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-glucan
1 Department of Applied and Bioapplied Chemistry, Graduate School of Engineering, Osaka City University, Sugimoto 3-3-138, Sumiyoshi-ku, Osaka 558-8585, Japan
2 Division of Molecular Science, Graduate School of Science and Technology, Kobe University, 1-1 Rokkodaicho, Nada-ku, 657-8501, Kobe, Japan
3 Department of Chemical Science and Engineering, Faculty of Engineering, Kobe University, 1-1 Rokkodaicho, Nada-ku, 657-8501, Kobe, Japan
Correspondence
Masayuki Azuma
azuma{at}bioa.eng.osaka-cu.ac.jp
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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haploid was partially restored by the addition of 0·6 M sorbitol to the growth medium. Rot1p is predicted to contain 256 amino acids, to have a molecular mass of 29 kDa, and to possess a transmembrane domain near its C-terminus. Candida albicans and Schizosaccharomyces pombe have Rot1p homologues with high identity that also have predicted transmembrane domains. To explore the role of Rot1p, the phenotypes of the rot1 haploid were analysed. Deletion of ROT1 caused cell aggregation and an abnormal morphology. Analysis of the cell cycle showed that rot1 cells are delayed at the G2/M phase. The rot1 cells were resistant to K1 killer toxin and hypersensitive to SDS and hygromycin B, suggesting that they had cell wall defects. Indeed, greatly reduced levels of alkali-soluble and -insoluble 1,6-
-glucan, and increased levels of chitin and 1,3--glucan, were found in rot1 cells. Furthermore, the phenotypes of rot1 cells resemble those of disruption mutants of the KRE5 and BIG1 genes, which show greatly reduced levels of cell wall 1,6--glucan. Incorporation of glycosylphosphatidylinositol (GPI)-dependent cell wall proteins in big1 and rot1 cells was examined using a GFP–Flo1 fusion protein. GFP fluorescence was detected both on the cell surface and in the culture medium, suggesting that, in these mutants, mannoproteins may become only weakly bound to the cell wall and some of these proteins are released into the medium. Electron microscopic analyses of rot1 and big1 cells showed that the electron-dense mannoprotein rim staining was more diffuse and paler than that in the wild-type, and that the outer boundary of the cell wall was irregular. A big1rot1 double mutant had a growth rate similar to the corresponding single mutants, suggesting that Rot1p and Big1p have related functions in 1,6--glucan synthesis.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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-glucan, 1,6--glucan, mannoproteins and chitin, and it is essential for maintaining cell shape and integrity. The wall structure can dynamically change to adapt to extracellular conditions and different states of the fungal life cycle (Orlean, 1997
). 1,3--Glucan accounts for about 40 % of the cell wall dry weight; it is synthesized at the cell surface and consists of linear chains with an average length of 1500 glucose units. 1,6--Glucan, which constitutes approximately 10 % of the cell wall dry weight, consists of polymers with an average length of 350 glucose units; it plays a critical role in the cell wall architecture by anchoring mannoproteins to other cell wall components (Kollar et al., 1997; Kapteyn et al., 1996). This linkage of 1,6--glucan to mannoproteins is thought to be mainly through mannose residues of the glycosylphosphatidylinositol (GPI) core structure (Shahinian & Bussey, 2000).
Many genes involved in 1,6--glucan biosynthesis have been identified based on resistance to K1 killer toxin, which kills yeast following binding to a 1,6--glucan-containing cell surface receptor (Al-Aidroos & Bussey, 1978; Boone et al., 1990; Brown et al., 1993), and on hypersensitivity to Calcofluor white (Ram et al., 1994; Lussier et al., 1997). Their role in 1,6--glucan biosynthesis is supported by the fact that null mutations of these genes lead to a reduction in the level of 1,6--glucan. These gene products and their functional homologues are located along the secretory pathway, including the endoplasmic reticulum (ER), the Golgi apparatus, the cytoplasm and the cell surface, suggesting that 1,6--glucan is synthesized along the secretory pathway and completed at the cell surface (Shahinian & Bussey, 2000). Among those gene products, Kre5p is particularly worth noting. The Kre5p protein is a soluble N-glycoprotein located in the ER (Meaden et al., 1990; Levinson et al., 2002), and mutation of the Kre5p gene greatly reduces formation of cell wall 1,6--glucan, leading to severe growth defects or lethality (Meaden et al., 1990; Levinson et al., 2002; Shahinian et al., 1998; Azuma et al., 2002). The biological activity of this protein is still unknown, but it is reported to have a limited but significant sequence similarity with UDP-glucose : glycoprotein glucosyltransferases (UGGT) from Drosophila melanogaster (Parker et al.; 1995) and Schizosaccharomyces pombe (Fernandez et al., 1996). Recently, we have found that deletion of BIG1 leads to an approximately 95 % reduction in cell wall 1,6--glucan (Page et al., 2003). In addition, Big1p is an N-glycosylated integral membrane protein with a type I topology, it is located in the ER, and some phenotypes of a big1 mutant resemble those of a kre5 mutant. Although a Big1p homologue is also found in Candida albicans (Azuma et al., 2002), its biochemical function has not yet been defined.
