Glycosylation defects activate filamentous growth Kss1 MAPK and inhibit osmoregulatory Hog1 MAPK
2009; Springer Nature; Volume: 28; Issue: 10 Linguagem: Inglês
10.1038/emboj.2009.104
ISSN1460-2075
AutoresHuiyu Yang, Kazuo Tatebayashi, Katsuyoshi Yamamoto, Haruo Saito,
Tópico(s)Ubiquitin and proteasome pathways
ResumoArticle16 April 2009free access Glycosylation defects activate filamentous growth Kss1 MAPK and inhibit osmoregulatory Hog1 MAPK Hui-Yu Yang Hui-Yu Yang Division of Molecular Cell Signaling, Institute of Medical Sciences, The University of Tokyo, Shirokanedai, Minato-ku, Tokyo, Japan Department of Biophysics and Biochemistry, Graduate School of Science, The University of Tokyo, Hongo, Bunkyo-ku, Tokyo, Japan Search for more papers by this author Kazuo Tatebayashi Kazuo Tatebayashi Division of Molecular Cell Signaling, Institute of Medical Sciences, The University of Tokyo, Shirokanedai, Minato-ku, Tokyo, Japan Department of Biophysics and Biochemistry, Graduate School of Science, The University of Tokyo, Hongo, Bunkyo-ku, Tokyo, Japan Search for more papers by this author Katsuyoshi Yamamoto Katsuyoshi Yamamoto Division of Molecular Cell Signaling, Institute of Medical Sciences, The University of Tokyo, Shirokanedai, Minato-ku, Tokyo, Japan Frontier Research Initiative, Institute of Medical Sciences, The University of Tokyo, Shirokanedai, Minato-ku, Tokyo, Japan Search for more papers by this author Haruo Saito Corresponding Author Haruo Saito Division of Molecular Cell Signaling, Institute of Medical Sciences, The University of Tokyo, Shirokanedai, Minato-ku, Tokyo, Japan Department of Biophysics and Biochemistry, Graduate School of Science, The University of Tokyo, Hongo, Bunkyo-ku, Tokyo, Japan Search for more papers by this author Hui-Yu Yang Hui-Yu Yang Division of Molecular Cell Signaling, Institute of Medical Sciences, The University of Tokyo, Shirokanedai, Minato-ku, Tokyo, Japan Department of Biophysics and Biochemistry, Graduate School of Science, The University of Tokyo, Hongo, Bunkyo-ku, Tokyo, Japan Search for more papers by this author Kazuo Tatebayashi Kazuo Tatebayashi Division of Molecular Cell Signaling, Institute of Medical Sciences, The University of Tokyo, Shirokanedai, Minato-ku, Tokyo, Japan Department of Biophysics and Biochemistry, Graduate School of Science, The University of Tokyo, Hongo, Bunkyo-ku, Tokyo, Japan Search for more papers by this author Katsuyoshi Yamamoto Katsuyoshi Yamamoto Division of Molecular Cell Signaling, Institute of Medical Sciences, The University of Tokyo, Shirokanedai, Minato-ku, Tokyo, Japan Frontier Research Initiative, Institute of Medical Sciences, The University of Tokyo, Shirokanedai, Minato-ku, Tokyo, Japan Search for more papers by this author Haruo Saito Corresponding Author Haruo Saito Division of Molecular Cell Signaling, Institute of Medical Sciences, The University of Tokyo, Shirokanedai, Minato-ku, Tokyo, Japan Department of Biophysics and Biochemistry, Graduate School of Science, The University of Tokyo, Hongo, Bunkyo-ku, Tokyo, Japan Search for more papers by this author Author Information Hui-Yu Yang1,2,‡, Kazuo Tatebayashi1,2,‡, Katsuyoshi Yamamoto1,3 and Haruo Saito 1,2 1Division of Molecular Cell Signaling, Institute of Medical Sciences, The University of Tokyo, Shirokanedai, Minato-ku, Tokyo, Japan 2Department of Biophysics and Biochemistry, Graduate School of Science, The University of Tokyo, Hongo, Bunkyo-ku, Tokyo, Japan 3Frontier Research Initiative, Institute of Medical Sciences, The University of Tokyo, Shirokanedai, Minato-ku, Tokyo, Japan ‡These authors contributed equally to this work *Corresponding author. Institute of Medical Sciences, University of Tokyo, 4-6-1 Shirokanedai, Minato-ku, Tokyo 