Inositol acylation of glycosylphosphatidylinositols in the pathogenic fungus Cryptococcus neoformans and the model yeast Saccharomyces cerevisiae. Biochem. J. 340: 25-­‐32. Fujita, M., Jigami, Y., 2008 Lipid remodeling of GPI-­‐anchored proteins and its function. Biochim. Biophys. Acta 1780: 410-­‐420. Kostova, Z., Yan, B. C., Vainauskas, S., Schwartz, R., Menon, A. K., et al. 2003 Comparative importance in vivo of conserved glutamates in the EX7E-­‐motif retaining glycosyltransferase Gpi3p, the UDP-­‐GlcNAc-­‐binding subunit of the first enzyme in glycosylphosphatidylinositol assembly. Eur. J. Biochem. 270: 4507-­‐4514. Losev, E., Papanikou, E., Rossanese, O. W., Glick, B. S., 2008 Cdc1p is an endoplasmic reticulum-­‐localized putative lipid 2+
phosphatase that affects Golgi inheritance and actin polarization by activating Ca signaling. Mol. Cell. Biol. 28: 3336–3343. 26 SI P. Orlean Murakami, Y., Siripanyapinyo, U., Hong, Y., Kang, J. Y., Ishihara, S., Nakakuma, H., et al., 2003 PIG-­‐W is critical for inositol acylation but not for flipping of glycosylphosphatidylinositol-­‐anchor. Mol. Biol. Cell 14: 4285-­‐4295. 2+
Paidhungat, M., Garrett, S., 1998 Cdc1 and the vacuole coordinately regulate Mn homeostasis in the yeast Saccharomyces cerevisiae. Genetics 148: 1787–1798. Reggiori, F., Canivenc-­‐Gansel, E., Conzelmann, A., 1997 Lipid remodeling leads to the introduction and exchange of defined ceramides on GPI proteins in the ER and Golgi of Saccharomyces cerevisiae. EMBO J. 16: 3506-­‐3518. Sevlever, D., Mann, K. J., Medof, M. E., 2001, Differential effect of 1,10-­‐phenanthroline on mammalian, yeast, and parasite glycosylphosphatidylinositol anchor synthesis. Biochem. Biophys. Res. Commun. 288: 1112-­‐1118. Shishioh, N., Hong, Y., Ohishi, K., Ashida, H., Maeda, Y., et al., 2005 GPI7 is the second partner of PIG-­‐F and involved in modification of glycosylphosphatidylinositol. J. Biol. Chem. 280: 9728-­‐9734. Sipos, G., Reggiori, F., Vionnet, C., Conzelmann, A., 1997 Alternative lipid remodelling pathways for glycosylphosphatidylinositol membrane anchors in Saccharomyces cerevisiae. EMBO J. 16: 3494-­‐3505. Sobering, A. K., Romeo, M. J., Vay, H. A., Levin, D. E., 2003 A novel Ras inhibitor, Eri1, engages yeast Ras at the endoplasmic reticulum. Mol. Cell. Biol. 23: 4983-­‐49890. Storey, M. K., Wu, W. I., Voelker, D. R., 2001 A genetic screen for ethanolamine auxotrophs in Saccharomyces cerevisiae identifies a novel mutation on Mcd4p, a protein implicated in glycosylphosphatidylinositol anchor synthesis. Biochim. Biophys. Acta. 1532: 234-­‐247. Toh-­‐e, A., Oguchi, T., 2002 Genetic characterization of genes encoding enzymes catalyzing addition of phospho-­‐ethanolamine to the glycosylphosphatidylinositol anchor in Saccharomyces cerevisiae. Genes Genet. Syst. 77: 309-­‐322. P. Orlean 27 SI Vashist, S., Kim, W., Belden, W. J., Spear, E. D., Barlowe, C., et al., 2001 Distinct retrieval and retention mechanisms are required for the quality control of endoplasmic reticulum protein folding. J. Cell Biol. 155: 355-­‐368. Zhong, X., Malhotra, R., Guidotti, G., 2003 ATP uptake in the Golgi and extracellular release require Mcd4 protein and the +
vacuolar H -­‐ATPase. J. Biol. Chem. 278: 33436-­‐33444. 28 SI P. Orlean File S5 Sugar nucleotide transport This Supporting File contains additional information related to Biosynthesis of Wall Components Along the Secretory Pathway, Sugar nucleotide transport. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. GDP-­‐Man transport: The GDP-­‐Man transporter, Vrg4/Vig4. This protein forms homodimers (Abe et al. 1999; Gao and Dean, 2000), shows a wide distribution in the Golgi, and contains a GALNK motif involved in GDP-­‐Man binding (Gao et al. 2001). Gda1 and Ynd1. Evidence these proteins have partially overlapping functions is as follows. i) Deletion of either GDA1 or YND1 impacts mannosylation of N-­‐ and O-­‐glycans, ii) high-­‐level expression of YND1 corrects some of gda1Δ’s glycosylation defects, and iii) gda1Δ ynd1Δ double mutants have a synthetic phenotype and show growth and cell wall defects (Gao et al. 1999). However, gda1Δ ynd1Δ double mutants are viable and capable of some mannosylation of N-­‐ and O-­‐linked glycans, indicating that GDP-­‐Man can enter the Golgi in their absence, and suggesting there may be a mechanism for GDP exit independent of GDP hydrolysis (D’Alessio et al. 2005). GMP generated upon Man-­‐P transfer to glycoproteins could also be a source of antiporter, but it is not a significant one because because the glycans made gda1Δ or gda1Δ ynd1Δ strains are not affected by disruption of MNN4 or MNN6 (Jigami and Odani, 1999; D’Alessio et al. 2005). Other sugar nucleotide transport activities: Transport activities for UDP-­‐Glc, UDP-­‐GlcNAc, and UDP-­‐Gal also occur in S. cerevisiae (Roy et al. 1998; 2000 Castro et al. 1999), and there are eight further candidate transporters (Dean et al. 1997; Esther et al. 2008), a couple of which have been associated with these transport activities. Some of the transporters may have specificity for more than one sugar nucleotide. In the case of UDP-­‐Glc, transport activity was present in the ER (Castro et al. 1999), but the responsible protein for that activity has yet to identified, although broad specificity Yea4 and Hut1 (see below) may transport UDP-­‐Glc (Esther et al. 2008). One possible need for UDP-­‐Glc transport into the ER might be for a glucosylation reaction at an early stage of β1,6-­‐glucan assembly (Section VI). The Hut1 protein is a candidate for the UDP-­‐Gal transporter (Kainuma et al. 2001), but whether that is Hut1’s primary role in vivo is unclear because galactose has not been detected on S. cerevisiae glycans. Yea4 was characterized as an ER-­‐localized UDP-­‐GlcNAc transporter and its deletion impacts chitin synthesis (Roy et al. 2000; Section V). Of the other P. Orlean 29 SI transporter homologs, Hvg1 resembles Vrg4 most closely, but hvgΔ cells have neither a mannosylation nor a GDP-­‐Man transport defect (Dean et al. 1997). The roles of the other proteins in sugar nucleotide transport, if any, is unknown. One or more transporters may supply the Golgi GlcNAc-­‐T Gnt1 with its substrate (Section IV.1.c.ii). Literature Cited D'Alessio, C., Caramelo, J. J., Parodi, A. J., 2005 Absence of nucleoside diphosphatase activities in the yeast secretory pathway does not abolish nucleotide sugar-­‐dependent protein glycosylation. J. Biol. Chem. 280: 40417-­‐40427. Gao, X. D., Dean, N., 2000 Distinct protein domains of the yeast Golgi GDP-­‐mannose transporter mediate oligomer assembly and export from the endoplasmic reticulum. J. Biol. Chem. 275: 17718-­‐17727. Gao, X. D., Nishikawa, A., Dean, N., 2001 Identification of a conserved motif in the yeast Golgi GDP-­‐mannose transporter required for binding to nucleotide sugar. J. Biol. Chem. 276: 4424-­‐4432. 30 SI P. Orlean File S6 Chitin This Supporting File contains additional information and discussion related to Biosynthesis of Wall Components at the Plasma Membrane, Chitin. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Septum formation: Phenotypes of chs1Δ chs2Δ chs3Δ triple mutants. chs1Δ chs2Δ chs3Δ strains grew very slowly but acquired a suppressor mutation that conferred a growth rate as fast as that of a chs2Δ mutant, although over a third of suppressed or unsuppressed cells in a culture were dead (Schmidt, 2004). Membranes from the triple mutants had no detectable chitin synthase activity. Unsuppressed triple mutants formed chains of up to eight cells that appeared to be connected by “cytoplasmic stalks”, whereas suppressed strains formed shorter chains. Nuclear division continued in the mutant, but in some cells, nuclear segregation was unsuccessful. Ultrastructural analysis showed that in both suppressed and unsuppressed mutants, a bulky remedial septum arises upon thickening of the lateral walls in the mother cell-­‐bud neck region. The suppressor was not identified, but its effect was to allow the remedial septa to be formed more efficiently. The phenotypes of the triple chitin synthase mutants indicate that although it is possible for S. cerevisiae to grow without chitin, Chs3-­‐dependent chitin synthesis is nonetheless important for remedial septum formation in chs2Δ cells. Chitin synthase biochemistry: Directionality and mechanism of extension of β1,4-­‐linked polysaccharide chains. Although the bacterial chitin synthase homologue NodC extends chito-­‐oligosaccharides at their non-­‐reducing ends (Kamst et al. 1999), both reducing-­‐ and non-­‐reducing end extension has been reported for Chs-­‐related vertebrate Class I hyaluronate synthases (Weigel and DeAngelis, 2007), and extension by insertion of Glc at the reducing end of a glycan chain has also been proposed for a bacterial cellulose synthase (Han and Robyt, 1998). The latter mechanism was suggested to involve a lipid pyrophosphate intermediate. However, no evidence has been obtained for any lipid-­‐linked intermediate in chitin synthesis. The growing glycan chain may be extruded through the plasma membrane through a pore made up by a bundle of transmembrane helices, which occur towards the C-­‐