BIG1 was first described as a multicopy suppressor of the synthetic lethality of a rot1-1 rot2-1 double mutant. Mutations in ROT1 (reversal of TOR2) and ROT2 cause cell wall defects and suppress the loss of TOR2, an essential phosphatidylinositol-kinase-like protein kinase (Bickle et al., 1998). ROT2 was identified based on the similarity of its protein sequence with that of the 
subunit of mammalian glucosidase II (Trombetta et al., 1996). In addition, a rot2 mutant has a partial 1,6--glucan defect (Simons et al., 1998). However, like Big1p, the functional role of Rot1p remains unknown. ROT1 is essential for growth, with rot1 ascospores germinating and arresting growth within one cell cycle (Bickle et al., 1998). In the disruption consortium BY4741 strain series used in this work, ROT1 is also classified as an essential gene (Winzeler et al., 1999).
Here, to examine whether the presence of Rot1p is necessary for formation of normal levels of cell wall 1,6--glucan, we begin an analysis of rot1 mutants.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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and 2. Wild-type strains used were S. cerevisiae BY4741 (MATa), BY4742 (MAT) and BY4743 (MATa/) (Brachmann et al., 1998). Deletion strains were obtained from the Saccharomyces Genome Deletion Consortium (Winzeler et al., 1999). Haploid rot1 mutants were obtained by dissection of the heterozygous diploid strains from the Consortium on medium containing 0·6 M sorbitol. The mating type and the nutrient requirement of the haploid mutants obtained were analysed, and strains isogenic with the wild-type were taken and used for this study. Media for yeast growth were as described by Bussey et al. (1982). YPD is yeast complex medium; YNB is a synthetic medium, which was supplemented with appropriate nutrients. Sorbitol (0·6 or 1·0 M) was added to those media when osmotic support was needed; the resulting media were called YPDS or YNBS. Yeast mating, sporulation, and tetrad analysis were performed as described by Sherman et al. (1982). To determine mating type the tester strains MC75 and MC76 were used. Yeast transformation was carried out using the one-step transformation method (Chen et al., 1992).
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). Alkaline phosphatase and T4 DNA ligase were purchased from NIPPON GENE. Restriction endonucleases were purchased from NIPPON GENE and Takara Bio. The vector pMB2 (provided by M. N. Hall; Bickle et al., 1998), which is YCplac33 (Gietz et al., 1995) containing a 1·4 kb SacI–EcoRI fragment bearing ROT1, was cut with SacI and EcoRI. The resulting 1·4 kb ROT1 fragment was inserted into SacI- and EcoRI-cut pRS426 (Christianson et al., 1992), generating the construct pRS426-ROT1. DNA sequencing of ROT1 (pRS426-ROT1) was performed with an ABI377 DNA sequencer at Takara Bio. The plasmid pMUF-GFP, bearing the gene for the GFP–Flo1 fusion protein, was constructed by first amplifying the GFP gene from pEGFP (Clontech) using PCR with nGFPfw and nGFPrv primers (Table 2). The amplified fragment was inserted into the NcoI and XhoI sites of pIF318 (Sato et al., 2002), and the resulting plasmid was named pWI3-GFP-Flo318. Next, pWI3-GFP-Flo318 was cut with PvuII. The resulting fragment contained the gene encoding the prepro--factor leader region, GFP, and the C-terminal region of Flo1p (318 aa) under the control of the 5'-upstream region of the isocitrate lyase gene from Candida tropicalis (UPR-ICL), which is induced by glucose exhaustion (Sato et al., 2002). The fragment was inserted into the PvuII site of pRS406-2µm, which was constructed by introducing part of the 2 µm ori from pWI3 (Kanai et al., 1996) into the ArtII site of pRS406 (Sikorski & Hieter, 1989).