108-8639, Japan. Tel.: +81 3 5449 5505; Fax: +81 3 5449 5701; E-mail: [email protected] The EMBO Journal (2009)28:1380-1391https://doi.org/10.1038/emboj.2009.104 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The yeast filamentous growth (FG) MAP kinase (MAPK) pathway is activated under poor nutritional conditions. We found that the FG-specific Kss1 MAPK is activated by a combination of an O-glycosylation defect caused by disruption of the gene encoding the protein O-mannosyltransferase Pmt4, and an N-glycosylation defect induced by tunicamycin. The O-glycosylated membrane proteins Msb2 and Opy2 are both essential for activating the FG MAPK pathway, but only defective glycosylation of Msb2 activates the FG MAPK pathway. Although the osmoregulatory HOG (high osmolarity glycerol) MAPK pathway and the FG MAPK pathway share almost the entire upstream signalling machinery, osmostress activates only the HOG-specific Hog1 MAPK. Conversely, we now show that glycosylation defects activate only Kss1, while activated Kss1 and the Ptp2 tyrosine phosphatase inhibit Hog1. In the absence of Kss1 or Ptp2, however, glycosylation defects activate Hog1. When Hog1 is activated by glycosylation defects in ptp2 mutant, Kss1 activation is suppressed by Hog1. Thus, the reciprocal inhibitory loop between Kss1 and Hog1 allows only one or the other of these MAPKs to be stably activated under various stress conditions. Introduction Under optimal growth conditions, the budding yeast Saccharomyces cerevisiae grows as round-shaped, single cells. On nitrogen starvation or in the presence of poor carbon sources, however, diploid cells of certain strains (such as Σ1278 h) initiate a fungus-like filamentous growth (FG) (Gimeno et al, 1992). Haploid cells also undergo a related phenomenon whereby the elongated cells invade solid media (Roberts and Fink, 1994). These responses, which we will refer to collectively as FG, are considered as an adaptive mechanism that allows normally sessile yeast to forage for scarce nutrients or to evade harmful waste products. An FG response requires at least three distinct protein kinases: the cAMP-dependent protein kinase Tpk2, the 5′-AMP-dependent protein kinase Snf1, and the MAP kinase (MAPK) Kss1 (Chen and Thorner, 2007). This article focuses on the signalling pathway that activates Kss1, namely, the FG MAPK pathway, whose core is a three-kinase cascade composed of the Ste11 MAPKKK, the Ste7 MAPKK and the Kss1 MAPK (Roberts and Fink, 1994). Another MAPK cascade, termed the high osmolarity glycerol (HOG) MAPK pathway, is activated by, and required for adaptation to, an increased external osmolarity (Brewster et al, 1993; Hohmann, 2002). The HOG MAPK pathway is composed of two functionally redundant upstream signalling branches that regulate the common Hog1 MAPK (Maeda et al, 1994, 1995). Of these branches, the SHO1 branch is closely related to the FG MAPK pathway. The SHO1 branch is initiated by an interplay between the membrane anchor protein Sho1 and the putative osmosensor membrane proteins Msb2 and Hkr1, leading to activation of Ste11, that is, the same MAPKKK that is involved in the FG MAPK pathway (Maeda et al, 1995; Tatebayashi et al, 2006, 2007). In the HOG pathway, however, activated Ste11 activates the Pbs2 MAPKK, which then activates the Hog1 MAPK (Posas and Saito, 1997) (see Figure 7A). Activated Hog1 governs a series of adaptive responses to high osmolarity, including temporarily arrest of the cell-cycle progression, readjustment of the transcription and translation patterns, and synthesis and intracellular retention of the compatible osmolyte glycerol (Bilsland-Marchesan et al, 2000; Teige et al, 2001; Hohmann, 2002; Escote et al, 2004; O'Rourke and