terminus of chitin synthases (Delmer, 1999; Guerriero et al. 2010; Merzendorfer, 2011; Carpita, 2011). Separate proteins might mediate chitin translocation, but no candidates have been identified. With non-­‐reducing end extension, a nascent chitin chain would be extruded into the cell wall reducing end first, which would be compatible with the formation of linkages between P. Orlean 31 SI chitin and non-­‐reducing ends of β-­‐glucans (see Cross-­‐linkage of chitin to β1,6-­‐ and β1,3-­‐glucan; Kollar et al. 1995, 1997; Cabib and Duran, 2005; Cabib, 2009). The stereochemical challenge in formation of β1,4-­‐linked polysaccharides. Each sugar in a β1,4-­‐linked polymer is rotated by about 180° relative to its neighbor, which presents the synthase with a steric challenge, because with successive rounds of addition of a β1,4-­‐linked GlcNAc, the new acceptor 4-­‐OH would alternate between two positions relative to incoming substrate and catalytic residues. Various ways of overcoming this, without invoking movements of the enzyme or the acceptor glycan, have been considered. The first possibility, that UDP-­‐di-­‐N-­‐acetylchitobiose is the donor, has been ruled out by the finding that yeast membranes make no chitin when supplied with synthetic UDP-­‐GlcNAc2 (Chang et al. 2003). The second possibility is that β1,4-­‐linked polysaccharide synthases have two UDP-­‐sugar binding sites that orient the monosaccharides such that neither enzyme nor polymer needs to rotate, then catalyzes two glycosyltransfers (Saxena et al. 1995; Guerriero et al. 2010; Carpita, 2011). Evidence supportive of a two active site mechanism came from the finding that a bivalent UDP-­‐GlcNAc analog consisting of two tethered uridine mimetics, envisaged to bind in both active sites, was a better inhibitor than the monomeric analog (Yaeger and Finney, 2004). The observation that the NodC protein, Chs1, and Chs2 all synthesize odd-­‐ as well as even-­‐numbered chito-­‐ooligosaccharides in vitro (Kang et al. 1984; Yabe et al. 1998; Kamst et al. 1999) is consistent with extension by addition with single GlcNAcs, but extension of GlcNAc, GlcNAc3, or GlcNAc5 by two GlcNAcs at a time would also generate odd-­‐numbered chito-­‐oligosaccharides, if these oligosaccharides are indeed used as primers. Third, it is possible that a chain is extended by a dimeric synthase whose subunits alternately add GlcNAcs, as discussed for cellulose synthase (Carpita, 2011). Consistent with this notion, a two-­‐hybrid analysis indicated that Chs3 can interact with itself (DeMarini et al. 1997). The molecular weight of purified native Chs1 was estimated to be around 570,000, approximately consistent with a tetramer, but the authors noted the result may have been due to protein aggregation (Kang et al. 1984). 14
In vitro properties of yeast chitin synthases. Chitin synthase assays typically detect the transfer of [ C]GlcNAc from 14
UDP[ C]GlcNAc to insoluble chitin that is then collected on filters, but a high-­‐throughput method that relies on product binding to immobilized wheat germ agglutinin has also been described (Lucero et al. 2002). Of the two procedures, the filtration method would not detect chito-­‐oligosaccharides (Yabe et al. 1998). CS I, CS II, and CS III activities differ in their pH optima and their responses to divalent cations (Sburlati and Cabib, 1986; Orlean, 1987; Choi and Cabib, 1994). The three chitin synthase activities have Kms for UDP-­‐GlcNAc in the range of 0.5-­‐1.3 mM (Kang et al. 1984; Sburlati and Cabib, 1986; Orlean, 1987; Uchida et al. 1996). At low substrate concentrations relative to Km (0.03-­‐0.1 mM), purified Chs1 and membranes from cells overexpressing CHS2 make chito-­‐oligosaccharides (Kang et al. 1984; Yabe et al. 1998). Whether these are bona fide chitin 32 SI P. Orlean synthase products whose formation reflects low rates of chain extension, or whether the oligosaccharides are generated by chitinase activity on longer nascent chains is not clear (Kang et al. 1984). Effects of free GlcNAc and chitin oligosaccharides on chitin synthesis. S. cerevisiae’s three chitin synthases are all stimulated up to a few fold in vitro by high concentrations of free GlcNAc (e.g. 32 mM; Sburlati and Cabib, 1986; Orlean, 1987). Neither the mechanistic basis nor the physiological relevance of this are clear, but possible explanations are that GlcNAc serves as a primer or allosteric activator in the chitin synthetic reaction. Results of a kinetic analysis of the chitin synthase activity in wild type membranes led to the proposal that GlcNAc participates along with UDP-­‐GlcNAc in a two substrate reaction with an ordered mechanism in which UDP-­‐GlcNAc binds first (Fähnrich and Ahlers, 1981). Consistent with the idea that GlcNAc serves as a primer or co-­‐substrate, the bacterial NodC chitin synthase homologue incorporates free GlcNAc at the reducing end of chito-­‐
oligosaccharide chains that are extended at their non-­‐reducing end by GlcNAc transfer from UDP-­‐GlcNAc (Kamst et al. 1999). However, were free GlcNAc to serve as a co-­‐substrate or activator of chitin synthases in vivo, there would have to be a mechanism to generate it, for example from GlcNAc-­‐1-­‐P or GlcNAc-­‐6-­‐P (see Precursors and Carrier Lipids) or by turnover of GlcNAc-­‐containing molecules. Growing chitin chains presumably serve as acceptors for further GlcNAc addition, but such a primer function has not been shown using short oligosaccharides. NodC did not use short chito-­‐oligosaccharides as GlcNAc acceptor from UDP-­‐GlcNAc (Kamst et al. 1999), nor did purified Chs1 elongate chitotetraose into insoluble chitin in the presence of UDP-­‐GlcNAc (Kang et al. 1984). However, inclusion of 1 mM GlcNAc5 and GlcNAc8 in assays of membrane preparations expressing predominantly Chs1 led to about a 1.25-­‐fold increase in incorporation of GlcNAc into chitin from UDP-­‐GlcNAc in the presence of free GlcNAc (Becker et al. 2011), suggesting a primer function for longer chito-­‐oligosaccharides. The initiation and early elongation steps in chitin synthesis clearly still need to be defined. S. cerevisiae’s chitin synthases and auxiliary proteins: Chitin synthase classes. Fungal chitin synthases can be classified into five to seven classes on the basis of amino acid sequence similarity, with S. cerevisiae Chs1, Chs2, and Chs3 being assigned to Classes I, II, and IV respectively (Roncero, 2002; Ruiz-­‐Herrera et al. 2002; Van Dellen et al. 2006; Merzendorfer, 2011). Members of the other classes are found in filamentous fungi. S. cerevisiae’s chitin synthases show most amino acid sequence divergence in their amino terminal halves, and these non-­‐
homologous regions may make interactions with proteins involved in regulation or trafficking of the individual synthases (Ford et al. 1996). Deletion analyses have shown that amino acids in Chs3’s hydrophilic C-­‐terminal region are also important for function (Cos et al. 1998). P. Orlean 33 SI Chitin synthase I: Activity of N-­‐terminally truncated Chs1. N-­‐terminally truncated forms of Chs1 lacking up to 390 amino acids show a gradual lowering of both specific activity and their ability to be activated by trypsin (Ford et al. 1996). Chitin synthase II and proteins impacting its localization and activity: Detection of Chs2’s activity. Studies of Chs2 enzymology use membranes from strains overexpressing the protein because the activity of genomically encoded Chs2 in membranes of cells grown in minimal medium is negligible (Nagahashi et al. 1995). The high amounts of in vitro activity obtained by overexpressing Chs2 indicate that levels of Chs2 activity are not tightly limited by endogenous activating or regulatory proteins, in contrast to Chs3. Effects of proteolysis on wild type and truncated forms of Chs2. Although endogenously activated, processed forms of Chs2 have not been identified, trypsin treatment of partially purified, full-­‐size and N-­‐terminally truncated Chs2 generated a range of discrete protein fragments. The smallest of these, a 35 kDa protein containing the amino acid sequences proposed to be involved in catalysis, was suggested to be sufficient for catalysis, although the instablity of this form prevented its purification to test this notion (Uchida et al. 1996). Some 220 amino terminal amino acids of Chs2 are dispensable for in vivo function (Ford et al. 1996), and moreover, Chs2 versions lacking these amino terminal amino acids have higher in vitro activity than the full-­‐length protein, and this activity is stimulated by trypsin (Uchida et al. 1996; Martínez-­‐Rucobo et al. 2009). Other truncated forms of Chs2, or forms with amino acid substitutions, also vary in their extent of activation by trypsin (Ford et al. 1996; Uchida et al. 1996). It has been noted that amino acid deletions or substitutions in Chs2 could