O-mannosylation is a crucial protein modification in eukaryotes that is initiated by the essential family of protein O-mannosyltransferases (PMTs). Here we demonstrate that in the model yeast Saccharomyces cerevisiae rhodanine-3-acetic acid derivatives affect members of all PMT subfamilies. Specifically, we used OGT2468 to analyse genome-wide transcriptional changes in response to general inhibition of O-mannosylation in baker's yeast. PMT inhibition results in the activation of the cell wall integrity (CWI) pathway. Coinciding, the mitogen-activated kinase Slt2p is activated in vivo and CWI pathway mutants are hypersensitive towards OGT2468. Further, induction of many target genes of the unfolded protein response (UPR) and ER-associated protein degradation (ERAD) is observed. The interdependence of O-mannosylation and UPR/ERAD is confirmed by genetic interactions between HAC1 and PMTs, and increased degradation of the ERAD substrate Pdr5p* in pmtΔ mutants. Transcriptome analyses further suggested that mating and filamentous growth are repressed upon PMT inhibition. Accordingly, in vivo mating efficiency and invasive growth are considerably decreased upon OGT2468 treatment. Quantitative PCR and ChIP analyses suggest that downregulation of mating genes is dependent on the transcription factor Ste12p. Finally, inhibitor studies identified a role of the Ste12p-dependent vegetative signalling cascade in the adaptive response to inhibition of O-mannosylation.
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Glycosylation is an essential and abundant protein modification (Lehle et al., 2006). The majority of secreted and membrane proteins in biological systems are glycosylated and glycan chains influence a large number of biological processes (Van den Steen et al., 1998).
Protein O-mannosylation represents a conserved modification among eukaryotes and some bacteria (Lommel and Strahl, 2009). O-mannosyl glycans are short linear oligosaccharides linked via α-glycosidically bound mannose to serine (Ser) and threonine (Thr) residues (Willer et al., 2003). Biosynthesis is initiated in the lumen of the endoplasmic reticulum (ER) by the transfer of mannose from dolichyl phosphate-activated mannose to Ser or Thr residues of proteins (Lehle et al., 2006). Further chain elongation takes place in the Golgi apparatus using nucleotide activated sugars as donors. Defects in the assembly of O-mannosyl glycans are of fatal consequences in diverse organisms ranging from yeasts to human (Lommel and Strahl, 2009). For instance, in human lack of O-mannosylation results in severe congenital muscular dystrophies with neuronal migration defects (Messina et al., 2008).
The initial O-mannosyltransfer reaction is catalysed by an essential family of dolichyl phosphate-d-mannose:protein O-mannosyltransferases (PMTs) that is conserved from yeast to humans (Lommel and Strahl, 2009). PMTs have been identified and extensively characterized in baker's yeast. In Saccharomyces cerevisiae, the PMT family is highly redundant (Pmt1p–Pmt6p) (Strahl-Bolsinger et al., 1993; Gentzsch and Tanner, 1996) and is subdivided into the PMT1, PMT2 and PMT4 subfamilies (Girrbach et al., 2000). In wild-type (WT) yeast cells O-mannosylation is predominantly catalysed by distinct heteromeric Pmt1p–Pmt2p and homomeric Pmt4p–Pmt4p complexes (Girrbach and Strahl, 2003) that exhibit different substrate specificities and mannosylate different protein substrates in vivo (Gentzsch and Tanner, 1997; Hutzler et al., 2007). The simultaneous knockout of particular combinations of PMT family members in S. cerevisiae results in lethality (e.g., pmt1Δpmt2Δpmt4Δ) demonstrating that protein O-mannosylation is an essential protein modification (Gentzsch and Tanner, 1996). Further, the characterization of viable single and double pmtΔ mutants revealed that O-mannosylation serves a variety of different functions; it is required for stability, sorting and localization of proteins thereby affecting protein function and being indispensable for cell wall integrity, cell polarity and morphogenesis (Lehle et al., 2006). In addition, O-mannosylation is crucial for processing of aberrantly folded proteins in the ER (Harty et al., 2001; Nakatsukasa et al., 2004; Hirayama et al., 2008).
For an improved and comprehensive understanding of protein O-mannosylation it is crucial to understand global changes in gene expression upon loss of O-mannosylation. Viable mutants lacking individual PMTs commonly adapt to decreased amounts of O-mannosyl glycans by evolving compensatory mechanisms, like the upregulation of other PMT family members and the formation of alternative PMT complexes (Girrbach and Strahl, 2003). In addition, more general compensatory changes were observed in glycosylation mutants that allow near-normal growth of cells, like the upregulation of chitin synthesis (Kapteyn et al., 1999) or the activation of metabolic and stress response pathways (Cullen et al., 2000; Klebl et al., 2001; Cantero et al., 2007). Thus, long-term adaptation mechanisms of pmtΔ mutants might actually differ from immediate global changes in gene expression upon loss of O-mannosylation. Recently, rhodanine-3-acetic acid derivatives have been described as potent inhibitors of Candida albicans Pmt1p (Orchard et al., 2004). These compounds turned out to specifically inhibit CaPmt1p activity in vitro and in vivo (Orchard et al., 2004; Cantero et al., 2007). The exact mode of action of the CaPmt1p inhibitors is not understood. However, these compounds provide strong means to study the physiological impact of protein O-mannosylation.
In this study we demonstrate that in contrast to C. albicans, in baker's yeast rhodanine-3-acetic acid derivatives affect the members of all three PMT subfamilies. Taking advantage of this circumstance, we use OGT2468 to inhibit O-mannosylation and present for the first time a genome-wide transcriptional analysis of the response to a general inhibition of O-mannosylation in a eukaryotic organism. Biological meaning of this response is provided.