Herskowitz, 2004). Although both the FG and the HOG MAPK pathways involve Ste11, each controls completely different, and mutually incompatible responses. Naturally, osmostress normally activates only the Hog1 MAPK (Posas and Saito, 1997). However, osmostress can activate the FG MAPK pathway in mutant cells, such as pbs2Δ or hog1Δ, in which the Hog1 MAPK cannot be activated (O'Rourke and Herskowitz, 1998; Davenport et al, 1999). In other words, activated Hog1 inhibits undesirable cross-talk activation of the FG MAPK pathway by osmostress in normal cells. In the pathogenic Candida albicans yeast, its Hog1 homolog inhibits, or raises the activation threshold of, the filamentation-inducing Cek1 MAPK (the Candida homolog of Kss1) by various natural inducers, thereby affecting its virulence (Eisman et al, 2006). Thus, inhibition of Kss1 by Hog1 is a conserved feature of the yeast MAPK pathways. It is, however, unknown whether or how an inhibition of cross-talk in the reverse orientation occurs, that is, does activation of the FG pathway suppress the HOG pathway? Using a novel approach based on the use of glycosylation defective mutants, we show here that activation of the FG-specific Kss1 MAPK in fact suppresses a concomitant activation of the HOG-specific Hog1 MAPK. When activation of Kss1 is blocked by mutations, such as ste7Δ or kss1Δ, the Hog1 MAPK is activated by glycosylation defects. Kss1 inhibits Hog1 indirectly through the Ptp2 protein phosphatase. Thus, the FG MAPK pathway and the HOG MAPK pathway suppress each other so that only the relevant pathway is activated by a specific stimulus. Results A combination of O- and N-glycosylation defects induces FUS1-lacZ expression To investigate the possible cross-regulation between the FG and the HOG pathways, a rapid and reliable method of activating the FG MAPK cascade is necessary. The commonly used method, namely the nutritional limitation, is inadequate, as it takes days to effect significant FG responses. A more promising approach is to use mutants that are defective in protein glycosylation. It has been shown that mutants of genes that are required for general glycosylation, such as OCH1, PMI40 and DPM1, constitutively activate an FG-like response (Lee and Elion, 1999; Cullen et al, 2000). However, because mutants of these genes are severely defective in protein glycosylation, they are either very sick or nonviable. It was also reported that the inhibitor of N-glycosylation tunicamycin moderately activated the FG MAPK pathway in rich media after 16 h (Cullen et al, 2000). As we will describe in this article, however, tunicamycin alone could not appreciably activate the FG MAPK pathway, at least under our experimental conditions. We, therefore, examined the possibility that a combination of separate, and more limited, defects in O-glycosylation and N-glycosylation might induce the FG response more robustly. We found that the commonly used reporters for the FG response, such as FRE-lacZ and FG(TyA)-lacZ, were unsuitable for our purposes because they had high baseline expression levels without stimulation and relatively moderate increases on stimulation (Mosch et al, 1996; Madhani and Fink, 1997; Davenport et al, 1999; Cullen et al, 2004). We, therefore, used the FUS1-lacZ reporter, which had much lower baseline expression levels and larger increases on stimulation (Cullen et al, 2000). Furthermore, all strains used in this study carried the ssk2Δ ssk22Δ double mutation unless otherwise specified, to eliminate any potential influence from the SLN1 branch of the HOG pathway. To inhibit O-glycosylation, we disrupted the genes that encode a family of protein O-mannosyltransferases (Pmt) (Gentzsch and Tanner, 1996; Girrbach and Strahl, 