YEASTBOOK

CELL SIGNALING & DEVELOPMENT

Architecture and Biosynthesis of the Saccharomycescerevisiae Cell WallPeter 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 b1,6-glucans, a small amount of chitin, and many different proteins that may bear N- and O-linked glycans and a glycolipid anchor. Thesecomponents become cross-linked in various ways to form higher-order complexes. Wall composition and degree of cross-linking varyduring 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, glycosylphos-phatidylinositol membrane anchors, b1,3- and b1,6-linked glucans, and chitin; the reactions that cross-link wall components; and thepossible functions of enzymatic and nonenzymatic cell wall proteins.

TABLE OF CONTENTS

Abstract 775

Introduction 777

Wall Composition and Architecture 777Polysaccharides 778

Chitin: 778b-Glucans: 778Cross-links between polysaccharides: 779

Cell wall mannoproteins 779GPI proteins: 780Mild alkali-releasable proteins: 780Disulfide-linked proteins: 780

Strategies to identify CWP 780

Cell wall phenotypes 781

Precursors and Carrier Lipids 781Sugar nucleotides 781

Dolichol and dolichol phosphate sugars 781Dolichol phosphate synthesis: 781

Continued

Copyright © 2012 by the Genetics Society of Americadoi: 10.1534/genetics.112.144485Manuscript received May 17, 2012; accepted for publication August 6, 2012Supporting information is available online at http://www.genetics.org/lookup/suppl/doi:10.1534/genetics.112.144485/-/DC1.1Address 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: p-orlean@illinois.edu

Genetics, Vol. 192, 775–818 November 2012 775

CONTENTS, continued

Dol-P-Man and Dol-P-Glc synthesis: 781

Biosynthesis of Wall Components Along the Secretory Pathway 781N-Glycosylation 782

Assembly and transfer of the Dol-PP-linked precursor oligosaccharide: 782Steps on the cytoplasmic face of the ER membrane: 782Transmembrane translocation of Dol-PP-oligosaccharides: 782Lumenal steps in Dol-PP-oligosaccharide assembly: 783Oligosaccharide transfer to protein: 783

N-glycan processing in the ER and glycoprotein quality control: 783Mannan elaboration in the Golgi: 784

Formation of core-type N-glycan and mannan outer chains: 784Mannan side branching and mannose phosphate addition: 784

O-Mannosylation 785Protein O-mannosyltransferases in the ER: 785Extension and phosphorylation of O-linked manno-oligosaccharide chains: 785Importance and functions of O-mannosyl glycans: 785

GPI anchoring 785GPI structure and proteins that receive GPIs: 785

GPI structure: 785Identification of GPI proteins: 786

Assembly of the GPI precursor and its attachment to protein in the ER: 786Steps on the cytoplasmic face of ER membrane: 786Lumenal steps in GPI assembly: 787GPI transfer to protein: 788

Remodeling of protein-bound GPIs: 788

Sugar nucleotide transport 789GDP-Man transport: 789Other sugar nucleotide transport activities: 789

Biosynthesis of Wall Components at the Plasma Membrane 789Chitin 789

Septum formation: 789Chitin synthase biochemistry: 790S. cerevisiae’s chitin synthases and auxiliary proteins: 791

Chitin synthase I: 791Chitin synthase II and proteins impacting its localization and activity: 791Chitin synthase III and proteins impacting its localization and activity: 792

Chitin synthesis in response to cell wall stress: 793Chitin synthase III in mating and ascospore wall formation: 794

b1,3-Glucan 794Fks family of b1,3-glucan synthases: 794Roles of the Fks proteins in b1,3-glucan synthesis: 794Rho1 GTPase, a regulatory subunit of b1,3-glucan synthase: 795

b1,6-Glucan 795In vitro synthesis of b1,6-glucan 795

Proteins involved in b1,6-glucan assembly 796ER proteins: 796

Homologs of the UGGT/calnexin protein quality control machinery: 796Fungus-specific ER chaperones required for b1,6-glucansynthesis: 796

More widely distributed secretory pathway proteins: 797Kre6 and Skn1: 797Kre9 and Knh1: 797

Plasma membrane protein Kre1: 797How might b1,6-glucan be made?: 797

Continued

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CONTENTS, continued

Remodeling and Cross-Linking Activities at the Cell Surface 797Order of incorporation of components into the cell wall 797

Incorporation of GPI proteins into the wall 798

Incorporation of PIR proteins into the wall 798

Cross-linkage of chitin to b1,6- and b1,3-glucan 799

Cell Wall-Active and Nonenzymatic Surface Proteins and Their Functions 799Known and predicted enzymes 799

Chitinases: 799b1,3-glucanases: 799

Exg1, Exg2, and Ssg/Spr1 exo-b1,3-glucanases: 799Bgl2, Scw4, Scw10, and Scw11 endo-b1,3-glucanases: 800Eng1/Dse4 and Eng2/Acf2 endo-b1,3-glucanases: 800

Gas1 family b1,3-glucanosyltransferases: 800Yapsin aspartyl proteases: 800

Nonenzymatic CWPs 801Structural GPI proteins: 801

Sps2 family: 801Tip1 family: 801Sed1 and Spi1: 801Ccw12: 801Other nonenzymatic GPI proteins: 802Flocculins and agglutinins: 802

Non-GPI-CWP: 802PIR proteins: 802Scw3 (Sun4): 803Srl1: 803

What Is Next? 803

THE wall gives Saccharomyces cerevisiae its morphologiesduring budding growth, pseudohypha formation, mat-

ing, and sporulation; it preserves the cell’s osmotic integrity;and it provides a scaffold to present agglutinins and floccu-lins to other yeast cells. The wall consists of mannoproteins,b-glucans, and a small amount of chitin, which becomecross-linked in various ways. Wall composition and organi-zation vary during growth and development. During thebudding cycle, deposition of chitin is tightly controlled,and expression of certain hydrolases involved in cell separa-tion is daughter cell-specific. The wall can be weakened, andthe cell consequently stressed, by treatment with polysaccha-ride binding agents such as Calcofluor White (CFW), CongoRed, sodium dodecyl sulfate (SDS), aminoglycoside antibio-tics, and b-glucanase preparations or by mutational loss ofcapacity to make a wall component. Such stresses commonlyactivate 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 rolein maintenance of a normal wall. From the results of a surveyof deletion strains for cell wall phenotypes, De Groot et al.(2001) estimated that �1200 genes, not counting essentialones, impact the wall. Most of the effects, however, are in-

direct, and the number of genes that encode enzymes directlyinvolved in biosynthesis or remodeling of the wall, or non-enzymatic wall proteins, is now �180 (see Supporting Infor-mation, Table S1). This review covers these proteins, withemphasis on the wall of vegetative cells during the buddingcycle and in response to stress. Wall synthetic activities will becovered in the context of their cellular localization, startingwith precursors in the cytoplasm, proceeding along the secre-tory pathway from the endoplasmic reticulum (ER) to theplasma membrane, and culminating with the events outsidethe plasma membrane that generate covalent cross-links be-tween wall components. Additional information about indi-vidual proteins and the phenotypes of strains lacking them ispresented 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 cellwall 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 vege-tative 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 fromtransmission electron micrographs and by using an atomicforce microscope to detect surface accessibility of “molecularrulers” consisting of versions of the plasma membrane sensorWsc1 with different lengths (Dupres et al. 2010; Yamaguchiet al. 2011). The wall’s major components are b1,3- andb1,6-linked glucans, mannoproteins, and chitin, which canbe covalently joined to form higher-order complexes. Theb1,6-glucan, although quantitatively a minor componentof the wall, has a central role in cross-linking wall compo-nents (Kollar et al. 1997). Some mannoproteins have or arepredicted to have enzymatic activity as hydrolases or cross-linkers; others may have structural roles or mediate “socialactivity” by serving as mating agglutinins or flocculins.Among the latter, Flo1 and Flo11 promote formation of ex-tensive 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 ofvegetative cells reveal two layers. The outer one is electron-dense, has a brush-like surface (Osumi et al. 1998) (Babaet al. 1989; Osumi et al. 1998); Kapteyn et al. 1999a; Hagenet al. 2004; Yamaguchi et al. 2011), and can be removedby proteolysis (Kopecka et al. 1974; Zlotnik et al. 1984); ittherefore consists mostly of mannoproteins. The inner layer,more electron transparent, is microfibrillar and b-glucanase-digestible, indicating that its major components are glucans.The relative thicknesses of the two layers and their apparentorganization can be altered in cell wall mutants.

Relative amounts and localization of individual wallcomponents vary depending on cell cycle or developmentalstage, growth phase, nutritional conditions, and wall stressesimposed by hypo-osmolarity, mutational loss of wall bio-synthetic activities or wall proteins, or drug treatment.Variations in wall composition and organization impact theextent to which the wall is a barrier to export of soluble,secreted proteins to the medium. Some proteins can beretained by the wall outside the plasma membrane in theperiplasmic space; in the case of Suc2, this is due to the abilityof the protein to form large multimers (Orlean 1987). Thebarrier function of the wall is dependent on growth phase andcultural conditions, with the walls of growing cells beingmore porous (De Nobel and Barnett 1991). Native glycopro-teins such as Cts1, as well as many heterologously expressedsoluble glycoproteins with masses up to 400 kDa, can passthrough the wall of logarithmically growing cells to the me-dium, whereas walls of stationary-phase cells are less porous(De Nobel et al. 1990; Kuranda and Robbins 1991). Therelatively high porosity of walls of logarithmic-phase cellscould reflect a lower degree of cross-linking, but the dissolu-tion of septal material that occurs when dividing cells sepa-rate could also release wall proteins to the medium (seeOrder of incorporation of components into the cell wall). Per-spectives 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 compo-nents and strategies for isolating them are as follows.

Polysaccharides

Wall polysaccharides are typically separated into threefractions defined on the basis of their solubility in alkaliand acid (Fleet 1991). These fractions contain differing rel-ative amounts of b1,3- and b1,6-linked glucans and mannan(Magnelli et al. 2002) and also differ in whether and to whatextent the glucans are cross-linked to chitin, which deter-mines their solubility in alkali. Determination of Man-to-Glcratios in total acid hydrolysates of walls has been useful inassessing the impact of mutations on wall composition (Ramet al. 1994; Dallies et al. 1998). Digestion of isolated wallsand wall fractions with linkage-specific glycosidases hasbeen used to quantify wall components and determine thefine structure of b1,6-glucan (Boone et al. 1990; Magnelliet al. 2002; Aimanianda et al. 2009), as well as to generateoligosaccharides for structural analysis and characterizationof linkages between polymers (Kollar et al. 1995, 1997).

Chitin: This polymer of b1,4-linked GlcNAc contributes only1–2% of the dry weight of the wall of unstressed wild-typecells. Chitin is normally deposited in a ring in the neck be-tween a mother cell and its emerging bud, in the primarydivision septum, and in the lateral walls of newly separateddaughter cells. Chitin can be visualized in situ by stainingwith CFW, which reveals that most of it is present in divisionsepta and bud scars. Chitin in lateral walls and in divisionsepta can also be detected by immunoelectron microscopy(Shaw et al. 1991). Chitin levels are typically determinedafter extraction of walls with acid and alkali or hot SDS,followed by acid or enzymatic hydrolysis and quantificationof GlcNAc (Kang et al. 1984; Orlean et al. 1985; Dallies et al.1998; Magnelli et al. 2002). The average length of chitin inb-glucanase-digested septa is �110 GlcNAc residues (Kanget al. 1984). However, chitin occurs in three different andpolydisperse forms in the wall: in addition to free chitin,some is bound to b1,3-glucan and present mainly in theneck between mother and daughter cell, whereas a lesseramount, found in lateral walls, is bound to b1,6-glucan,which is in turn linked to mannan and b1,3-glucan (Cabiband Duran 2005; Cabib 2009). Chitin levels increase in re-sponse to mating pheromones (Schekman and Brawley1979; Orlean et al. 1985; see Sugar nucleotides) and delo-calized chitin in lateral walls can increase to as much as 20%of the wall in S. cerevisiae mutants mounting the cell wallstress 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 Chitinsynthesis in response to cell wall stress).

b-Glucans: b-linked glucans compose 30–60% of the dryweight of the wall and can be separated into three fractionsthat contain both b1,3 and b1,6 linkages. The major frac-tion, which makes up �35% of the dry weight of the wall, isan acid- and alkali-insoluble b1,3-glucan with a degree ofpolymerization of �1500 and b1,3-linked glucan side chainsinitiated at branching b1,6-linked glucoses that represent

778 P. Orlean

�3% of the whole polymer (Fleet 1991). The nonreducingends of b1,3-glucan chains in this fraction can be linked tochitin, 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 compo-sition to the alkali-insoluble b1,3-glucan, but soluble in al-kali because it is not cross-linked to chitin (Hartland et al.1994). A third fraction, making up �5% of the dry weight ofthe wall, can be released from alkali-insoluble glucan byextraction with acid or digestion with endo-b1,3-glucanase(Manners et al. 1973; Boone et al. 1990). This fraction isa b1,6-glucan with a degree of polymerization of 140, inwhich 14% of the b1,6-linked residues bear a side-branchingb1,3 Glc (Manners et al. 1973). A procedure involving serialdigestion with purified hydrolases has also been used toseparate and quantify b1,3- and b1,6-glucan, mannan, andchitin (Magnelli et al. 2002). The b1,6-glucan was releasedfrom the high-molecular-weight material remaining aftertreatment of walls with a mixture of b1,3-glucanase and chi-tinase by digestion with recombinant endo-b1,6-glucanase.The b1,6-glucan was therefore recovered as a mixture ofoligosaccharides whose major component was Glcb1,6Glc,and which also contained Glcb1,6Glcb1,6Glc and smalleramounts of Glcb1,3Glcb1,6Glc and Glcb1,6Glcb1,6Glc witha b1,3-Glc branching from its middle Glc (Magnelli et al.2002). The degree of b1,3 branching inferred from the oli-gosaccharide profile was similar to that reported by Mannerset al. (1973). This b1,6-glucan analysis would also includethe b1,6-glucan present in the alkali-soluble cell wall frac-tion, which is not included in procedures involving alkaliextraction. In another approach, b1,6-glucan was isolatedfollowing extraction of intact cells with hot SDS and mer-captoethanol, treatment with hot alkali under reducing con-ditions, and b1,3-glucanase digestion of the alkali-insolublematerial (Aimanianda et al. 2009). The b1,3-glucanase re-leasable material was a b1,6-glucan of 190–200 glucoseswith, on average, a b1,3-Glc or a b1,3-Glcb1,3-Glc side

branch on every fifth b1,6-linked glucose (Aimaniandaet al. 2009).

Cross-links between polysaccharides: Three types of link-ages between wall polysaccharides have been described(Figure 1). The first is a b1,4-linkage between the reducingend of a chitin chain and the nonreducing end of a b1,3-linked glucan (Kollar et al. 1995), and up to half of the chitinchains in the wall may be linked to b-glucan in this way.Because there is about one chitin-b-glucan linkage per 8000hexoses, these rare cross-links have a major impact on thesolubility of b-glucan (Kollar et al. 1995). The second linkageis between the reducing end of chitin and the nonreducingend of a b1,3-Glc that branches off b1,6-glucan (Kollar et al.1997; see Remodeling and Cross-Linking Activities at the CellSurface). The configuration of this linkage is either b1,2- orb1,4-. The two types of chitin-b-glucan linkage are found indifferent parts of the wall. In the third linkage, the reducingends 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-)linkedglycans, O-linked manno-oligosaccharides, and often a gly-cosylphosphatidylinositol (GPI) as well. The N-linked gly-cans can be extended with an outer chain of 50 or morea1,6-linked Man that is extensively decorated with shorta1,2-Man side branches terminated in a1,3-Man. Phospho-diester-linked mannoses can also be attached to a1,2-linkedresidues. 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 ofthree groups. The first contains those with the potential toparticipate in wall construction as hydrolases or trans-glycosidases. The second contains nonenzymatic aggluti-nins, flocculins, or b1,3-glucan cross-connectors (Klis et al.2006, 2010; Dranginis et al. 2007; Goossens and Willaert

Figure 1 Wall components and cross-links be-tween them. (A) Reducing end of chitin linked toa side-branching b1,3-Glc on b1,6-glucan. (B) Re-ducing end of chitin linked to a nonreducing endof b1,3-glucan. (C) Reducing end of b1,3-glucanchain linked to a side-branching b1,6-Glc on b1,3-glucan. (D) Reducing end of GPI glycan (possiblythe a1,4-Man) to internal Glc in b1,6-glucan (link-age to nonreducing end of b1,6-glucan is also pos-sible). (E) Ester linkages between b1,3-Glc andg-carboxyl groups of glutamates in PIR protein in-ternal repeats. (F) Disulfide link between CWP.Chemical treatments used to release CWP areindicated.

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2010). Most, if not all the proteins in these two groups areglycosylated. Proteins that are covalently attached to cell wallglycan are referred to as CWP (Yin et al. 2005) and fall intothe subgroups below. The third group consists of single-passplasma membrane proteins with short C-terminal cytoplasmicdomains and long Ser/Thr-rich extracellular regions. Theseinclude Wsc1, Wsc2, and Wsc3, which also have N-terminalcysteine-rich domains, as well as Mtl1 and Mid2. These aremechanosensors that detect cell wall stress and activate theCWI pathway (Rodicio and Heinisch 2010; Levin 2011). CWPand cell wall-active enzymes are discussed in Cell Wall-Activeand Nonenzymatic Surface Proteins and Their Functions.

GPI proteins: These receive a GPI that initially anchors themin the outer face of the plasma membrane, but many thenbecome cross-linked to b1,6-glucan via a remnant of the GPI(Gonzalez et al. 2009). Results to date suggest that the GPIis cleaved between its GlcN residue and Man, whereuponthe mannose’s reducing end is glycosidically linked to a non-reducing 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 towhich the GPI-CWP is attached is in turn linked to b1,3-glucan and chitin (Kapteyn et al. 1996; Van der Vaartet al. 1996; Kollar et al. 1997; Fujii et al. 1999; Figure 1).Some wall-bound GPI proteins may retain enzymatic activ-ity, whereas others may have a structural role (Yin et al.2005). GPI-CWP are released by treatment with hydrogenfluoride (HF)/pyridine, which cleaves the phosphodiesterof 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 signal-anchor sequence, and this, and signals for wall anchorage ofGPI-CWP, are discussed in Lumenal steps in GPI assembly andin Incorporation of GPI proteins into the cell wall. At least oneGPI-CWP, Cwp1, can additionally be linked to the wall viaan alkali-labile linkage (Kapteyn et al. 2001).

Mild alkali-releasable proteins: These include four proteinswith internal repeats (PIR proteins), which have multiplecopies 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 arereleased by mild alkali or b1,3-glucanase (Mrša et al. 1997).PIR proteins have no GPI attachment sequence and are notlinked to b1,6-glucan; rather, they are ester-linked to b1,3-glucan via side chains of amino acids in the repeat sequences(Ecker et al. 2006; see Incorporation of PIR proteins into thecell wall). Because PIR proteins can form several linkages tob1,3-glucan, they could interconnect glucans. Single PIRrepeats are also present in certain GPI-CWP (see Incorpora-tion of PIR proteins into the cell wall), and additional proteinslacking PIR sequences can be also extracted with alkali orb1,3-glucanase (Yin et al. 2005; see Cell Wall-Active andNonenzymatic Surface Proteins and Their Functions).