OGT2468 is a general inhibitor of PMTs in S. cerevisiae
The analysis of rhodanine-3-acetic acid derivatives (e.g., OGT1458, OGT1576, OGT2078 and OGT2468; Orchard et al., 2004) effects on S. cerevisiae revealed that the inhibitors affect in vitro activity not only of ScPmt1p, but of several PMT1 and PMT2 subfamily members (Table S1), in contrast to the effect reported on C. albicans. Enzymatic activities of other ER-resident glycosyltransferases such as dolichol-phosphate-mannose synthase are not affected (data not shown). Thus, we further characterized in vivo effects, thereby focusing on the most potent inhibitor OGT2468. In liquid culture, WT S. cerevisiae strain SEY6210 was treated with increasing amounts of OGT2468 (final concentrations 0.05–10 µM). Determination of the number of survivors over time revealed inhibition of growth that was dosage dependent. However, OGT2468 concentrations above 1 µM did not further decrease the number of survivors significantly during periods of incubation between 3 and 5 h (Fig. S1A). Based on these results, for further phenotypic characterization 1 µM OGT2468 was applied for 3 h. Light microscopy analysis of WT cells treated with OGT2468 revealed that cell morphology and separation is affected (Fig. 1A). Cells aggregated and formed huge clumps, a phenotype that is typical for viable single and double mutants of members of the PMT1 and PMT2 subfamilies (Strahl-Bolsinger et al., 1993; Gentzsch and Tanner, 1996). In addition, OGT2468 treatment changed the regular axial budding pattern of haploid yeast cells into uni- and bipolar budding pattern (Fig. 1A, image detail), a phenotype that was described for pmt4Δ mutants (Sanders et al., 1999). To further elucidate the specificity of PMT inhibition, in vivo O-mannosylation of different PMT substrate proteins was analysed (Fig. 1B). It turned out that Pmt1p/Pmt2p substrate proteins [FUSw/oTMZZ (Hutzler et al., 2007)] and Pmt4p substrates [FUSwTMZZ (Hutzler et al., 2007)] were hypo-glycosylated upon OGT2468 treatment (Fig. 1B).
To further corroborate that in S. cerevisiae OGT2468 is not a Pmt1p-specific inhibitor, OGT2468 sensitivity of cells lacking representatives of any of the three PMT subfamilies (pmt1Δ, pmt2Δ and pmt4Δ) or all PMT1 subfamily members (pmt1Δpmt5Δ) was characterized. Cells were grown in liquid medium containing increasing amounts of inhibitor as detailed in Experimental procedures. Although with some variations all pmtΔ mutants are hypersensitive to the inhibitor compared with WT cells (Fig. 1C). Again, S. cerevisiae OGT2468 sensitivity differs from C. albicans, where growth of pmt4Δ but not pmt1Δ mutants is affected by rhodanine-3-acetic acid derivatives (Cantero et al., 2007).
In summary these data demonstrate that in S. cerevisiae rhodanine-3-acetic acid derivatives affect not only Pmt1p, but members of all three PMT subfamilies. Thus they are excellent tools to inhibit protein O-mannosylation in general.
Global transcriptional response of WT yeast to inhibition of protein O-mannosylation
We took advantage of PMT inhibitors to analyse the short-term transcriptional response upon inhibition of protein O-mannosylation by the bias of genome-wide transcriptional profiling of S. cerevisiae WT strain SEY6210 treated with OGT2468. Mid-log-phase cells were grown in the presence of 0.1 µM OGT2468, 1 µM OGT2468 and DMSO 1% (OGT2468 solvent), respectively, for 3 h and their transcriptional profiles were characterized using Affymetrix GeneChips as described in Experimental procedures. The effect of the inhibitor on gene expression was evaluated comparing the transcriptional profiles of OGT2468-treated versus control non-treated cells. Microarray analyses revealed that 167 genes were induced and 64 genes repressed upon treatment with 1 µM OGT2468. The majority of these genes were also affected upon treatment with 0.1 µM OGT2468, yet with a clear dosage-dependent response. Complete data sets of genes with statistically significant regulation of transcript levels are included in Tables S2 (ratios of transcript levels ≥ 2.0) and S3 (ratios of transcript levels ≤ 0.5).
Functional classification of the response based on SGD and YPD Proteome databases showed that the upregulated genes cover a variety of physiological processes (Table 1). Among them we found a variety of functional clusters, namely cell wall biogenesis and metabolism; stress response and transport; signal transduction and protein fate that were significantly induced by PMT inhibition (Table 1 and Table S2). Further statistical analysis using the web-based tool Genecodis (http://genecodis.dacya.ucm.es/) (Carmona-Saez et al., 2007) confirmed these categories with P-values highly significant (data not shown). Similarly, specific functional subgroups could be extracted from the data set of repressed genes (Table 2 and Table S3) and two of these categories were further confirmed by Genecodis, namely mating (P-value 0) and sporulation (P-value 6.5E-3). In addition, a group of genes involved in pseudohyphal growth (TEC1, MUC1, PGU1; P-value 6.5E-3) was also identified.
Table 1. Functional categories and number of genes induced in response to OGT2468 and in pmtΔ mutants.
Table 2. Functional categories and number of genes repressed in response to OGT2468 and in pmtΔ mutants.
Global transcriptional response of pmt1Δpmt2Δ and pmt1Δpmt4Δ mutants
To further recognize crucial transcriptional responses in consequence of decreased O-mannosylation and compare short- and long-term responses, we performed genome-wide transcriptional profiling of viable pmt1Δpmt2Δ and pmt1Δpmt4Δ double mutants (Gentzsch and Tanner, 1996), where the major protein O-mannosyltransferases were deleted. Transcriptional profiling using Affymetrix GeneChips is described in Experimental procedures. Complete functional data sets of genes statistically significantly induced and repressed in these mutants are included in Tables S2 and S3.
Transcription of 204 genes and of 432 genes was induced in pmt1Δpmt2Δ and pmt1Δpmt4Δ mutants respectively (ratios of transcript levels ≥ 2.0). This indicates that transcriptional induction is enhanced in pmtΔ mutants compared with the transcriptional response upon immediate inhibition of O-mannosylation by OGT2468. Further, 28 genes were repressed in pmt1Δpmt2Δ and 54 genes in pmt1Δpmt4Δ mutants (ratios of transcript levels ≤ 0.5).
Functional classification of the induced and repressed genes in the pmtΔ mutants mainly revealed the functional clusters previously identified for the OGT2468 response (Tables 1 and 2). The number of genes related to metabolism significantly increased in both pmtΔ mutants (Table 1). In contrast, similar numbers of cell wall-related genes were affected in pmtΔ mutants and in response to OGT2468 (Table 1). Thirty-eight per cent of those genes were affected in all three conditions, and 69% in at least two of the conditions analysed (Fig. 2B, Tables S2 and S3). Comparison of the transcriptional profiles of the pmtΔ mutants revealed that transcription of 139 genes (67%) of the total genes upregulated in pmt1Δpmt2Δ is also induced in the pmt1Δpmt4Δ strain (Fig. 2A). Out of the 167 genes induced by OGT2468 treatment, 108 (65%) were also induced either in pmt1Δpmt2Δ, pmt1Δpmt4Δ or both mutants (Fig. 2 and Table S2), whereas 71 genes were induced in all three conditions (Fig. 2, Table S2 and Fig. S2). The functional catalogue of the co-induced response mainly includes genes related to cell wall maintenance, signal transduction, stress, metabolism and genes of unknown function as well as several genes related to mannosylation, protein fate and transport (Fig. 2B).