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). An amplification of the 910 bp fragment was found. To verify the deletion, the complementation of the growth defect by plasmid pRS426-ROT1 was examined.
FACS analysis.
Cells were pre-cultured at 30 °C in 4 ml YPDS for 1 day, then 0·2 ml of the culture was transferred to 4 ml YPDS and cells were cultured to exponential phase. Hydroxyurea (0·4 ml of 1·0 M solution) was added to the culture solution and incubated for 4 h. The cells were washed with YPDS three times to remove hydroxyurea, resuspended in 10 ml YPDS and cultured at 30 °C for 2–9 h. Cells were fixed in 70 % cold ethanol at –20 °C overnight, washed with 0·2 M Tris/HCl (pH 7·5) and treated with 1 mg RNase A ml–1 (Sigma) at 37 °C overnight. Cells were washed with 0·2 M Tris/HCl (pH 7·5), and resuspended in 100 µl Na-PI solution (0·05 mg propidium iodide ml–1, 1·0 mg sodium citrate ml–1, 0·58 mg sodium chloride ml–1); 10 µl of 2·0 mg propidium iodide ml–1 was then added to the solution. Samples were allowed to stand at room temperature for 30 min and diluted with 900 µl 0·2 M Tris/HCl buffer and 10 µl of 2·0 mg propidium iodide ml–1. After the cell suspensions had been briefly sonicated, samples were analysed using a Becton Dickinson FACSCalibur and Cell Quest. Flow cytometric analysis was performed using FACSCalibur (Becton Dickinson). Event rate was maintained at 300 cells s–1 and data for 20 000 events were collected.
Phenotypes of rot1 cells.
Assays for K1 killer toxin sensitivity were carried out as previously described (Bussey, 1991). Yeast strains were grown on YPD+0·6 M sorbitol at 30 °C for 18 h. Cells were suspended at approximately 1x107 cells ml–1 in 100 µl of a sterilized solution of 1·0 M sorbitol, and 5 µl of this suspension was added to 5 ml medium (1 % Difco yeast extract, 2 % peptone, 1 % agar, 0·001 % methylene blue, 0·6 M sorbitol and 1x Halvorson medium buffered at pH 4·7) kept at 45 °C. The medium was quickly poured into Petri dishes (60x15 mm). After the agar had gelled and attained room temperature, 5 µl K1 killer toxin (1000x stock diluted 1 : 10) was spotted onto the centre of the medium. The plate was incubated at 18 °C overnight, followed by 24–48 h at 30 °C, after which the death zone was measured and photographed.
Drug sensitivity was determined by spotting diluted yeast cultures onto agar media containing various drugs (Ram et al., 1994; Lussier et al., 1997). Cells were cultured in liquid YPD+1·0 M sorbitol overnight at 30 °C. The cell density was adjusted to 0·5 (optical density at 600 nm), and 2 µl drops of a set of 1 : 10 serial dilutions were spotted onto agar plates. We used YPD+1·0 M sorbitol agar medium containing the following drug concentrations: hygromycin B, 1–100 µg ml–1; or SDS, 0·0005–0·005 %. Cells were cultured at 30 °C for 72 h, and the growth was then observed.
Alkali-soluble 1,6--glucan assay.