Disulfide-linked proteins: Various proteins can be releasedfrom the walls of living cells with sulfhydryl reagents,

indicating that they are directly attached via disulfides orretained behind a network of disulfide-linked proteins(Orlean et al. 1986; Cappellaro et al. 1998; Moukadiriet al. 1999; Moukadiri and Zueco 2001; Insenser et al.2010). Disulfide-linked mannoproteins create a barrier thatprotects wall polysaccharides from externally added glyco-sylhydrolases, making mercaptoethanol and protease pre-treatment necessary for spheroplasting with lytic enzymes(Zlotnik et al. 1984). Furthermore, the ability of the cyste-ine-rich domain of Wsc1 to form disulfide cross-links is im-portant for this mechanosensor in forming clusters and infunctioning in CWI signaling (Heinisch et al. 2010; Dupreset al. 2011).

Strategies to identify CWP

Biochemistry and bioinformatics have been used to identifyCWP. Because proteins can be associated with the wall indifferent ways, different treatments are necessary to releasethem. Separation and identification of individual CWP canbe complicated by their heavy and heterogeneous glycosyl-ation. CWP can be released from the wall by treatment withb1,3- and b1,6-glucanases (Van der Vaart et al. 1995; Mršaet al. 1997; Shimoi et al. 1998). In one approach, labeling ofintact cells with a membrane-impermeable biotinylation re-agent, followed successively by SDS and mercaptoethanolextraction 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 withSDS, mercaptoethanol, NaCl, and EDTA, were then treatedwith HF/pyridine or mild alkali, and the CWP released wereidentified by mass spectrometry. Additional CWP were iden-tified following proteolytic digestion of walls, the two pro-cedures yielding 19 CWP, including GPI and PIR proteinsand alkali-releasable proteins without PIR sequences (Yinet al. 2005, 2007). These studies led to the estimate thata dividing haploid cell contains �2 · 106 covalently at-tached CWPs and the suggestion that CWP form a denselypacked surface layer (Yin et al. 2007). A strategy that alsopermitted identification of noncovalently associated surfaceproteins used treatment of intact cells with dithiothreitolfollowed by two-dimensional electrophoretic separation, ordirect proteolytic digestion and isolation of peptides, andthen mass spectrometric protein fingerprinting (Insenseret al. 2010). The 99 proteins so identified included CWPand glycosylhydrolases, as well as proteins associated withintracellular functions. The presence in the wall of proteinsconsidered cytosolic raises the possibility that they reach thewall via a nonconventional export pathway (Nombela et al.2006; Insenser et al. 2010). However, mercaptoethanol canmake the plasma membrane permeable to cytosolic proteins(Klis et al. 2007).

Bioinformatics has been used identify proteins likely toreceive a GPI anchor; hence, members of the major class ofCWP. In silico surveys for GPI attachment sequences revealthat the S. cerevisiae proteome contains 60–70 potential GPI

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proteins, which often contain Ser/Thr-rich stretches (Caroet 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 sensitivityto hypo-osmotic stress, which can be tested on half-strengthyeast extract peptone medium (Valdivia and Schekman2003); sensitivity or resistance to CFW and Congo Red; sen-sitivity to aminoglycosides, b1,3-glucan synthase inhibitors,caffeine, SDS, and K1 killer toxin; and sensitivity to b1,3-glucanase 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 andGlc-1-P by the essential UDPGlc pyrophosphorylase Ugp1(Daran et al. 1995). Impairment of UDP-Glc synthesis ulti-mately impacts formation of cell wall b-glucans, althoughcells with no more than 5% of the activities of the phospho-glucomutases and Ugp1 that generate UDP-Glc are unaf-fected in growth and viability (Daran et al. 1997). GDP-Man is formed from Fru-6-P by the successive actions ofphosphomannose isomerase (Pmi40), phosphomannomutase(Sec53), and GDP-Man pyrophosphorylase (Psa1/Srb1/Vig9),which are all encoded by essential genes, and loss of any ofthese enzyme activities leads to severe glycosylation and se-cretion defects (Hashimoto et al. 1997; Orlean 1997; Yodaet al. 2000). Elevated expression of GDP-Man pyrophosphor-ylase, which presumably increases GDP-Man levels, correctsthe N-glycosylation defects in alg1 and alg2 mutants and themannosylation and GPI synthetic defects in dpm1 cells (Janiket al. 2003). GDP-Man transport into the Golgi lumen is dis-cussed 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 gluta-mine:Fru-6-P amidotransferase Gfa1 (Watzele and Tanner1989), N-acetylation of GlcN-6-P by Gna1 (Mio et al.1999), conversion of GlcNAc-6-P to GlcNAc-1-P by theGlcNAc phosphate mutase Agm1/Pcm1 (Hofmann et al.1994), and formation of UDP-GlcNAc by the pyrophosphor-ylase Uap1/Qri1 (Mio et al. 1998). Deficiencies in theseenzymes lead to formation of short chains of undivided cells,swelling, and eventual lysis, a phenomenon known as glu-cosamineless death (Ballou et al. 1977; Mio et al. 1998,1999). Glucosamine supply is highly regulated and impactschitin levels, which increase in response to mating phero-mones and cell wall stress (File S1).

Dolichol and dolichol phosphate sugars

Dolichol phosphate synthesis: Yeast dolichols contain 14–18 isoprene units (Jung and Tanner 1973). Biosynthesis of

dolichol (Schenk et al. 2001a; Grabinska and Palamarczyk2002) starts with extension of trans farnesyl-PP by succes-sive addition of cis-isoprene units by the homologous cis-prenyltransferases Rer2 and Srt1 (Sato et al. 1999; Schenket al. 2001b). Rer2 is dominant and makes dolichols with10–14 isoprene units, whereas dolichols made by Srt1 incells lacking Rer2 contain 19–22 isoprenes. rer2D strainshave severe defects in growth and in N- and O-glycosylation(Sato et al. 1999). The next two steps are likely the removalof the two phosphates from dehydrodolichyl diphosphate byunknown enzymes. The a-isoprene unit of the polyprenol isthen reduced, and Dfg10 is responsible for much of thisactivity (Cantagrel et al. 2010; File S1). Dolichol is likelynext phosphorylated by the CTP-dependent Dol kinaseSec59 (Heller et al. 1992).

Dol-PP generated on the lumenal side of the ER mem-brane after transfer of the N-linked oligosaccharide toprotein is dephosphorylated to Dol-P and Pi on that side ofthe membrane by the phosphatase Cwh8/Cax4 (Van Berkelet al. 1999; Fernandez et al. 2001). CWH8-disruptants havean N-glycosylation defect and a growth defect that is par-tially suppressed by high-level expression of RER2, SEC59,and the lipid phosphatase gene LPP1. Cwh8 likely has a role inrecycling of Dol-PP for use in new rounds of N-glycosylationon the cytoplasmic face of the ER membrane.

Dol-P-Man and Dol-P-Glc synthesis: Dol-P-Man and Dol-P-Glc are the donors in the lumenal glycosyltransfers thatoccur in protein O-mannosylation and the assembly path-ways for the Dol-PP-linked precursor in N-glycosylation andthe GPI anchor precursor glycolipid. Dol-P-Man is formedupon transfer of Man from GDP-Man to Dol-P by the Dol-P-Man synthase Dpm1 (Orlean et al. 1988; Orlean 1990).Temperature-sensitive dpm1 mutants have cell wall defects,consistent with a general block of glycosylation and GPIanchoring, and these phenotypes are suppressed by high-level expression of RER2, which presumably elevates Dol-Plevels (Orlowski et al. 2007).

Dol-P-Glc is formed from UDP-Glc and Dol-P. Deletion ofthe synthase gene, ALG5, is not lethal, and the disruptantsshow no obvious growth defects (Te Heesen et al. 1994).Because Dol-P-Man and Dol-P-Glc are used in lumenal reac-tions, and because spontaneous transmembrane transloca-tion of these glycolipids is not favored energetically, theirtranslocation may be protein-mediated. Assays for Dol-P-Man flipping have been reported (Haselbeck and Tanner1982; Sanyal and Menon 2010), but a protein involvedhas yet to be identified. One possibility is that the Dol-P-Man and Dol-P-Glc-utilizing transferases are their own flip-pases (Burda and Aebi 1999).

Biosynthesis of Wall Components Along theSecretory Pathway

Cell surface proteins can be modified with N-glycans,O-linked manno-oligosaccharides, and a GPI anchor as they

S. cerevisiae Cell Wall 781

transit the secretory pathway. Initial attachment of thesestructures occurs in the ER lumen, and the glycans aremodified in the Golgi before the glycoproteins are depositedin the plasma membrane or secreted from the cell, where-upon many become cross-linked to wall polysaccharides.

N-Glycosylation

N-glycosylation involves preassembly of a branched 14-sugar oligosaccharide on the carrier Dol-PP in the ERmembrane and then transfer of the oligosaccharide toselected asparagines in the ER lumen (Burda and Aebi1999; Helenius and Aebi 2004; Lehle et al. 2006; Larkinand Imperiali 2011). The first 7 sugars are transferredfrom sugar nucleotides on the cytosolic side of the ER mem-brane, and the remainder from Dol-P on the lumenal side(Figure 2).

Assembly and transfer of the Dol-PP-linked precursoroligosaccharide: Steps on the cytoplasmic face of the ERmembrane: These steps are (i) transfer of GlcNAc-1-P fromUDP-GlcNAc to Dol-P by Alg7, the target of the N-glycosylationinhibitor tunicamycin (Barnes et al. 1984), (ii) transfer of b1,4-

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 a1,6Man 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). Theseproteins 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 theER membrane is somehow translocated into the lumen(Burda and Aebi 1999; Helenius and Aebi 2002), and Rft1is a candidate for the flippase (Helenius et al. 2002). Strainsdeficient in Rft1 accumulate Dol-PP-GlcNAc2Man5, but retainAlg3 Man-T activity and are unaffected in O-mannosylationor in GPI assembly, ruling out deficiences in Dol-P-Man sup-ply to the lumen. Furthermore, high level expression ofRFT1 partially suppresses the growth defect of alg11D andleads to increased levels of lumenal Dol-PP-GlcNAc2Man6-7and an increase in glycosylation of the reporter carboxypepti-dase Y, consistent with enhanced flipping of the suboptimal

Figure 2 Assembly of the Dol-PP-linked precursor oligosaccharide in N-glycosylation, its transfer to protein, and subsequent glycan processing. Residuesadded at the cytoplasmic face of the ER membrane originate from sugar nucleotides, whereas Dol-P sugars generated at the cytoplasmic face of themembrane are the donors in lumenal transfers. Symbols are adaptations of those used by the Consortium of Glycobiology Editors in Essentials inGlycobiology (Varki and Sharon 2009).

782 P. Orlean

substrate Dol-PP-GlcNAc2Man3 (Helenius et al. 2002). How-ever, although the above evidence is consistent with Rft1being the flippase, depletion of Rft1 did not lead to loss offlipping activity measured in vitro (Frank et al. 2008; Rushet al. 2009; File S2).

Lumenal steps in Dol-PP-oligosaccharide assembly: Dol-PP-GlcNAc2Man5 is extended by four Man and three Glc on thelumenal side of the ER membrane using Dol-P-Man and Dol-P-Glc as donors. Alg3 adds the sixth, a1,3-Man to the a1,6Man of Dol-PP-GlcNAc2Man5 (Aebi et al. 1996; Sharma et al.2001), Alg9 then transfers an a1,2-linked Man to the Manadded by Alg3 (Burda et al. 1999; Cipollo and Trimble2002), and Alg12 next adds the eighth, a1,6-Man to theMan added by Alg9 (Burda et al. 1999). Alg9 acts again toadd the ninth Man, a1,2-linked Man to the Man added byAlg12 (Frank and Aebi 2005). Two a1,3-linked Glc are suc-cessively added by Alg6 and Alg8 to extend the arm of theheptasaccharide ending in the a1,2-linked Man transferredby Alg11, and finally, Alg10 adds an a1,2-Glc (Stagljar et al.1994; Reiss et al. 1996; Burda and Aebi 1998). The six Dol-P-sugar-utilizing transferases are members of a family ofmultispanning membrane proteins that includes Man-T in-volved in GPI biosynthesis (Oriol et al. 2002).

Oligosaccharide transfer to protein: GlcNAc2Man9Glc3 istransferred from Dol-PP to asparagines by the oligosacchar-yltransferase complex (OST) (Knauer and Lehle 1999a; Yanand Lennarz 2005a; Kelleher and Gilmore 2006; Lehle et al.2006; Weerapana and Imperiali 2006; Lennarz 2007; Larkinand Imperiali 2011). Acceptor asparagines occur in thesequon Asn-X-Ser/Thr, where X can be any amino acid exceptPro. Mass spectrometric analyses of wall-derived peptidesrevealed that 85% of sequons were completely occupied, withpreferential usage Asn-X-Thr over Asn-X-Ser sites (Schulzand Aebi 2009). Analyses of protein-linked N-glycans inmutants defective in the elaboration of the Dol-PP-linkedprecursor indicate that structures smaller than GlcNAc2-Man9Glc3 can be transferred in vivo.

Yeast OST consists of Stt3, Ost1, Ost2, Wbp1, Swp1,Ost4, Ost5, and either of the paralogues Ost3 or Ost6. Thefirst five are encoded by essential genes. Two OST com-plexes can be formed, containing either Ost3 or Ost6(Schwarz et al. 2005; Spirig et al. 2005; Yan and Lennarz2005b). The Ost3-containing complex is about four times asabundant as the Ost6-containing one (Spirig et al. 2005).Genetic interaction studies and coimmunoprecipitation andchemical cross-linking experiments suggest the existence ofthree OST subcomplexes: (i) Swp1-Wbp1-Ost2, (ii) Stt3-Ost4-Ost3, and (iii) Ost1-Ost5 (Karaoglu et al. 1997; Reisset al. 1997; Spirig et al. 1997; Knauer and Lehle 1999b; Kimet al. 2003; Li et al. 2003; Kelleher and Gilmore 2006; FileS2). OST complexes themselves may function as dimers(Chavan et al. 2006).

Stt3 is the catalytic subunit of OST. It can be chemicallycross-linked to peptides derivatized with photoactivatablegroups (Yan and Lennarz 2002; Nilsson et al. 2003), andits 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 witha CXXC motif common to proteins involved in disulfide bondshuffling during oxidative protein folding (Kelleher andGilmore 2006; Schulz et al. 2009), and the proteins likelyform transient disulfide bonds with nascent proteins andpromote efficient glycosylation of Asn-X-Ser/Thr sites bydelaying oxidative protein folding (Schulz and Aebi 2009;Schulz et al. 2009). The Swp1p, Wbp1p, and Ost2p subcom-plex may confer the preference of OST for GlcNAc2Man9Glc3(Pathak et al. 1995; Kelleher and Gilmore 2006), Ost4 isrequired for recruitment of Ost3 and Ost6 to OST and alsointeracts 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 theCWI pathway via an interaction between Pkc1 or compo-nents of the PKC pathway with Stt3 (Park and Lennarz2000; Chavan et al. 2003a; File S2).

N-glycan processing in the ER and glycoprotein qualitycontrol: Protein-linked GlcNAc2Man9Glc3 is processed toglycans that are recognized by mechanisms that monitorcorrect protein folding and permit export from the ER orensure degradation if the protein misfolds (Herscovics1999; Aebi et al. 2010). Processing proceeds by removal ofthe a1,2-linked Glc by glucosidase I, Gls1/Cwh41 (Romeroet al. 1997), and then of the two a1,3-linked Glc by solubleglucosidase II, a heterodimer of catalytic Gls2/Rot2 andGtb1 (Trombetta et al. 1996; Wilkinson et al. 2006; Quinnet al. 2009; Figure 2). ER mannosidase I, Mns1, removesan a1,2 Man to generate GlcNAc2Man8 (Jakob et al. 1998;Herscovics 1999), and, if correctly folded, proteins bearingthis glycan are exported from the ER. Un- or misfolded pro-teins are bound by protein disulfide isomerase Pdi1, some ofwhich is in complex with Mns1 homolog Htm1, which trimsthe glycan to a GlcNAc2Man7 (Clerc et al. 2009; Gauss et al.2011; File S2). Misfolded proteins with GlcNAc2Man7 aretargeted to the cytosol for destruction by the ER-associatedprotein degradation (ERAD) system (Helenius and Aebi2004). They are bound by the lectin Yos9 (Buschhorn et al.2004; Bhamidipati et al. 2005; Kim et al. 2005; Szathmaryet al. 2005) and in turn directed to the HRD-ubiquitin ligasecomplex of Hrd1 and Hrd3 for retrotranslocation to the cy-toplasm (Bays et al. 2001; Deak and Wolf 2001; Gauss et al.2006), where they are deglycosylated by peptide N-glycanasePng1 (Suzuki et al. 2000; Hirayama et al. 2010).

In mammals and Schizosaccharomyces pombe, followingglucosidase II action, UDP-Glc:glycoprotein glucosyltransfer-ase (UGGT) adds back an a1,3-Glc, allowing the monoglu-cosylated N-glycans to interact with the lumenal lectindomains of calnexin or calreticulin (Parodi 1999; Carameloand Parodi 2007; Aebi et al. 2010). This interaction retainspartially folded or misfolded proteins in the ER and buysthem time to fold properly and be deglucosylated. Properly

S. cerevisiae Cell Wall 783

folded proteins are no longer recognized by UGGT andexported to the Golgi, whereas persistent misfolders are re-moved by ERAD. In S. cerevisiae, however, this quality con-trol mechanism does not operate because UGGT activity isnot detectable, and although the S. cerevisiae ER proteinKre5 is a sequence homolog of S. pombe UGGT, expressionof the S. pombe UGGT cannot rescue the growth defect ofkre5 mutants. However, kre5, as well as glucosidase I and IImutants and mutants in the calnexin homolog Cne1, are de-fective in b1,6-glucan synthesis, indicating roles for S. cerevisiaehomologs of players in the UGGT/calnexin quality controlsystem in b1,6-glucan synthesis (Jiang et al. 1996; Shahinianet al. 1998; Simons et al. 1998; see b1,6-Glucan).

Mannan elaboration in the Golgi: N-linked glycans onproteins are extended with a Man10-14 core-type structure orwith 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). Themannoses all originate from GDP-Man and are transferredby 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 inthe cis-Golgi by Och1, which transfers an a1,6-Man to thea1,3-Man of the N-glycan that had been added by Alg2(Nakayama et al. 1997). OCH1 deletion is lethal in somestrain backgrounds, and och1D strains have severe growthdefects, highlighting the importance of mannan.

Synthesis of the poly-a1,6-mannan backbone is carriedout in the cis-Golgi by two protein complexes: Man-Pol I,see containing homologs Mnn9 and Van1, and Man-Pol II,

containing Mnn9, Anp1, Hoc1, and related Mnn10 andMnn11 (Hashimoto and Yoda 1997; Jungmann and Munro1998; Jungmann et al. 1999; File S2). M-Pol I acts first, withits Mnn9 subunit adding the first a1,6-Man to the Och1-derived Man, upon which 10–15 a1,6-Man are added inVan1-requiring reactions (Stolz and Munro 2002; Rodionovet al. 2009). This a1,6 backbone is further elongated with40–60 a1,6-Man by M-Pol II, whose Mnn10 and Mnn11subunits are responsible for the majority of the a1,6-Man-T activity (Jungmann et al. 1999). Hoc1’s role is unclear.