Forty-seven genes are upregulated only in mutant pmt1Δpmt2Δ, and 274 genes are specifically overexpressed in mutant pmt1Δpmt4Δ (Fig. 2A), indicating a broader response spectrum and adaptation mechanism attributable to the absence of Pmt4p. Genes related to transport, stress, protein fate as well as meiosis/sporulation are significantly upregulated in this mutant compared with pmt1Δpmt2Δ and OGT2468-treated cells (Table 1 and Table S2). Among the 68 genes that are upregulated exclusively in both pmtΔ mutants (Fig. 2A and Table S2), 29% of them are related to metabolism but only 13% to stress and signal transduction, and 1% to cell wall biogenesis. Genes related to metabolism were analysed using the Cytoscape plug-in ClueGO that integrates gene ontology (GO) terms as well as KEGG/BioCarta pathways and creates a functionally organized GO/pathway term network (Bindea et al., 2009). The analysis revealed that the elevated number of induced genes in pmtΔ mutants is mainly due to auxotrophies complemented in the course of PMT1, PMT2 and PMT4 disruption (data not shown). Further, it cannot be excluded that part of the transcriptional responses of the pmtΔ mutants are related to slow growth phenotypes (Gentzsch and Tanner, 1996).
Regarding the repression response, 17 genes (60% of total genes downregulated in pmt1Δpmt2Δ) were repressed in both pmtΔ mutants (Fig. 2A and B). Out of the 64 genes repressed by OGT2468 treatment, 23 were repressed in pmt1Δpmt2Δ or/and pmt1Δpmt4Δ mutants, the functional catalogue of this cluster being mainly related to mating and filamentation (Fig. 2B). Eleven of these genes were downregulated in all three conditions, 72% of those related to mating (Fig. 2B and Table S3). Genecodis revealed that the functional categories with P-values highly significative within this cluster were cell adhesion (P-value 9.6E-8) and sexual reproduction (P-value 1.2E-7), both processes crucial for mating ability.
Comparison of the response to OGT2468 with other genome-wide transcriptional analyses
To gain additional insight into the impact of inhibition of O-mannosylation, we compared the OGT2468 transcriptional response to transcriptional profiles based on block of N-glycosylation (Travers et al., 2000), inhibition of disulphide bond formation (Travers et al., 2000), osmotic stress (O'Rourke and Herskowitz, 2004), heat shock, oxidative stress (Hughes et al., 2000) and cell wall stress (Garcia et al., 2004). Moreover, genome-wide transcriptional profiles of cell wall-related mutants gas1Δ, fks1Δ, kre6Δ, mnn9Δ, knr4Δ (Lagorce et al., 2003) and those from yeast cells with a constitutive activation of the cell wall integrity (CWI) pathway (Roberts et al., 2000) were included. Hierarchical clustering revealed three clusters of genes that are induced upon OGT2468 treatment, in pmtΔ and cell wall mutants as well as in response to cell wall stress conditions (Fig. 3, clusters 1, 3 and 6). These gene clusters include key elements necessary for cell wall structure and remodelling such as CRH1, PST1, CWP1, PIR3, DFG5, GFA1, GSC2, FKS2 and regulatory components like SLT2 and MLP1. These genes are also induced in cells with constitutive activation of the CWI pathway, suggesting a crucial role for this pathway in the PMT inhibition response regulation. Further, inhibition of O-mannosylation, of N-glycosylation and interference with ER protein folding induce a specific set of genes including genes related to protein glycosylation (PMT3, OCH1, MNT4, MNN4) and protein fate (ERO1, YPT53, MPD1, EUG1, LMS1) (Fig. 3, clusters 4 and 5). Interestingly, the majority of the genes in cluster 5 are almost not affected in any of the other conditions analysed. Hierarchical clustering of the 71 genes co-induced in response to OGT2468 and pmtΔ mutants with the above mentioned transcriptional responses confirmed the clusters identified (Fig. S2).
Most of the genes repressed by OGT treatment and in pmtΔ mutants are related to mating (Table 2). A good part of these genes are repressed particularly under constitutive activation of the CWI pathway, but not in response to cell wall stress and in cell wall mutants (Fig. 3, cluster 7). In contrast, cluster 8 depicts a set of genes that are repressed in most of the conditions analysed (Fig. 3, cluster 8).
In summary, hierarchical clustering suggested that defective O-mannosylation predominantly affects cell wall- and ER stress-associated processes as well as mating and filamentation.
Inhibition of O-mannosylation activates the CWI pathway
In S. cerevisiae, defects in cell wall integrity are sensed at the plasma membrane and eventually result in the activation of the protein kinase C. Pkc1p turns on the CWI pathway including the mitogen-activated protein (MAP) kinase Slt2p, which leads to a number of cellular responses. One of those is the transcriptional activation of a variety of genes that have been implicated in cell wall assembly and structure (reviewed in Levin, 2005).
To confirm the transcriptional induction of genes related to cell wall integrity in decreased O-mannosylation conditions we quantified relative mRNA levels of some selected genes by quantitative RT-PCR (qRT-PCR). The transcriptional change upon OGT2468 treatment was calculated as the ratio of the transcript levels of treated cells versus non-treated cells. In agreement with the microarray data, GFA1, GSC2, CRH1 and SRL3, all of them previously reported as CWI pathway-dependent (Jung and Levin, 1999; Garcia et al., 2004), were specifically induced in the presence of OGT2468 (Fig. 4A). Furthermore, the induction of genes like CRH1 and GFA1 was, at least partially, dependent on Rlm1p, a transcription factor regulated by the MAPK Slt2p (data not shown).
To further explore induction of the CWI pathway, we analysed the activation of Slt2p in response to OGT2468. Slt2p phosphorylation was determined by Western blot in WT cells after treatment with OGT2468 (1 µM) using a phospho-specific p44/42 MAP kinase antibody. As shown in Fig. 4B, inhibition of O-mannosylation activated Slt2p, whereby the amount of active Slt2p increased over time from 1 to 4 h of OGT2468 treatment. In addition, we analysed whether the induction of this compensatory response is crucial for cells to survive when O-mannosylation is on the decline. Thus, we characterized the growth of cells lacking different elements of the CWI pathway in the presence of increasing amounts of OGT2468. As expected, strains deleted in the MAPK Slt2p (slt2Δ), the MAPKKK (bck1Δ) or the phosphotyrosine-specific protein phosphatase (ptp2Δ), acting on Slt2p, were hypersensitive to OGT2468 (Fig. 4C). In combination these data show that the CWI pathway is activated in response to OGT2468 suggesting that inhibition of O-mannosylation immediately affects cell wall integrity.