Yeast strains were pre-grown on YPD+0·6 M sorbitol plates for 2 or 3 days, and then cells were transferred to YPD+0·6 M sorbitol liquid medium (25 ml) with a toothpick and cultured overnight. The cells were harvested by centrifugation for 10 min at 1860 g, washed with 1·0 M sorbitol (10 ml), and resuspended in 500 µl water. Glass beads were added to the cell suspension, and the mixtures were vortexed five times for 30 s, with intervals (at least 5 min) on ice, and lysates removed from the beads. The protein levels in lysates were determined using the Bradford assay. Lysates containing 8 µg total cell wall protein were brought up to 50 µl with water, and 50 µl NaOH (1·5 M) was added to each, followed by incubation for 1 h at 75 °C. After removal of the alkali-insoluble components by centrifugation (5 min, 11 000 g), a 1 : 2 serial dilution of the alkali-soluble fractions was spotted on Hybrid-C nitrocellulose membrane (Amersham). The immunoblotting was carried out in TBST (10 mM Tris/HCl, pH 8·0/150 mM NaCl/0·05 % Tween 20) containing 5·0 % non-fat dried milk powder using a 1 : 2000 dilution of the affinity-purified rabbit anti-1,6--glucan primary antibody (Kollar et al., 1997) and a 1 : 2000 dilution of horseradish peroxidase–goat anti-rabbit secondary antibody (Amersham). The glucan signal on the membrane was visualized following development with a chemiluminescence detection kit.
Alkali-insoluble -glucan assay.
Alkali-insoluble -glucan levels were determined as described by Kapteyn et al. (1999). Cells were cultured in 50 ml flasks containing 10 ml YPD+1·0 M sorbitol, harvested by centrifugation (10 min, 2600 g) and washed with 1·0 M sorbitol (25 ml). The cells were broken with glass beads on ice, and the cell wall fraction was collected by centrifugation (15 min, 2600 g). Alkali-insoluble -glucans were extracted three times in 3 % (w/v) NaOH at 75 °C for 1 h. The pellet was washed twice in 0·1 M Tris/HCl (pH 7·5), washed in 0·01 M Tris/HCl (pH 7·5), resuspended in 0·01 M Tris/HCl (pH 7·5) containing 1,3--glucanase (Zymolyase 100T, 1·0 mg ml–1) and incubated at 37 °C overnight. After dialysis, the 1,6--glucan was collected and quantified. The total alkali-insoluble glucan was measured as the hexose content before dialysis. The alkali-insoluble 1,3--glucan level was calculated by subtraction of the 1,6--glucan content from total glucan.
Fluorescence microscopy.
Chitin was visualized by Calcofluor white staining. Cells grown on solid medium (YPD+1·0 M sorbitol) for 48 h were suspended in 1·0 M sorbitol, stained with Calcofluor white M2R (Sigma) at 0·1 mg ml–1, washed with 1·0 M sorbitol, and observed with a fluorophotomicroscope (Olympus BX50-34FLAD/PM30). The images were recorded with a Coolsnap camera (Nippon Roper). Mannan was visualized by staining with concanavalin A–fluorescein isothiocyanate (ConA–FITC, Sigma). Cells grown on YPD+0·6 M sorbitol for 48 h were fixed with 3·7 % formaldehyde for 30 min and washed with PBS. ConA–FITC (0·1 mg) was added to the cell suspension (1·0 ml). The suspension was incubated at room temperature for 10 min, washed with PBS, and observed.
Observation of GFP–Flo1p.
To evaluate cell surface proteins in rot1 cells, the fluorescence of a fusion protein of GFP and the C-terminal part of Flo1p (a GPI-anchored protein located in cell wall) (Bony et al., 1997) was analysed. Cells with pMUF-GFP, bearing the gene encoding the fusion protein, were cultured in YNB (without uracil)+0·6 M sorbitol containing 50 mM HEPES (pH 7·2) at 30 °C. Cells were washed, resuspended in the same medium and observed with a fluorophotomicroscope. The cultures were also centrifuged, and the fluorescence intensities of the supernatant and the pellet (cells) resuspended in a volume of the same medium equal to the culture were measured (excitation, 488 nm; emission, 510 nm) with a Shimadzu RF-5000 fluorometer.
SEM and TEM images.