Core-type N-glycans are formed when an a1,2-Man is addedto the Och1-derived Man, blocking elongation of an a1,6 man-nan chain. The protein(s) involved have not been identified,but presumably either they, or M-Pol I, can tell from the contextof an N-glycan which type of structure it is to bear (Lewis andBallou 1991; Stolz and Munro 2002; Rodionov et al. 2009).Core-type structures are completed when that a1,2-Man, aswell as the two other terminal a1,2-Man on the Man8GlcNAc2structure, receives a1,3 mannoses from Mnn1.

Mannan side branching and mannose phosphate addition:Branching of the a1,6 mannan backbone is initiated by theMnn2 a1,2-Man-T, and Mnn5 adds a second a1,2-Man(Rayner and Munro 1998). Mnn2 and Mnn5 make up oneof two Mnn1 subfamilies (Lussier et al. 1999). Five membersof 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 subse-quent a1,2-mannoses to mannan side branches (Lussieret al. 1996, 1997a, 1999).

Core-type glycans and mannan can be modified with Man-Pon a1,2-linked mannoses. Mnn6/Ktr6, a Ktr1 subfamily

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 theOch1 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 chainis 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-typeglycans are formed when an a1,2-linked Man is added to the Och1-derived a1,6-Man. Symbols are as used in Figure 2.

784 P. Orlean

member, is mostly responsible for transferring Man-1-Pfrom GDP-Man, generating GMP (Wang et al. 1997; Jigamiand Odani 1999; File S2). Mnn4 is also involved in Man-Paddition but does not resemble glycosyltransferases andmay be regulatory (Odani et al. 1996). Levels of mannanphosphorylation are highest in the late log and stationaryphases, when MNN4 expression is elevated (Odani et al.1997). Terminal a1,2 mannoses and Man-1-Ps can be cap-ped with a1,3-Man, added by Mnn1 (Ballou 1990; Yip et al.1994).

O-Mannosylation

Many yeast proteins are modified on extracytoplasmic Ser orThr residues with linear manno-oligosaccharides. The firstMan is attached in a-linkage in the lumen of the ER, and upto four further Man are added by Man-T that act mostly inthe Golgi.

Protein O-mannosyltransferases in the ER: The first Man istransferred from Dol-P-Man (Strahl-Bolsinger et al. 1999;Lehle et al. 2006; Lommel and Strahl 2009). Consistent withthe requirement for Dol-P-Man, O-mannosylation of the modelprotein 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 membrane-spanning domains with conserved residues important forcatalysis and for interactions with acceptor peptides locatedin the first lumenal loop (Strahl-Bolsinger and Scheinost1999; Girrbach et al. 2000; Lommel et al. 2011).

Pmts function as hetero- or homodimers, and the pairs thatare formed are determined by membership of a subunit inone of three Pmt subfamilies. Pmt1 family members Pmt1 andPmt5 can form heterodimers with members of the Pmt2 fam-ily (which also contains Pmt3 and Pmt6), for example, Pmt1-Pmt2 and Pmt5-Pmt3 dimers, which are the most prevalentcomplexes (Girrbach and Strahl 2003). Pmt4, the lone repre-sentative of the third family, forms homodimers.

Analyses of O-mannosylation of individual proteins in pmtDstrains reveal that the different Pmt complexes have specificityfor different protein substrates (File S3). Substrates for Pmt4need to be attached to the membrane by a transmembranedomain or a GPI anchor and have an adjacent, lumenal Ser/Thr-rich domain, whereas Pmt1/Pmt2 substrates can be solu-ble or membrane-associated (Hutzler et al. 2007).

Because PMTs modify Ser and Thr, N-linked glycosylationsites are also potential targets, and this is the case with Cwp5.This protein contains a single sequon, NAT, that is normallyO-mannosylated by Pmt4, but which receives an N-linkedglycan in pmt4D cells (Ecker et al. 2003). O-mannosylation,therefore, normally precedes the action of OST on Cwp5 andmay control N-glycosylation of this protein, and perhapsothers as well.

Extension and phosphorylation of O-linked manno-oligosaccharide chains: The Ser- or Thr-linked Man is ex-tended 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 iscarried out by the Ktr1 subfamily members Ktr1, Ktr3, andKre2 and extension of the trisaccharide chain with oneor 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 modifiedwith Man-1-P by Mnn6 with the involvement of the regula-tor Mnn4 (Nakayama et al. 1998).

Importance and functions of O-mannosyl glycans: Noindividual PMT deletion is lethal, but strains lacking certaincombinations of three Pmts, such as pmt1D pmt2D pmt4D orpmt2D pmt3D pmt4D, are inviable, even with osmotic sup-port, indicating that yeast must carry out some minimumlevel of O-mannosylation to be viable or that one or moreessential proteins need to be O-mannosylated (Gentzsch andTanner 1996; Lommel et al. 2004). Moreover, strains lackingother combinations of Pmts, such as the pmt2D pmt3D andpmt2D pmt4D double nulls or the pmt1D pmt2D pmt3D tri-ple null, are osmotically fragile, indicating impaired wallassembly (Gentzsch and Tanner 1996). Analyses of pmtmutants show that O-mannosylation can be important forfunction of individual O-mannosylated proteins (File S3).

The phenotypes of pmtmutants are mimicked by treatmentwith the rhodanine-3-acetic acid derivative OGT1458, whichinhibits PMT activity in vitro (Orchard et al. 2004; Arroyo et al.2011). OGT1458 was used to analyze genome-wide transcrip-tional changes in response to inhibition of O-mannosylation.Consistent with the importance of O-mannosylation in wallconstruction and protein stability, consequences of defectiveO-mannosylation were activation of the CWI pathway andthe unfolded protein response (Arroyo et al. 2011). Further-more, certain genes involved in N-linked mannan outer chainassembly were upregulated. This, together with the findingthat PMT gene transcription is elevated when N-glycosylationis inhibited by tunicamycin (Travers et al. 2000), suggeststhat the N- and O-linked glycans of cell wall mannoproteinscan compensate for one another to some extent (Arroyoet al. 2011).

GPI anchoring

GPI structure and proteins that receive GPIs: GPI structure:S. cerevisiae GPI anchors have the core structure protein CO-NH2-CH2-CH2-PO4-6-Mana1,2Mana1,6Mana1,4GlcNa1,6-myoinositol phospholipid. In addition, the third, a1,2-Man,bears a fourth a1,2-Man that is added during precursor as-sembly, and this Man may receive another a1,2- or a1,3-linked Man in the Golgi (Fankhauser et al. 1993). Thea1,4- and a1,6-linked Man are also modified with Etn-P attheir 29- and 69-OHs, respectively, and the 2-OH of inositol istransiently modified with palmitate (Orlean and Menon2007; Pittet and Conzelmann 2007) (Figure 5). The lipidmoiety, initially diacylglycerol, is remodeled to a diacylgly-cerol with C26, acyl chains, or, in many cases, to a ceramide(Conzelmann et al. 1992; Fankhauser et al. 1993).

S. cerevisiae Cell Wall 785

Identification of GPI proteins: Biochemical demonstrationsof a GPI on a yeast protein are rare, and the criterion ofrelease of a protein by treatment with Ptd-Ins-specificphospholipase C (PI-PLC) is unreliable because althoughprotein-bound GPIs are mostly sensitive to PI-PLC, thistreatment does not always render the protein aqueoussoluble in the commonly used Triton X-114 fractionationprocedure (Conzelmann et al. 1990). Many GPI proteinsbecome covalently linked to wall polysaccharide, and re-lease from walls by treatment with HF/pyridine is a cluethat the protein had received a GPI (see GPI proteins; Yinet al. 2005). The presence of a GPI is usually inferred fromthe results of in silico analyses of a protein’s sequence.

Features of a likely GPI protein are a hydrophobicN-terminal secretion signal and a C-terminal GPI signal-anchor sequence that includes the amino acid residue, v, towhich the GPI will be amide-linked. Amino acids N-terminalto v are designated v(2), and those C-terminal, are desig-nated v(+). Proceeding from the C-terminal amino acid ofthe unprocessed protein, the signal anchor signal consists of(i) a variable stretch of hydrophobic amino acids capableof spanning the membrane; (ii) a spacer region of moder-ately 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) thev residue itself, generally G, A, S, N, D, or C; and (v)a stretch of some 10 amino acids that may form a flexiblelinker region and whose relative polarity may influenceplasma membrane or wall localization of the protein(Nuoffer et al. 1991, 1993; see Incorporation of GPI proteinsinto the cell wall). Some C-terminal sequences may containalternative candidates for the v and v+2 amino acids. Ev-

idence that a predicted GPI attachment sequence is func-tional can be obtained by fusing the sequence to the Cterminus of a reporter protein and testing whether the re-porter becomes expressed at the plasma membrane or in thewall (Hamada et al. 1998a).

Assembly of the GPI precursor and its attachment toprotein in the ER: At least 21 proteins are involved in GPIprecursor synthesis and attachment to protein (Figure 5).Eighteen are encoded by essential genes, and mutants lack-ing any of the other noncatalytic proteins or GPI side-branching enzymes have severe growth defects. Additionalinformation about GPI synthetic proteins and phenotypesassociated with deficiencies in them is given in File S4.

Steps on the cytoplasmic face of ER membrane: GPI assemblystarts with transfer of GlcNAc from UDP-GlcNAc to PI. Acomplex of at least six proteins (GPI-GnT) is involved, ofwhich Gpi3 is catalytic because it can be labeled with a photo-activatable UDP-GlcNAc analog (Kostova et al. 2000). GlcNActransfer occurs at the cytoplasmic face of the ER membrane(Vidugiriene and Menon 1993; Watanabe et al. 1996; Tiedeet al. 2000). Essential Gpi2, Gpi15, and Gpi19 (Leidich et al.1995; Yan et al. 2001; Newman et al. 2005), and nonessentialGpi1 and Eri1 (Leidich and Orlean 1996; Sobering et al.2004), are also required for GlcNAc-PI synthesis. ERI1 andGPI1 null mutants are temperature-sensitive. The mammalianorthologs of these proteins form a complex (Watanabe et al.1998; Tiede et al. 2000; Eisenhaber et al. 2003; Murakamiet al. 2005), and the yeast proteins likely also do, for Eri1 andGpi19 associate with Gpi2 (Sobering et al. 2004; Newmanet al. 2005). Roles of the noncatalytic subunits are unclear.

Figure 4 Biosynthesis of O-linked glycans. (A) Addition ofa-Man by protein O-mannosyltransferases in the ER lu-men. Pmt4 homodimers act on membrane proteins orGPI proteins. Representative Pmt heterodimers are shown.(B) Extension of O-linked manno-oligosaccharides in theGolgi. Ktr1 family members have a collective role in addinga1,2-linked mannoses, and Mnt1 family members adda1,3-linked mannoses. The dominant Man-T active ateach step are shown in boldface type. Man-P may beadded to saccharides with two a1,2-linked Man.

786 P. Orlean

Ras2, in its GTP-bound form, can also join GPI-GnT(Sobering et al. 2004). Membranes from ras2D cells have8- to 10-fold higher in vitro GPI-GnT activity than wild-typemembranes, whereas membranes from cells expressingconstitutively active Ras2-Val19 have almost undetectableactivity. These findings indicate that Ras2-GTP is a negativeregulator of GPI-GnT, and, depending on the degree towhich the GTPase is activated, this could permit abouta 200-fold range of GlcNAc-PI synthetic activity.

Once formed, GlcNAc-PI is de-N-acetylated at the cyto-plasmic face of the ER membrane by Gpi12 (Vidugiriene andMenon 1993; Watanabe et al. 1999). GlcN-PI is the precursorlikely to be translocated to the lumenal side of the ER mem-brane. Its flipping has been reconstituted in rat liver micro-somes, but the protein involved is unknown (Vishwakarmaand Menon 2005).

Lumenal steps in GPI assembly: The inositol ring in GlcN-PIis next acylated on its 2-OH, making the glycolipid resistantto cleavage by PI-specific phospholipase C. The reaction usesacyl CoA as donor (Costello and Orlean 1992), and the acylchain transferred in vivo is likely palmitate. Gwt1, the acyl-transferase, was identified in a screen for resistance to 1-[4-butylbenzyl] isoquinoline, which inhibits surface expressionof GPI proteins (Tsukahara et al. 2003; Umemura et al.2003). Disruption of GWT1 is lethal or leads to slow growthand temperature sensitivity, depending on the strain back-ground (Tsukahara et al. 2003). The inositol acyl chain mayprevent GPIs from being translocated back to the cytoplas-mic side of the ER membrane (Sagane et al. 2011), be im-portant 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 GPI-Man-T I-IV, and the first three Man are concurrentlymodified with Etn-P by Etn-P-T I, II, and III. Dol-P-Mandonates the mannoses because the dpm1 mutant accumu-

lates GlcN-(acyl)PI (Orlean 1990). The first, a1,4-linkedMan (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 sterolmetabolism. ARV1 disruptants are impaired in ER-to-Golgitransport of GPI proteins and accumulate GlcN-(acyl)PIin vitro (but not in vivo), although they are not defective inin 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 becauseexpression of both GPI14 and PBN1 is necessary to comple-ment mammalian cell lines defective in Pbn1’s mammalianhomolog Pig-X, and co-expression of PIG-X and the gene forGpi14’s mammalian homolog, PIG-M, partially rescues thelethality of gpi14D (Ashida et al. 2005; Kim et al. 2007).Furthermore, Pbn1 depletion leads to accumulation of someof the ER form of the GPI protein Gas1, a phenotype of GPIprecursor 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). Gpi18-deficient cells accumulate both a Man1-GPI with Etn-P esteri-fied to its Man and an unmodified Man1-GPI, suggesting thatGPI-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 andMan-4 (Canivenc-Gansel et al. 1998; Sütterlin et al. 1998;Grimme et al. 2001). Smp3-dependent addition of Man-4 isessential because addition of this residue precedes additionof the Etn-P that subsequently becomes linked to protein(Grimme et al. 2001).

As the GPI glycan is extended, Etn-P moieties are addedto the 2-OH of Man-1 and to the 6-OH of Man-2 and Man-3

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 toGlcN-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 andMcd4 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 alsobeen detected in radiolabeling experiments with certain late-stage GPI assembly mutants.

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(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, andGpi13, respectively, which are 830- to 1100-amino-acid pro-teins predicted to have 10–14 transmembrane domains anda large lumenal loop containing sequences characteristicof the alkaline phosphatase superfamily that are importantfor function (Benachour et al. 1999; Gaynor et al. 1999;Galperin and Jedrzejas 2001; File S4). GPI-Etn-P-T-II andIII also require small, hydrophobic Gpi11 for activity. mcd4mutants accumulate unmodified Man1 and Man2-GPI(Wiedman et al. 2007; Scarcelli et al. 2012), suggesting thatboth structures can serve as Etn-P acceptors. From this, andbecause Gpi18-depleted cells accumulate Etn-P-modifiedMan1-GPI (Fabre et al. 2005), it seems that both Mcd4and Gpi18 can use Man1-GPI as acceptor and then modifythe GPI that the other has acted on (Figure 5; File S4). Etn-Ptransfer to Man-1 and GPI-dependent processing of Gas1 areinhibited by the terpenoid lactone YW3548 (Sütterlin et al.1997, 1998). The Etn-P on Man-1 may enhance the ability ofGpi10 to add Man-3, promote export of GPI proteins fromthe ER, and be necessary for remodeling of the lipid moietyto ceramide (Zhu et al. 2006).

Gpi7 is the catalytic subunit of GPI-Etn-P-T-II, and GPI7nulls, which are viable but temperature-sensitive, accumulatea Man4-GPI with Etn-P on Man-1 and Man-3 (Benachouret al. 1999). Essential Gpi11 was implicated at this stepbecause Gpi11-deficient cells have similar GPI precursor ac-cumulation profiles to gpi7D (Taron et al. 2000). The Etn-Pon Man-2 enhances transfer of GPIs to protein, ER-to-Golgitransport of GPI proteins, GPI lipid remodeling to ceramide,transfer of GPI proteins to the wall, and targeting of certainGPI-anchored proteins in daughter cells (Benachour et al.1999; Toh-E and Oguchi 1999; Richard et al. 2002; Fujitaet al. 2004).

Gpi13 is the catalytic subunit of GPI-Etn-P-T-III. The ma-jor GPI accumulated upon Gpi13 depletion is a Man4-GPIwith 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 reac-tion because a gpi11-Ts mutant also accumulates a Man4-GPI with its Etn-P on Man-1 (K. Willis and P. Orlean, un-published results), and human Gpi11 interacts with andstabilizes human Gpi13 (Hong et al. 2000).

GPI transfer to protein: Man4-GPIs bearing three Etn-Psare transferred to proteins with a C-terminal GPI signal-anchor sequence in a transamidation reaction in which theamino group of the Etn-P on Man-3 acts as nucleophile. Fiveessential membrane proteins are involved: Gaa1, Gab1,Gpi8, Gpi16, and Gpi17 (Hamburger et al. 1995; Benghezalet al. 1996; Fraering et al. 2001; Ohishi et al. 2000, 2001;Hong et al. 2003; Grimme et al. 2004). Gpi18 is catalyticbecause it resembles cysteine proteases and mutation ofpredicted active site residues eliminates its function (Meyeret al. 2000). The five transamidase subunits form a complexitself consisting of two subcomplexes: one containing Gaa1,Gpi8, and Gpi16, and the other, Gab1 and Gpi17 (Fraering

et al. 2001; Grimme et al. 2004; Zhu et al. 2005). Roles forthe noncatalytic subunits include recognition of the peptideand glycolipid substrates (Signorell and Menon 2009), and,in the case of Gab1 and Gpi8, possible interactions with theactin cytoskeleton (Grimme et al. 2004; File S4)

Remodeling of protein-bound GPIs: Following GPI transferto 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 moietyby lipase-related Bst1 (Tanaka et al. 2004; Fujita et al.2006a). Next, the sn-2 acyl chain of the diacylglycerol isremoved by the ER membrane protein Per1 to generatea lyso-GPI (Fujita et al. 2006b), whereupon a C26:0 acylchain is transferred to the sn-2 position by Gup1 in the ERmembrane (Bosson et al. 2006). Modifications of the GPIlipid by Bst1, Per1, and Gup1 are necessary for efficienttransport of GPI proteins from the ER to the Golgi (File S4).

Many GPIs are next remodeled by replacement of theirdiacylglycerol with ceramide by Cwh43 (Martin-Yken et al.2001; Ghugtyal et al. 2007; Umemura et al. 2007). Ceramideremodeling requires prior action of Bst1, and, because per1Dand gup1D strains show defects in remodeling, the exchangereaction likely takes place after the first three lipid modificationsteps. The mechanism could involve a phospholipase-like re-action that replaces diphosphatidic acid with ceramide phos-phate or diacylglycerol with ceramide (Ghugtyal et al. 2007;Fujita and Kinoshita 2010). Ceramide remodeling is notobligatory because certain GPI proteins, such as Gas1, reachthe plasma membrane with a diacylglycerol-based anchor(Fankhauser et al. 1993). Moreover, ceramide remodelingdoes not seem to be required for incorporation of GPI pro-teins into the wall (Ghugtyal et al. 2007).

Further GPI processing events may be the removal of theEtn-P moieties from Man-2 and Man-1. This is inferred fromthe fact that mammalian PGAP5, which removes the side-branching Etn-P from Man-2 (Fujita et al. 2009), has twohomologs in yeast: ER-localized Ted1 and Cdc1. Export ofGas1 is retarded in ted1D cells, and genetic interactionsconnect TED1 and CDC1 with processing and export ofGPI proteins (Haass et al. 2007). Because Etn-P side chainsare important for ceramide remodeling, they are likely re-moved after Cwh43 has acted.