Inhibition of O-mannosylation activates the unfolded protein response
Compounds that interfere with protein folding in the ER such as tunicamycin, an inhibitor of protein N-glycosylation, or the reducing agent DTT which prevents disulphide bond formation, lead to the accumulation of unfolded proteins in the ER, induction of the unfolded protein response (UPR) and eventually, ER-associated protein degradation (ERAD) (Ron and Walter, 2007). The transcriptional programme of UPR includes genes involved in a wide range of cellular processes like protein folding, protein degradation, lipid biosynthesis and cell wall construction (Travers et al., 2000). Comparison of the transcriptional responses to OGT2468, tunicamycin and DTT revealed the presence of many co-induced genes (see Fig. 3, clusters 4 and 5) that are specifically induced by these treatments. Included in these clusters are many target genes of the UPR response (Travers et al., 2000). Genes involved in protein folding (FKB2, JEM1, LHS1, ERO1, EUG1 and MPD1), glycosylation (MCD4 and PMT3), lipid metabolism (SCS3) and ERAD (DER1 and HRD1) were induced as consequence of decreasing protein O-mannosylation.
These data indicate that inhibition of O-mannosylation specifically interferes with ER homeostasis while non-glycosylated, misfolded proteins are accumulating. Thus, UPR might be crucial when protein O-mannosylation is decreased. To further test this assumption, genetic interactions between HAC1, a key regulator of the UPR, and PMTs were analysed. Deletion of HAC1 in pmt2Δ, pmt1Δpmt2Δ as well as in pmt4Δ mutants resulted in synthetic lethality, whereas overexpression of HAC1 partially complements the temperature-sensitive phenotype of mutant pmt1Δpmt4Δ (Fig. S3). To verify that UPR and ERAD are enhanced in pmtΔ mutants, the ERAD substrate Pdr5p* (pdr5-26; Plemper et al., 1998) was analysed. As shown in Fig. 5, degradation of Pdr5p* is enhanced in pmtΔ mutants, confirming interdependence of O-mannosylation and UPR/ERAD.
Inhibition of O-mannosylation affects invasive growth
The transcriptional response to OGT2468 included the repression of genes that are also regulated by the filamentous growth MAPK pathway (Madhani and Fink, 1998), such as TEC1[transcription factor of the filamentation pathway (Gavrias et al., 1996)] and MUC1[cell surface flocculin, required for invasive growth (Lambrechts et al., 1996)]. Further, comparison of the OGT2486-repressed genes with the genome-wide transcriptional response of mutant tec1Δ (Madhani and Fink, 1998) identified PGU1 (endopolygalacturonase) and five additional targets of the filamentation pathway, including YHR177W, MMP1, IRC10, PRM6 and YJL218W (Table 3).
Table 3. Genes repressed in response to OGT2468 and in mutant tec1Δ.
To test whether this response was functionally relevant in vivo, we evaluated the ability of the haploid invasive growth competent WT strain MLY40 to invade the agar when growing in the presence of increasing concentrations of OGT2468. As shown in Fig. 6A, MLY40 cells were able to induce invasive growth and penetrate into the agar as deduced from colonies detected after mechanical washing off cells that appear at the surface. In contrast, dose-dependent inhibition of invasive growth was observed in the presence of OGT2468 (Fig. 6A). These data confirm the physiological relevance of the transcriptional downregulation of genes affecting invasive growth in response to OGT2468 in vivo.
Inhibition of O-mannosylation affects mating
Transcription of many genes related to the pheromone response pathway (Dohlman and Slessareva, 2006), including genes encoding for mating pheromones, membrane and cell wall proteins, the sensor Ste2p, the MAPK Fus3p and several genes whose transcription is regulated through this pathway was severely reduced in OGT2468-treated cells (Fig. 3, cluster 7; Table S3). Furthermore, many of these genes are also repressed in the pmtΔ mutants (Figs 2B and 3; Table S3). To evaluate the functional impact of this response in vivo we analysed the ability of yeasts to mate in the presence of OGT2468. Quantitative mating assays were carried out as described in Experimental procedures. As shown in Fig. 6B, in the presence of 0.25 µM OGT2468 mating efficiency was reduced to ∼25% and further decreased with increasing amounts of inhibitor, confirming the physiological impact of the transcriptional downregulation of genes involved in mating.
Ste12p is crucial for the transcriptional regulation of mating genes by OGT2468
Mating pheromone-regulated gene expression and filamentation are both dependent on the transcriptional regulator Ste12p (Madhani and Fink, 1998). To elucidate the impact of Ste12p on the observed decrease in transcription of genes controlled by the mating pathway upon PMT inhibition, qRT-PCR and chromatin immunoprecipitation (ChIP) were carried out.
For qRT-PCR analyses, a WT strain expressing genomically a haemagglutinin (HA)-epitope tagged Ste12p (Ste12pHA) and a ste12Δ mutant were grown in the presence or absence of OGT2468. Total RNA was isolated and transcriptional changes upon OGT2468 treatment of selected mating-relevant genes (FIG1, PRM3, FUS1, AGA1 and KAR4) were determined. Figure 7A shows that the repression of mating-relevant genes observed by microarray analyses was highly reproducible by qRT-PCR. The repression was highly dependent on Ste12p, since the transcript levels of FIG1, PRM3, FUS1, AGA1 and KAR4 did not significantly change in the ste12Δ mutant upon OGT2468 treatment (Fig. 7A). These results indicate a direct contribution of Ste12p to the transcriptional repression of mating relevant genes upon inhibition of protein O-mannosylation.
Ste12p commonly functions as a transcriptional activator. Thus, Ste12p-dependent repression of mating genes is most probably due to reduced binding of Ste12p to Pheromone Response Elements (PREs) (Yuan and Fields, 1991). To test this hypothesis, ChIP assays were conducted. Thereby, cells expressing Ste12pHA, or WT cells as a control, were treated with OGT2468 or DMSO. Subsequently, DNA–protein interactions were stabilized by formaldehyde cross-linking and Ste12pHA was immunoprecipitated. The precipitate was analysed by semi-quantitative and quantitative PCR for the presence and abundance of the PRE containing promoter region of a representative mating-relevant gene, AGA1 (Fig. 7B and C). As shown in Fig. 7C, the amount of AGA1 promoter DNA that co-purified with Ste12pHA was remarkably reduced upon OGT2468 treatment compared with untreated cells. These data indicate that binding of Ste12pHA to the AGA1 promoter was diminished under conditions of defective O-mannosylation. Quantitative PCR (qPCR) experiments confirmed the decrease in binding of Ste12pHA to box1 of the AGA1 promoter as a consequence of the OGT2468 treatment (Fig. 7D).
Together, these results show that the repression of genes controlled by the mating pathway upon PMT inhibition depends on the presence of Ste12p. In response to OGT2468, Ste12p is released from PREs in promoter regions of mating-relevant genes thus abolishing basal transcription.