2003). Disruption of any of the six PMT genes (PMT1–PMT6), individually, had no significant effect on expression of FUS1-lacZ (Figure 1). Inhibition of N-glycosylation by tunicamycin, in wild-type cells, also did not induce FUS1-lacZ expression. In clear contrast, tunicamycin treatment of pmt4Δ mutant cells strongly induced FUS1-lacZ expression (Figure 1). Tunicamycin treatment of other pmt single mutants did not induce FUS1-lacZ expression. Furthermore, disruption of one or two additional PMT genes in the pmt4Δ background (e.g. pmt3/4/5Δ or pmt4/5/6Δ) did not have any stronger effect than pmt4Δ alone (Supplementary Figure S1). Thus, FUS1-lacZ expression is induced when N-glycosylation and a specific type of O-glycosylation is inhibited. Figure 1.Induction of the FUS1-lacZ reporter gene by glycosylation defects. The indicated yeast mutant cells carrying the pFUS1-lacZ plasmid were grown exponentially and were treated with (+tun) or without (−tun) tunicamycin (final conc. 25 μg/ml) for the indicated times. Activity of β-galactosidase in cell extracts was normalized using cell densities and expressed as Miller units (Miller, 1972). The strains used, and their full genotypes, are listed in Supplementary Tables S1 and S2, respectively. Download figure Download PowerPoint Induction of FUS1-lacZ expression by glycosylation defects requires signalling elements of the FG pathway We next examined whether induction of FUS1-lacZ expression by glycosylation defects is due to the activation of the FG pathway, because the FUS1-lacZ reporter can be induced also by the mating MAPK pathway. To determine which pathway is activated by the glycosylation defects, we disrupted, in the pmt4Δ background, genes that are specific for, or shared between, the mating and FG pathways. The results of an extensive survey are summarized in Figure 2A, which is a compilation of data from several different experiments. Although the mean value of the positive control strain (pmt4Δ) was somewhat variable from experiment to experiment (the range is between 34.3 and 61.9), only a typical control result is shown to save space. Thus, the exact values may not be directly comparable, but the following general conclusion is statistically valid. Figure 2.Signalling elements required for induction of FUS1-lacZ expression by glycosylation defects. (A) Induction of the FUS1-lacZ reporter gene by glycosylation defects. The indicated yeast mutant cells carrying the pFUS1-lacZ plasmid were grown exponentially, treated with (+tun) or without (−tun) tunicamycin for 8 h, and assayed for β-galactosidase activity as in Figure 1. (B) The indicated mutants were assayed as in (A). (C) Exponentially growing cells of the indicated genotypes were treated with 25 μg/ml tunicamycin for the indicated times. Cell extracts were probed for phosphorylated (activated) forms of Fus3 (P-Fus3), Kss1 (P-Kss1) and Mpk1 (P-Mpk1) by immunoblotting using anti-phospho-p44/42 antibody. Download figure Download PowerPoint From these data, two components in FUS1-lacZ expression are discernible: the basal component seen without tunicamycin treatment, and the tunicamycin-induced component. On the basis of the differential effect of the mutants on these two components, we could classify the mutants into four groups. The first group (ste4Δ and ste5Δ) abolished only the basal component without affecting the induced component. The second group, composed of ste20Δ, ste50Δ, ste11Δ, ste7Δ, ste12Δ and the fus3Δ kss1Δ double mutation, abolished both the basal and induced components. The third group, composed of msb2Δ, opy2Δ and ptp2Δ, affected only the induced component. Finally, the fourth group (tec1Δ, fus1Δ, sho1Δ, hkr1Δ, hog1Δ, yps1Δ and ptp3Δ) affected neither the basal nor