Finally, a fifth, a1,2- or a1,3-linked Man can be added toMan-4 of protein-bound GPIs (Fankhauser et al. 1993).This modification is made to 20–30% of GPI proteins andoccurs in the Golgi, but none of the many Golgi Man-T seemsto be involved (Sipos et al. 1995; Pittet and Conzelmann2007). On reaching the plasma membrane, the GPIs onmany proteins become cross-linked to b1,6-glucan (see In-corporation of GPI proteins into the cell wall), and these GPI-CWP play structural or enzymatic roles in the wall (see CellWall-Active and Nonenzymatic Surface Proteins and TheirFunctions).

No individual GPI protein is essential in unstressedwild-type cells, so the lethality of mutations blocking GPI

788 P. Orlean

anchoring may be due to the collective effects of retardingER exit and plasma membrane or wall anchorage of multipleproteins. Consistent with this, temperature-sensitive GPIanchoring mutants grown at semipermissive temperaturehave aberrant morphologies and shed wall proteins into themedium (Leidich and Orlean 1996; Vossen et al. 1997).

Sugar nucleotide transport

GDP-Man transport: Cytoplasmically generated GDP-Manused by Golgi Man-T is transported into the Golgi lumen byVrg4/Vig4. GMP, generated from GDP formed in Man-Treactions by GDPase activity, serves as antiporter. Vrg4/Vig4 is essential, and vrg4 mutants are defective in manno-sylation of N- and O-linked glycans and mannosyl inositol-phosphoceramides (Dean et al. 1997; Abe et al. 1999).

Two homologous Golgi proteins, Gda1 and Ynd1, haveGDP-hydrolyzing activity. Gda1 has the highest activity to-ward GDP (Abeijon et al. 1989), and, consistent with GMP’srole as antiporter, rates of in vitro GDP-Man import intoGolgi vesicles from gdaD cells are fivefold lower than thoseof vesicles from wild-type cells (Berninsone et al. 1994).Ynd1 is a broader specificity apyrase (Gao et al. 1999) thathas a partially overlapping function with Gda1, and bothYnd1 and Gda1 are necessary for full elongation of N- andO-linked glycans (Gao et al. 1999; File S5).

Other sugar nucleotide transport activities: Transportactivities for UDP-Glc, UDP-GlcNAc, and UDP-Gal also occurin S. cerevisiae (Roy et al. 1998, 2000; Castro et al. 1999), andthere are eight more candidate transporters (Dean et al.1997; Esther et al. 2008) whose functions are unclear. UDP-Glc transport activity is present in the ER (Castro et al. 1999),and one possible need for it might be for a glucosylation re-action 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; FileS6). Hut1 is a candidate UDP-Gal transporter (Kainuma et al.2001), although galactose has not been detected on S. cere-visiae glycans. Both Hut1 and Yea4 may have broader speci-ficity and transport UDP-Glc (Esther et al. 2008).

Biosynthesis of Wall Components at the PlasmaMembrane

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 theplasma membrane although they originate from the roughER. The pathways for trafficking and activation of Chs2 andChs3 involve different sets of auxiliary proteins that ensurethe correct spatial and temporal localization of chitin syn-thesis during septation.

Septum formation: Factors determining the site at whicha bud will be formed, and the proteins that recruit andorganize the participants in septum formation, includingseptins and an actin–myosin contractile ring, are reviewedby Cabib et al. (2001), Cabib (2004), Roncero and Sanchez(2010), and Bi and Park (2012). Two chitin-containingstructures are made during bud emergence and septum for-mation (Figure 7). The first is a ring deposited in the wallaround the base of the emerging bud. This chitin is formedby 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 formedby centripetal synthesis of chitin by Chs2 in the neck regionbetween mother cell and bud (Shaw et al. 1991). Uponclosure, the septum separates the plasma membranes of

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 Gup1transfers a C26:0 acyl chain to the sn-2 position. Cwh43 can replace diphosphatidic acid with ceramide phosphate (shown here) or diacylglycerol withceramide. 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-linkedMan 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, andthe reducing end of the GPI remnant transferred to b1,6-glucan. Symbols are as used in Figure 1 and Figure 5.

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the two cells, accomplishing cytokinesis. In budding wild-type cells, the primary septum is thickened on both sides bydeposition of a secondary septum that normally containschitin, b1,3-glucan, b1,6-glucan, and covalently cross-linkedmannoprotein (Rolli et al. 2009), resulting in a three-layeredstructure (Shaw et al. 1991).

Chs2 and Chs3 have important roles in septation andcytokinesis although in the absence of Chs2 or Chs3, or in-deed of all three chitin synthases, cytokinesis can still takeplace. In chs2D mutants, the primary septum is missing, anda thick, amorphous septum is formed that contains chitinmade by Chs3 (Shaw et al. 1991; Cabib and Schmidt2003). chs3D mutants form a three-layered septum, butthe neck region between mother cell and bud is elongated(Shaw et al. 1991). chs2D chs3D and chs1D chs2D chs3Dstrains grow very slowly on osmotically supported medium(Sanz et al. 2004; Schmidt 2004; File S6). The triplemutants, however, acquired a suppressor mutation thateliminated the need for osmotic support and conferreda growth rate as fast as that of a chs2D mutant althoughover a third of suppressed and unsuppressed cells in a cul-ture were dead (Schmidt 2004).

For mother and daughter cells to separate, septalmaterial must be degraded, a process that results fromsecretion of chitinase Cts1 (Kuranda and Robbins 1991),endo-b1,3-glucanases Eng1/Dse4 and Scw11 (Cappellaroet al. 1998; Colman-Lerner et al. 2001; Baladron et al.2002; see Known and predicted enzymes), and possibly addi-tional activities from the daughter cell’s side of the septum.Daughter cell-specific expression of these enzymes is underthe control of the transcription factor Ace2 (Colman-Lerneret al. 2001).

Chitin synthase biochemistry: Chs1, Chs2, and Chs3 useUDP-GlcNAc as donor and are members of GT family 2 ofprocessive inverting glycosyltransferases, which includeshyaluronate and cellulose synthases. Yeast’s chitin synthasesare predicted to have three to five transmembrane helicestoward their C termini, and Chs3 likely has two more trans-membrane domains nearer its N terminus (Jimenez et al.2010; Merzendorfer 2011). Amino acid residues importantfor catalysis lie in a large cytoplasmic domain containingthe 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; Merzendorfer2011). An additional motif, (S/T)WG(X)T(R/K), predictedto be extracellularly oriented (Merzendorfer 2011), lies nearthe protein’s C terminus (Cos et al. 1998; Merzendorfer 2011).

The molecular mechanism of chitin synthesis is not yetclear. By analogy with bacterial NodC, which synthesizeschito-oligosaccharides, and with nonfungal chitin synthases,chain extension would be at the nonreducing end (Kamstet al. 1999; Imai et al. 2003). This topic, and the issue ofhow the synthases overcome the steric challenge that eachsugar in a b1,4-linked polymer is rotated by �180� relativeto its neighbor, are discussed further in File S6.

Chitin made in vitro by CS I or CS III contains, on aver-age, 115–170 GlcNAc residues (Kang et al. 1984; Orlean1987). Chitin synthases presumably make chitin chains witha range of lengths, and the range would be predicted to shiftto shorter chains as UDP-GlcNAc concentration drops belowKm, resulting in lowered rates of chain extension. Indeed,purified Chs1 and membranes from cells overexpressingChs2 make chito-oligosaccharides at low substrate concen-trations (Kang et al. 1984; Yabe et al. 1998). Chitin madein vivo is polydisperse (Cabib and Duran 2005), and in-creased chitin chain lengths are seen in fks1D and gas1Dmutants and CFW-treated cells, which mount the chitinstress response, whereas shorter chains were made ina strain expressing a Chs4 variant with lower in vitro CSIII activity (Grabinska et al. 2007). However, GlcN treat-ment, which stimulates chitin synthesis in vivo (Bulik et al.2003; see Sugar nucleotides), had little effect on polymerchain length (Grabinska et al. 2007).

S. cerevisiae’s three chitin synthases are all stimulated upto a few fold in vitro by high concentrations of GlcNAc

Figure 7 Roles of chitin synthases II and III in chitin deposition duringbudding growth. (A) Chitin synthase III synthesizes a chitin ring (blue)around the base of the emerging bud. (B) The plasma membrane inva-ginates and chitin synthase II synthesizes the primary septum (red). Nochitin 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 primaryseptum, and chitin synthase III starts synthesizing lateral wall chitin inthe bud (blue). (D) After cell separation, the bud scar (which is formedfrom the chitin ring made by Chs3), most of the primary septum made byChs2, as well as secondary septal material deposited on the mother cellside, remain on the mother cell. The birth scar on the daughter cellcontains residual chitin from the primary septum as well as secondaryseptal material. (E and F) Chitinase digestion of the primary septum fromthe daughter-cell side facilitates cell separation, and lateral wall chitinsynthesis continues as the daughter cell grows. Figure is adapted fromCabib and Duran (2005).

790 P. Orlean

(Sburlati and Cabib 1986; Orlean 1987). Possible explana-tions are that GlcNAc serves as a primer or allosteric activa-tor 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 detectablein vitro only after pretreatment of membranes or extensivelypurified Chs1 with trypsin (Duran and Cabib 1978; Kanget al. 1984; Orlean 1987). Proteolytically activated Chs1has the highest in vitro activity of the chitin synthasesassayed in membranes from wild-type cells (Sburlati andCabib 1986; Orlean 1987), although Chs1 does not contrib-ute measurably to chitin synthesis in vivo, even in the ab-sence of Chs2 and Chs3 (Shaw et al. 1991). Although trypsinactivation may mimic the effect of an endogenous activatingprotease, neither such an activator, nor an active, processedform of Chs1, have been identified.

Levels of protease-elicited Chs1 activity are the same inmembranes from logarithmically growing and stationary-phase cells (Orlean 1987), and levels of Chs1 show littlechange during the cell division cycle (Ziman et al. 1996).CHS1 transcription and in vitro CS I activity increase in re-sponse to mating factors, but elevated in vitro activity isdetectable only after trypsin activation (Schekman andBrawley 1979; Orlean 1987; Appeltauer and Achstetter1989). However, Chs1 does not contribute to pheromone-induced chitin synthesis (Orlean 1987).

chs1D cultures contain the occasional lysed bud, a phenotypemore pronounced in acidic medium but partially alleviatedwhen Cts1 chitinase is also deleted (Cabib et al. 1989). Twoexplanations, which are not mutually exclusive, are that Chs1may repair wall damage due to overdigestion of chitin by Cts1or that Chs1 participates in septum synthesis and makes chitinduring growth in acidic medium (Cabib et al. 1989; Bulawa1993). Chs1 promotes wall association of at least one proteinbecause small amounts of the GPI protein Gas1 are releasedinto the medium from chs1D cells (Rolli et al. 2009).

Although the contribution of Chs1 to chitin synthesis issmall, a wider role for the protein emerged from an analysisof the networks of genes that interact synthetically withCHS1 and CHS3 (Lesage et al. 2005). Most of the 57 genesin the CHS1 interaction network fell into two sets. One setcontained genes that, when mutated, impact cell integrity orthat themselves interact with genes involved in b1,3-glucansynthesis, indicating a role for Chs1 in buffering the wallagainst changes impacting its robustness. The other set con-tained genes involved in budding and in endocytic proteinrecycling, which in turn may impact Chs2 function, suggest-ing that Chs1 also buffers against deficiencies in Chs2. TheCHS1-interacting genes were mostly distinct from the genesin the network that impacts Chs3 function, and, moreover,mutations in CHS1 itself or in the genes in the CHS1 inter-action set do not trigger the Chs3-dependent chitin stressresponse. Chs1 and Chs3 therefore have distinct functionsand one does not buffer against defects in the other (Lesageet al. 2005).

Chitin synthase II and proteins impacting its localizationand activity: Chs2 makes no more than 5% of the chitin inbudding cells. Activity of endogenous Chs2 is detectableonly in membranes from growing cells and can be stimu-lated by treatment with trypsin (Sburlati and Cabib 1986)although, in some studies, membrane preparations as wellas partially purified Chs2 have significant in vitro activitywithout prior trypsin treatment, raising the possibility thatfull-size Chs2 makes chitin (Uchida et al. 1996; Oh et al.2012). A soluble fraction from growing yeast cells, whichstimulates Chs2 activity two- to fourfold but which must itselfbe pretreated with trypsin, has been described (Martínez-Rucobo et al. 2009). An endogenously activated, processedform of Chs2 has not been identified (File S6).

Levels of CHS2 expression and localization of the proteinare coordinated with synthesis of the primary septum (Fig-ure 8). CHS2 message levels peak just prior to primary sep-tum formation at the G2/M phase (Pammer et al. 1992; Choet al. 1998; Spellman et al. 1998), and levels of Chs2 and CSII activity then peak as the primary septum is made (Pammeret al. 1992; Choi et al. 1994a; Chuang and Schekman 1996).Upon completion of cytokinesis, levels of Chs2 and its mes-sage drop, indicating that both turn over rapidly.

Temporal and spatial localization of Chs2 is impacted atat least two stages by protein kinases. Chs2 is synthesized inthe ER during metaphase, but its release from the ER iscoordinated with exit of the cell from mitosis and triggeredupon inactivation of mitotic kinase by Sic1 (Zhang et al.2006). The mitotic kinase Cdk1 likely acts directly onChs2, which contains four CDK1 phosphorylation sites nearits N terminus, because mutation of the target Ser residuesto Glu leads to retention of Chs2 in the ER, whereas chang-ing the serines to Ala leads to constitutive release of themutant Chs2 even in the presence of high Cdk1 activity(Teh et al. 2009). Timed release of Chs2 from the ER afterchromosome separation and exit of the cells from mitosis istriggered by dephosphorylation of the Cdk1 sites by theCdc14 phosphatase, the terminal component of the mitoticexit network (MEN) cascade (Chin et al. 2012).

Exit of Chs2 from the ER and its delivery to the plasmamembrane at the mother cell–bud junction is effected byCOPII vesicles (Chuang and Schekman 1996; VerPlankand Li 2005; Zhang et al. 2006). Localization of Chs2 atthe bud neck, correct formation of the primary septum,and removal of Chs2 at the end of cytokinesis depend onphosphorylation of Chs2 by the mitotic exit kinase Dbf2, alsoa component of MEN (Oh et al. 2012). Inn1 and Cyk3,whose localization to the division site is also regulated byMEN, are also involved in activation of Chs2 for primaryseptum formation (Nishihama et al. 2009; Meitinger et al.2010; Oh et al. 2012). Overexpression of CYK3 leads to in-creased deposition of chitin at the division site in chs1Dchs3D 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 onChs2 is unclear and Inn1’s role in Chs2 activation is unknown.

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Additional phosphorylation sites are present in Chs2’s N-ter-minal domain (Martínez-Rucobo et al. 2009), but their rolesare unclear.

Chs2 resides at the site of primary septum formation foronly 7–8 min (Roh et al. 2002a; Zhang et al. 2006). Theprotein is degraded upon endocytosis and delivery to thevacuole (Chuang and Schekman 1996; Schmidt et al.2002; VerPlank and Li 2005), and optimal endocytic turnoverof Chs2 requires components of the endosomal sorting com-plexes required for transport (ESCRT) pathway (McMurrayet al. 2011).

Chitin synthase III and proteins impacting its localizationand activity: Chitin synthase III is responsible for thesynthesis of .90% of the chitin in unstressed vegetativecells, for the additional chitin made in the chitin stress re-sponse and in response to mating pheromones, and for thesynthesis of the chitin that is de-N-acetylated to chitosanduring ascospore wall formation. Cells deficient in CS IIIactivity are resistant to CFW (Roncero et al. 1988). Chs3 isthe transferase, but its function depends on its regulatedtransport from the ER to the plasma membrane, its removalfrom the plasma membrane and sequestration in intracellu-lar vesicles called chitosomes, and its remobilization fromchitosomes to the plasma membrane. A number of proteinsare required for regulated Chs3 trafficking and for enzymeactivity (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 frac-tions from logarithmically growing wild-type cells withoutprior treatment with trypsin and is trypsin sensitive (Orlean1987). CS III activity determined in this way is presumablyeither due to constitutively active Chs3 or to an endogenouslyactivated form of the protein. A pool of trypsin-activatableCS III was detected in detergent-treated membranes fromchs1D chs2D cells or from cells lacking Chs4, an activatorof CS III (Choi et al. 1994b; Trilla et al. 1997). The latterfinding, together with the observation that overexpression ofChs4 lowers the extent to which trypsin activates CS III, sug-gested that trypsin treatment might mimic Chs4-dependentprocessing of Chs3. However, because no endogenously pro-cessed forms of Chs3 have been detected (Santos andSnyder 1997; Cos et al. 1998), and because Chs4 does notresemble any protease, the apparent zymogenicity of Chs3in chs4D may be an artifact (Reyes et al. 2007).

Levels of CHS3 mRNA and Chs3 vary little during thebudding cycle (Choi et al. 1994a; Chuang and Schekman1996; Cos et al. 1998), indicating that CS III is regulated atthe post-translational level. Sequences involved include theC-terminal extracellular region containing the motif (S/T)WG(X)T(R/K), which is required for in vitro CS III activityand chitin synthesis in vivo (Cos et al. 1998).

A number of proteins interact with Chs3 as it transits thesecretory pathway (Figure 9). Chs3 is palmitoylated in theER by Pfa4 (Lam et al. 2006; Montoro et al. 2011). pfa4Dmutants are CFW-resistant and accumulate Chs3 in the ER,indicating a role for palmitoylation in Chs3 export (Lam et al.

2006). Chs3 has two palmitoylation sites in a cytoplasmic do-main N-terminal to the proposed catalytic residues (Meissneret al. 2010). Exit of Chs3 from the ER also requires Chs7, anER chaperone with six or seven transmembrane domains(Trilla et al. 1999) that interacts with Chs3. The effects ofCHS7 deletion on chitin levels and CSIII activity are almostas severe as those of CHS3 deletion (Trilla et al. 1999). Chs3aggregates in the ER in chs7D cells (Lam et al. 2006), andChs7 is a limiting factor in export of Chs3 because simulta-neous overexpression of CHS3 and CHS7 leads to elevatedCSIII activity, whereas overexpression of CHS3 alone does not.Neither Pfa4 nor Chs7 is required for exit of Chs1 and Chs2from the ER (Trilla et al. 1999; Lam et al. 2006). ER-membraneproteins Rcr1 and Yea4 also impact Chs3-dependent chitinsynthesis in ways that are unclear (File S6).

Transport of new Chs3 from the trans-Golgi to the plasmamembrane, as well as Chs3 cycling from chitosomes to theplasma membrane, requires the peripheral Golgi membraneproteins Chs5 and Chs6 (Santos and Snyder 1997; Santos

Figure 8 Trafficking and regulation of Chs2. Cell cycle-regulated expres-sion of CHS2 peaks at the G2-M phase transition, and Chs2 is synthesizedat the ER. Phosphorylation of Chs2 by Cdk1 retains Chs2 in the ER. Uponchromosomal separation, Cdc14-dependent dephosphorylation of Chs2allows release of the protein from the ER and its transit to the mothercell–bud junction. Inn1 and Cyk3, localized at the division site, are in-volved in Chs2 activation. After primary septum formation is complete,Chs2 is endocytosed and degraded. Localization, function, and subse-quent removal of Chs2 when the primary septum is complete dependon phosphorylation by Dbf2. Figure is adapted from Lesage and Bussey(2006).