Mutants of the STE vegetative pathway are hypersensitive to OGT2468
Ste12p is the central transcription factor not only of the mating and filamentation pathways, but also of the STE vegetative (SVG) pathway. This pathway involves elements of HOG, pheromone and invasive growth pathways (Ste20p/Ste50p–Ste11p–Ste7p–Kss1p–Ste12p) and has been proposed to promote cell integrity under vegetative growth conditions (Lee and Elion, 1999). Furthermore, the SVG pathway is activated in mutants affected in different steps of N-linked glycosylation as well as under treatments that prevents N-linked glycosylation, like tunicamycin treatment (Cullen et al., 2000). Based on these findings and our results, we were prompted to characterize the effect of OGT2468 on the growth of strains deficient in different elements of this pathway. A mutant lacking the transcription factor STE12 was highly susceptible to OGT2468 treatment (Fig. 8A). In addition, we observed conditional lethality between ste12Δ and mutant pmt2Δpmt4Δ (data not shown). Further, deletion of SHO1, STE7, STE11, STE20 and KSS1 rendered strains hypersensitive towards OGT2468 (Fig. 8B). However, growth of strains with deletions in elements of the mating pathway not related to the SVG pathway such as STE2 or STE4 (Cullen et al., 2000) is not affected by OGT2468 (Fig. 8B). In combination these findings support a role of the SVG pathway in response to defective protein O-mannosylation.
In fungi, O-mannosylation is an essential protein modification, thus PMTs are considered as targets for the development of antifungal drugs (Strahl-Bolsinger et al., 1999). The first inhibitors described were rhodanine-3-acetic acid derivatives that specifically block the Pmt1p isoform of C. albicans (e.g. OGT2599 and OGT2371) (Orchard et al., 2004; Cantero et al., 2007). In this study, specification of the previously uncharacterized rhodanine-3-acetic acid derivative OGT2468 in S. cerevisiae revealed that it affects not only ScPmt1p, but members of all PMT subfamilies, PMT1, PMT2 and PMT4 in vivo. Varying aspects could underlie these qualities such as increased membrane permeability, intracellular concentration/accumulation or effective dose, to take only a few examples. OGT2468 enabled us to analyse for the first time the genome-wide transcriptional response upon general inhibition of protein O-mannosylation in the model yeast S. cerevisiae.
In yeast, the only types of protein glycosylation are N-glycosylation and O-mannosylation, both initiated in the ER (Lehle et al., 2006). Transcriptome analyses and hierarchical clustering identified major stress regulatory pathways – the CWI pathway and the UPR – that are activated in response to OGT2468, and mostly also in the pmtΔ mutants analysed (Fig. 3 and Fig. S2). The vast majority of these genes is also induced in response to elimination of protein N-glycosylation by the inhibitor tunicamycin (Travers et al., 2000; Lecca et al., 2005) and in N-glycosylation mutants (Klebl et al., 2001; Lagorce et al., 2003; Cullen et al., 2006) (Fig. 3 and Fig. S2) suggesting an overlapping/compensatory impact of N- and O-glycosylation on cell wall integrity and ER homeostasis in yeast.
Protein O-mannosylation and cell wall integrity
The cell wall is an essential structure in yeast consisting of β-glucan (∼50%), mannoproteins (∼50%) and chitin (∼2%) (Klis et al., 2006). Cell wall mannoproteins are highly N-glycosylated, O-mannosylated or both. Especially the high degree of O-linked and high-mannose type N-linked carbohydrate chains and their modification by negatively charged phosphate groups determine cell wall permeability (Klis et al., 2006). In addition to this joint performance, glycosylation determines specific features of individual proteins that are crucial for cell wall biogenesis and/or structure (Strahl-Bolsinger et al., 1999). Thus, O- and N-glycosylation mutants show various cell wall-related phenotypes (Ballou et al., 1991; Nakayama et al., 1992; Strahl-Bolsinger et al., 1993; Gentzsch and Tanner, 1996).
Our study identified a compensatory response between O-mannosylation and N-glycosylation, particularly the formation of high mannose outer chains, suggesting that O- and N-linked glycans of cell wall mannoproteins can at least partially compensate for each other. Upon OGT2468 treatment and in pmtΔ mutants transcription of OCH1, KTR2 and MNT4 is induced (Fig. 3 and Table S2). These genes encode mannosyltransferases that specifically function in the biosynthesis of N-linked high mannose carbohydrate chains. OCH1 encodes an α1,6-mannosyltransferase that initiates poly-mannose outer chain elongation of N-linked oligosaccharides of glycoproteins in the Golgi apparatus (Nakayama et al., 1992). KTR2 and MNT4 encode Golgi α1,2- and α1,3-mannosyltransferases, respectively, that act in the further assembly of high mannose chains (Lussier et al., 1999; Romero et al., 1999). This is in agreement with Travers et al. (2000) who reported that inhibition of N-glycosylation by tunicamycin enhances transcription of PMTs and KTR1 (Golgi α1,2-mannosyltransferase involved in extension of O-linked glycan chains; Lussier et al., 1999).
Defects in glycosylation lead to the activation of the CWI pathway (see below). During vegetative growth and periods of environmental stress, the CWI pathway is induced to compensate for cell wall defects (Levin, 2005). Cell wall stress is sensed by the highly glycosylated plasma membrane sensors Wsc1p and Mid2p. These proteins activate (via Rom2p and Rho1p) the protein kinase C which in turn activates a MAPK cascade consisting of Bck1p (MAPKKK), Mkk1p and Mkk2p (MAPKKs) and the MAPK Slt2p/Mpk1p. Ultimately, stimulation of Slt2p leads inter alia to the transcriptional activation of a variety of genes that have been involved in cell wall assembly and structure such as chitin and glucan synthases (Levin, 2005). Genome-wide expression profiling of an N-glycosylation mutant bearing cell wall proteins with truncated N-linked high mannose glycan chains (mnn9Δ) and of cells treated with tunicamycin resulted in the induction of the CWI pathway transcriptional profile (Travers et al., 2000; Lagorce et al., 2003). Further, tunicamycin treatment leads to the activation of the MAPK of the CWI pathway, Slt2p (Chen et al., 2005; Cohen et al., 2008). We show here that inhibition of O-mannosylation by OGT2468 treatment also activates Slt2p (Fig. 4B). In consequence, the transcriptional CWI pathway response is induced to preserve cell wall integrity (Fig. 3, gene clusters 1, 3 and 6, and Table S2). In addition, the CWI pathway is constitutively activated in pmt1Δpmt2Δ and pmt1Δpmt4Δ mutants (Fig. 3, gene clusters 1, 3 and 6, and Table S2). Consistent with these data, a basal activation of the CWI pathway occurs also in pmt2Δpmt4Δ cells, as indicated by a constitutively increased level of Slt2p phosphorylation (Lommel et al., 2004). Hence, the cell wall chitin content is significantly increased in pmtΔ mutants (Gentzsch and Tanner, 1996).