the induced expression levels. Genes in groups 1 and 2, whose disruption abolishes the basal components, are known to be involved in the mating pathway. On the other hand, genes in the groups 2 and 3, whose disruption abolishes the tunicamycin-induced components, are known to be required for FG response, with the notable exception of PTP2. Any role of the Ptp2 protein tyrosine phosphatase (PTP) in the FG MAPK pathway has not been known or suspected. Later in this article, we will explain why inactivation of the PTP2 gene suppresses FUS1-lacZ expression. Tec1 is required for the full FG response but is dispensable for induction of the FUS1-lacZ reporter (Madhani and Fink, 1997). Thus, in general, there is a good concordance between the genes that are required for the FUS1-lacZ expression induced by glycosylation defects and those that have been shown to be involved in the FG MAPK pathway. Glycosylation defects induce characteristic FG responses We examined whether glycosylation defects do indeed induce the FG response. A hallmark of the FG response is activation of the Kss1 MAPK, as opposed to the preferential activation of the Fus3 MAPK by the mating pathway (Chen and Thorner, 2007). As shown in Figure 2B, disruption of KSS1 completely abolished tunicamycin-induced FUS1-lacZ expression, whereas disruption of FUS3 only moderately reduced it, indicating that the Kss1 MAPK is mainly responsible for FUS1-lacZ expression by glycosylation defects. We then examined whether glycosylation defects activate Kss1 and/or Fus3 by probing for activation-associated phosphorylation of these kinases. The combination of a pmt4Δ mutation and tunicamycin treatment activated Kss1 but not Fus3 (Figure 2C). Disruption of STE7, which encodes the direct activator of Kss1, completely inhibited Kss1 phosphorylation, whereas disruption of STE4, which encodes an upstream signalling element in the mating pathway, had no effect. The preferential activation of Kss1 by glycosylation defects is thus consistent with activation of the FG pathway. Tunicamycin also activates the cell-wall integrity Mpk1 MAPK pathway (Bonilla and Cunningham, 2003; Cohen et al, 2008). Phosphorylation of Mpk1, however, occurs very slowly after tunicamycin treatment and is independent of Kss1/Fus3 (Figure 2C, upper panel). As there was no apparent connection between Mpk1 and the FUS1-lacZ response under study, we did not analyse Mpk1 any further. We also examined whether glycosylation defects induce FG. As tunicamycin inhibits cell growth, however, it cannot be used to induce FG. Therefore, as an alternative to tunicamycin, we used 2-butanol that is known to accelerate FG in liquid culture (Lorenz et al, 2000; Andersson et al, 2004). When the parental (PMT4+) yeast strain was grown in SC medium containing 1% 2-butanol, no FG was observed over a 16-h period (Supplementary Figure S2). In clear contrast, pmt4Δ mutant cells became elongated and formed filaments. The pmt4Δ msb2Δ double mutant cells did not form any filaments. Thus, we conclude that O-glycosylation defects caused by pmt4Δ significantly lower the threshold of other stimuli for induction of the FG response. Opy2 and Msb2 are O-glycosylated by Pmt4 Next, we investigated the potential Pmt4 target(s) whose underglycosylation might activate the FG response. Pmt4 is unique among the protein O-mannosyltransferases in that it specifically glycosylates Ser/Thr-rich extracellular segments in membrane-attached proteins (Hutzler et al, 2007). This fact suggests that two membrane proteins required in the FG response, Opy2 and Msb2, are both potential substrates of Pmt4. Indeed, Opy2 has been shown to be glycosylated by Pmt4 (Hutzler et al, 2007). Opy2 has two potential glycosylated Ser-rich (SR) regions, SR1 (aa 2–22) and SR2 (aa 75–84), in its extracellular domain (Figure 3A). The full-length, GFP-tagged Opy2 protein (Opy2-FL-GFP) isolated from PMT4+ cells was significantly larger than the same protein isolated from pmt4Δ cells (Figure 3B, compare lanes 1 and 5), indicating Pmt4-dependent glycosylation of Opy2. Deletion of the SR2 region did not reduce the Pmt4-dependent glycosylation (Figure 3B, lanes 3 and 7). In contrast, Opy2-ΔSR1-GFP had the same size in PMT4+ and pmt4Δ cells (Figure 3B, lanes 2 and 6), indicating that Pmt4 modifies only the SR1 region of Opy2. Figure 3.Glycosylation of Opy2 by Pmt4, and its role in the FG response. (A) Schematic diagrams of the Opy2 constructs used in (B–D). FL, full-length; SR, Ser-rich; TM, transmembrane segment. (B) C-terminally GFP-tagged Opy2 constructs were generated using a single-copy vector with the GAL1 promoter (p414GAL1). Expression of Opy2-FL-GFP, or the indicated deletion constructs, was induced by 2% galactose for 4 h in PMT4+ or pmt4Δ host cells. Total cell extracts were probed with an anti-GFP antibody by immunoblotting. The horizontal arrow indicates cleavage products. The estimated cleavage site is shown in (A) by an upright arrow. (C) The full-length OPY2 gene (with the OPY2 promoter) carried on a single-copy vector (pRS414) or its deletion constructs were individually expressed in pmt4Δ ste4Δ opy2Δ host cells that also carried the pFUS1-lacZ reporter plasmid. Cells were treated with (+tun) or without (−tun) tunicamycin (final conc. 25 μg/ml) for 8 h, before extracts were prepared for β-galactosidase assays. (D) GST-tagged Ste50 (GST-Ste50), or GST alone, was constitutively expressed using pFP122 or the vector p426TEG1, and expression of GFP-tagged Opy2-FL (Opy2-FL-GFP), or Opy2ΔC-GFP, was induced by 2% galactose for 4 h using the expression constructs based on the p414GAL1 vector, in pmt4Δ ste50Δ opy2Δ host cells. GST-Ste50 (or GST) was immunoprecipitated by glutathione-Sepharose beads (bottom panel), and co-precipitated Opy2-GFP protein was detected by immunoblotting (top panel). Expression levels of the Opy2-GFP proteins are shown in the middle panel. Download figure Download PowerPoint Msb2 is also a glycosylated transmembrane protein (Cullen et al, 2004). Its extracellular domain contains an extended Ser/Thr rich (STR) region (aa 52–956) that is likely to be O-glycosylated (Hutzler et al, 2007) (see Figure 4A). The full-length HA-tagged Msb2 (Msb2-FL-HA) isolated from pmt4Δ cells migrated slightly faster in SDS–PAGE than the same molecule obtained from PMT4+ cells, indicating a contribution of Pmt4-dependent glycosylation. However, both molecules migrated at positions much higher than the calculated molecular weight of Msb2-FL-HA (ca. 150 kDa), reflecting the presence of Pmt4-independent glycosylation as well (Figure 4B). To clarify the roles of individual Pmt enzymes in Msb2 glycosylation, we used Msb2-ΔSTR1-HA and Msb2-ΔSTR2-HA for further analyses (see Figure 4A). The smaller size of these constructs allowed easier evaluation of the effect of pmt mutations. Both proteins, when expressed in pmt1Δ, pmt2Δ or pmt4Δ mutant cells, were smaller than when expressed in PMT+, pmt3Δ, pmt5Δ or pmt6Δ cells (Figure 4C and D), indicating that Pmt1, Pmt2 and Pmt4 contribute to Msb2 O-glycosylation. The fact that pmt1Δ and pmt2Δ reduced Msb2 glycosylation to a similar extent is consistent with the earlier observation that Pmt1 and Pmt2 form a functional heterodimer (Girrbach and Strahl, 2003). More important, the size reduction of Msb2 in pmt4Δ cells is quite different from that in pmt1Δ and pmt2Δ cells, clearly indicating that Pmt4 contributes to a distinct subset of Msb2 O-glycosylation. The Msb2-ΔSTR3-HA mutant