792 P. Orlean

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 transientlybind Chs3 to promote its incorporation into secretoryvesicles (Sanchatjate and Schekman 2006; Trautwein et al.2006; Wang et al. 2006). Although Chs5 and Chs6 act ina complex, the two have different impacts on Chs3 activityand transport. chs5D and chs6D mutants make 25 and 10%of wild-type amounts of chitin, respectively, but whereaschs5D membranes lack in vitro CS III activity, this activityis normal in chs6D membranes (Bulawa et al. 1993; Santoset al. 1997). This may be because, in chs5D cells, Chs3 accu-mulates in late Golgi vesicles (Santos and Snyder 1997),whereas, in chs6D mutants, it collects in chitosomes, whereit 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 Chs3and Crh2 would ensure colocalization of these proteins forefficient cross-linking of chitin to b1,3-glucan.

At the plasma membrane, Chs4 (Csd4/Skt5) interactswith Chs3 (DeMarini et al. 1997; Ono et al. 2000; Meissneret al. 2010) and has two roles apparently specific to Chs3.chs4D mutants lack in vitro CS III activity and make verylittle chitin (Bulawa 1993; Trilla et al. 1997). Overexpres-sion of CHS4, but not CHS3, raises in vitro CS III activity(Bulawa 1993; Trilla et al. 1997; Ono et al. 2000) as well aslevels of Chs3 in the plasma membrane (Reyes et al. 2007),suggesting that Chs4 is an activator of CS III. Stimulation ofCS III by Chs4 requires a region of Chs4 to bind Chs3 be-cause the ability of truncated forms of Chs4 to elicit CS IIIactivity correlates with the ability of Chs4 fragments to in-teract with Chs3 in a two-hybrid analysis (Ono et al. 2000;Meissner et al. 2010). Chs4 has a C-terminal farnesylationsite (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 Chs3localization on the mother cell’s plasma membrane at thesite of formation of the chitin ring prior to bud emergence.There, it interacts with the scaffold protein Bni4, whichin turn associates with the septins (DeMarini et al. 1997;Kozubowski et al. 2003; Sanz et al. 2004). Absence of Bni4leads to mislocalized deposition of chitin (DeMarini et al.1997; Kozubowski et al. 2003; Sanz et al. 2004), and Chs4is absent from the base of buds in small-budded cells (Sanzet al. 2004). In contrast to chs4D mutants, chitin synthesisand 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 membranejust before formation of the chitin ring at the site of budemergence and reside there in a ring at the base of thebud in many cells with small buds, then become scarcelydetectable in cells with medium-sized buds, only to reap-pear, in a Bni4-independent manner, at both sides of theneck in cells with large buds prior to cytokinesis (Chuangand Schekman 1996; DeMarini et al. 1997; Santos andSnyder 1997; Kozubowski et al. 2003; Sanz et al. 2004).In between, Chs3 is retrieved from the membrane to chito-somes 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 Chs6-dependent manner (Ziman et al. 1998; Wang et al. 2006).Chs3’s itinerary is consistent with the overall order of eventsin yeast cytokinesis.

Chitin synthesis in response to cell wall stress: Cells withmutations affecting the formation of b-glucan, mannan,O-linked glycans, and GPI anchors respond by depositingadditional chitin—as much as 10 times more than in wild-type cells—in their lateral walls in apparent compensationfor compromised cell integrity (Gentzsch and Tanner 1996;Kapteyn et al. 1997, 1999a; Popolo et al. 1997; Dallies et al.

Figure 9 Overview of Chs3 trafficking. Chs3, synthesized in the ER,requires palmitoylation by Pfa4 and association with Chs7 to exit theER. In the trans-Golgi, Chs3 association with exomer components Chs5and Chs6 facilitates incorporation of Chs3 into secretory vesicles for de-livery to the plasma membrane at the site of chitin ring formation. Local-ization and activation of Chs3 depends on association with Chs4, whoseassociation with the septin ring is mediated in turn via an interaction withBni4. In cells with medium-sized buds, Chs3 is retrieved from the plasmamembrane and sequestered in chitosomes in an endocytic processdepending on End4 and later recruited back to the neck region ina Chs6-dependent manner. During the cell wall stress response, Rho1and Pkc1 trigger mobilization of Chs3 from chitosomes to the plasmamembrane for synthesis of extra chitin in the lateral wall. Figure is adap-ted from Lesage and Bussey (2006).

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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; Lesageet al. 2005). This chitin stress response, which is accompa-nied by increased precursor supply (see Precursors and Car-rier 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 theCHS genes, but, rather, an altered distribution of Chs3,which was seen in the plasma membrane of buds of gas1Dand fks1D cells, and Chs4 was also delocalized (Garcia-Rodriguez et al. 2000; Valdivieso et al. 2000; Carotti et al.2002; Valdivia and Schekman 2003). Interestingly, gas1Dsuppressed the lysed bud phenotype of chs1D, suggestingthat the chitin stress response also repaired weakened budwalls (Valdivieso et al. 2000). The Chs3 making the stressresponse originates from chitosomes, and its translocation tothe plasma membrane is regulated by Rho1 and Pkc1, whichact early in the CWI pathway that triggers the chitin stressresponse (Valdivia and Schekman 2003).

Chitin synthase III in mating and ascospore wall forma-tion: Chitin synthase III is responsible for the extra chitinmade in response to mating pheromones and for formationof the chitosan of ascospore walls. MATa cells treated witha-factor show a three- to fourfold increase in chitin, which islaid down diffusely in the shmoo (Schekman and Brawley1979). Chs3 is necessary because no extra chitin is made inpheromone-treated chs3D cells, and the response is eitherabolished or much smaller in chs5D, chs6D, and chs4D cells,indicating that the machinery for trafficking and activationof 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 localizedat the periphery of the mating projection, and it remainsthere because it is not subject to endocytic turnover as it isin budding cells (Santos and Snyder 1997; Sacristan et al.2012). Although the extra chitin synthesis in response toa-factor is presumably driven by the increased amountof UDP-GlcNAc made during the pheromone response(Orlean et al. 1985; Bulik et al. 2003), the mechanism be-hind pheromone-stimulated chitin synthesis by Chs3 is unclear.Levels of Chs3 increase sixfold upon a-factor treatment (Coset al. 1998), but neither CHS3 transcription nor levels ofchitin synthase III activity are elevated (Orlean 1987; Choiet al. 1994a). Factors that might limit total Chs3 activitymight include prevention of the mobilization of the proteinto the plasma membrane in shmoos or interference withinteractions between Chs3 and regulatory proteins (Choiet al. 1994a).

The chitosan of the ascospore wall is initially synthesizedas chitin by Chs3 (Pammer et al. 1992) and is then de-N-acetylated by chitin deacetylases Cda1 and Cda2, of which Cda2has the dominant role (Mishra et al. 1997; Christodoulidouet al. 1999). From the sporulation defects in mutants in proteinsinvolved in Chs3 trafficking in vegetative cells, Chs6 and Chs7,

but not Chs5, have as-yet-undefined roles in ascospore matura-tion (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 arereviewed by Neiman (2011).

b 1,3-Glucan

De novo b1,3-glucan synthetic activity is associated withmembers of the Fks family, of which Fks1 and Fks2 requirethe soluble Rho1 GTPase as a regulatory subunit. In vitroactivity is membrane-associated, uses UDP-Glc as donor, isstimulated by GTP via Rho1, and yields a product with achain length of 60–80 glucoses (Shematek et al. 1980; Kangand Cabib 1986; Drgonová et al. 1996; Mazur and Baginsky1996; Qadota et al. 1996). In vitro b1,3-glucan synthaseactivity is inhibited by acylated cyclic hexapeptides of theechinocandin group and by papulocandins, acylated deriva-tives 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 alsocontains proteins implicated in callose sythesis in plants(Verma and Hong 2001). Fks1 has an N-terminal cytoplas-mic domain that is followed by 6 transmembrane helices,a large cytoplasmic domain, and then 10 transmembranehelices (Inoue et al. 1995; Mazur et al. 1995; Qadota et al.1996; Dijkgraaf et al. 2002; Okada et al. 2010). Three func-tional domains, mutations in which separately affect in vivoand in vitro b1,3 glucan synthetic activity, as well as cellpolarity and endocytosis, have been distinguished (Okadaet al. 2010; File S7). The phenotypes of fks1 mutants mayin part reflect the involvement of the protein in processesother than b1,3-glucan biosynthesis. For example, muta-tions in both FKS1 and FKS2 result in lowered b1,6-glucansynthesis (Dijkgraaf et al. 2002). Fks1 is localized to theplasma membrane at sites of polarized growth and cell wallremodeling throughout the cell cycle, and this localizationcoincides with that of actin patches (Qadota et al. 1996;Dijkgraaf et al. 2002; Utsugi et al. 2002). Fks1 transits thesecretory pathway because it accumulates intracellularly invesicular transport mutants and its activity is sensitive tophytosphingosine levels in the ER (El-Sherbeini and Clemas1995; Abe et al. 2001; File S7).

Roles of the Fks proteins in b1,3-glucan synthesis: The Fksproteins show a degree of specialization. Deletion of FKS1leads to slow growth, a 75% reduction in b1,3-glucan, andlow in vitro b-1,3-glucan synthase activity, whereas thein vitro activity of fks2D membranes is nearly that of wild-type membranes and the disruptants have no defect in veg-etative growth (Inoue et al. 1995; Mazur et al. 1995). Al-though this suggests that Fks1 is the major contributor tob1,3-glucan synthesis in budding cells, fks1D fks2D nullmutants are inviable, indicating that Fks1 and Fks2 haveoverlapping functions (Inoue et al. 1995; Mazur et al.

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1995). Consistent with this, overexpression of either FKS1or FKS2 can partially correct the defects caused by deletingthe other of the two genes (Mazur et al. 1995; Dijkgraafet al. 2002), and, furthermore, the two proteins colocalizein sites of polarized growth in budding cells, although Fks1is the most abundant (Dijkgraaf et al. 2002).

FKS1 and FKS2 show different expression patterns. FKS1is expressed during budding growth and transcript levelspeak in the late G1 and early S phases (Mazur et al. 1995;Ram et al. 1995; Lesage and Bussey 2006). FKS2 mRNA, incontrast, cannot be detected in budding cultures grown inglucose, but appears when glucose becomes depleted; whencells are grown on acetate, glycerol, or galactose; when cellsare treated with a-factor or Ca2+; in fks1 mutants andmutants 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 Bussey2006). Induction of FKS2 is mediated via the PKC CWIand 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 dip-loids have a severe defect in this process (Mazur et al. 1995;Huang et al. 2005; Ishihara et al. 2007). Homozygous fks3Dfks3D diploids also form abnormal spores, indicating a rolefor Fks3 in ascopore wall formation, although Fks3’s role insporulation does not overlap with Fks2’s. It was proposedthat Fks2 is primarily responsible for synthesis of b1,3-glucanin the ascospore wall and that Fks3, rather than functioningas a synthase, modulates glucan synthesis during ascosporewall formation (Ishihara et al. 2007; File S7).

After their export through the plasma membrane, b1,3-glucan chains can be cross-linked to chitin by Crh1 and Crh2(Cabib 2009), and the polymer can be extended through theaction of Gas1 family b1,3-glucanosyltransferases (Mouynaet al. 2000), and side-branching b1,6-linked glucoses as wellas PIR proteins may be attached (Ecker et al. 2006; see In-corporation 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-dependentchitin synthesis (Garcia-Rodriguez et al. 2000; Valdiviesoet al. 2000; Carotti et al. 2002), and fks1D shows syntheticinteractions with chs3D, chs4D, chs5D, chs6D, and chs7D(Osmond et al. 1999; Lesage et al. 2004), but correct syn-thesis of other wall constituents is also necessary whenb1,3-glucan synthesis is compromised (Lesage et al.2004). Analyses of the genome-wide responses to FKS1deletion revealed upregulation of a “cell wall compensa-tory cluster” of 79 coregulated genes whose products in-clude a range of proteins involved in wall synthesis andremodeling (Terashima et al. 2000; Lagorce et al. 2003).An overlapping set of genes, whose products function inthe biosynthesis of chitin, b1,6-glucan, and mannan, aswell as in the function of the secretory pathway and inmaintenance of cell polarity, was identified in an analysisof 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, consis-tent with differential expression of FKS2 and FKS3 (Lesageet al. 2004).

Rho1 GTPase, a regulatory subunit of b1,3-glucan synthase:The essential Rho1 GTPase, which activates Pkc1 in the CWIpathway and is required for cell cycle progression and po-larization of growth (Drgonová et al. 1999; Levin 2011), hasa distinct role as a regulatory subunit of b1,3-glucan syn-thetic complexes containing Fks proteins. Evidence for this isthat (i) Fks1 and Rho1 colocalize and coimmunoprecipitate,(ii) membranes from a temperature-sensitive rho1 mutanthave a thermolabile b1,3-glucan synthase activity that canbe corrected by adding back purified Rho1, (iii) membranesfrom cells expressing a consitutively active rho1 allele haveGTP-independent b1,3-glucan synthase activity, and (iv) in-activation of Rho1 by ADP ribosylation eliminates thein vitro b1,3-glucan synthase activity of membranes fromfks1D and fks2D strains (Drgonová et al. 1996; Mazur andBaginsky 1996; Qadota et al. 1996). Moreover, there arerho1 mutations that affect regulation of b1,3-glucan synthe-sis, but not other Rho1 functions, and the amino acids af-fected are different from those whose mutation causes cellcycle and polarization defects (Saka et al. 2001; Roh et al.2002b). The amino acid changes in the b1,3-glucan synthesis-specific rho1 mutants might impact binding to Fks proteins,but the interacting domains on the regulatory and catalyticsubunits have not been defined. The Rho1-Fks interaction atthe cytoplasmic face of the plasma membrane, as well asactivation of b1,3-glucan synthesis, requires Rho1 to be ger-anylgeranylated at its C terminus (Inoue et al. 1999).

b1,6-Glucan

Mutations in genes with products localized along the secretorypathway impact formation of b1,6-glucan (Shahinian andBussey 2000; Lesage and Bussey 2006), but the biochemistryof b1,6-glucan synthesis is unclear. In vitro synthesis of b1,6-glucan is hard to detect, and no fungal enzyme has yet beenshown to catalyze formation of a b1,6-glucosidic linkage usingUDP-Glc as donor, although the linkage can be generated bythe 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 thispolymer (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 brancheson average every fifth Glc (see b-glucans), it could be gen-erated by a processive, UDP-Glc-dependent b1,6-glucan syn-thase and then branched or by assembly of shorter repeatunits, whose glucoses originate from UDP-Glc. Detection ofUDP-Glc-dependent formation of b1,6-glucan is complicatedby the fact that UDP-Glc is also the donor in the synthesis ofb1,3-glucan, glycogen, and glucolipids.

S. cerevisiae Cell Wall 795

Two assays of the formation of b1,6-glucan using UDP-Glc as donor have been described. In the first, formation ofb1,6-glucanase-sensitive polymer by membranes was de-tected by dot-blot assay using an anti-b1,6 glucan antibody(Vink et al. 2004). The reaction was distinguished fromb1,3-glucan synthase because membranes from kre5 mu-tants, which make little b1,6-glucan but have normalb1,3-glucan synthetic capability, made little b1,6-glucanin vitro but had nearly wild-type b1,3-glucan synthase ac-tivities. Comparisons of the activities of wild-type and b1,6-glucan synthesis-defective strains revealed that levels ofb1,6-glucan formed de novo correlated with the reductionin b1,6-glucan synthesis in vivo. It was proposed that thedot-blot assay measured b1,6-glucan chain extension andthat higher rates of Glc transfer reflected the presence ofmore acceptor (Vink et al. 2004). The reaction was stimu-lated by GTP and higher b1,6-glucan synthetic activity wasdetected in membranes from cells overexpressing Rho1GTPase, suggesting that b1,6-glucan synthase, like b1,3-glucansynthase, is Rho1-dependent (Vink et al. 2004).

In the second approach, formation of b1,6-glucan wasmeasured in cells permeablized by osmotic shock and incu-bated with radiolabeled UDP-Glc (Aimanianda et al. 2009).The insoluble, radiolabeled b1,6-glucan formed was chem-ically identical to the branched b1,6-linked glucan isolatedfrom cell walls, and radioactivity was distributed throughoutthe in situ product, indicating that de novo polymerization ofb1,6-glucan had occurred (Aimanianda et al. 2009). Consis-tent with their severe in vivo defects in b1,6-glucan synthe-sis, permeabilized kre5 and kre9 mutants showed no in situb1,6-glucan synthetic activity, but made b1,3-glucan. Theb1,6-glucan synthetic activity in permeabilized cells wasnot stimulated by GTP. However, because b1,3-glucan syn-thesis mutants make less b1,6-glucan, and vice versa, for-mation of the two polymers may be coordinated in anotherway (Dijkgraaf et al. 2002; Aimanianda et al. 2009; see TheFks family of b1,3-glucan synthases).

Proteins involved in b1,6-glucan assembly

Mutants defective in b1,6-glucan synthesis were identifiedin screens for resistance to K1 killer toxin, which uses b1,6-glucan as its receptor (Hutchins and Bussey 1983), and inscreens for CFW sensitivity (Ram et al. 1994; Lussier et al.1997b; Orlean 1997; Shahinian and Bussey 2000; Pagéet al. 2003). In these mutants, levels of alkali-insoluble b1,6-glucan were lowered to different extents, and the propor-tions of b1,6- and b1,3-linked Glc residues in the alkali-insoluble glucan fraction were often altered. The finding thatthe proteins implicated in b1,6-glucan assembly were local-ized in the ER, Golgi, or plasma membrane, together withdemonstrations of epistasis relationships and genetic interac-tions, led to the notion of a secretory pathway-based pathwayfor b1,6-glucan elaboration (Boone et al. 1990; reviewed byOrlean 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

are indirect. Proteins affecting the formation of b1,6-glucanwill be discussed in the order of their location along thesecretory pathway.

ER proteins: Homologs of the UGGT/calnexin protein qualitycontrol machinery: Four homologs of proteins involved in theUGGT/calnexin protein quality control system (see N-glycanprocessing in the ER and glycoprotein quality control) are re-quired for formation of normal amounts of b1,6-glucan(Jiang et al. 1996; Abeijon and Chen 1998; Shahinianet al. 1998; Simons et al. 1998). These are diverged UGGThomologs Kre5, Gls1/Cwh41, Gls2/Rot2, and Cne1, ofwhich Kre5 has the most important role because kre5mutants make no more than 5% of normal amounts of b1,6-glucan (Meaden et al. 1990; Montijn et al. 1999; Levinsonet al. 2002; Aimanianda et al. 2009). The contributions ofthe glucosidases and calnexin are likely indirect ones inmaintaining normal levels of unknown components of theb1,6-glucan assembly machinery in the secretory pathway(Shahinian et al. 1998; Lesage and Bussey 2006). The es-sential function of Kre5 is other than as a UGGT in proteinquality control because kre5D remained lethal in an alg8Dgls2D background in which all N-glycans stayed monoglu-cosylated, thereby bypassing the need for UGGT activity(Shahinian et al. 1998). Kre5 could be a glucosyltransferasewith a specialized role in quality control of b1,6-glucanassembly proteins (Levinson et al. 2002; Herrero et al.2004; Lesage and Bussey 2006), or it could glucosylatethe GPI glycan of future GPI-CWPs to generate a signalor attachment point for subsequent transfer to b1,6-glucan(Shahinian and Bussey 2000). S. cerevisiae has the neces-sary ER UDP-Glc transport activity to supply the donor(Castro et al. 1999).