perturb interactions with native mechanisms for activation and localization of the protein (Ford et al. 1996). Chitin synthase III and proteins impacting its localization and activity: Relationship between Pfa4 and Chs7 and their roles in Chs3 exit from the ER. Chs3 interacts with Chs7 and is palmitoylated by Pfa4. The Chs3-­‐Chs7 interaction also occurs in pfa4Δ cells, though to a slightly reduced extent, and Chs3 can still be palmitoylated, likewise to a lesser extent, in chs7Δ cells, indicating that Chs3 palmitoylation is not obligatory for Chs3 recognition by Chs7 (Lam et al. 2006). Pfa4 does not palmitoyate Chs7. It seems that Pfa4 and Chs7 act in parallel, though not wholly independently, to promote folding of Chs3 prior to the synthase’s exit from the ER. These roles of Pfa4 and Chs7 are specific to Chs3, for neither is required for exit of Chs1 and Chs2 from the ER (Trilla et al. 1999; Lam et al. 2006). Rcr1 and Yea4 in Chs3-­‐dependent chitin synthesis. These proteins have both been localized to the ER membrane. Rcr1 has a slight negative regulatory effect on Chs3-­‐dependent chitin synthesis. High copy RCR1 confers resistance to Congo Red, a dye that binds chitin (as well as β1,3-­‐glucan (Kopecká and Gabriel, 1992)), whereas rcr1Δ cells showed slightly increased 34 SI P. Orlean sensitivity to Congo Red and CFW (Imai et al. 2005). Wild type cells overexpressing RCR1 have 70% of the chitin in control cells, and rcr1Δ cells make 115% of wild type levels of chitin. However, RCR1 overexpression affects neither the amount nor localization of Chs3, Chs5, and Chs7, nor do Rcr1 and Chs7 physically interact (Imai et al. 2005). The role of Rcr1 in Chs3-­‐
dependent chitin synthesis is therefore not clear, but the protein has also been reported to act after the ER and have a role in an endosome-­‐vacuole pathway that impacts trafficking of plasma membrane nutrient transporters (Kota et al. 2007). The second ER membrane protein, Yea4, was identified through its homology to the Kluyveromyces lactis UDP-­‐GlcNAc transporter (Roy et al. 2000). Membrane vesicles from cells overexpressing Yea4 have 8-­‐fold elevated levels of UDP-­‐GlcNAc transport activity, consistent with Yea4’s function as a transporter (Roy et al. 2000). yea4Δ cells contain 65% of wild type levels of chitin, implicating Yea4 in chitin synthesis, but whether and how Yea4’s transport activity contributes to this process is unclear. Role of exomer in transport of wall related proteins other than Chs3. Exomer has roles in polarized transport of other wall related proteins to the cell surface. Thus, transport of Fus1, which promotes cell fusion during mating, requires Chs5 for transport to the shmoo tip (Santos and Snyder, 2003), along with the ChAPs Bch1 and Bus7, but not Chs6 (Barfield et al. 2009). Further, much of the GPI-­‐anchored chitin-­‐β1,3-­‐glucan cross-­‐linker Crh2 (see Cross-­‐linkage of chitin to β1,6-­‐ and β1,3-­‐glucan) fails to reach sites of polarized growth and accumulates intracellularly in chs5Δ, although another GPI-­‐protein, Cwp1, was unaffected (Rodriguez-­‐Pena et al. 2002). Co-­‐transport of Chs3 and Crh2 would ensure colocalization of these proteins for efficient cross linking of nascent chitin to β1,3-­‐glucan. Role of Chs4 farnesylation in the activation and localization of Chs3. Chs4 has a C-­‐terminal farnesylation site (Bulawa et al. 1993; Trilla et al. 1997) that is used (Grabinska et al. 2007) and the consensus of studies of the importance of the prenyl group is that the modification has roles in Chs4 function and localization. Mutants expressing a non-­‐farnesylatable Cys to Ser variant of Chs4 make one third of normal amounts of chitin, have lower in vitro CS III activity, and show CFW resistance (Grabinska et al. 2007; Meissner et al. 2010). In two of three studies, the prenylation site mutant of Chs4 was found in the cytoplasm, suggesting that lipidation is important for membrane localization of the protein (Reyes et al. 2007; Meissner et al. 2010). Chs4 reaches the plasma membrane in mutants affected in Chs3 transport, indicating it is transported there independently of Chs3 (Reyes et al. 2007), but two sets of findings raise the possibility that Chs3 interacts with Chs4 at the level of the ER. First, two-­‐hybrid analyses established that cytoplasmic domains of Chs3 and the ER-­‐localized CAAX protease Ste24 interact. Second, ste24Δ cells exhibit moderate CFW resistance, chitin content is reduced, and less Chs3 was localized at the bud neck. Vice versa, high-­‐copy expression of STE24 leads to CFW sensitivity and some increase in cellular chitin (Meissner et al. 2010). Chs4 localization, though, was not affected in ste24Δ, nor was an interaction detected between Chs4 and Ste24. It was P. Orlean 35 SI suggested that Chs3 recruits farnesylated Chs4 in the ER for processing by Ste24, and that the modification contributes to subsequent correct localization of Chs3 and activation of CS III (Meissner et al. 2010). Chitin synthase III in mating and ascospore wall formation: Regulation of Chs3 during chitosan synthesis. The Chs4 homologue Shc1, which is 43% identical to Chs4 but expressed only during sporulation, has a role in chitosan synthesis, because homozygous shc1Δ shc1Δ diploids make ascospores with very little chitosan (Sanz et al. 2002). Shc1 and Chs4 are functionally related because when Shc1 is expressed in vegetative cells, it can activate CS III, and when Chs4 is overexpressed in shc1Δ shc1Δ diploids, it partially corrects the sporulation defect (Sanz et al. 2002). However, although Shc1 serves as CS III activator in chs4Δ cells, it does so without properly localizing Chs3 to septins as Chs4 does in vegetative cells, likely because it cannot interact with Bni4 (Sanz et al. 2002). Haploid chs4Δ shc1Δ cells do not show a synthetic growth defect, indicating they are not an essential redundant pair, and indeed, analyses of the SHC1 genetic interaction network suggests Shc1 may have additional roles distinct from those of Chs4 that are not directly related to chitin synthesis (Lesage et al. 2005). Sporulation-­‐specific kinase Sps1, regulates mobilization of Chs3 as well as sporulation-­‐specific β1,3-­‐glucan synthase Fks2/Gsc2 (see β1,3-­‐glucan) to the prospore membrane (Iwamoto et al. 2005). Literature Cited Barfield, R. M., Fromme, J. C., Schekman, R., 2009 The exomer coat complex transports Fus1p to the plasma membrane via a novel plasma membrane sorting signal in yeast. Mol. Biol. Cell 20: 4985-­‐4996. Becker, H.F., Piffeteau, A., Thellend, A. 2011 Saccharomyces cerevisiae chitin biosynthesis activation by N-­‐acetylchitooses depends on size and structure of chito-­‐oligosaccharides. BMC Res. Notes. 4: 454. Carpita, N. C., 2011 Update on mechanisms of plant cell wall biosynthesis: how plants make cellulose and other (1→4)-­‐β-­‐D-­‐
glycans. Plant Physiol. 155: 171-­‐184. Chang, R., Yeager, A. R. Finney, N. S., 2003 Probing the mechanism of a fungal glycosyltransferase essential for cell wall biosynthesis. UDP-­‐chitobiose is not a substrate for chitin synthase. Org. Biomol. Chem. 1: 39-­‐41. 36 SI P. Orlean Choi, W. J., Cabib, E., 1994 The use of divalent cations and pH for the determination of specific yeast chitin synthetases. Anal. Biochem. 219: 368-­‐372. Delmer, D. P., 1999 Cellulose biosynthesis: exciting times for a difficult field of study. Annu. Rev. Plant Physiol. Plant Mol. Biol. 50: 245-­‐276. Fähnrich, M., Ahlers, J. 1981 Improved assay and mechanism of the reaction catalyzed by the chitin synthase from Saccharomyces cerevisiae. Eur. J. Biochem. 121: 113-­‐118. Ford, R. A., Shaw, J. A., Cabib, E., 1996 Yeast chitin synthases 1 and 2 consist of a non-­‐homologous and dispensable N-­‐terminal region and of a homologous moiety essential for function. Mol. Gen. Genet. 252: 420-­‐428. Imai, K., Noda, Y., Adachi, H., Yoda, K., 2005 A novel endoplasmic reticulum membrane protein Rcr1 regulates chitin deposition in the cell wall of Saccharomyces cerevisiae. J. Biol. Chem. 280: 8275-­‐828. Kopecká, M., Gabriel, M., 1992 The influence of congo red on the cell wall and (1-­‐3)-­‐β-­‐D-­‐glucan microfibril biogenesis in Saccharomyces cerevisiae. Arch Microbiol. 158: 115-­‐126. Guerriero, G., Fugelstad, J., Bulone, V. 2010 What do we really know about cellulose biosynthesis in higher plants? J. Integr. Plant Biol. 52: 161-­‐175. Iwamoto, M. A., Fairclough, S. R., Rudge, S. A., Engebrecht, J., 2005 Saccharomyces cerevisiae Sps1p regulates trafficking of enzymes required for spore wall synthesis. Eukaryot. Cell 4: 536-­‐544. Kota, J., Melin-­‐Larsson, M., Ljungdahl, P. O., Forsberg, H., 2007 Ssh4, Rcr2 and Rcr1 affect plasma membrane transporter activity in Saccharomyces cerevisiae. Genetics 175: 1681-­‐1694. P. Orlean 37 SI Lucero, H. A., Kuranda M. J., Bulik, D. A., 2002 A nonradioactive, high throughput assay for chitin synthase activity. Anal. Biochem. 