Cells were cultured on YPDS plates at 30 °C for 2 days. Specimens were fixed by the freeze-substitution method with minor modifications (Baba & Osumi, 1987). The cells were mounted on the copper meshes to form a thin layer and plunged into liquid propane cooled with liquid N2. Frozen cells were transferred to anhydrous acetone containing 2·5 % OsO4, cooled in a solid CO2/acetone bath, and kept in the bath for 48 h. The solution including cells was incubated at –20 °C for 2 h, at 4 °C for 2 h, and subsequently at room temperature for 2 h. After washing the cells with anhydrous acetone three times, cells for SEM were incubated in anhydrous acetone/isoamyl acetate (1 : 1, v/v) for 20 min, and then in 100 % isoamyl acetate for 20 min. The cells were dried with a critical point dryer (Hitachi), coated with a gold layer and observed with a Hitachi Natural SEM S-3500N scanning electron microscope. Cells for TEM were embedded using ERL4206-Quentol 653 (Nissin EM). Ultrathin sections were prepared with an ULTRACUT N microtome (Reichert-Nisei); they were stained with uranyl acetate and lead citrate and viewed with a Hitachi H-7000 electron microscope.
Genetic interaction.
Crossing of rot1 and big1 cells was carried out. The rot1 and big1 cells were mixed on a YPDS plate, incubated at 30 °C for 6 h, transferred to a YNBS plate (not containing Met and Lys) to select only heterozygous diploid cells, and cultured at 30 °C for 4 or 5 days. The heterozygous diploid cells that grew on the selective medium were transferred to a GNA plate, cultured for 1 day, transferred to a SPO plate to make spores, and incubated at room temperature for 7 or 9 days. These cells were treated with 0·5 mg Zymolyase 100T ml–1. The spores in individual asci were dissected onto YPDS plates and incubated at 30 °C for 6 days. Genotypes of resulting spores were determined by growth on media containing G418 and by PCR analysis using primers KanB, ScROT1-A and ScBIG1-A (Table 2).
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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), an additional thymidine at position 758 was found that was not reported in the database. We sequenced the 3'-end of ROT1, and found that our sequence matched that in the database (data not shown). Therefore, in the current studies, we used the sequences in the SGD database.
The PSORT and Sosui programs predicted the presence of a transmembrane domain between residues 238 and 256 in Rot1p. Although a signal peptide was not found using either of these two programs, another program, SignalP, predicted an N-terminal signal peptide. Rot1p also has a consensus sequence for GPI-modified proteins, suggesting that it remains attached to the plasma membrane (Caro et al., 1997, De Groot et al., 2003). On the other hand, Huh et al. (2003) predicted that Rot1p may be localized in the ER, based on analyses of a collection of yeast strains expressing Rot1p tagged with GFP at the C-terminus (Huh et al., 2003). However, further work is required to determine the subcellular location of Rot1p more precisely.
According to a BLASTP search and the C. albicans genome sequence at the Stanford University Genome Center (http://genome-www.stanford.edu/), Rot1p has homologues in Sz. pombe and C. albicans. Because these homologues have high identities with Rot1p, we refer to them here as SzRot1p (43 % identity) and CaRot1p (53 % identity), respectively. The presence of a transmembrane domain near the C-terminus and an N-terminal signal peptide was predicted in both SzRot1p and CaRot1p using PSORT and Sosui.
Isolation of a haploid rot1 mutant
The rot1-1 mutant has an altered content of chitin (188 % of wild-type), suggesting the participation of the ROT1 gene product in cell wall synthesis (Bickle et al., 1998). To understand its role, we performed a phenotypic analysis of ROT1 by isolating a haploid rot1 mutant. The heterozygous rot1/ROT1 mutant was sporulated, and spores were dissected on YPD containing 0·6 M sorbitol. After 6 days of culture, the rot1 mutant produced small colonies on this medium; it was unable to produce colonies on YPD without sorbitol. A representative tetrad from the rot1/ROT1 heterozygous diploid is shown in Fig. 1. The colony size was similar or slightly smaller than the big1 colonies (data not shown). To test if the haploid mutants obtained were indeed disruptants with an insertion of the G418 resistance gene, cells were picked and transferred to medium containing G418. The mutants grew, although poorly, on YPD containing sorbitol and G418. The poor growth of the mutants was complemented by transformation with the plasmid pRS426-ROT1 (data not shown). Confirmation of the disruption of ROT1 was also obtained by PCR analysis.
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cells cells. Cell aggregation was observed when rot1 cells were cultured in liquid YPD+0·6 M sorbitol medium. The rot1 cells were round and large compared to the wild-type. This may be due to the cells staying in isotropic growth for a longer period. The cells were sensitive to hypo-osmotic conditions, bursting when they were transferred from a sorbitol solution to distilled water (data not shown). These results suggested that the deletion mutant has a weak cell wall.