N-glycosylation is important for wild-type levels of b1,6-glucan to be made. For example, stt3 mutants have a severedefect in b1,6-glucan synthesis and are synthetically lethalwith kre5 and kre9 (Chavan et al. 2003b). This may reflecta requirement for N-glycosylation of one or more b1,6-glu-can synthetic proteins or for an N-glycan to serve as acceptorfor initiation of a b1,6-glucan chain (Lesage and Bussey2006). Interestingly, mutations such as och1 andmnn9, whichaffect 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 main-tained between these two polymers.

Fungus-specific ER chaperones required for b1,6-glucansynthesis: Mutations in genes encoding the ER-localized,fungus-specific membrane proteins Rot1, Big1, and Keg1 allcause a b1,6-glucan synthetic defect. ROT1 and BIG1 nullmutants grow only with osmotic support, and even thenvery slowly, and in this and in the severity of their b1,6-glucan synthetic defect—a 95% reduction—they resemblekre5D strains (Bickle et al. 1998; Azuma et al. 2002; Machiet al. 2004). Levels of b1,3-glucan and chitin are elevated inrot1D and big1D. The b1,6-glucan defect in keg1D cells is

796 P. Orlean

similar to that in kre6D—about a 50% reduction (Nakamataet al. 2007).

Rot1, Big1, and Keg1 are small proteins that show nosimilarity to one another or to carbohydrate-active enzymes(Lesage and Bussey 2006). They seem to function as ERchaperones with varying degrees of importance for the stabil-ity of proteins involved in b1,6-glucan synthesis and may insome cases cooperate. Observations supporting this notionand indicating a relationship to Kre5 are discussed in File S8.

More widely distributed secretory pathway proteins: Kre6and Skn1: Kre6 and Skn1 are homologous type 2 membraneproteins in GH family 16 of b-1,6/b-1,3-glucanases (Henrissatand Davies 1997; Montijn et al. 1999). kre6D cells make halfnormal amounts of b1,6-glucan, whereas skn1D cells makeb1,6-glucan normally and have no growth defect. Expressedat high copy, Skn1 restores almost normal levels of b1,6-glucan to kre6D cells, and kreD skn1D double mutants areinviable or very slow growing, depending on the strain back-ground, and make no more than 10% of normal amounts ofb1,6-glucan (Roemer et al. 1993). From this, Kre6 and Skn1seem to be functional homologs, with Kre6 normally havingthe dominant role in b1,6-glucan synthesis. As hydrolases ortransglycosylases, Kre6 and Skn1 could act on a structurethat serves as a precursor or acceptor in elaboration of b1,6-glucan or on a glycoprotein involved in b1,6-glucan synthe-sis (Lesage and Bussey 2006), but enzyme activity has yet tobe demonstrated for these proteins. Much of Kre6 is ER-localized, where it interacts with Keg1, but the protein isalso detectable in the Golgi, in secretory vesicles, and atthe plasma membrane at sites of polarized growth (Liet al. 2002; Nakamata et al. 2007; Kurita et al. 2011; FileS8). Localization of Skn1 has not been explored in detail.Skn1 also has a role in the formation of mannosyl diinosi-tolphosphoryl ceramide [M(IP)2C], because skn1D, but notkre6D strains, is defective in M(IP)2C (Thevissen et al. 2005;File S8).

Kre6 has been implicated in the mode of action of a pyr-idobenzimidazole derivative identified in a screen for inhib-itors of cell wall incorporation of a reporter GPI-CWP(Kitamura et al. 2009). Because cells treated with this com-pound showed lowered incorporation of radiolabeled Glcinto a b1,6-glucan fraction, and because a resistant mutanthad an amino acid substitution in Kre6, it was proposed thatthe compound is an inhibitor of b1,6-glucan synthesis andthat Kre6 is its likely target (Kitamura et al. 2009).

Kre9 and Knh1: Kre9 and Knh1 are 30 kDa, soluble, fun-gus-specific, O-mannosylated proteins that are secreted intothe 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 slowgrowing and show an 80% reduction in b1,6-glucan, andthe residual b1,6-glucan in them has about half the molec-ular mass as the wild-type polymer and is altered in its pro-portion 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 andb1,6-glucan defects of kre9D, but the kre9 skn1 combinationis synthetically lethal (Dijkgraaf et al. 1996). kre9D, but notknh1D, is also synthetically lethal with kre1D (see below)and kre6D, but not with skn1D. KRE9’s genetic interactionsindicate that its product has a pleiotropic impact on b1,6-glucan formation, although its effects must be exerted afterKre5’s because the kre5D kre9D double null has the samephenotype as kre5D (Dijkgraaf et al. 1996; Shahinian andBussey 2000). Neither Kre9 nor Knh1 shows similarity toproteins of known function. If they are not enzymes, Kre9and Knh1 may serve to anchor b1,6-glucan in the cell wall(Lesage and Bussey 2006), but this must be reconciled withthe finding that kre9 mutants have no UDP-Glc-dependentb1,6-glucan synthetic activity (Aimanianda et al. 2009).

Plasma membrane protein Kre1: Kre1, a GPI protein, func-tions at the plasma membrane or in the wall. kre1D cellsmake 40% of wild-type levels of b1,6-glucan, but this glucanis smaller and its b1,3 side branches are not extended(Boone et al. 1990; Roemer and Bussey 1995). GPI attach-ment is necessary for Kre1’s function and cell surface local-ization (Breinig et al. 2004). The hydrophilic portion of Kre1shows no similarity to known enzymes. Kre1 has a structuralrole and becomes cross-linked to other wall proteins (Breiniget al. 2004), and it also serves as a receptor for K1 killertoxin (File S8).

How might b1,6-glucan be made?: Obstacles to identifyingthe b1,6-glucan synthase gene might be an inherent difficultyin obtaining hypomorphic alleles of an essential synthasegene or the existence of multiple redundant synthase geneswhose individual mutation gives no phenotype (Lesage andBussey (2006). Furthermore, there are no precedents in otherorganisms that could be exploited in bioinformatics-basedapproaches to b1,6-glucan synthesis. Although b1,6-glucanis widely distributed in the Fungi (Lesage and Bussey2006), it is very rare elsewhere. The bacterium Actinobacillussuis makes a lipopolysaccharide containing a b1,6-glucan ho-mopolymer (Monteiro et al. 2000), but the proteins involvedin its formation are unknown. Some bacteria have GT2 familytransferases that make polymers of b1,6-linked GlcNAc(Gerke et al. 1998; Itoh et al. 2008), but these enzymes re-semble the S. cerevisiae Chs proteins. If b1,6-glucan is indeedformed directly from UDP-Glc, the b1,6-glucosyltransferasewould represent a new GT family. Further possibilities arethat a known yeast GT may also form b1,6-glucosidic linkagesusing UDP-Glc as donor or that b1,6-glucan is generatedsolely by transglycosylation.

Remodeling and Cross-Linking Activities at the CellSurface

Order of incorporation of components into the cell wall

CWPs delivered by the secretory pathway meet up with chitinand b-glucans at the outer face of the plasma membrane and

S. cerevisiae Cell Wall 797

undergo cross-linking reactions that incorporate them intothe wall. The order in which wall components are assembledhas been inferred from analyses of the material formedwhen spheroplasts regenerate their walls and from the wallcompositions of mutants unable to make a particular com-ponent (Kreger and Kopecká 1976; Roh et al. 2002b). Thestarting component is b1,3-glucan, which is necessary forincorporation of both b1,6-glucan and mannoproteins. Be-cause b1,6-glucan was still attached to b1,3-glucan whenGPI anchoring was inhibited (Roh et al. 2002b), and becauseincorporation of GPI-CWPs is lowered in b1,6-glucan syn-thesis mutants (Lu et al. 1995; Kapteyn et al. 1997), GPI-CWPs are likely incorporated after b1,6-glucan. Because chi-tin became detectable in the walls of daughter cells onlyafter cytokinesis (Shaw et al. 1991), it was concluded thatchitin is the last component to be incorporated into the wall(Roh et al. 2002b). The sequence b1,3-glucan/b1,6-glucan/mannoprotein must be able to accommodatechanges in expression or assembly of individual componentsdictated by the cell cycle, cell wall stress, mating, or sporu-lation, as well as remodeling of individual polysaccharides.For example, a compensatory incorporation of PIR proteindirectly attached to b1,3-glucan is seen in b1,6-glucan syn-thesis mutants (Kapteyn et al. 1999b).

The model for the order of incorporation of wallcomponents needs to be reconciled with the model fora bilayered wall, during whose formation CWPs are pro-pelled to the cell surface, leaving polysaccharides nearer theplasma membrane. Furthermore, surface CWP may not beretained at the surface of wild-type cells. Thus, wild-typediploids expressing a Sag1-GFP fusion released a significantbasal level of that glycoprotein into the medium (Gonzaleset al. 2010). CWP may therefore routinely be shed duringvegetative growth, perhaps upon digestion of the wall be-tween mother cell and bud, along with secreted proteinssuch as chitinase and invertase (Kuranda and Robbins1991). Cross-linking and remodeling reactions will be de-scribed next, and hydrolases of known or unknown func-tions, as well as nonenzymatic wall proteins are discussedin Cell Wall-Active and Nonenzymatic Surface Proteins andTheir Functions.

Incorporation of GPI proteins into the wall

The v(2) region of a GPI protein (see Identification of GPIproteins) influences whether the protein will be retained inthe plasma membrane in lipid-anchored form or whether itcan be transferred into the wall (Caro et al. 1997; Hamadaet al. 1998a, 1999; De Sampaïo et al. 1999; Frieman andCormack 2004). If this region includes two basic aminoacids, the protein will be mostly retained in the plasmamembrane (Caro et al. 1997; Frieman and Cormack 2003),but if basic residues are absent or replaced with hydrophobicones, the predominant location is the wall (Hamada et al.1998b, 1999; Frieman and Cormack 2003). However, havingtwo basic amino acids in the v(2) region does not guaranteemembrane localization because the additional presence of

a longer stretch of amino acids rich in Ser and Thr will overridethe dibasic motif and shift the protein to the wall (Friemanand Cormack 2004). Furthermore, not all wall-anchored GPIproteins have the amino acids suggested to promote incor-poration into the wall (De Groot et al. 2003). In general, GPIproteins are partitioned between the membrane and wallto varying extents, and none may be restricted to only onelocation (Gonzales et al. 2009).

The nature of a GPI-protein’s mode of cell surface attach-ment can be critical. Ecm33, which is required for growth atelevated temperature (see Sps2 family), occurs mainly asa plasma membrane-anchored GPI protein, and this locali-zation is required for in vivo function. Replacement of v(2)amino acids v-1-13 of Ecm33 with the corresponding aminoacid sequences from wall-localized proteins resulted in in-creased cross-linking of Ecm33 to the wall, but also in loss ofthe protein’s ability to support growth at high temperature(Terashima et al. 2003).

The lipid-to-wall transfer reaction could be a one-steptransglycosylation in which the GPI glycan is cleaved and itsreducing end transferred to b1,6-glucan, or it could involveseparate GPI cleavage and transglycosylation steps. Candi-dates for cross-linkers are Dfg5 and Dcw1, an essential, re-dundant pair of homologous GPI proteins that resemble ana1,6-endomannanase and are in GH Family 76 (Kitagakiet al. 2002). Single dfg5D and dcw1D mutants are viable,although dcw1D is hypersensitive to Zymolyase, but thecombination of dfg5D and dcw1D is lethal (Kitagaki et al.2002). Depletion of Dfg5 or Dcw1 by repressing their ex-pression in the double-null background led to cell enlarge-ment, delocalized chitin deposition, and secretion of a GPI-CWP protein into the growth medium (Kitagaki et al. 2002).dcw1D was also recovered in a screen for impaired cross-linking of GPI proteins (Gonzalez et al. 2010). The defectscaused by loss of Dfg5 and Dcw1, together with the proteins’resemblance to an a-endomannanase, are consistent withtheir having a role in GPI cleavage and/or transglycosyla-tion. Homozygous DFG5 nulls are defective in filamentousgrowth (Mosch and Fink 1997).

GPI-CWP can be used to display heterologous proteins onthe yeast cell surface (Schreuder et al. 1993; Van der Vaartet al. 1997; Gai and Wittrup 2007; Shibasaki et al. 2009). Inone such system, heterologous proteins are fused to theAga2 subunit of the a-mating agglutinin (see Flocculinsand 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 arerequired for the alkali-labile linkage that joins these proteinsto b1,3-glucan (see Wall Composition and Architecture). De-letion of all PIR sequences from Pir1 and Pir4 leads to re-lease of these proteins from the cells (Castillo et al. 2003;Sumita et al. 2005), indicating that the repeats are necessaryfor wall association. The more repeats, the stronger thebinding: deletion of increasing numbers of Pir1’s repeats

798 P. Orlean

led to release of increasing amounts of Pir1 into the medium(Sumita et al. 2005).

Studies of Pir4/Ccw5, which has one PIR sequence andneeds it for cell wall anchorage, revealed that the alkali-labile linkage was an ester between the g-carboxyl groupof glutamate and the hydroxyl groups of b1,3-glucan. Thelinkage was generated in a transglutaminase reaction withQ74 in the PIR sequence SQIGDGQ74[V/I]QAT[T/S] (Eckeret al. 2006). In addition to substitutions of Q74, individualmutations of Q69, D72, and Q76 also resulted in loss of wallanchorage of the protein, indicating that these residues haveroles in the reaction. No transglutaminase has yet been iden-tified, but Ecker et al. (2006) point out that, because the freeenergy of hydrolysis of the amide is high enough to driveformation of the ester linkage, the PIR proteins could cata-lyze their own attachment to b1,3-glucan.

The glucan attachment sequence of PIR repeats is alsofound in the GPI-CWP Tip1, Tir1, Cwp1, and Cwp2 (Van derVaart et al. 1995). In the case of Cwp1, the PIR repeat maybe used as an additional wall anchorage point because theprotein is attached to the wall by both an alkali-labile anda GPI-dependent linkage (Kapteyn et al. 2001). Like GPI-CWP, PIR proteins can be used as carriers to direct surfaceexpression of heterologous proteins fused to them (Andréset 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 thereducing ends of chitin chains and both the nonreducing endof b1,3-glucose side branches on b1,6-glucan and the non-reducing ends of b1,3-glucan chains (Cabib et al. 2007; Cabib2009), and their homolog Crr1 likely does so during asco-spore wall assembly. These proteins are in GH family 16, andCrh2 and Crr1 also have a chitin-binding module (Rodriguez-Pena et al. 2000; Cabib et al. 2008). Crh1 and Crh2 are GPIproteins (Caro et al. 1997; Hamada et al. 1998a) whose lo-calization matches that of Chs3. Crh1-GFP fusions are detect-able at the site of bud emergence and later in the neck regionbetween mother cell and bud, and Crh2-GFP is seen in theneck region throughout the budding cycle, as well as in thelateral wall (Rodriguez-Pena et al. 2000, 2002). Single crh1and crh2 null mutants show Calcofluor White and Congo Redsensitivity, phenotypes enhanced in the double null, suggest-ing that Crh1 and Crh2 have a common wall-related function.Single crh mutants have a higher ratio of alkali soluble- toalkali-insoluble glucan, and this ratio is higher still in crh1Dcrh2D, indicating a role for Crh1 and Crh2 in linking b-glucanand chitin (Rodriguez-Pena et al. 2000).

Evidence that Crh1 and Crh2 are transglycosylases camefrom elegant studies by Cabib and coworkers, who used fluo-rescent, sulforhodamine-conjugated b1,3-gluco-oligosaccharidesas acceptors and showed that they became cross-linked tochitin in bud scars and the lateral walls of live cells (Cabibet al. 2008). Fluorescent labeling was very weak in crh1Dcrh2D cells or in cells lacking Chs3, which makes the chitinnormally bound to b1,3- and b1,6-glucan (Cabib and Duran

2005). The entire process of chitin polymerization and cross-linking could be reconstituted in detergent permeabilized,protease-treated cells. Cross-linking of fluorescent b1,3-gluco-oligosaccharides depended on the addition of UDP-GlcNAc (Cabib et al. 2008). Interestingly, the nascent chitinwas generated in situ by Chs1, which is highly active inpermeabilized cells, rather than by Chs3.

CRR1 shows sporulation-specific expression. Crr1-GFPfusions are localized on the surface of ascospores, and ho-mozygous crr1D diploids have ascospore wall abnormalities,with irregular deposition of the outer dityrosine and chito-san layers over the inner b-glucan layer (Gómez-Esqueret al. 2004). Ascopores from Crr1-deficient diploids showincreased sensitivity to heat shock and lytic enzymes, andthese defects are exacerbated when the chitin deacetylasesCda1 and Cda2 are also absent. These findings suggesta role for Crr1 in generating cross-links between the b-glucanand chitosan or chitin during ascospore wall maturation(Gómez-Esquer et al. 2004).

Cell Wall-Active and Nonenzymatic Surface Proteinsand Their Functions

Secreted, membrane, or wall proteins with known or con-jectured roles in wall biogenesis, adhesion, and nutrition aresurveyed here. The primary division is according to whetherproteins have or are likely to have enzymatic activity orwhether they are nonenzymatic, structural proteins. Bothgroups contain GPI proteins. Cell wall proteins have beenreviewed by Klis et al. (2002, 2006), De Groot et al. (2005),Lesage and Bussey (2006), and Gonzalez et al. (2009), glyco-sylhydrolases by Adams (2004), agglutinins by Dranginiset al. (2007), and flocculins by Goossens and Willaert(2010). Additional information about these proteins is pre-sented 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 (Kurandaand Robbins 1991). Cts1 is periplasmic, but much of it issecreted into the medium of cells grown in rich medium(Correa et al. 1982; Kuranda and Robbins 1991). Cts1 hasa key role in cell separation because cts1D strains form aggre-gates of cells that remain joined at their chitin-containingsepta, a phenotype mimicked when cells are treated withthe chitinase inhibitor dimethyallosamidin (Kuranda andRobbins 1991). Cts1’s chitin-binding domain contributes tothe enzyme’s localization in the septal region because Cts1truncations lacking it only partially complement the cts1Dseparation defect (Kuranda and Robbins 1991). Cts2 mayhave 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 EXG1and EXG2 have no overt defects, although exg1D cells haveslightly elevated levels of b1,6-glucan, and EXG1 overex-pressers lower amounts of that polymer, suggesting rolesfor Exg1 and Exg2 in b-glucan remodeling (Jiang et al.1995; Lesage and Bussey 2006). Ssg1/Spr1 is a sporulation-specific protein (File S9).