305: 97-­‐105. Nan, N. S., Robyt, J. F. 1998. The mechanism of Acetobacter xylinum cellulose biosynthesis: direction of chain elongation and the role of lipid pyrophosphate intermediates in the cell membrane. Carbohydrate Res. 313: 125-­‐133. Santos, B., Snyder, M., 2003. Specific protein targeting during cell differentiation: polarized localization of Fus1p during mating depends on Chs5p in Saccharomyces cerevisiae. Eukaryot. Cell 2: 821–825. Van Dellen, K. L., Bulik, D. A., Specht, C. A., Robbins, P. W., Samuelson, J. C., 2006 Heterologous expression of an Entamoeba histolytica chitin synthase in Saccharomyces cerevisiae. Eukaryot. Cell. 5: 203-­‐206. Weigel, P. H., DeAngelis, P. L., 2007 Hyaluronan synthases: a decade-­‐plus of novel glycosyltransferases. J. Biol. Chem. 282: 36777-­‐36781. Yaeger, A.R., Finney, N. S., 2004 The first direct evaluation of the two-­‐active site mechanism for chitin synthase. J. Org. Chem. 69: 613-­‐618. 38 SI P. Orlean File S7 β 1,3-­‐glucan This Supporting File contains additional information and discussion related to Biosynthesis of Wall Components at the Plasma Membrane, β 1,3-­‐glucan. The subheadings used in the main text are retained, and new subheadings are underlined. Fks family of β 1,3-­‐glucan synthases: Identification of Fks1, Fks2, and Fks3. Fks1 (Cwh53/Etg1/Gsc1/Pbr1) was identified in screens for hypersensitivity to the calcineurin inhibitors FK506 and cyclosporin A and to CFW, for resistance to echinocandin and papulocandin, and following purification of β1,3-­‐glucan synthase activity (reviewed by Orlean, 1997 and Lesage and Bussey, 2006). Cross-­‐hybridization with FKS1 and copurification with Fks1 led to identification of Fks2/Gsc2, which is 88% identical to Fks1 (Inoue et al. 1995; Mazur et al. 1995). The S. cerevisiae proteome also contains Fks3, which is 55% identical to Fks1 and Fks2 (Dijkgraaf et al. 2002). The Fks proteins are assigned to GT Family 48, and a strong case can be made for them being processive β1,3-­‐glucan synthases themselves, although roles as glucan exporters cannot yet be excluded (Mazur et al. 1995; Dijkgraaf et al. 2002; Lesage and Bussey, 2006). Functional domains of Fks1. Fks1 is predicted to have an N-­‐terminal cytoplasmic domain of some 300 amino acids that is followed by six transmembrane helices, a second cytoplasmic domain of about 600 amino acids, then 10 transmembrane helices (Inoue et al. 1995; Mazur et al. 1995; Qadota et al. 1996; Dijkgraaf et al. 2002; Okada et al. 2010). Three functional domains have been distinguished (Okada et al. 2010). Amino acids important for β1,3 glucan synthesis in vivo are located in the first cytoplasmic domain. Mutations here have little impact on in vitro activity and do not affect the protein’s interaction with Rho1, but cells have a lowered β1,3 glucan content. Mutations in the second cytoplasmic domain that lie close to the C-­‐
terminus of the sixth helix lead to a loss of cell polarity as well as defects in endocytosis, but have little effect on in vitro and in vivo b-­‐glucan synthesis, and this part of Fks1 may interact with factors involved in cell polarity (Okada et al. 2010). Mutations in Fks1 in residues more distal to the sixth helix lead to low in vitro glucan synthase activity and large decreases in in vivo 14
incorporation of [ C]glucose into β1,3 glucan, suggesting that if Fks1 is a synthase, this part of the protein contains the catalytic site (Dijkgraaf et al. 2002; Okada et al. 2010). Fatty acid elongases and phytosphingosine and Fks1 function. The ER-­‐localized fatty acid elongase Elo2/Gns1 may impact Fks1 at the level of that organelle, because gns1 mutants, isolated on account of their resistance to a papulocandin analogue, have very low in vitro β1,3-­‐glucan synthase activity (el-­‐Sherbeini and Clemas, 1995) and accumulate P. Orlean 39 SI phytosphingosine in the ER membrane (Abe et al. 2001). Phytosphingosine inhibits β1,3 glucan synthase in vitro, leading to the idea that this sphingolipid synthetic intermediate is a negative regulator of β1,3-­‐glucan synthesis at the level of the ER (Abe et al. 2001). Roles of the Fks proteins in β 1,3-­‐glucan synthesis Roles of Fks3 and Fks3 in sporulation. Fks2 is important in sporulation because fks2Δ fks2Δ diploids have a severe defect in this process (Mazur et al. 1995; Huang et al. 2005), and form disorganized ascospore walls with lower relative amounts of hexose in their alkali-­‐insoluble fraction and a lower alkali soluble β1,3-­‐glucan content (Ishihara et al. 2007). Homozygous fks3Δ fks3Δ diploids also form abnormal spores, indicating a role for the third Fks homologue in ascopore wall formation, but showed no alteration in the distribution of hexoses between alkali soluble-­‐ and insoluble fractions (Ishihara et al. 2007). However, the walls of ascospores formed in diploids lacking both Fks2 and Fks3 were more disorganized than those of ascospores made by fks2Δ fks2Δ diploids (Ishihara et al. 2007). Expression of FKS2 or FKS1 under the control of the FKS2 promoter, but not the FKS1 promoter, corrected the sporulation defect of homozygous fks1Δ fks2Δ diploids, suggesting that the function of Fks2 in sporulating diploids resembles that of Fks1 in vegetative cells. In contrast, overexpression of FKS3 did not suppress the phenotype of fks2Δ spores, and FKS1 or FKS2 overexpression does not correct the defect in fks3Δ spores, indicating Fks3’s function in sporulation does not overlap with that of Fks2. It was proposed that Fks2 is primarily responsible for synthesis of β1,3-­‐glucan in the ascospore wall, and that Fks3, rather than functioning as a synthase, modulates glucan synthesis by interacting with glucan synthase regulators such as Rho1 (Ishihara et al. 2007). 40 SI P. Orlean File S8 β1,6-­‐Glucan This Supporting File contains additional information and discussion related to β 1,6-­‐Glucan. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Proteins involved in β 1,6-­‐glucan assembly ER proteins: Fungus-­‐specific ER chaperones required for β1,6-­‐glucan synthesis: Evidence for the chaperone function of Rot1, Big1, and Keg1 in β1,6-­‐glucan synthesis. Rot1, Big1, and Keg1, which do not resemble known carbohydrate-­‐active enzymes, seem unlikely to catalyze formation of β1,6-­‐glucan (Lesage and Bussey, 2006). Rather, they seem to function as ER chaperones with varying degrees of importance for the stability of proteins involved in β1,6-­‐glucan synthesis, and in some cases, they may cooperate. Observations supporting this notion, and indicating a relationship to Kre5, are as follows. Analyses of levels of β1,6-­‐glucan synthesis-­‐related proteins in a rot1-­‐Ts mutant indicate that Kre6 has the strongest dependence on Rot1 for stability, although Kre5 and Big1 show appreciable dependence as well (Takeuchi et al. 2008). Keg1, a protein essential for growth in osmotically supported medium, physically interacts with Kre6 in the ER membrane, and a keg1-­‐Ts mutant is suppressed at high copy by ROT1, though not BIG1; however, a physical interaction between Keg1 and Rot1 could not be detected (Nakamata et al. 2007). Because the big1Δ rot1Δ double mutant has the same growth rate as each single mutant, it was suggested that Rot1 and Big1 impact β1,6-­‐glucan synthesis in the same way, and possibly function in the same compartment or even in a complex (Machi et al. 2004). However, although rot1, big1, and kre5 mutations individually all lower β1,6-­‐glucan levels to the same extent, the kre5 big1 double mutant, but apparently not a kre5 rot1 strain (Lesage and Bussey, 2006), shows a reduced growth rate and lowered β1,6-­‐glucan content compared with each single mutant, suggesting the function of Rot1 is partly distinct from that of Kre5 (Azuma et al. 2002; Lesage and Bussey, 2006). Indeed, the non-­‐conditional rot1-­‐1 mutant shows a synthetic growth and N-­‐glycosylation defect in combination with ost3Δ (though not ost6Δ), as well as a partial defect in O-­‐mannosylation of the chitinase Cts1, indicating a wider role for Rot1 in glycosylation (Pasikowska et al. 2012). More widely distributed secretory pathway proteins: Kre6 and Skn1: P. Orlean 41 SI Localization and transport of Kre6. Recent studies indicate that much of Kre6 is ER-­‐localized, where it interacts with Keg1, but Kre6 is also detectable in secretory vesicles and at the plasma membrane at sites of polarized growth (Nakamata et al. 2007; Kurita et al. 2011). In addition to Kre6’s lumenal domain, the protein’s cytoplasmic tail is important for Kre6’s function in β1,6-­‐glucan assembly and its transport to the plasma membrane (Li et al. 2002; Kurita et al. 2011). A truncated form of Kre6 lacking its 230 N-­‐terminal amino acids failed to be localized to the plasma membrane, and did not correct the β1,6-­‐glucan synthetic defect of kre6Δ, although it appeared stable (Kurita et al. 2011). It was concluded that transport of Kre6 to the plasma membrane is necessary for the protein to fulfill its role in β1,6-­‐glucan synthesis (Kurita et al. 2002). Localization of Skn1 has not been explored in detail. Skn1 and plant defensin resistance. skn1Δ, but not kre6Δ strains, are defective in M(IP)2C synthesis and resistant to a plant defensin that interacts with this sphingolipid to exert its antifungal activity (Thevissen et al. 2005). Defensin-­‐susceptibility is unconnected with cellular β1,6-­‐glucan content because other β1,6-­‐glucan synthesis mutants are defensin-­‐sensitive (Thevissen et al. 2005). Plasma membrane protein Kre1: Kre1 as receptor for K1 killer toxin. Membrane anchored Kre1 has an additional role as receptor for K1 killer toxin. Spheroplasts of kre1Δ cells are resistant to this toxin, but expression of the C-­‐terminal 63 amino acids of Kre1 was sufficient to make spheroplasts, but not intact cells, toxin sensitive again, leading to the proposal that Kre1’s GPI-­‐modified C-­‐terminus serves as the membrane receptor for K1 toxin after initial toxin binding to β1,6-­‐glucan (Breinig et al. 2002). Literature Cited Breinig, F., Tipper D. J., Schmitt, M. J., 2002 Kre1p, the plasma membrane receptor for the yeast K1 viral toxin. Cell 108: 395-­‐