Protein O-mannosylation and ER homeostasis
Nascent secretory and membrane proteins synthesized at the rough ER are translocated, potentially glycosylated and soon begin to fold. N-linked glycans play a crucial role in the folding and degradation of glycoproteins in the ER (Helenius and Aebi, 2004). Among other ER stress conditions, drugs that interfere with N-linked glycosylation (e.g., tunicamycin) or the formation of disulphide bonds (e.g., DTT) induce the accumulation of unfolded proteins in the ER thereby triggering a protective response termed the UPR (Kaufman, 1999; Ron and Walter, 2007). In yeast, unfolded proteins sequester the molecular chaperone Kar2p from the ER membrane-localized kinase/endoribonuclease Ire1p, resulting in its activation. As a consequence, Ire1p removes an intron harbouring translational attenuator of the mRNA for the transcription factor Hac1p. Spliced HAC1 mRNA (HAC1i) is translated and Hac1ip activates transcription of genes encoding ER-resident chaperones and folding helpers (Kaufman, 1999). Once protein folding fails, ERAD is induced to remove aberrant proteins from the ER (Ron and Walter, 2007).
Expression profiles of cells treated with tunicamycin or DTT identified induction of many target genes of UPR and ERAD (Travers et al., 2000). We show here that inhibition of O-mannosylation triggers the activation of a series of genes that are also highly induced upon ER stress conditions but hardly in response to cell wall stress or in cell wall mutants (Fig. 3, clusters 4 and 5), suggesting an important function of protein O-mannosylation for ER homeostasis in yeast. Several lines of evidence support that O-linked glycans bear complex functions in the ER: (i) O-mannosylation by Pmt4p is required for solubilization of heterologously expressed human β-amyloid precursor protein (Murakami-Sekimata et al., 2009), (ii) O-mannosylation plays a role for proteasome-dependent degradation of the ERAD substrate Gas1*p (Hirayama et al., 2008), (iii) O-mannosylation overall is enhanced upon tunicamycin treatment of yeast cells (Harty et al., 2001), (iv) Pmt2p acts on aberrant proteins (Harty et al., 2001; Nakatsukasa et al., 2004), (v) PMT3 is specifically induced upon tunicamycin, DTT as well as OGT2468 treatment (Fig. 3, cluster 4), (vi) HAC1 deletion is lethal in a pmtΔ mutant background (Fig. S3A), (vii) Hac1ip overexpression suppresses the temperature-sensitive phenotype of mutant pmt1Δpmt4Δ (Fig. S3B), and (viii) degradation of the ERAD substrate Pdr5p* is enhanced in pmtΔ mutants (Fig. 5). Due to its complexity, the precise function of O-mannosylation for ER homeostasis is not defined yet and remains a challenging question for the future.
Interrelation between ER and cell wall stress triggered by O-mannosylation
Recently, it has been shown that the slt2Δire1Δ double mutant is as sensitive to cell wall stress as the single mutant slt2Δ, thus, displaying epistasis (Scrimale et al., 2009). Based on these and other findings a linear pathway was suggested connecting cell wall stress signals and UPR (Krysan, 2009). However, ER stress activates UPR and CWI signalling pathways through parallel circuits, since the slt2Δire1Δ mutants display synergistic hypersensitivity to ER stress relative to the single mutants (Chen et al., 2005), and Hos2p/Set3p deacetylase complex was shown to mediate transmission of the ER stress signal to Slt2p (Cohen et al., 2008).
Our transcriptome analyses suggest that an immediate response to inhibition of O-mannosylation is initiation of the UPR, in a CWI pathway-independent manner. Expression of UPR target genes is induced in response to OGT2468 treatment, but most of these genes are not activated by constitutive active CWI pathway (GAL-PKC1-R398A) as well as cell in wall mutants (Fig. 3, clusters 4 and 5). UPR is essential for survival of yeast cells when O-mannosylation is blocked. This is demonstrated by the fact that hac1Δpmt2Δ, hac1Δpmt4Δ as well as hac1Δpmt1Δpmt2Δ mutants are not viable, even when osmotically stabilized and grown at lowered temperatures (Fig. S3). Since many cell wall proteins are highly O-mannosylated, decreased O-mannosylation results in misprocessed cell wall proteins and, consequently, the CWI pathway is activated to buffer the cell against defects in cell wall biosynthesis.
Protein O-mannosylation and mating/filamentation
Haploid yeast cells mate to form diploid cells in response to opposite mating type pheromones. In response to nitrogen starvation and other signals, yeast cells undergo a haploid invasive (haploid cells) or pseudohyphal (diploid cells) growth. MAPK pathways regulating these processes, namely the yeast pheromone response and the filamentation pathway, respectively, share various elements (Schwartz and Madhani, 2004).
Here we show that inhibition of O-mannosylation results in a common repression of mating (Fig. 3, cluster 7, and Table S3) and filamentation (Table 3) genes, at least partially in a STE12-dependent manner (Fig. 7A). Further, expression of key transcription factors of the filamentation pathway, TEC1 (2.8-fold repression) and to a minor extent also STE12 (pheromone and filamentation pathway; 1.7-fold repression) is affected. Consistent with these data are the mating and haploid invasive growth defects observed upon OGT2468 treatment (Fig. 6). Interestingly, it has been reported that induction of the UPR represses filamentous growth in yeast, further supporting our findings (Schroder et al., 2000). Very recently Yang et al. (2009) reported that the filamentation pathway-specific MAPK Kss1p is activated by tunicamycin treatment of mutant pmt4Δ. We did not find enhancement of KSS1 transcription in response to OGT2468 treatment (data not shown). However, strain backgrounds used in the study of Yang and co-workers are substantially different to the WT strain used here.
Mating and filamentation pathways activate the transcriptional regulator Ste12p that is also crucial for the SVG pathway. The SVG pathway involves elements of the osmolarity response, the pheromone and filamentation pathways (Sho1p–Ste20p–Ste11p–Ste7p–Kss1p–Ste12p) (Lee and Elion, 1999; Chen and Thorner, 2007). This pathway is activated in mutants affected in different steps of N-linked glycosylation as well as in response to tunicamycin (Cullen et al., 2000; Klebl et al., 2001).