Bgl2, Scw4, Scw10, and Scw11 endo-b1,3-glucanases:These are GPI-less secretory proteins. Bgl2 has endo-b1,3-glucanase activity in vitro (Mrša et al. 1993), but it can alsocreate a b1,6 linkage between the reducing end that it gen-erates by cleaving a b1,3-gluco-oligosaccharide and the non-reducing end of another b1,3-glucan chain (Goldman et al.1995), and so could function as a b1,3-glucan branchingenzyme. No enzymatic activity has been shown for Scw4,Scw10, or Scw11, although mutation of predicted catalyticresidues in Scw10 abolished in vivo function (Sestak et al.2004). Scw4, Scw10, and Bgl2 are wall-associated via disul-fides (Cappellaro et al. 1998), but some Scw4 and Scw10can 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; Kalebinaet al. 2003; Sestak et al. 2004), and bgl2D walls have elevatedlevels of alkali-soluble glucan (Sestak et al. 2004). Strainslacking both Scw4 and Scw10 are CFW-hypersensitive andmorphologically abnormal, have doubled chitin content andincreased alkali-soluble glucan, and show alterations inb1,3-glucan structure and in cross-linking of mannoproteinsto the wall (Cappellaro et al. 1998; Sestak et al. 2004).The growth and morphological defects of scw4D scw10Dare exacerbated by deletion of CHS3 or FKS2 (Sestak et al.2004). From the phenotypes of strains expressing differ-ent relative amounts of Bgl2 and Scw10, it was proposedthat levels of Bgl2 and Scw10 need to be balancedto ensure wall stability (Klebl and Tanner 1989; Sestaket al. 2004; File S9). Cells lacking Scw11 have a separationdefect, and, consistent with this, Scw11 is a daughter cell-specific protein (Cappellaro et al. 1998; Colman-Lerneret al. 2001).

Eng1/Dse4 and Eng2/Acf2 endo-b1,3-glucanases: Theserelated proteins have endo-b1,3-glucanase activity in vitro,but different localizations. Eng1 is a GPI protein (Baladronet al. 2002; De Groot et al. 2003), whereas Eng2 is likelyintracellular. Mutants lacking one or both proteins makenormal walls, but eng1D cells have a separation defect, con-sistent with Eng1’s localization to the daughter side of theseptum (Colman-Lerner et al. 2001; Baladron et al. 2002).ENG2 expression increases during sporulation, althougheng2D diploids are not defective in that process (Baladronet al. 2002). Surprisingly, loss of multiple exo- and endo-b1,3-glucanases is not catastrophic because cells lackingExg1, Exg2, Eng1, Eng2, and Bgl2 grow well and show onlythe eng1D separation defect (Cabib et al. 2008).

Gas1 family b1,3-glucanosyltransferases: This family hasfive members, all of which have GPI attachment sites(Fankhauser et al. 1993; Caro et al. 1997; Popolo and Vai1999; De Groot et al. 2003). Gas1, Gas3, and Gas5 can alsobe covalently linked to the wall (De Sampaïo et al. 1999; Yinet al. 2005). Gas proteins have b1,3-glucanosyltransfer activ-ity: they cleave b1,3-glucosidic linkages within b1,3-glucanchains and then transfer the newly generated reducing endof the cleaved glycan to the nonreducing end of anotherb1,3-glucan molecule, thereby extending the recipientb1,3-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 intheir walls (Ram et al. 1995; Popolo et al. 1997; Valdiviesoet al. 2000). gas1D cells release b1,3-glucan to the medium(Ram et al. 1998), and Gas1’s b1,3-glucan elongase activitymay therefore be necessary for incorporation of b1,3-glucaninto the wall. In addition, analyses of the synthetic interac-tion network of gas1D revealed that survival in the absenceof Gas1 requires correct assembly of b1,6-glucan (Tomishigeet al. 2003; Lesage et al. 2004). Gas1 is detectable in thelateral wall, in the chitin ring in small-budded cells, andnear the primary septum and remains in the bud scar aftercell separation, and its localization is dependent on the pres-ence of its GPI-attachment sequence (Rolli et al. 2009).

Gas3 and Gas5 likely have wall-related functions in veg-etative cells (File S9). GAS2 and GAS4 are expressed onlyduring sporulation, and diploids lacking both Gas2 and Gas4have a severe sporulation defect. The inner glucan layer ofthe wall of double homozygous gas2 gas4 null spores wasdisorganized and detached from chitosan, suggesting thatthe b1,3-glucanosyltransferase activity of Gas2 and Gas4generates b1,3-glucan chains that associate optimally withchitosan (Ragni et al. 2007a).

Yapsin aspartyl proteases: GPI-anchored aspartyl proteasesof the yapsin family have roles in the turnover of CWPs andwall-localized enzymes. Yps1, Yps2/Mkc7, Yps3, and Yps6are mostly plasma membrane-associated, whereas Yps7 ispredicted to be wall anchored (Krysan et al. 2005; Gagnon-Arsenault et al. 2006). Yapsins cleave their substratesC-terminally to Lys or Arg or pairs of these residues (Olsenet al. 1998; Komano et al. 1999) and themselves undergoproteolytic processing to generate active enzyme (File S9).

Individual yapsin null mutants are sensitive to variouswall-disrupting agents, and loss of multiple YPS genes leadsto osmotically remedial, temperature-sensitive lysis defects,findings that indicate that the yapsins are involved in wallmaintenance (Krysan et al. 2005). Walls from yps1D yps2Dand yps1D yps2D yps3D yps6D yps7D null mutants showedlowered b1,3- and b1,6-glucan and elevated chitin levels,whereas mannan levels were unchanged, with the wallalterations being most pronounced in the quintuple mutant(Krysan et al. 2005). The b-glucan defects were due to

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decreased incorporation of these polymers into the wall,because synthesis of the two b-glucans was normal in thedeletion strains. These findings suggest that yapsins act onwall hydrolases and transglycosidases, thereby regulatingactivity of the latter, and hence, incorporation of glucansinto the wall (Krysan et al. 2005). Support for this camefrom identification of Gas1, Pir4, and Msb2 as Yps1 sub-strates (Gagnon-Arsenault et al. 2008; Vadaie et al. 2008).In addition to degrading or shedding proteins during wallremodeling, yapsins also have roles in mediating release ofaberrantly folded or overexpressed GPI proteins that induceER stress (Miller et al. 2010).

Nonenzymatic CWPs

Structural GPI proteins: There are three families of GPI-CWP and several individual GPI-CWP that do not resembleknown enzymes. Strains lacking one or more of these GPI-CWP have wall defects, and expression of some of theseproteins can vary with cell cycle stage or be induced duringmating or sporulation in response to cell wall stress or whenoxygen levels are low. In general, GPI-CWPs have a collectiverole in maintaining cell wall stability (Lesage and Bussey2006; Ragni et al. 2007c).

Sps2 family: This group comprises Ecm33, Pst1, Sps2, andSps22 (Caro et al. 1997). Of these, Ecm33 has an importantrole in vegetative walls. ecm33D cells are temperature-sensitiveand sensitive to various wall-perturbing agents, have a dis-organized wall with a thin or absent mannoprotein layer,and shed b1,6-glucan-linked mannoproteins and Pir2 thatis possibly linked to b1,3-glucan more than normal in themedium (Lussier et al. 1997b; Pardo et al. 2004). pst1D cellshave no obvious phenotype, but ecm33D pst1D double nullsshow exacerbated sensitivity to various wall stresses.Ecm33’s and Pst1’s functions partially overlap, but the pro-teins are not fully redundant because overexpression ofPST1 only weakly suppresses the ecm33D defects (Pardoet al. 2004). Sps2 and Sps22 are a redundant pair requiredfor normal ascospore wall formation. Diploids lacking themform spores with abnormal b-glucan, chitosan, and dityro-sine layers (Coluccio et al. 2004). Sps2 and Sps22 likely actat 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, andDan1/Ccw13 are mostly small, Ser- and Ala-rich GPI-CWPthat show differential expression during the cell cycle andduring aerobic or anaerobic growth and can be localizeddifferently on the cell surface. Cwp2 also contains a PIR re-peat and so could be linked to b1,3-glucan (Klis et al. 2010).

Cwp1, Cwp2, Tip1, and Tir1 have roles in the vegetativewall. Deletion of their genes individually leads to CFW hy-persensitivity (Van der Vaart et al. 1995), and cwp1D cwp2Ddouble mutants show increased permeability to DNA-bindingagents relative to the single nulls (Zhang et al. 2008). Inaddition, the walls of cwp2D and cwp1D cwp2D cells arethinner 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, andTir1 are expressed during the S-to-G2 transition, Cwp2 beingfound in small-to-medium-sized buds (Caro et al. 1998;Smits et al. 2006). Localization of Cwp2 and Tip1 is deter-mined by the timing of their expression in the cell cycle(Smits et al. 2006; File S9). Tip1 and Tip2 are also heat-and cold-shock-inducible, and Tir1 and Tir4 are induced bycold shock (Kowalski et al. 1995; Abramova et al. 2001).

CWP1 and CWP2 are downregulated upon shift to anaer-obic conditions, whereas Tip1, Tir1, Tir2, Tir3, Tir4, Dan1/Ccw13, and Dan4 are induced (Abramova et al. 2001). Ofthese, the Dan proteins are strongly repressed by oxygen.Strains lacking Tir1, Tir3, or Tir4 do not grow under anaer-obic conditions. Shift to anaerobiosis therefore leads toremodeling of the wall (Abramova et al. 2001), althoughit is not clear how the anaerobically induced CWPs permitanaerobic growth.

Sed1 and Spi1: These are two related, Ser/Thr-rich GPI-CWP whose expression is induced by nutrient limitation andstress. Sed1 is releasable from walls by treatment withb-glucanases or proteases (Shimoi et al. 1998). Associationof Sed1 with the wall is dependent on Kre6 (Bowen andWheals 2004), consistent with anchorage involving b1,6-glucan. SED1 expression is induced in the stationary phase,a time when the wall becomes thicker and more resistant tolytic enzymes (De Nobel et al. 1990). Consistent with a pro-tective role in stationary-phase walls, sed1D cells becomemore sensitive to Zymolyase in that growth phase (Shimoiet al. 1998). Elevated Sed1 expression is also part of thecompensatory response made by cells lacking multipleGPI-CWP (Hagen et al. 2004; File S9). Expression of SPI1is induced by weak organic acids, and Sps1 is a major con-tributor to the b1,3-glucanase resistance that arises in re-sponse to this stress (Simoes et al. 2003). Low external pHalso leads to formation of new alkali-labile linkages betweenGPI-CWPs and b1,3-glucan (Kapteyn et al. 2001).

Ccw12: Ccw12 is a small, heavily glycosylated, Ser/Thr-rich GPI-CWP with two C-terminal repeats of an amino acidsequence critical for its function (File S9). Ccw12 is releas-able by b1,3-glucanases (Mrša et al. 1999), but also has thepotential to make disulfide cross-links because it has a three-Cys motif found in several S. cerevisiae flocculins (see Floc-culins and agglutinins) and in wall proteins of other yeasts(Klis et al. 2010). Ccw12 is likely abundant because its genehas a very high codon adaptation index (Klis et al. 2010).Ccw12 has a major role in the wall, because cells lacking itare hypersensitive to CFW and other wall stressing agentsand 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; Shankarnarayanet al. 2008). Man-to-Glc ratios in ccw12D cells are unchanged,but levels of alkali-soluble relative to alkali-insoluble glucanare higher, indicating altered organization and cross-linkageof wall components (Ragni et al. 2007c). Ccw12 localizes tosites of active wall synthesis, including the future bud site,

S. cerevisiae Cell Wall 801

the septum, the lateral walls of enlarging daughter cells, aswell as the tips of mating projections, but then turns over,suggesting that it may stabilize walls as daughter cells andthat mating projections are being formed (Ragni et al.2011). Loss of Ccw12 alone activates the CWI pathway-me-diated chitin stress response (Ragni et al. 2007c, 2011; seeChitin synthesis in response to cell wall stress), but deletion ofadditional GPI-CWP genes forces cells over a threshold thatleads to triggering of a new compensatory response to lossof multiple GPI-CWP that depends on Sed1 and the non-GPI-CWP Srl1 (see File S7).

Other nonenzymatic GPI proteins: Ccw14 (Ssr1/Icwp) isa b1,3-glucanase-extractable, Ser-rich GPI-CWP that has beenlocalized to the inner cell wall (Moukadiri et al. 1997; Mršaet al. 1999; File S9). The protein has an eight-Cys-containingCFEM domain found in various fungal surface proteins(Kulkarni et al. 2003; De Groot et al. 2005) and, hence, a po-tential for disulfide formation. CCW14/SSR1 null mutantshave no obvious growth defects, but show increased sensitiv-ity to CFW, Congo Red, and Zymolyase. Overexpression ofCCW14/SSR1 also leads to increased CFW and Congo Redsensitivity, although not Zymolyase sensitivity, suggesting thatlevels of Ccw14/Ssr1 relative to one or more other wall com-ponents need to be balanced (Moukadiri et al. 1997).

Dse2 and Egt2, which are unrelated to one another, aredaughter cell-specific proteins with roles in cell separation.In haploids, Dse2 is concentrated in regions connectingmother and daughter cells (Colman-Lerner et al. 2001; Doolinet al. 2001), and Egt2 is localized to the septum (Fujitaet al. 2004). Of these two GPI proteins (Hamada et al.1998a; Terashima et al. 2002; De Groot et al. 2003), Egt2’slocalization also depends on Gpi7 (Fujita et al. 2004). dse2Dhaploids show no defects, but homozygous DSE2 nulls showunipolar budding and form chains of cells. egt2D cells haveseparation defects similar to those of eng1D cells, a pheno-type exacerbated in the double null, indicating that the twoproteins 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 expres-sion is induced by iron limitation, and the proteins normallyretain iron bound to ferrichrome because Zymolyase treat-ment of FIT-deletion mutants releases less iron from cells(Protchenko et al. 2001). Fit1 localizes to the wall, whereit, Fit2, and Fit3 concentrate siderophore iron and facilitatesubsequent uptake of the metal, highlighting a role of thewall in nutrient acquisition (Protchenko et al. 2001).

Flocculins and agglutinins: GPI-CWP involved in cell–celladhesion are the related Flo1, Flo5, Flo9, Flo10, and Flo11/Muc1 flocculins, the Aga1 and Fig2 pair, and the a-agglutininSag1 (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 composedof 1100–1500 amino acids. Major features are N-terminal

PA14 domains that bind a-mannosides and mediate adhe-sion to adjacent cells, a central Ser/Thr-rich domain that isorganized in repeat sequences and heavily glycosylated,and two or three conserved three-Cys repeats toward theirC termini (Verstrepen and Klis 2006; Goossens and Willaert2010; Klis et al. 2010; Veelders et al. 2010; Goossens et al.2011). In addition, the Ser/Thr-rich domains of Flo1 andFlo11/Muc1 have short sequences enriched in Ile, Thr, andVal that are predicted to form intramolecular b-sheet-like interactions or amyloids, and both a soluble, GPI-lessportion of Flo11/Muc1 and a Flo1-derived form fibrillarb-aggregates in vitro (Ramsook et al. 2010). Amyloid forma-tion correlates with flocculation in vivo, for cells expressingFlo1 and Flo11/Muc1 that had been induced to flocculate inthe presence of Ca2+ stained more brightly with an amyloid-binding dye, and amyloid formation may be part of themechanism by which these proteins promote cell aggrega-tion (Ramsook et al. 2010). FLO1, FLO5, FLO9, and FLO10are not expressed in laboratory strains such as S288C be-cause of a mutation in the transcriptional activator Flo8.However, activation of individual FLO genes confers the abil-ity 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, in-vasion of agar by haploids, and biofilm development (Loand Dranginis 1998; Guo et al. 2000; Reynolds and Fink2001; Dranginis et al. 2007; Bojsen et al. 2012).

Related Aga1 and Fig2 function in mating and localize tothe mating projection (Erdman et al. 1998; Guo et al. 2000;Jue and Lipke 2002). Aga1 is a component of a-agglutininthat displays the Aga2 subunit, which is disulfide linkedto it, and which confers binding specificity to a-agglutininSag1 (Orlean et al. 1986; Roy et al. 1991; Cappellaro et al.1994; Shen et al. 2001). Fig2, which like Aga1 is expressedin both mating types, is required for formation of matingprojections 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 halfthat may hold up the N-terminal, Aga2-binding portionof the protein at the cell surface. Sag1’s N-terminal regioncontains three sequential domains that resemble variableimmunoglobulin-like folds (Chen et al. 1995; Shen et al.2001), the most C-terminal of which contains amino acids nec-essary for Aga2 binding (Wojciechowicz et al. 1993; Cappellaroet al. 1994; De Nobel et al. 1996).

Non-GPI-CWP: PIR proteins: Expression and localization ofPir1 (Ccw6), Pir2 (Ccw7/Hsp150), Pir3 (Ccw8), and Pir4(Ccw5/Cis3) is regulated during cell cycle progression andin response to stress. PIR1, PIR2, and PIR3 show peaks ofexpression in early G1, whereas PIR4 expression is highest inG2 (Spellman et al. 1998). PIR2 is also induced by heatshock, treatment with CFW or Zymolyase, and nitrogen lim-itation (Russo et al. 1993; Toh-e et al. 1993; Yun et al. 1997;

802 P. Orlean

Boorsma et al. 2004). Consistent with their upregulationupon wall stress, all four PIR genes show elevated expres-sion in an mpk1 mutant that constitutively activates the pro-tein kinase C-dependent CWI pathway, an effect eliminatedin mutants lacking the PKC pathway’s target transcriptionfactor, Rlm1 (Jung and Levin 1999).

PIR proteins localize to different parts of the surface ofbudding cells (Sumita et al. 2005; File S9). Pir1 and Pir2 arefound at bud scars of both haploids and diploids, Pir1 beinglocalized inside the chitin ring. Some Pir1 and Pir2 and mostPir3 are also present in lateral walls (Yun et al. 1997). Pir4has been reported be uniformly distributed at the cell sur-face or restricted to growing buds (Moukadiri et al. 1999;Sumita et al. 2005).

Strains lacking individual PIR proteins have subtlegrowth defects, but as more PIR genes are deleted, disrup-tants show a progressive increase in sensitivity to CFW,Congo Red, and heat shock, and cells become larger andirregularly shaped (Toh-e et al. 1993; Mrša and Tanner1999). pir1D pir2D pir3D pir4D mutants show a loss of vi-ability that is suppressed in osmotically supported medium(Teparic et al. 2004). These findings suggest a collective rolefor PIR proteins in maintenance of a normal wall. How theseproteins contribute is unclear, because the carbohydrate com-position of the quadruple PIR disruptant’s wall is unaltered,and the relative amounts of alkali-soluble and -insoluble glu-can and chitin show modest changes (Teparic et al. 2004;Mazan et al. 2008). PIR proteins, however, impact permeabil-ity of the wall because the pir1D pir2D pir3D mutant is hy-persensitive to membrane-active tobacco osmotin, whereasoverexpression of PIR1, PIR2, or PIR3 confers osmotin resis-tance on walled cells but not spheroplasts (Yun et al. 1997).The effects of PIR protein levels on wall permeability areconsistent with the role of these proteins in cross-linkingb1,3-glucans (see Mild alkali-releasable proteins).

Scw3 (Sun4): Haploids lacking this soluble cell wallprotein (Cappellaro et al. 1998) are larger than wild-typecells and have a separation defect and thickened septa(Mouassite et al. 2000). Scw3/Sun4 is a member of theSUN family of proteins, of which Sim1 and Uth1 are alsoreleased from cell walls by dithiothreitol treatment (Velourset al. 2002). Uth1 and Scw3/Sun4 additionally localize tomitochondria (Velours et al. 2002), but the significance ofthis distribution of the SUN proteins is unclear. The bio-chemical function of the SUN proteins is unknown as theyshow no similarity to known enzymes (File S9).