405. Pasikowska, M., Palamarczyk, G., Lehle, L. (2012) The essential endoplasmic reticulum chaperone Rot1 is required for protein N-­‐ and O-­‐glycosylation in yeast. Glycobiology 22: 939-­‐947. 42 SI P. Orlean Takeuchi, M., Kimata, Y., Kohno, K., 2008 Saccharomyces cerevisiae Rot1 is an essential molecular chaperone in the endoplasmic reticulum. Mol. Biol. Cell 19: 3514-­‐3525. P. Orlean 43 SI File S9 Cell Wall-­‐Active and Nonenzymatic Surface Proteins and Their Functions This Supporting File contains additional information and discussion related to Cell Wall-­‐Active and Nonenzymatic Surface Proteins and Their Functions. The subheadings used in the main text are retained, and new subheadings are underlined. Literature cited in this File but not In the main text is listed at the end of the File. Known and predicted enzymes Chitinases: S. cerevisiae’s two chitinases, Cts1 and Cts2, are both members of GH Family 18, but of the two, Cts1 resembles plant-­‐
type chitinases, whereas the predicted Cts2 protein is more similar to the bacterial chitinase subfamily (Hurtado-­‐Guerrero and van Aalten, 2007). Cts1 has endochitinase activity, a pH optimum of 2.5, and is more active on nascent than on preformed chitin (Correa et al. 1982). The structure of the catalytic domain, which has chitinase activity on its own, has been determined (Hurtado-­‐Guerrero and van Aalten, 2007). Little is known about Cts2, but because CTS2 complements a defect in the sporulation-­‐specific chitinase of Ashbya gossypii (Dünkler et al. 2008), Cts2 may have a role in sporulation. β 1,3-­‐glucanases: Exg1, Exg2 and Ssg/Spr1 exo-­‐β1,3-­‐glucanases: These proteins are members of GH Family 5 and were originally characterized biochemically as exo-­‐β1,3-­‐glucanases (Larriba et al. 1995). Exg1 is a soluble cell wall protein released upon treatment with dithiothreitol (Cappellaro et al. 1998), whereas Exg2 may normally be membrane-­‐ or wall-­‐anchored because it has a potential GPI attachment site (Caro et al. 1997), whose deletion results in release of the protein into the medium (Larriba et al. 1995). Single or double null mutants in EXG1 and EXG2 have no obvious defects, although exg1Δ cells have slightly elevated levels of β1,6 glucan and EXG1 overexpressers lower amounts of that polymer. This, together with the finding that the Exg proteins can act on the β1,6-­‐glucan pustulan in vitro (Nebreda et al. 1986), raises the possibility that Exg1 and Exg2 have roles in β-­‐glucan remodeling (Jiang et al. 1995; Lesage and Bussey, 2006). Ssg1/Spr1 is a sporulation-­‐specific protein. Its mRNA is expressed late in sporulation, and homozygous null diploids show a delay in the onset of ascus formation (Muthukumar et al. 1993; San Segundo et al. 1993). Bgl2, Scw4, Scw10 endo-­‐β1,3-­‐glucanases:
These proteins are members of GH Family 17. Scw4, Scw10, and Bgl2 can be extracted from the wall with dithiothreitol (Capellaro et al. 1998), suggesting wall association via disulfides. However, a population of Scw4 and Scw10 44 SI P. Orlean resists extraction by hot SDS and β-­‐mercaptoethanol, and is released instead by mild alkali or by β1,3-­‐glucanase digestion, indicating a covalent linkage to β1,3-­‐glucan (Yin et al. 2005). However, Scw4 and Scw10 lack PIR sequences. Purified Bgl2 binds both β1,3-­‐glucan and chitin (Klebl and Tanner, 1989), but whether these non-­‐covalent interactions represent an additional mode of wall association, or reflect an enzyme-­‐substrate interaction, is unexplored. Levels of Bgl2 and Scw10 need to be balanced in order to ensure cell wall stability (Sestak et al. 2004). This proposal is based on the findings that deletion of BGL2 in the scw4Δ scw10Δ background (but not of SCW11, EXG1, CRH1, or CRH2) alleviated many of the phenotypes of that double mutant, that overexpression of BGL2 is lethal in a wild type background, and that high level expression of SCW10 in bgl2Δ significantly increases the strain’s CFW sensitivity (Klebl and Tanner, 1989; Sestak et al. 2004). Bgl2 and Scw10 may also contribute to compensatory responses to mutationally induced wall stress, because BGL2 and SCW10, as well as EXT1 and CRH1, are upregulated in mnn9, kre6, mnn9, and gas1 mutants (Lagorce et al. 2003). What Bgl2 and Scw10’s precise biochemical roles are, and how they antagonize one another, are intriguing questions. Eng1/Dse4 and Eng2/Acf2 endo-­‐β1,3-­‐glucanases: These two related proteins are members of GH family 81. ENG1 expression is highest at the M to G1-­‐phase transition and shut down during sporulation. Eng1 localizes to the daughter side of the septum, consistent with a hydrolytic role during cell separation (see Septum formation; Baladron et al. 2002). Eng2 recognizes β1,3-­‐glucans of at least five residues and releases trisaccharides from the non-­‐reducing end of the substrate, but has no detectable transglycosidase activity (Martín-­‐Cuadrado et al. 2008). Gas1 family β 1,3-­‐glucanosyltransferases: Domain organization and mechanism of Gas proteins. Gas1 and its four paralogues, Gas2, Gas3, Gas, 4, and Gas5 (Popolo and Vai, 1999), are members of the GH Family 72. The catalytic domain of Gas proteins lies in their N-­‐terminal half, and in the case of Gas1 and Gas2, is followed by a cysteine-­‐rich domain that is a member of the CBM43 group of carbohydrate binding modules. The other Gas proteins lack this module but have a serine and threonine-­‐rich sequence instead, and Gas1 has both (Popolo and Vai, 1999). The biochemical activity of Gas proteins was first defined for the Aspergillus fumigatus Gas1 homologue, Gel1, but S. cerevisiae Gas1, Gas2, Gas4, and Gas5 all proved to carry out the same reaction in vitro (Mouyna et al. 2000; Carotti et al. 2004; Ragni et al. 2007b; Mazan et al. 2011). The proteins have β1,3-­‐glucanosyltransfer or “elongase” activity, which involves cleavage of a β1,3 glucosidic linkage within a β1,3-­‐glucan chain, then transfer of the newly generated reducing end of the P. Orlean 45 SI cleaved glycan to the non-­‐reducing end of another β1,3 glucan molecule, thus extending the acceptor β1,3-­‐glucan chain (Mouyna et al. 2000). The structure of a soluble form of Gas2 in complex with β1,3-­‐gluco-­‐oligosaccharides revealed the presence of two oligosaccharide binding sites and led to a base-­‐occlusion hypothesis for how transglycosylation could be favored over hydrolysis. In the hypothesized mechanism, one binding site is occupied by the donor glucan, which is hydrolyzed with formation of an enzyme-­‐oligosaccharide intermediate, whereupon the other, acceptor, site is transiently filled by the second product of the hydrolysis reaction. Occupancy of the acceptor site has the effect of occluding the catalytic base on the enzyme, preventing any incoming water molecule from being activated for nucleophilic attack on the enzyme-­‐saccharide intermediate. The gluco-­‐oligosaccharide in the acceptor site is then displaced by a longer and tighter binding acceptor glucan with concomitant formation of the new β1,3-­‐glucosidic linkage (Hurtado-­‐Guerrero et al. 2009). In the case of Gas1 and Gas2, the cysteine-­‐rich domain is necessary for catalytic activity, being required for proper folding of the catalytic domain, for substrate binding, or for both (Popolo et al. 2008). This domain, however, is not necessary for activity of Gas4 or Gas5, which lack it, and, because Gas4 and Gas5 generate profiles of oligosaccharides from β1,3-­‐gluco-­‐
oligosaccharide substrates that are different from those released by Gas1 and Gas2, it is possible that the cysteine-­‐rich domain influences cleavage site preference (Ragni et al. 2007b). Nonetheless, expression of Gas4, but not Gas2, in a gas1Δ strain fully complemented the gas1Δ growth defect in media with a pH of 6.5 or above (Ragni et al. 2007a).