The results presented here suggest that the SVG signalling cascade is induced upon inhibition of O-mannosylation. Although transcription of SHO1, STE20, STE11, STE7 and KSS1 is not enhanced upon OGT2468 treatment (data not shown), mutants lacking any of these components were found to be hypersensitive to PMT inhibition (Fig. 8). During normal growth conditions, the SVG pathway is inhibited by Fus3p, the MAPK of the pheromone response pathway (Lee and Elion, 1999). FUS3 mRNA levels are attenuated 2.3-fold upon PMT inhibition (Table S3), thus block of the SVG pathway could be released. Displacement of shared components of the mating and filamentation pathways towards the SVG pathway might further contribute to SVG pathway activation. This rearrangement might affect promoter selection of the transcription factor Ste12p as suggested by ChIP analysis (Fig. 7B–D). Indeed, a condition-dependent redistribution of Ste12p across the genome was reported. Chou et al. (2006) demonstrated that differential interactions of Ste12p confer specificity to the response to pheromone or nutrient limitation. Similarly, Zeitlinger et al. (2003) demonstrated that the programme-specific distribution of Ste12p depends on partner transcription factors and MAPK signalling.
So far, the exact role of Ste12p in compensating O-mannosylation defects is not clear. However, Ste12p is crucial in the absence of O-mannosylation. This is supported by the finding that ste12Δ mutant is highly susceptible to PMT inhibition (Fig. 8A). In silico analyses revealed that 35% of all genes regulated upon PMT inhibition contain putative Ste12p binding sites (data not shown), implying a key role of Ste12p and the SVG pathway in the adaptive response to inhibition of O-mannosylation.
Yeast strains and culture conditions
Saccharomyces cerevisiae strains used in this study are listed in Table S3.
JHY4 (STE12-3×HA) and LBY10 (ste12Δ): To create a PCR template for genomic C-terminal HA-tagging the KanMX resistance cassette was excised from pFA6a-GFP-kanMX6 (Wach et al., 1997) using EcoRI/BglII, blunted by T4 polymerase and ligated into pRS426-HA, which was cut with SacI and blunted. The tagging construct was amplified by PCR on the resulting plasmid pJH24 using primers 838 and 839. Further, to delete STE12 a G418 cassette flanked by sequences homologous to the up- and downstream regions of the STE12-coding sequence was amplified on genomic DNA isolated from strain MLY216a (Lorenz and Heitman, 1997). Purified PCR fragments were transformed into S. cerevisiae WT strain SEY6210. Integrants were selected on YPD plates containing 400 µg ml−1 G418. Expression of Ste12pHA was confirmed by Western blotting. Deletion of STE12 was confirmed by PCR using primers 339 and 340. Primer sequences are available upon request.
Yeast strains were grown in YPD or SC dropout medium at 30°C (Treco and Winston, 2008). Transformations were performed following the method of Gietz et al. (1992). For microarray, qPCR, ChIP and other PMT inhibitor experiments (unless otherwise stated), yeast cells were grown to an optical density (OD600) of 0.1. Cultures were split up and treated with OGT2468 at final concentrations of 0.1 µM or 1 µM, and DMSO (final concentration 1%) for 3 h. Cells were collected by centrifugation and further processed. Stock solutions (10 mM) of the rhodanine-3-acetic acid derivatives OGT2305, OGT2078 and OGT2468 [gift of Dr C. Stubberfield, Celltech, UK (Orchard et al., 2004)] were prepared in DMSO.
Total RNA was extracted from 3.8 × 108S. cerevisiae cells after mechanical disruption following the instructions of RNeasy Midi kit manufacturer (QIAGEN, Hilden, Germany). Concentration of RNA was estimated at 260 nm and sample quality was checked using RNA Nano Labchips in a Bioanalyzer 2100B (Agilent Technologies, Santa Clara, CA).
Microarray experiments and data analyses
PMT inhibitor microarrays. Biological triplicates of these experiments were carried out. Double-stranded cDNA was synthesized from 5 µg of total RNA using One-cycle cDNA Synthesis Kit (Affymetrix, Santa Clara, CA). After purification using the ‘GeneChip Sample Cleanup Module’ (Affymetrix), cDNA was used as template for in vitro transcription to obtain biotin-labelled cRNA. The obtained cRNA was fragmented and hybridized to the Affymetrix GeneChip® Yeast Genome 2.0 array for 16 h a 45°C. Hybridized microarrays were washed and stained with a streptavidin-phycoerythrin conjugate in a GeneChip® Fluidics Station 450. All these procedures were carried out as suggested by the manufacturer. Hybridized cRNA was finally identified by the fluorescence signal in a GeneChip® 3000 scanner. The files generated from the scanning (.CEL) were converted to gene expression signals using GCRMA software (Bioconductor) (Wu et al., 2004) (99:909; http://www.bioconductor.org/packages/2.0/bioc/html/gcrma.html). For each experimental condition, three microarray experiments corresponding to three mRNA replicates were processed and analysed. Fold changes between experimental conditions were calculated as a quotient between the mean of the gene expression signals. Genes with fold change ≥ 2 or ≤ 0.5 were included to further analysis. Statistical analysis was performed with the Cyber-t software (http://cybert.microarray.ics.uci.edu/) (Baldi and Long, 2001). Those values with a Bayesian P-value < 0.01 were considered as significant and the corresponding genes considered for further analysis.
pmtΔ microarrays. For the analysis of differential gene expression of pmt1Δpmt2Δ and pmt1Δpmt4Δ mutants, cell cultures and RNA isolations were performed in six independent biological replicates. RNA isolations of three biological replicates were combined and analysed in two independent experiments. RNA isolation, labelling, hybridization, washing and staining steps were performed as mentioned above but using GeneChip® Yeast Genome S98 arrays. After scanning, numerical data were obtained with GCOS software. Data from pmtΔ mutants and strain SEY6210 were compared in pairs obtaining four comparisons for each mutant. Genes with at least two present as ‘detection call’ and four increase or decrease values as ‘signal call’ were selected for further analysis. The signal log ratio was obtained as the mean of the signal log ratio obtained in each of the four comparisons; genes with signal ratio ≥ 2 or ≤ 0.5 were selected for further analysis.
Strain JHY4 (STE12-3×HA) was treated with OGT2468 as described above. In vivo DNA–protein cross-linking, harvesting, lysis of yeast cells, immunoprecipitation, DNA isolation, semi-quantitative PCR analyses and gel electrophoresis were carried out following the protocol of Hecht et al. (1999). Ste12pHA was precipitated using monoclonal HA antibodies (16B12; Covance) and Protein A SepharoseTM CL-4B (Amersham Biosciences).