Srl1: This small Ser/Thr-rich protein is involved in thecompensatory response to loss of multiple GPI-CWP (Hagenet al. 2004; File S9). It rescues the lysis defects of strainsdefective in the function of the “regulation of Ace2 and po-larized morphogenesis” (RAM) signaling network when over-expressed (Kurischko et al. 2005), and some of it is tightlyassociated with the wall and released by b1,3-glucanase(Terashima et al. 2002). Slr1 localizes to the periphery ofsmall buds (Shepard et al. 2003). srl1D mutants have noobvious morphological defects and show modest Calcofluor

White sensitivity at 22�, but are hypersensitive to this agentat 37� (Kurischko et al. 2005). Mutants defective in RAMfunction are also suppressed by overexpression of Sim1 (seeabove) and Ccw12 (Kurischko et al. 2005), and slr1D andccw12D show a strong genetic interaction in the RAM-defective background. The srl1D ccw12D strain is CFW hy-persensitive at both 22� and 37�, and at 22�, but not at 37�,resembles mating pheromone-treated wild-type cells (Kurischkoet al. 2005). Srl1 and Ccw12 have been proposed to haveparallel functions in activation of a CWI pathway that oper-ates when RAM signaling is defective (Kurischko et al.2005).

What Is Next?

The biosynthesis of most individual yeast wall componentsis now understood in much detail and involves conservedpathways such as N-glycosylation and GPI anchoring andenzymes represented in other organisms, such as chitin andb1,3-glucan synthases. In contrast, b1,6-glucan formationand cross-linkage to GPI proteins and cross-linking of chitinto b-glucans are clearly restricted to certain yeasts and fila-mentous fungi, and the enzymes implicated in the latterprocesses, as well as certain CWP, are signatures of fungalcell walls, whose evolution of has been reviewed by Ruiz-Herrera and Ortiz-Castellanos (2010) and Xie and Lipke(2010).

Much of the work necessary to take both the conservedand the yeast-specific aspects of wall biogenesis to the nextlevel must be biochemical and analytical. These efforts willinvolve charting new biochemical territory, such as de-termining how b1,6-glucan is made and defining the func-tions of wall-active proteins such as Ccw12, Ecm33, Kre1,and Kre9, which have key roles in wall biogenesis, but showno resemblance to proteins of known function and may notbe enzymes. Other biochemical challenges are the mecha-nism and activation of chitin and b1,3-glucan synthases, themechanism and conjectured flippase activities of the multispan-ning glycosyltransferases of the dolichol, O-mannosylation, andGPI pathways, the functions of the Etn-P side branches onGPIs, and the biochemical activities of predicted enzymessuch as the Dcw1/Dfg5 pair, Kre5, Kre6, Scw4, and Scw10.The latter efforts require application of high-resolution tech-niques to analyze the fine structure and linkages of cell wallglycans (Magnelli et al. 2002; Aimanianda et al. 2009),which should highlight the reactions for which biochemistsneed to develop assays and screen mutants.

With the identification of so many proteins involved in cellwall biogenesis, and with ever-improving knowledge of wallcomposition, we can look forward to deepening our un-derstanding of the complexities of yeast cell wall biogenesis.

Acknowledgments

I thank my students for their many contributions to GPI andwall biosynthesis. I also acknowledge the contributions of

S. cerevisiae Cell Wall 803

the late Yoshifumi Jigami to our field. I am grateful to threereviewers for their helpful comments. Work in my laboratoryhas been supported by grant GM-46220 from the NationalInstitutes of Health and by a Burroughs Wellcome ScholarAward in Molecular Pathogenic Mycology.

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Communicating editor: J. Thorner

818 P. Orlean

GENETICSSupporting Information

http://www.genetics.org/lookup/suppl/doi:10.1534/genetics.112.144485/-/DC1

Architecture and Biosynthesis of the Saccharomycescerevisiae Cell Wall

Peter Orlean

Copyright © 2012 by the Genetics Society of AmericaDOI: 10.1534/genetics.112.144485

P.  Orlean   1  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  2  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.  

 

P.  Orlean   3  SI  

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  4  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.  

P.  Orlean   5  SI  

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  6  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  

P.  Orlean   7  SI  

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  8  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  

P.  Orlean   9  SI  

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  10  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  

P.  Orlean   11  SI  

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  12  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  

 

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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  

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Chi,  J.  H.,  Roos,  J.,  Dean,  N.,  1996  The  OST4  gene  of  Saccharomyces  cerevisiae  encodes  an  unusually  small  protein  required  for  

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Conde,   R.,   Cueva,   R.,   Larriba,   G.,   2007   Rsc14-­‐controlled   expression   of   MNN6,   MNN4   and   MNN1   regulates  

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1336-­‐1340.  

P.  Orlean   13  SI  

 

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Gao,  X.  D.,  Moriyama,  S.,  Miura,  N.,  Dean,  N.,  Nishimura,  S.,  2008  Interaction  between  the  C  termini  of  Alg13  and  Alg14  

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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  

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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  14  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.  

 

P.  Orlean   15  SI  

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:  

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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  16  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  Gas1G291R,  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).  

P.  Orlean   17  SI  

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  18  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.  

 

P.  Orlean   19  SI  

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  20  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).  

   

P.  Orlean   21  SI  

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  22  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,  

but  still  capable  of  making  Ptd-­‐Etn  by  decarboxylation  of  Ptd-­‐Ser,  do  not  incorporate  [3H]Etn  into  protein-­‐bound  GPIs  or  into  a  

Man2-­‐GPI  precursor  that  otherwise  receives  Etn-­‐P  on  Man-­‐1.  However,  radioactivity  supplied  as  [3H]Ser  is  incorporated  into  the  

Man2-­‐GPI  after  formation  and  decarboxylation  of  Ptd-­‐[3H]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  

involvement   in   formation   or   cleavage   of   a   phosphodiester.   This   domain   is   important   for   function,   because   the   G227E  

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  

et  al.  2001)  and  Zn2+  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).  

P.  Orlean   23  SI  

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-­‐P301L   variant,   but   not   G227E,   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  24  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  

P.  Orlean   25  SI  

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  26  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  

temperature-­‐sensitivity   is   suppressed   by  Mn2+,   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  

phosphatase  that  affects  Golgi  inheritance  and  actin  polarization  by  activating  Ca2+  signaling.  Mol.  Cell.  Biol.  28:  3336–3343.  

 

P.  Orlean   27  SI  

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.  

 

Paidhungat,  M.,  Garrett,  S.,  1998    Cdc1  and  the  vacuole  coordinately   regulate  Mn2+  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  28  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.    

P.  Orlean   29  SI  

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  30  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.  

 

 

     

P.  Orlean   31  SI  

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  32  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).  

In  vitro  properties  of  yeast  chitin  synthases.  Chitin  synthase  assays  typically  detect  the  transfer  of  [14C]GlcNAc  from  

UDP[14C]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  

P.  Orlean   33  SI  

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  34  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  

P.  Orlean   35  SI  

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  36  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  

 

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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.  

 

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Choi,  W.  J.,  Cabib,  E.,  1994    The  use  of  divalent  cations  and  pH  for  the  determination  of  specific  yeast  chitin  synthetases.  Anal.  

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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.  

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Lucero,   H.   A.,   Kuranda  M.   J.,   Bulik,   D.   A.,   2002     A   nonradioactive,   high   throughput   assay   for   chitin   synthase   activity.   Anal.  

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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.  

     

P.  Orlean   39  SI  

File  S7  

β1,3-­‐glucan  

This  Supporting  File  contains  additional  information  and  discussion  related  to  Biosynthesis  of  Wall  Components  at  the  Plasma  

Membrane,  β1,3-­‐glucan.    The  subheadings  used  in  the  main  text  are  retained,  and  new  subheadings  are  underlined.  

 Fks  family  of  β1,3-­‐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  

incorporation  of  [14C]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  40  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  β1,3-­‐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).  

 

 

     

P.  Orlean   41  SI  

File  S8  β1,6-­‐Glucan  

This  Supporting  File  contains  additional  information  and  discussion  related  to  β1,6-­‐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  β1,6-­‐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  42  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.  

 

P.  Orlean   43  SI  

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  44  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.  

β1,3-­‐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  

P.  Orlean   45  SI  

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  β1,3-­‐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  46  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  

P.  Orlean   47  SI  

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  48  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  

P.  Orlean   49  SI  

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  50  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  

P.  Orlean   51  SI  

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  

 

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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.  

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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  

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P.  Orlean   53  SI  

Table  S1      Proteins  involved  in  cell  wall  biogenesis  in  Saccharomyces  cerevisiae   Process  or   Protein  name   Activity  or  Function                 CAZy  Family1  protein  type    

Precursor  supply  

Ugp1     UDPGlc  pyrophosphorylase  

Pmi40     phosphomannose  isomerase  

Sec53     phosphomannomutase  

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  

Vrg4/Vig4   GDP-­‐Man  transporter  

Gda1     GDPase  

Ynd1     Apyrase  

N-­‐glycosylation  

Alg7     UDP-­‐GlcNAc:  Dol-­‐P  GlcNAc-­‐1-­‐P-­‐T  

    Alg13  +  Alg14   UDP-­‐GlcNAc:  Dol-­‐PP-­‐GlcNAc β1,4-­‐GlcNAc-­‐T               GT1  

P.  Orlean  54  SI  

Alg1     GDP-­‐Man:  Dol-­‐PP-­‐GlcNAc2  β1,4-­‐Man-­‐T                 GT33  

Alg2     GDP-­‐Man:  Dol-­‐PP-­‐GlcNAc2Man α1,3-­‐Man-­‐T  and  GDP-­‐Man:  Dol-­‐PP-­‐GlcNAc2Man2 α1,6-­‐Man-­‐T     GT4  

Alg11     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  

Htm1/Mnl1   ER-­‐degradation  enhancing  a-­‐mannosidase-­‐like  protein             GH47  

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   55  SI  

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  

O-­‐mannosylation  

Pmt1     Dol-­‐P-­‐Man:  protein:  O-­‐Man-­‐T;  Pmt1  family               GT39  

Pmt2     Dol-­‐P-­‐Man:  protein:  O-­‐Man-­‐T;  Pmt2  family               GT39  

P.  Orlean  56  SI  

Pmt3     Dol-­‐P-­‐Man:  protein:  O-­‐Man-­‐T;  Pmt2  family               GT39  

Pmt4     Dol-­‐P-­‐Man:  protein:  O-­‐Man-­‐T;  specific  for  membrane  proteins           GT39  

Pmt5     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

GPI  anchoring  

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  

P.  Orlean   57  SI  

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  58  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  

β -­‐1,6  glucan  formation  

    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  

Glycosidases,  cross-­‐linking  enzymes,  and  proteases  

Cts1     Endo-­‐chitinase                     GH18  

Cts2     Chitinase                       GH18  

Exg1/Bgl1   Major  exo-­‐β-­‐1,3-­‐glucanase  of  the  cell  wall;  soluble             GH5  

P.  Orlean   59  SI  

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  

GPI-­‐CWP  

Ecm33     Sps2  family;  structural/non-­‐enzymatic  

Pst1     Sps2  family;  structural/non-­‐enzymatic  

Sps2     Sps2  family;  structural/non-­‐enzymatic;  required  for  ascospore  wall  formation  

P.  Orlean  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   61  SI  

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  

  1CAZy  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   1  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  2  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.  

 

P.  Orlean   3  SI  

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  4  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.  

P.  Orlean   5  SI  

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  6  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  

P.  Orlean   7  SI  

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  8  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  

P.  Orlean   9  SI  

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  10  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  

P.  Orlean   11  SI  

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  12  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  

 

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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  

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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.  

 

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1336-­‐1340.  

P.  Orlean   13  SI  

 

Dan,   N.,   Lehrman,  M.   A.,   1997   Oligomerization   of   hamster   UDP-­‐GlcNAc:dolichol-­‐P   GlcNAc-­‐1-­‐P   transferase,   an   enzyme   with  

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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  

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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  

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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  14  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  

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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.  

 

P.  Orlean   15  SI  

Yan,  A.,  Ahmed,  E.,  Yan,  Q.,  Lennarz,  W.  J.,  2003  New  findings  on  interactions  among  the  yeast  oligosaccharyl  transferase  

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Yan,  A.,  Wu.  E.,  Lennarz,  W.  J.,  2005  Studies  of  yeast  oligosaccharyl  transferase  subunits  using  the  split-­‐ubiquitin  system:  

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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  16  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  Gas1G291R,  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).  

P.  Orlean   17  SI  

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    

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does  not  abolish  nucleotide  sugar-­‐dependent  protein  glycosylation.  J.  Biol.  Chem.  280:  40417-­‐40427.  

P.  Orlean  18  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.  

 

P.  Orlean   19  SI  

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  20  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).  

   

P.  Orlean   21  SI  

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  22  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,  

but  still  capable  of  making  Ptd-­‐Etn  by  decarboxylation  of  Ptd-­‐Ser,  do  not  incorporate  [3H]Etn  into  protein-­‐bound  GPIs  or  into  a  

Man2-­‐GPI  precursor  that  otherwise  receives  Etn-­‐P  on  Man-­‐1.  However,  radioactivity  supplied  as  [3H]Ser  is  incorporated  into  the  

Man2-­‐GPI  after  formation  and  decarboxylation  of  Ptd-­‐[3H]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  

involvement   in   formation   or   cleavage   of   a   phosphodiester.   This   domain   is   important   for   function,   because   the   G227E  

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  

et  al.  2001)  and  Zn2+  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).  

P.  Orlean   23  SI  

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-­‐P301L   variant,   but   not   G227E,   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  24  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  

P.  Orlean   25  SI  

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  26  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  

temperature-­‐sensitivity   is   suppressed   by  Mn2+,   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  

phosphatase  that  affects  Golgi  inheritance  and  actin  polarization  by  activating  Ca2+  signaling.  Mol.  Cell.  Biol.  28:  3336–3343.  

 

P.  Orlean   27  SI  

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.  

 

Paidhungat,  M.,  Garrett,  S.,  1998    Cdc1  and  the  vacuole  coordinately   regulate  Mn2+  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  28  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.    

P.  Orlean   29  SI  

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  30  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.  

 

 

     

P.  Orlean   31  SI  

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  32  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).  

In  vitro  properties  of  yeast  chitin  synthases.  Chitin  synthase  assays  typically  detect  the  transfer  of  [14C]GlcNAc  from  

UDP[14C]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  

P.  Orlean   33  SI  

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  34  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  

P.  Orlean   35  SI  

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  36  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  

 

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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.  

 

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Choi,  W.  J.,  Cabib,  E.,  1994    The  use  of  divalent  cations  and  pH  for  the  determination  of  specific  yeast  chitin  synthetases.  Anal.  

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Delmer,  D.  P.,  1999    Cellulose  biosynthesis:  exciting  times  for  a  difficult  field  of  study.  Annu.  Rev.  Plant  Physiol.  Plant  Mol.  Biol.  

50:  245-­‐276.  

 

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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.  

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Lucero,   H.   A.,   Kuranda  M.   J.,   Bulik,   D.   A.,   2002     A   nonradioactive,   high   throughput   assay   for   chitin   synthase   activity.   Anal.  

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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.  

     

P.  Orlean   39  SI  

File  S7  

β1,3-­‐glucan  

This  Supporting  File  contains  additional  information  and  discussion  related  to  Biosynthesis  of  Wall  Components  at  the  Plasma  

Membrane,  β1,3-­‐glucan.    The  subheadings  used  in  the  main  text  are  retained,  and  new  subheadings  are  underlined.  

 Fks  family  of  β1,3-­‐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  

incorporation  of  [14C]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  40  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  β1,3-­‐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).  

 

 

     

P.  Orlean   41  SI  

File  S8  β1,6-­‐Glucan  

This  Supporting  File  contains  additional  information  and  discussion  related  to  β1,6-­‐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  β1,6-­‐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  42  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.  

 

P.  Orlean   43  SI  

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  44  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.  

β1,3-­‐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  

P.  Orlean   45  SI  

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  β1,3-­‐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  46  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  

P.  Orlean   47  SI  

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  48  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  

P.  Orlean   49  SI  

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  50  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  

P.  Orlean   51  SI  

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  

 

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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  

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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  

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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.  

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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-­‐

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Skory,  C.  D.,  Freer,  S.  N.,  1995  Cloning  and  characterization  of  a  gene  encoding  a  cell-­‐bound,  extracellular  β-­‐glucosidase  in  the  

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P.  Orlean   53  SI  

Table  S1      Proteins  involved  in  cell  wall  biogenesis  in  Saccharomyces  cerevisiae   Process  or   Protein  name   Activity  or  Function                 CAZy  Family1  protein  type    

Precursor  supply  

Ugp1     UDPGlc  pyrophosphorylase  

Pmi40     phosphomannose  isomerase  

Sec53     phosphomannomutase  

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  

Vrg4/Vig4   GDP-­‐Man  transporter  

Gda1     GDPase  

Ynd1     Apyrase  

N-­‐glycosylation  

Alg7     UDP-­‐GlcNAc:  Dol-­‐P  GlcNAc-­‐1-­‐P-­‐T  

    Alg13  +  Alg14   UDP-­‐GlcNAc:  Dol-­‐PP-­‐GlcNAc β1,4-­‐GlcNAc-­‐T               GT1  

P.  Orlean  54  SI  

Alg1     GDP-­‐Man:  Dol-­‐PP-­‐GlcNAc2  β1,4-­‐Man-­‐T                 GT33  

Alg2     GDP-­‐Man:  Dol-­‐PP-­‐GlcNAc2Man α1,3-­‐Man-­‐T  and  GDP-­‐Man:  Dol-­‐PP-­‐GlcNAc2Man2 α1,6-­‐Man-­‐T     GT4  

Alg11     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  

Htm1/Mnl1   ER-­‐degradation  enhancing  a-­‐mannosidase-­‐like  protein             GH47  

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   55  SI  

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  

O-­‐mannosylation  

Pmt1     Dol-­‐P-­‐Man:  protein:  O-­‐Man-­‐T;  Pmt1  family               GT39  

Pmt2     Dol-­‐P-­‐Man:  protein:  O-­‐Man-­‐T;  Pmt2  family               GT39  

P.  Orlean  56  SI  

Pmt3     Dol-­‐P-­‐Man:  protein:  O-­‐Man-­‐T;  Pmt2  family               GT39  

Pmt4     Dol-­‐P-­‐Man:  protein:  O-­‐Man-­‐T;  specific  for  membrane  proteins           GT39  

Pmt5     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

GPI  anchoring  

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  

P.  Orlean   57  SI  

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  58  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  

β -­‐1,6  glucan  formation  

    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  

Glycosidases,  cross-­‐linking  enzymes,  and  proteases  

Cts1     Endo-­‐chitinase                     GH18  

Cts2     Chitinase                       GH18  

Exg1/Bgl1   Major  exo-­‐β-­‐1,3-­‐glucanase  of  the  cell  wall;  soluble             GH5  

P.  Orlean   59  SI  

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  

GPI-­‐CWP  

Ecm33     Sps2  family;  structural/non-­‐enzymatic  

Pst1     Sps2  family;  structural/non-­‐enzymatic  

Sps2     Sps2  family;  structural/non-­‐enzymatic;  required  for  ascospore  wall  formation  

P.  Orlean  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   61  SI  

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  

  1CAZy  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).