Localization of Gas1. Gas1 fused to GFP but retaining its N-­‐ and C-­‐terminal signal sequences is detectable in the lateral wall, in the chitin ring in small-­‐budded cells, and near the primary septum, and remains in the bud scar after cell separation (Rolli et al. 2009). Gas1 localization to the chitin ring and bud scars was abolished in cells lacking the chitin-­‐β1,3-­‐glucan cross-­‐
linkers Crh1 and Crh2, suggesting that Gas1 anchorage to chitin was dependent on linkage of a Gas1-­‐β1,6-­‐glucan-­‐β1,3-­‐glucan complex to chitin (Rolli et al. 2009). Consistent with this, Gas1 was shed into the medium from chs3Δ cells, which are unable to make the chitin known to be cross-­‐linked to β-­‐glucan (Cabib and Duran, 2005). Because the released Gas1 was not significantly larger than Gas1 in lysates of wild type cells (Rolli et al., 2009), the β1,6-­‐glucan-­‐β1,3-­‐glucan presumed to link the protein to chitin must be quite small. Some Gas1 was also released from chs2Δ cells, suggesting that localization of Gas1 near the primary septum requires Chs2-­‐dependent chitin synthesis (Rolli et al. 2009). However, because the chitin made by Chs2 is free of cross-­‐
links (Cabib and Duran, 2005), its association with Gas1 would be indirect. Cell-­‐associated Gas1 was distributed throughout the remedial septum made in chs2Δ cells (Section V.1.a). Intriguingly, Gas1 was also shed from chs1Δ cells, though at reduced levels when the medium was buffered to lower chitinase activity. Amounts and localization of cell-­‐associated Gas1 appeared 46 SI P. Orlean unchanged, however, presumably because Chs2 and Chs3 still make chitin. Nonetheless, this observation indicates that Chs1 or its product contribute to wall association of some Gas1 (Rolli et al. 2009). Functions of Gas2, Gas3, Gas4, and Gas5. The following findings indicate that Gas5 and Gas3 have wall-­‐related functions in vegetative cells. GAS5 is expressed during vegetative growth but repressed during sporulation, and gas5Δ strains are Calcofluor White sensitive (Caro et al. 1997). Purified Gas3 is inactive (Ragni et al. 2007b), and gas3Δ strains make no genetic interactions with strains with single or double deletions in other GAS genes (Rolli et al. 2010). Moreover, Gas3 cannot substitute for Gas1, but overexpression in gas1Δ of wild type GAS3 or a gas3 mutant encoding catalytically inactive Gas3 exacerbated the gas1Δ growth defect, indicating that high levels of Gas3 are toxic (Rolli et al. 2010). Gas2 and Gas4 have overlapping functions in ascospore wall assembly. Their genes are expressed only during sporulation, and although diploids homozygous for single GAS2 or GAS4 deletions sporulate normally, diploids lacking both Gas2 and Gas4 have a severe sporulation defect (Ragni et al. 2007a). The inner glucan layer of the spore wall from by double homozygous gas2 gas4 nulls was disorganized and detached from chitosan, and dityrosine, though present, was less abundant and diffusely distributed. The absence of β1,3-­‐glucanosyltransferase activity may result in shorter β1,3-­‐glucan chains that are more loosely associated with chitosan. Gas2 and Gas4 likely need to be GPI anchored to fulfill their key roles in ascospore wall formation, which in part explains the severe sporulation defect of homozygous gpi1/gpi1 and gpi2/gpi2 diploids (Leidich and Orlean, 1996). Because such diploids lack dityrosine, additional GPI-­‐proteins must normally be involved in ascospore wall assembly. Yapsin aspartyl proteases: Yapsin processing. Yapsins are synthesized as zymogens and undergo proteolytic processing to generate a mature active enzyme. The steps include removal of a propeptide and excision of an internal segment flanked by basic amino acids that separates the enzyme’s two catalytic domains, which remain disulfide-­‐linked (Gagnon-­‐Arsenault et al. 2006, 2008). In the case of Yps1, the propeptide removal and excision steps are likely autocatalytic at an environmental pH of 3, but involve other proteases, including yapsins, at pH 6 (Gagnon-­‐Arsenault et al. 2008). Cell wall phenotypes of yapsin-­‐deficient strains. Strains lacking individual yapsin genes are sensitive to various cell wall disrupting agents, though their sensitivity profiles differ. For example, yps7Δ is the only yps null hypersensitive to CFW, but yps1Δ the only mutant sensitive to the β1,3-­‐glucan synthase inhibitor caspofungin (Krysan et al. 2005). The quintuple yps1Δ yps2Δ yps3Δ yps6Δ yps7Δ null mutant is viable, but undergoes osmotically remedial lysis at 30°C, as does the yps1Δ yps2Δ P. Orlean 47 SI yps3Δ triple deletion strain, and to a slightly lesser extent, the yps1Δ yps2Δ double null (Krysan et al. 2005). The temperature-­‐
sensitive lysis phenotype of strains lacking multiple yapsins is consistent with a role for these proteins when cell walls are stressed, and indeed, expression of YPS1, YPS2, YPS3, and YPS6 is upregulated under such conditions (Garcia et al. 2004; Krysan et al. 2005). Non-­‐enzymatic CWPs Structural GPI proteins: Sps2 family: Ecm33. Mannan outer chains produced by ecm33Δ cells are slightly smaller than normal, although O-­‐mannosylation and core-­‐type N-­‐glycans are not affected. Epitope-­‐tagged Pst1 is most abundant at the surface of buds, but Ecm33’s localization is uncertain because tagging Ecm33 abolishes its in vivo function (Pardo et al. 2004). Ecm33 occurs in both plasma membrane and wall-­‐anchored forms, but must retain its GPI anchor and plasma membrane localization for in vivo function (see Incorporation of GPI proteins into the wall; Terashima et al. 2003; Yin et al. 2005). Expression of a minimal amount of GPI-­‐
anchored Ecm33 may be necessary for growth at high temperature, because the temperature-­‐sensitivity of mcd4, gpi7, gpi13 and gpi14 mutants is suppressed by overexpression of ECM33 (Toh-­‐e & Oguchi, 2002; A. Sembrano and P. Orlean, unpublished). Tip1 family: Localization of Cwp2 and Tip1 is influenced by the timing of their expression. A swap of the promoters of CWP2 and TIP1 caused these genes’ products to exchange their cellular location, indicating that the localization of Cwp2 and Tip1, and perhaps that of other CWPs, is influenced by the timing of their expression in the cell cycle (Smits et al. 2006). Cwp1, however, is localized to the birth scar in a manner that depends on normal septum formation, but, because neither Tip1 nor Cwp2 is targeted to the birth scar when expressed behind CWP1‘s promoter, additional CWP1 sequences are required for Cwp1 localization (Smits et al. 2006). Ccw12: Structural features of Ccw12. Ccw12 has a predicted mass of 13 kDa but migrates on denaturing polyacrylamide gels with an apparent molecular weight of a least 200 kDa. Elimination of Ccw12’s three N-­‐linked sites shows that N-­‐linked glycans are mostly responsible for this apparent size increase, but these modifications are not necessary for in vivo function, because Ccw12 lacking its N-­‐linked sites complements ccw12Δ phenotypes (Ragni et al. 2007c). O-­‐mannosylation contributes some 42 kDa to the apparent size of Ccw12 (Hagen et al. 2004). The protein is not obviously related to any known enzymes, but contains 48 SI P. Orlean two repeats of the sequence TTEAPKNGTSTAAP (Mrša et al. 1999). Deletion of one or both of these does not affect cross-­‐
linkage Ccw12 to the wall, but the repeats are nonetheless critical for in vivo function because proteins lacking them do not restore the growth and cell wall defects of ccw12Δ (Ragni et al. 2007c). Four sequences similar to the Ccw12 repeat are present in Sed1 (Mrša et al. 1999; Ragni et al. 2007c). Certain Tip1 family members and Slr1 also migrate in denaturing polyacrylamide gels with much higher molecular weights than would be expected (van der Vaart et al. 1995; Terashima et al. 2002). A new mechanism for compensating loss of multiple GPI-­‐CWP uncovered in ccw12Δ . Deletion of additional genes for GPI-­‐CWP in the ccw12Δ background uncovered a mechanism for compensating for loss of multiple GPI-­‐CWPs. Rather than showing an exacerbated phenotype, the ccw12Δ ccw14Δ double null was less sensitive to CFW compared with ccw12Δ, and the ccw12Δ ccw14Δ dan1Δ mutant showed wild type levels of sensitivity to CFW and nearly normal levels of chitin. Moreover, additional deletion of CWP1 and TIP1 had no further effect on CFW sensitivity, although walls of the quintuple mutant had a thicker inner glucan layer and a thinner but more ragged outer mannoprotein layer (Hagen et al. 2004). It seems that although loss of Ccw12 alone activates the CWI pathway-­‐mediated chitin stress response (Ragni et al. 2007c, 2011; see Chitin synthesis in response to cell wall stress), deletion of additional GPI-­‐CWP genes forces cells over a threshold that leads to triggering of a new compensatory response, whereupon the chitin response becomes less important. This new response depends on Sed1 and the non-­‐GPI-­‐CWP Srl1. Not only is their expression upregulated in the ccw12Δ ccw14Δ dan1Δ cwp1Δ tip1Δ strain, but deletion of either in the ccw12Δ ccw14Δ dan1Δ background reverts the strain to the high-­‐chitin phenotype of ccw12Δ (Hagen et al. 