protein, in which most of the STR region is absent, migrated closer to the calculated peptide molecular weight (ca. 50 kDa) (Figure 4E). Taken together, we conclude that Pmt4 plays a significant role in O-glycosylation of Msb2. Pmt1 and Pmt2 also glycosylate Msb2, but such glycosylation is either irrelevant or insufficient for the FG response. Figure 4.Pmt4 glycosylation of Msb2. (A) Schematic diagrams of the deletion constructs of Msb2 used in (B–F). Each construct contains two HA epitopes, one following amino acid position 48 and another at the C terminus. STR, Ser/Thr rich; HMH, Hkr1-Msb2 Homology domain; TM, transmembrane segment; FL, full-length. (B–E) Indicated Msb2-HA constructs, under the control of the GAL1 promoter in the single-copy vector p416GAL1, were induced by 2% galactose for 3 h in PMT+ or the indicated pmt mutant cells. Cell extracts were probed by immunoblotting with an anti-HA antibody. (F) Endo H treatment of Msb2. The Msb2-ΔSTR2-HA construct, under the control of the GAL1 promoter in the single-copy vector p416GAL1, was induced by 2% galactose for 3 h in PMT + cells and in the indicated pmt mutant cells. Cell extracts were prepared in Buffer N, immunoprecipitated with anti-HA antibody (3F10), treated with (+) or without (−) 0.4 U/ml endoglycosidase H (Endo H) at 37°C for 15 h, applied to SDS–PAGE, and probed with another anti-HA antibody (12CA5). Download figure Download PowerPoint Msb2 has a number of potential N-glycosylation sites in addition to its numerous O-glycosylation sites. As defects in Pmt4-dependent O-glycosylation can interfere with the N-glycosylation status of the same protein (Ecker et al, 2003), we estimated the degree of Msb2 N-glycosylation by endoglycosidase H (Endo H) treatment. To this end, the Msb2-ΔSTR2-HA mutant protein was expressed in PMT+, pmt1Δ and pmt4Δ host cells, and the size of the protein before and after Endo H treatment was compared by SDS–PAGE. As shown in Figure 4F, Endo H treatment reduced the size of Msb2-ΔSTR2-HA to a similar extent irrespective of the host pmt mutation. Thus, at least in these cases, O-glycosylation defects did not substantially affect the status of Msb2 N-glycosylation. In some cases, O-glycosylation is important for targeting proteins to the plasma membrane (Proszynski et al, 2004). Using YFP-fusion constructs, we examined whether removal of a substantial portion of the O-glycosylated STR region might affect the subcellular localization of Msb2. As shown in Supplementary Figure S3, both Msb2-FL-YFP and its deletion derivative Msb2-ΔSTR4-GFP are similarly localized on the plasma membrane, although a substantial amount of both constructs is also found in intracellular compartments, as observed earlier for Msb2-FL-YFP (Tatebayashi et al, 2007). Thus, removal of a substantial portion of the O-glycosylated STR region does not affect the cellular localization of the Msb2 protein. Glycosylation defects of Msb2, but not of Opy2, activate the FG MAPK pathway We then asked whether defective glycosylation of Opy2 or Msb2 induces the FG response. First, we examined the possibility that the glycosylated region in Opy2 is required for the FG response. However, an Opy2 mutant lacking the entire O-glycosylated domain (Opy2-ΔSR1) could fully support FUS1-lacZ expression (Figure 3C). In contrast, deletion of the cytoplasmic C-terminal region (aa 184–360) completely abolished FUS1-lacZ expression (Figure 3C). Although the full-length Opy2 protein binds to Ste50, Opy2-ΔC does not (Figure 3D). These results are consistent with the idea that the major function of Opy2 is to anchor Ste50 to the membrane, which is necessary for efficient signalling in the FG and HOG pathways (Wu et al, 2006; Tatebayashi et al, 2007). However, Opy2 