Immunoprecipitated DNAs were also quantified by quantitative real-time PCR using an ABI 7900HT Fast Real-Time PCR instrument (Applied Biosystems, Foster City, CA). For quantification, the fold enrichment (FE) at specific DNA regions was calculated using the Comparative Ct Method according to the protocol described in Aparicio et al. (2004). Occupancy was normalized relative to an intergenic region of chromosome V used as a control. Primer sequences will be provided upon request. The Ct of the input sample was subtracted from the Ct of the immunoprecipitated sample to calculate the ΔCt both in the control (ΔCtcont) and in the target DNA (ΔCtexp) for each condition. FE was calculated by using the following formula: FE = 2−[ΔCtexp − ΔCtcont].
First-strand cDNAs were synthesized from 1 µg of total RNA in 20 µl final volume, using the High Capacity cDNA Archive kit (Applied Biosystems) following the recommendations of the manufacturer. As control for genomic contamination, the same reactions were performed in the absence of reverse transcriptase.
For the analysis of ChIP samples, qPCR samples were prepared by adding 4.5 µl of input or immunoprecipitated DNA samples (diluted at 1/100 and 1/10), oligonucleotides and Power SYBR Green PCR Master Mix 1× (Applied Biosystems).
Real-time qPCR assays were carried out in an ABI 7900HT Fast Real-Time PCR instrument (Applied Biosystems, Foster City, CA) using standard PCR conditions. Quantification of FIG1, PRM3, FUS1, AGA1, KAR4, GFA1, SRL3, CRH1 and ACT1 was performed using probe numbers 74, 156, 69, 63, 20, 48, 39, 69 and 25, from the Universal Probe Library (Roche Applied Sciences) and the ‘TaqMan Universal PCR Master Mix No AmpErase UNG’ kit (Applied Biosystems). For GSC2 SYBRGreen assays were performed using ‘POWER SYBR Green PCR MASTER MIX’ (Applied Biosystems). All assays were performed in duplicate. Probes and primers specific for each gene were designed at the ‘Universal ProbeLibrary Assay Design Center’ (https://www.roche-applied-science.com/sis/rtpcr/upl/adc.jsp) with the sequences of the corresponding ORFs as stored in the Saccharomyces Genome Database (http://www.yeastgenome.org/). Primers sequences will be provided upon request. For quantification, the abundance of each gene was determined relative to the standard transcript of ACT1 and to the non-treated sample with the use of the Comparative Ct Method.
Preparation of cell extracts and Western blot analysis
PMT substrate proteins. FUSw/oTMZZ was expressed from plasmid pMS7.1 (Hutzler et al., 2007) and FUSwTMZZ from plasmid pMS7.2 (Hutzler et al., 2007) in strain SEY6210 and isogenic pmtΔ mutants. FUSw/oTMZZ was precipitated from the culture medium of 2 × 107 cells. To analyse FUSwTMZZ crude membranes were isolated from 2 × 106 cells. Protein and membrane isolations, SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and Western blot analysis were performed as previously described (Hutzler et al., 2007).
Phosphorylation of Slt2p. Strain BY4741 was grown to mid-log phase and treated with 1 µM OGT2468 or mock treated for the times indicated. Western blot analysis, including cell collection, cell lysis, fractionation of proteins by SDS-PAGE, and Western blotting were carried out as previously described (Bermejo et al., 2008).
Cycloheximide chase analysis of Pdr5p*. Strain SEY6210 and pmtΔ mutants expressing a HA-tagged version of Pdr5p* from plasmid pRP13/5-26 (Plemper et al., 1998) were grown to mid-log phase. Cultures were adjusted to ∼8 OD600 per ml and cycloheximide was added to a final concentration of 0.1 mg ml−1. At indicated times aliquots of cells were removed, crude membranes were prepared and proteins analysed by SDS-PAGE and Western blotting as described in Plemper et al. (1998). Anti-HA antibodies (dilution 1:5000; 16B12; Babco) and anti-Pmt6p serum (dilution 1:2000; Girrbach and Strahl, 2003) were used. Protein–antibody complexes were visualized by enhanced chemiluminescence using the Pierce SuperSignal West Pico substrate according to the manufacturer's instructions.
Analysis of OGT2468 sensitivity
Assays were performed in 96-well microtitre plates, preparing twofold serial dilutions of OGT2468 ranging from 64 µM to 0.01 µM in a final volume of 150 µl of YPD. Each well was inoculated with 104 yeast cells from exponentially growing cultures. Microtitre plates were incubated without agitation at 30°C and cell growth determined after 12 h, 24 h or 48 h by measuring the absorbance at 595 nm in a microplate reader (Model 680 ELISA Microplate Reader; Bio-Rad). For further details see also Fig. S1.
Invasive growth assay
Yeast strains MLY40 and BY4741 were tested for invasive growth by the plate washing assay. Cells were grown YPD plates for 2 days at 30°C. Fresh cells from these plates were resuspended in YPD and adjusted to an OD600 of 0.13. Five microlitres of this suspension, corresponding to 104 cells, was spotted onto YPD plates with different concentrations of OGT2468, grown for 3 days at 30°C and photographed. The plates were then rinsed with water; the surface was rubbed away by a gloved hand and photographed. Cells not invading the agar were washed away whereas those penetrating the surface of the agar remained visible after mechanical washing.
Quantitative mating assay
Yeast strains SEY6210 and SEY6211 were grown to mid-log phase. A total of 6.5 × 107 cells of each strain were collected and mixed in 0.6 ml of YPD. Fifty-microlitre aliquots of cell suspension were transferred to 96-well plates. One microlitre of DMSO and 1 µl of 0.2 µg µl−1, 0.05 µg µl−1 and 0.013 µg µl−1 OGT2468 were added. Mating mixes were incubated for 5 h at 30°C and then transferred into 1 ml of sterile water. Serial dilutions were plated on YPD plates (growth of haploid and diploid cells) and SC plates without adenine and tryptophan (growth of diploid cells) (Treco and Winston, 2008). Colonies were counted after 3 days at 30°C.
We thank C. Stubberfield (Celltech) for generously providing PMT inhibitors and S. Keller for excellent technical assistance. We are very grateful to U. Schermer for performing experiments shown in Fig. S3A; and J. Heitman and W. Tanner for generously providing yeast strains. We thank M. Lommel, J.M. Rodriguez-Peña and M. Büttner for many helpful discussions. This work was supported by the EU Grant FUNGWALL (EU-Project LSHB-CT-2004-511952) and Integrated Action Spain (HA2007-0055)/Germany (DAAD) to J.A. and S.S. and Grants BIO2007-67821, BIO2010-22146 and 920640 (UCM/BSCH) to J.A. This work was partially supported by the Deutsche Forschungs Gemeinschaft (SFB638, Project A18). S.S. is a member of CellNetworks – Cluster of Excellence (EXC81).