2004). In addition, the cell wall remodeling genes SCW10 and BGL2 are upregulated and CRH2 downregulated, suggesting that the response involves alterations of the structure of the β-­‐glucan layer (Hagen et al. 2004). More generally, the phenotypes of the multiple GPI-­‐CWP mutants indicate that GPI-­‐CWPs have a collective role in maintaining cell wall stability (Lesage and Bussey, 2006; Ragni et al. 2007c). Ccw12 and Slr1 also have parallel functions in a pathway that relieves defects in a polarized morphogenesis signaling network (see Slr1). Other non-­‐enzymatic GPI-­‐proteins: Ccw14/Ssr1/Icwp as an inner cell wall protein. A monoclonal antibody that recognizes Ccw14/Ssr1 on immunoblots does not detect the protein on intact cells, whereas it does have access to the glycoprotein in tunicamycin-­‐treated cells or in mnn1 mnn9 mutants (Moukadiri et al. 1997). Assuming that the antibody would have had access to its epitope on Ccw14/Ssr1 if the protein were at the surface of wild type cells, this finding is consistent with Ccw14/Ssr1 being a protein of the inner cell wall P. Orlean 49 SI (Moukadiri et al. 1997). Flocculins and agglutinins: Roles and interactions of Aga1 and Fig2 in mating. Deletion of FIG2 in MATa cells with the W303 background, but not MATa cells, increases the agglutinability of MATα cells, suggesting a role for Fig2 in attenuating agglutination of MATa cells (Erdman et al. 1998; Jue and Lipke, 2002). Both Aga1 and Fig2 have an additional, additive role in mating in MATα strains that is unconnected with Aga2, because simultaneous deletion of AGA1 and FIG2 in certain MATα sag1Δ backgrounds leads to a severe mating defect on solid medium, whereas individually deleting the AGA1 and FIG2 in those strain backgrounds does not (Guo et al. 2000). An explanation for the expanded roles for Aga1 and Fig2 in mating came from detection of heterotypic adhesive interactions between Aga1 and Fig2, and homotypic interactions between Fig2 and Fig2, which are mediated by WPCL and CX4C domains present in both proteins (Huang et al. 2009). Non-­‐GPI-­‐CWP: PIR proteins: PIR protein localization. Fusions of Pir1 and Pir2 with red fluorescent protein are found at bud scars of both haploid and diploid cells, with Pir1 being localized inside the chitin ring. This localization of Pir1 is independent of normal chitin ring and primary septum formation because the protein is still transported to the budding site in chs2Δ and chs3Δ cells, although in the absence of the chitin ring in chs3Δ, Pir1 no longer shows a ring-­‐like distribution (Sumita et al. 2005). Some Pir1 and Pir2, and most Pir3, are also present in lateral walls, where these proteins can be detected by immunoelectron microscopy using antibody to Pir3 (Yun et al. 1997). Pir4 has been reported to show a uniform distribution at the cell surface, but in one study, this distribution was restricted to growing buds (Moukadiri et al. 1999; Sumita et al. 2005). A Kex2 processing site in PIR proteins. The four PIR proteins contain a site for processing by the Kex2 protease, but although Kex2 acts on the PIR proteins in vivo, wall localization of these proteins is unaffected in kex2Δ, so the significance of this processing event is unclear (Mrša et al. 1997). Scw3 (Sun4): SUN proteins. Members of this family of highly glycosylated proteins have a common C-­‐terminal domain of some 250 amino acids in which the spacing of four cysteines is conserved (Velours et al. 2002). The SUN proteins other than Scw3/Sun4 (Sim1, Uth1, and Nca3) have been implicated in various cellular functions unrelated to the cell wall, but SUN family members have been assumed to be glucanases because they are homologous to Candida wickerhamii BglA, an additional protein 50 SI P. Orlean identified in a screen of a cDNA expression library for proteins that reacted with an antibody to a cell-­‐bound β-­‐glucosidase (Skory and Freer, 1995). However, glycosidase activity has not been verified for BglA and the SUN proteins show no homology to any carbohydrate active enzymes, making it doubtful they are glycosidases. Literature Cited Garcia, R., Bermejo, C., Grau, C., Perez, R., Rodriguez-­‐Pena, et al., 2004 The global transcriptional response to transient cell wall damage in Saccharomyces cerevisiae and its regulation by the cell integrity signaling pathway. J. Biol. Chem. 279: 15183-­‐15195. Huang, G., Dougherty, S. D., Erdman, S. E., 2009 Conserved WCPL and CX4C domains mediate several mating adhesin interactions in Saccharomyces cerevisiae. Genetics 182: 173-­‐189. Hurtado-­‐Guerrero, R., Schüttelkopf, A. W., Mouyna, I., Ibrahim, A. F. M., Shepherd, S., et al., 2009 Molecular mechanisms of yeast cell wall glucan remodeling. J. Biol. Chem. 284: 8461-­‐8469. Hurtado-­‐Guerrero, R., van Aalten, D. M., 2007 Structure of Saccharomyces cerevisiae chitinase 1 and screening-­‐based discovery of potent inhibitors. Chem. Biol. 14: 589-­‐599. Martín-­‐Cuadrado, A. B., Fontaine, T., Esteban, P. F., del Dedo, J. E., de Medina-­‐Redondo, M., et al., 2008 Characterization of the endo-­‐β-­‐1,3-­‐glucanase activity of S. cerevisiae Eng2 and other members of the GH81 family. Fungal Genet. Biol. 45: 542-­‐553. Muthukumar, G., Suhng, S. H., Magee, P. T., Jewell, R. D., Primerano, D. A., 1993 The Saccharomyces cerevisiae SPR1 gene encodes a sporulation-­‐specific exo-­‐1,3-­‐β-­‐glucanase which contributes to ascospore thermoresistance. J. Bacteriol. 175: 386-­‐
394. Nebreda, A. R., Villa, T. G., Villanueva, J. R., del Rey, F., 1986 Cloning of genes related to exo-­‐β-­‐glucanase production in Saccharomyces cerevisiae: characterization of an exo-­‐β-­‐glucanase structural gene. Gene 47: 245-­‐529. P. Orlean 51 SI Popolo, L., Ragni, E., Carotti, C., Palomares, O., Aardema, R., et al., 2008 Disulfide bond structure and domain organization of yeast β(1,3)-­‐glucanosyltransferases involved in cell wall biogenesis. J. Biol. Chem. 283: 18553-­‐18565. Rolli, E., Ragni, E., Rodriguez-­‐Peña, J. M., Arroyo, J., Popolo, L., 2010 GAS3, a developmentally regulated gene, encodes a highly mannosylated and inactive protein of the Gas family of Saccharomyces cerevisiae. Yeast 27: 597-­‐610. San Segundo, P., Correa, J., Vazquez de Aldana, C. R., del Rey, F., 1993 SSG1, a gene encoding a sporulation-­‐specific 1,3-­‐β-­‐
glucanase in Saccharomyces cerevisiae. J. Bacteriol. 175: 3823-­‐3837. Skory, C. D., Freer, S. N., 1995 Cloning and characterization of a gene encoding a cell-­‐bound, extracellular β-­‐glucosidase in the yeast Candida wickerhamii. Appl. Environ. Microbiol. 61: 518-­‐525. 52 SI P. Orlean Table S1 Proteins involved in cell wall biogenesis in Saccharomyces cerevisiae Process or protein type Protein name Activity or Function Ugp1 UDPGlc pyrophosphorylase Pmi40 phosphomannose isomerase Sec53 phosphomannomutase 1
CAZy Family Precursor supply Psa1/Srb1/Vig9 GDP-­‐Man pyrophosphorylase Gfa1 glutamine: Fru-­‐6-­‐P amidotransferase Gna1 GlcN-­‐6-­‐P N-­‐acetylase Agm1/Pcm1 GlcNAc phosphate mutase Uap1/Qri1 UDPGlcNAc pyrophosphorylase Rer2 cis-­‐prenyltransferase (Dol10-­‐14) Srt1 cis-­‐prenyltransferase (Dol19-­‐22) Dfg10 dehydrodolichol reductase Sec59 Dol-­‐kinase Cwh8/Cax4 Dolichyl pyrophosphate phosphatase Dpm1 GDP-­‐mannose:dolichyl-­‐phosphate Man-­‐T GT2 Alg5 UDP-­‐glucose:dolichyl-­‐phosphate Glc-­‐T GT2 Yea4 UDP-­‐GlcNAc transporter GT1 Vrg4/Vig4 GDP-­‐Man transporter Gda1 GDPase Ynd1 Apyrase Alg7 UDP-­‐GlcNAc: Dol-­‐P GlcNAc-­‐1-­‐P-­‐T N-­‐glycosylation Alg13 + Alg14 UDP-­‐GlcNAc: Dol-­‐PP-­‐GlcNAc β1,4-­‐GlcNAc-­‐T P. Orlean 53 SI 54 SI Alg1 GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2 β1,4-­‐Man-­‐T Alg2 Alg11 GT33 GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2Man α1,3-­‐Man-­‐T and GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2Man2 α1,6-­‐Man-­‐T GT4 GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2Man3 α1,2-­‐Man-­‐T and GDP-­‐Man: Dol-­‐PP-­‐GlcNAc2Man4 α1,2-­‐Man-­‐T GT4 Rft1 Candidate Dol-­‐PP-­‐oligosaccharide flippase Alg3 Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man5 α1,3-­‐Man-­‐T GT58 Alg9 Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man6 α1,2-­‐Man-­‐T and Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man8 α1,2-­‐Man-­‐T GT22 Alg12 Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man7 α1,6-­‐Man-­‐T GT22 Alg6 Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man9 α1,3-­‐Glc-­‐T GT57 Alg8 Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man9Glc α1,3-­‐Glc-­‐T GT57 Alg10 Dol-­‐P-­‐Man: Dol-­‐PP-­‐GlcNAc2Man9Glc2 α1,2-­‐Glc-­‐T GT59 Stt3 OST catalytic subunit GT66 Wbp1 OST subunit Swp1 OST subunit Ost1 OST subunit Ost2 OST subunit Ost3 OST subunit; cysteine oxidoreductase Ost6 OST subunit; cysteine oxidoreductase Gls1/Cwh41 ER glucosidase I (α1,2 exoglucosidase); indirectly affects β1,6-­‐glucan GH63 Gls2/Rot2 ER glucosidase II (α1,3 exoglucosidase α-­‐subunit); indirectly affects β1,6-­‐glucan GH31 Gtb1 ER glucosidase II (regulatory subunit) Mns1 ER α-­‐mannosidase I GH47 GH47 Htm1/Mnl1 