glycosylation, or lack of it, may not be a factor in the FG response. We then examined the possible role of the Msb2 glycosylation region in the FG response. When Msb2-ΔSTR4, in which a large segment of the glycosylated STR domain is deleted, was expressed in pmt4Δ host cells using the MSB2 own promoter, FUS1-lacZ was induced even in the absence of tunicamycin (Figure 5A and B). When less extensive deletion mutants, Msb2-ΔSTR7 and Msb2-ΔSTR8, were expressed, weaker induction of FUS1-lacZ was observed in the absence of tunicamycin. Expression of the least extensive deletion mutant, Msb2-ΔSTR5, did not induce FUS1-lacZ. Thus, there was a clear correlation between the extent of STR deletion and the extent of FUS1-lacZ induction in the absence of tunicamycin. All these Msb2 STR deletion mutants supported tunicamycin-induced FUS1-lacZ expression to similar extent. Thus, deletion of the glycosylated STR segment constitutively activated Msb2. Indeed, overexpression of Msb2-ΔSTR4 using an inducible GAL1 promoter activated the FG pathway even in glycosylation-proficient PMT4+ host cells, whereas expression of full-length Msb2 did not (Figure 5C). Thus, deletion of the STR region has a similar effect as glycosylation defects. In contrast, overexpression of glycosylation-defective Opy2-ΔSR1 did not activate the FG MAPK pathway at all, further supporting the earlier conclusion that glycosylation of Opy2 does not affect the FG MAPK pathway. Combining all the data, we conclude that deficiency of Pmt4-mediated O-glycosylation of Msb2 in the STR region is a part of signal that activates the FG MAPK pathway. Figure 5.Domains of Msb2 essential for the FG response. (A) Schematic diagrams of the Msb2 deletion constructs used in (B) and (C). STR, Ser/Thr rich; HMH, Hkr1-Msb2 Homology domain; TM, transmembrane segment; FL, full-length. (B) Induction of the FG reporter gene FUS1-lacZ by glycosylation defects. The full-length MSB2 gene, or its deletion constructs, carried on the single-copy vector pRS416 were individually expressed in pmt4Δ ste4Δ msb2Δ host cells in which the FUS1-lacZ reporter gene has been integrated. Cells were treated with (+tun) or without (−tun) tunicamycin (final conc. 25 μg/ml) for 8 h, before extracts were prepared for β-galactosidase assays. In HMH(Hkr1), the HMH domain of Msb2 was replaced by the homologous HMH domain from Hkr1. (C) Expression of the indicated Msb2 or Opy2 construct, placed under the GAL1 promoter, was induced by galactose for 2 h. Induction of the FG reporter gene FUS1-lacZ was assayed. Download figure Download PowerPoint Msb2 has seven potential N-glycosylation sites that conform to the N-X-T/S motif (X being any aa other than P) in its extracellular domain (Asn-30, Asn-859, Asn-885, Asn-945, Asn-1049, Asn-1088 and Asn-1175). To test whether the effect of tunicamycin can be accounted for by defective Msb2 N-glycosylation, we generated a mutant termed Msb2-6N/A in which six of the potential N-glycosylation sites were mutated to Ala (Asn-30, which is outside the STR region, was left unmodified). Expression of the Msb2-6N/A construct in pmt4Δ host cells at the endogenous level (i.e. from the MSB2 promoter in a single-copy plasmid) did not induce FUS1-lacZ expression any better than wild-type Msb2, either in the absence or presence of tunicamycin (Supplementary Figure S4). Likewise, overexpression of Msb2-6N/A in pmt4Δ host cells using the GAL1 promoter did not induce FUS1-lacZ expression any better than wild-type Msb2 (Supplementary Figure S5). These results suggest that N-glycosylation defects in Msb2 alone are not sufficient to account for the effect of tunicamycin, and that there might be (an)other
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