Bickle et al. (1998) described that rot1-null spores germinated and arrested growth within one cell cycle as medium- or large-budded cells. To examine cell cycle progression, the DNA content of rot1 cells was quantified by FACS. After arresting the cells in S phase with hydroxyurea, cell progression was restarted by removing the hydroxyurea. In wild-type cells there was a progression of (
G2/MG1) over a 4 h period (Fig. 2a). In both rot1 and big1 mutants (Fig. 2b and 2c, 0 h), the cell cycle was not completely arrested in S phase by hydroxyurea. This may be due to slow growth of the mutants. However, after removal of the hydroxyurea most of the cells arrested in G2/M for at least 4 h. These results suggested that rot1 cells are delayed at the G2/M phase.
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cells to SDS and hygromycin B, known indicators of cell surface defects. Compared to the wild-type, rot1 was hypersensitive to SDS and hygromycin B, and there was no growth in 0·003 % SDS or in 20 µg hygromycin B ml–1 (Fig. 3a). These phenotypes were similar to those of big1 and kre5, which were also resistant to K1 killer toxin. We therefore examined the sensitivity of rot1 cells to K1 killer toxin (Fig. 3b). We found that the rot1/ROT1 heterozygote was as sensitive as the parental strain, while the haploid deletion mutant was resistant to the toxin. Thus, the rot1 haploid mutant, as well as big1 and kre5, may be defective in cell wall 1,6--glucan synthesis.
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-glucan levels in the cell wall. The level of the alkali-soluble glucan was measured by immunodetection (Fig. 4a). We found that the rot1 mutant, like the big1 mutant (Azuma et al., 2002), has greatly reduced levels of alkali-soluble 1,6--glucan (
3 % of the wild-type). However, a weak signal was detected, indicating that residual 1,6--glucan remains. Analysis of alkali-insoluble 1,6--glucan levels indicated that the level of rot1 was 3 % or less of that in the wild-type (Fig. 4b). These results indicate that Rot1p is required for maintenance of normal 1,6--glucan levels. In contrast, based on Calcofluor white staining, rot1 has increased levels of alkali-insoluble 1,3--glucan (Fig. 4c) and chitin (Fig. 5a).
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-glucan defect on mannan in the outer layer of the cell wall because 1,6--glucan anchors mannoproteins to other cell wall components (Kollar et al., 1997; Kapteyn et al., 1996). The mannan layer in wild-type, big1, cwh41 and rot1 cells was examined by staining the cells with ConA–FITC and observing them by fluorescence microscopy. Cwh41p is a homologue of mammalian glucosidase I and has been reported to be an ER protein involved in 1,6--glucan synthesis. Disruption of the CWH41 gene leads to a 50 % reduction in the cell wall 1,6--glucan level (Jiang et al., 1996). We found no differences in the fluorescence intensities of wild-type, big1, cwh41, and rot1 cells (Fig. 5b).
Recently, a system to display proteins on the cell surface of S. cerevisiae was developed using the prepro--factor leader region and the C-terminal GPI-anchor attachment signal sequences of Flo1p, a native cell wall protein (Sato et al., 2002). To examine the effect of the 1,6--glucan defect on cell surface proteins, wild-type, big1, cwh41 and rot1 cells were transformed with pMUF-GFP (bearing the gene encoding the GFP–Flo1p fusion protein) and observed by fluorescence microscopy. We found that the fluorescence of GFP in big1, cwh41, and rot1 cells was similar to that in the wild-type (Fig. 5c).
Because of the role of 1,6--glucan in anchoring mannoproteins to the cell wall, we expected that disruption of Big1p or Rot1p would cause a release of the GFP–Flo1 fusion protein into the medium. We therefore examined the intensity of GFP fluorescence in the culture supernatant of the rot1, big1, cwh41, and wild-type cells (Fig. 6). After 72 h of culture, the fluorescence intensities of the supernatants from rot1 and big1 cells were approximately fivefold stronger than those in wild-type and the cwh41 cells. These results indicate that mannoproteins in rot1 and big1 cells fail to be correctly linked to the cell wall and are released to the medium.