ER-­‐degradation enhancing a-­‐mannosidase-­‐like protein Yos9 Lectin, recognizes α1,6-­‐Man on glucosidase II product, targets misfolded protein for ERAD Png1 Cytosolic peptide N-­‐glycanase Och1 Initiating α1,6-­‐Man-­‐T GT32 Mnn9 M-­‐Pol I α1,6-­‐Man-­‐T GT62 P. Orlean Van1 M-­‐Pol I α1,6-­‐Man-­‐T GT62 Mnn9 M-­‐Pol II α1,6-­‐Man-­‐T GT62 Anp1 M-­‐Pol II α1,6-­‐Man-­‐T GT62 Mnn10 M-­‐Pol II α1,6-­‐Man-­‐T GT34 Mnn11 M-­‐Pol II α1,6-­‐Man-­‐T GT34 Hoc1 M-­‐Pol II α1,6-­‐Man-­‐T GT32 Mnn2 α1,2-­‐Man-­‐T; Mnn1 subfamily; major role in mannan side chain branching GT71 Mnn5 α1,2-­‐Man-­‐T; Mnn1 subfamily; major role in mannan side chain branching GT71 Mnn4 Positive regulator of Man phosphorylation Mnn6/Ktr6 α-­‐Man-­‐P-­‐T; acts on N-­‐ and O-­‐glycans in Golgi GT15 Mnn1 α1,3-­‐Man-­‐T; acts on N-­‐ and O-­‐glycans in Golgi GT71 Kre2/Mnt1 α1,2-­‐Man-­‐T; acts on N-­‐ and O-­‐glycans in Golgi GT15 Ktr1 α1,2-­‐Man-­‐T; acts on N-­‐ and O-­‐glycans in Golgi GT15 Ktr2 α1,2-­‐Man-­‐T; acts on N-­‐glycans in Golgi GT15 Ktr3 α1,2-­‐Man-­‐T; acts on N-­‐ and O-­‐glycans in Golgi GT15 Yur1 α1,2-­‐Man-­‐T; acts on N-­‐glycans in Golgi GT15 Ktr4 Putative α-­‐ManT GT15 Ktr5 Putative α-­‐ManT GT15 Ktr7 Putative α-­‐ManT GT15 Gnt1 GlcNAc-­‐T GT8 Vrg4 GDP-­‐Man transporter Gda1 GDPase Ynd1 Apyrase Pmt1 Dol-­‐P-­‐Man: protein: O-­‐Man-­‐T; Pmt1 family GT39 Pmt2 Dol-­‐P-­‐Man: protein: O-­‐Man-­‐T; Pmt2 family GT39 O-­‐mannosylation P. Orlean 55 SI Pmt3 Dol-­‐P-­‐Man: protein: O-­‐Man-­‐T; Pmt2 family Pmt4 Pmt5 GT39 Dol-­‐P-­‐Man: protein: O-­‐Man-­‐T; specific for membrane proteins GT39 Dol-­‐P-­‐Man: protein: O-­‐Man-­‐T; Pmt1 family GT39 Pmt6 Dol-­‐P-­‐Man: protein: O-­‐Man-­‐T; Pmt2 family GT39 Mnt2 α1,3-­‐Man-­‐T; Mnn1 subfamily; acts on O-­‐glycans in Golgi GT71 Mnt3 α1,3-­‐Man-­‐T; Mnn1 subfamily; acts on O-­‐glycans in Golgi GT71
Gpi1 GPI-­‐Gnt subunit Gpi2 GPI-­‐Gnt subunit Gpi3 GPI-­‐Gnt subunit, UDP-­‐GlcNAc: Ptd-­‐Ins α1,6-­‐GlcNAc transferase GT4 Gpi15 GPI-­‐Gnt subunit Gpi19 GPI-­‐Gnt subunit Eri1 GPI-­‐Gnt subunit Ras2 Negative regulator of GPI-­‐Gnt Gpi12 GPI-­‐Ins-­‐deacetylase Gwt1 GPI-­‐Ins-­‐acyltransferase Gpi14 GPI-­‐ManT-­‐I: Dol-­‐P-­‐Man: GlcN-­‐Ptd-­‐(acyl)Ins α1,4-­‐Man-­‐T GT50 Pbn1 Putative subunit of GPI-­‐Man-­‐T-­‐I Arv1 Proposed to present GlcN-­‐(acyl)PI to Gpi14 Mcd4 GPI-­‐Etn-­‐P-­‐T-­‐I Gpi18 GPI-­‐ManT-­‐II: Dol-­‐P-­‐Man: Man-­‐GlcN-­‐Ptd-­‐(acyl)Ins α1,6-­‐Man-­‐T GT76 Pga1 GPI-­‐ManT-­‐II subunit Gpi10 GPI-­‐Man-­‐T-­‐III: Dol-­‐P-­‐Man: Man2-­‐GlcN-­‐Ptd-­‐(acyl)Ins α1,2-­‐Man-­‐T GT22 Smp3 GPI-­‐Man-­‐T-­‐IV: Dol-­‐P-­‐Man: Man3-­‐GlcN-­‐Ptd-­‐(acyl)Ins α1,2-­‐Man-­‐T GT22 Gpi13 GPI-­‐Etn-­‐P-­‐T-­‐III Gpi11 Subunit of GPI-­‐Etn-­‐P-­‐T-­‐II and GPI-­‐Etn-­‐P-­‐T-­‐III GPI anchoring 56 SI P. Orlean Gpi7 GPI-­‐Etn-­‐P-­‐T-­‐II Gpi8 GPI transamidase catalytic subunit Gaa1 GPI transamidase subunit Gab1 GPI transamidase subunit Gpi16 GPI transamidase subunit Gpi17 GPI transamidase subunit Bst1 GlcN-­‐(acyl)PI inositol deacylase Per1 Removes acyl chain at sn-­‐2 position of protein-­‐bound GPIs Gup1 MBOAT O-­‐acyltransferase, transfers C26 acyl chain to sn-­‐2 position of protein-­‐bound GPIs Cwh43 Replaces GPI diacylglycerol with ceramide Cdc1 Homologue of mammalian PGAP5; possible GPI-­‐Etn-­‐P phosphodiesterase Ted1 Homologue of mammalian PGAP5; possible GPI-­‐Etn-­‐P phosphodiesterase Chitin and chitosan synthesis Chs1 Chitin synthase I catalytic protein GT2 Chs2 Chitin synthase II catalytic protein GT2 Chs3 Chitin synthase catalytic subunit GT2 Cdk1 Mitotic protein kinase, phosphorylates Chs2 Cdc14 Phosphoprotein phosphatase, dephosphorylates Chs2 Dbf2 Mitotic exit kinase, phosphorylates Chs2 Inn1 Localized to mother cell-­‐bud junction with Chs2 and Cyk3, implicated in Chs2 activation Cyk3 Localized to mother cell-­‐bud junction with Chs2 and Inn1, implicated in Chs2 activation Pfa4 Protein acyltransferase, palmitoylates Chs3 Chs7 Chaperone required for ER exit of Chs3 Rcr1 ER protein, small negatve effect on Chs3-­‐dependent chitin synthesis Yea4 ER protein and UDP-­‐GlcNAc transporter, yea4Δ has 65% of wild type levels of chitin. Chs5 Exomer component, involved in Chs3 trafficking P. Orlean 57 SI Chs6 Exomer component, involved in Chs3 trafficking Chs4/Skt5 Prenylated protein that interacts with, activates, and anchors Chs3 to septin ring Bni4 Scaffold protein, tethers Chs3 and Chs4 to septins Shc1 Sporulation-­‐specific Chs4 homologue Cda1 Chitin de-­‐N-­‐acetylase Cda2 Chitin de-­‐N-­‐acetylase β-­‐1,3 glucan synthesis Fks1/Gsc1/Cwh53/ Etg1/Pbr1 Probable β1,3-­‐glucan synthase, major role in vegetative cells GT48 Fks2/Gsc2 Probable β1,3-­‐glucan synthase, stress-­‐induced, role in sporulation GT48 Fks3 Probable β1,3-­‐glucan synthase, role in sporulation GT48 Rho1 GTPase; activator of Fks1 and Fks2 Kre5 Diverged UDP-­‐Glc: glycoprotein Glc-­‐T homologue GT24 Rot1 Fungus-­‐specific ER chaperone Big1 Fungus-­‐specific ER chaperone Keg1 Fungus-­‐specific ER chaperone Kre6 Resembles β-­‐1,6/β-­‐1,3 glucanases GH16 Skn1 Sequence and functional Kre6 homologue; additional role in MIPC synthesis GH16 Kre9 Fungus-­‐specific O-­‐mannosylated protein Knh1 Kre9 homologue Kre1 GPI-­‐protein, secondary receptor for K1 killer toxin β-­‐1,6 glucan formation Glycosidases, cross-­‐linking enzymes, and proteases Cts1 Endo-­‐chitinase GH18 Cts2 Chitinase GH18 GH5 Exg1/Bgl1 58 SI Major exo-­‐β-­‐1,3-­‐glucanase of the cell wall; soluble P. Orlean Exg2 GPI-­‐anchored plasma membrane exo-­‐β1,3-­‐glucanase GH5 Ssg1/Spr1 Sporulation-­‐specific exo-­‐β-­‐1,3-­‐glucanase GH5 Bgl2 Endo-­‐β1,3-­‐glucanase; can make β1,6-­‐linked Glc side branch GH17 Scw4 Endo-­‐β1,3-­‐endoglucanase-­‐like GH17 Scw10 Endo-­‐β1,3-­‐endoglucanase-­‐like GH17 Scw11 Endo-­‐β1,3-­‐endoglucanase-­‐like GH17 Eng1/Dse4 Endo-­‐β1,3-­‐endoglucanase GH81 Eng2/Acf2 Endo-­‐β1,3-­‐endoglucanase GH81 Dcw1 GPI-­‐protein, resembles α1,6-­‐endomannanase GH76 Dfg5 GPI-­‐protein, resembles α1,6-­‐endomannanase; Dcw1 homologue GH76 Crh1 GPI-­‐protein, chitin β-­‐1,6/1,3-­‐glucanosyltransferase GH16 Crh2/Utr2 GPI-­‐protein, chitin β-­‐1,6/1,3-­‐glucanosyltransferase GH16 Crr1 GPI-­‐protein, chitin β-­‐1,6/1,3-­‐glucanosyltransferase; sporulation-­‐specific GH16 Gas1 GPI-­‐protein, β-­‐1,3-­‐glucanosyltransferase GH72 Gas2 GPI-­‐protein, β-­‐1,3-­‐glucanosyltransferase; sporulation specific GH72 Gas3 GPI-­‐protein, β-­‐1,3-­‐glucanosyltransferase GH72 Gas4 GPI-­‐protein, β-­‐1,3-­‐glucanosyltransferase; sporulation specific GH72 Gas5 GPI-­‐protein, β-­‐1,3-­‐glucanosyltransferase GH72 Yps1 GPI-­‐protein, yapsin aspartyl protease Yps2/Mkc7 GPI-­‐protein, yapsin aspartyl protease Yps3 GPI-­‐protein, yapsin aspartyl protease Yps6 GPI-­‐protein, yapsin aspartyl protease Ecm33 Sps2 family; structural/non-­‐enzymatic Pst1 Sps2 family; structural/non-­‐enzymatic Sps2 Sps2 family; structural/non-­‐enzymatic; required for ascospore wall formation GPI-­‐CWP P. Orlean 59 SI 60 SI Sps22 Sps2 family; structural/non-­‐enzymatic; required for ascospore wall formation Cwp1 Tip1 family Cwp2 Tip1 family Tip1 Tip1 family; anaerobically induced Tir1 Tip1 family; anaerobically induced Tir2 Tip1 family; anaerobically induced Tir3 Tip1 family; anaerobically induced Tir4 Tip1 family; anaerobically induced Dan1/Ccw13 Tip1 family; anaerobically induced Dan4 Tip1 family; anaerobically induced Sed1 Induced in stationary phase Spi1 Induced by stress with weak organic acids; related to Sed1 Ccw12 Major role in stabilizing walls of daughter cells walls and mating projections Ccw14/Ssr1 Inner cell wall protein Dse2 Daughter cell specific, role in cell separation Egt2 Daughter cell specific, role in cell separation Fit1 Iron binding Fit2 Iron binding Fit3 Iron binding Flo1 Flocculin Flo5 Flocculin Flo9 Flocculin Flo10 Flocculin Flo11/Muc1 Required for pseudohypha formation by diploids and agar invasion by haploids Aga1 MATa agglutinin subunit, disulfide-­‐linked to Aga2, which binds MATα agglutinin Sag1 Fig2 Aga1-­‐related adhesin P. Orlean Sag1 MATα agglutinin Non-­‐GPI-­‐CWP Pir1/Ccw6 “Protein with internal repeat”, ester-­‐linked via Glu (originally Gln in repeats) to β1,3-­‐glucan Pir2/Hsp150/Ccw7 “Protein with internal repeat”, ester-­‐linked via Glu (originally Gln in repeats) to β1,3-­‐glucan Pir3/Ccw8 “Protein with internal repeat”, ester-­‐linked via Glu (originally Gln in repeats) to β1,3-­‐glucan Pir4/Cis3/ Ccw5/Ccw11 One “internal repeat” sequence”, ester-­‐linked via Glu (originally Gln in repeats) to β1,3-­‐glucan Scw3/Sun4 Member of SUN family Srl1 Acts in parallel with Ccw12 in pathway operative when regulation of Ace2 and polarized morphogenesis are defective 1
CAZy glycosyltransferase (GT) and glycosylhydrolase (GH) families are defined in the Carbohydrate Active Enzymes database (http://www.cazy.org/) (Cantarel, B. L., Coutinho, P. M., Rancurel, C., Bernard, T., Lombard, V., et al., 2009 The Carbohydrate-­‐Active EnZymes database (CAZy): an expert resource for Glycogenomics. Nucleic Acids Res. 37: D233-­‐
238). P. Orlean 61 SI