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. However, considering the role of 1,6--glucan, we predicted that disruption of ROT1 would cause some changes in the outer mannoprotein layer. In support of this possibility, KRE5 mutants with a defect in 1,6--glucan synthesis have rough cell walls lacking the outer mannoprotein layer (Simons et al., 1998). To investigate the structure of the mannoprotein layer in rot1, we carried out electron microscopic analyses of rot1 and big1 cells. SEM images showed that the cell surface structures of rot1 and big1 are very rough and substantially different from those of wild-type cells (Fig. 7). TEM revealed that the outer layer of the S. cerevisiae cell wall is composed of mannoproteins, which have an electron-dense appearance in ultrathin sections on TEM images. The underlying layer, which is amorphous, is composed of 1,6--glucan and 1,3--glucan (Cid et al., 1995). In wild-type cells, the thickness of the wall around the cell was almost uniform, and the darkly stained outer layer of the mannoproteins was clearly observed. However, in many of the rot1 and big1 cells, the electron-dense mannoprotein rim staining was more diffuse and paler than that in the wild-type. Also, the outer boundary of the cell wall was irregular and the outer part of the cell wall was rough.
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cells resembled those of big1 cells, we tested whether a big1 rot1 double mutant would have a more severe phenotype than the corresponding single mutants. Growth of double mutants was compared to that of the single mutants (Fig. 8). Two tetrads of nonparental ditype (NPD) and 14 tetrads of tetrad ditype (TT) were analysed. In all colonies, the genotypes were verified by PCR analysis. Double mutant colonies were obtained in two NPD tetrads and nine TT tetrads. Although the colony size of the double mutant was a little variable compared to that of the single mutants, the average growth rate of the double mutant was similar to that of the single mutants. These results suggest that Rot1p and Big1p may have similar effects on the synthesis of 1,6--glucan.
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cells can be achieved with an osmotic support. In the current studies, we examined the function of Rot1p using a haploid rot1 mutant. This haploid mutant had greatly reduced levels of 1,6--glucan, suggesting that, like Big1p and Kre5p, Rot1p is required for 1,6--glucan synthesis. Clarification of the functions of these proteins will be necessary for a full understanding of the mechanism of 1,6--glucan synthesis. Previously, we reported that Big1p and Kre5p appear to have similar effects on 1,6--glucan synthesis (Azuma et al., 2002), and in the current studies, we show that rot1 mutants have properties similar to big1 mutants, with big1rot1 double mutants showing a similar growth rate to the single mutants. Therefore Rot1p, Big1p and Kre5p may have related effects on the synthesis of 1,6--glucan. Furthermore, it seems that Rot1p and Big1p not only have a similar function, but they may also reside in the same compartment or at the same location and might form a complex. In the near future, the subcellular location of Rot1p must be determined. Although rot1, big1 and kre5 mutants all have an osmoremedial phenotype, the restorations were partial, and growth, even on medium with osmotic support, was very slow compared to the wild-type (see Fig. 3a, A). Our analysis of the cell cycle also suggested that rot1 and big1 cells are delayed at the G2/M phase. Therefore, 1,6--glucan may play a significant role in maintaining not only the physical strength of the cell wall but also normal cell division.
Considering the role of 1,6--glucan in the cell wall, it was expected that big1 and rot1 mutants would have defects in anchoring cell surface mannoproteins. However, microscopic analyses using ConA–FITC and GFP–Flo1p showed no differences relative to the wild-type cells. In contrast, when a strain with defects in GPI anchor synthesis (mcd4 mutant) was observed as a negative control, there was little ConA–FITC and GFP–Flo1p fluorescence (unpublished results). In addition, the quantity of cell wall proteins obtained from rot1 cells was similar to or slightly more than that from the wild-type when the two cell types were cultivated to the same culture volume (data not shown). These results suggest that, even with this large defect in 1,6--glucan level, mannoproteins were transported to the cell surface and anchored to 1,3--glucan and chitin. In rot1 cells expressing pMUF-GFP, the fluorescence intensity in the supernatant was 77 % of that in the cells, suggesting that much of the GFP–Flo1p was released to the medium. Furthermore, electron microscopic observations indicate that the cell walls of rot1 and big1 mutants are very rough, that these strains do not have the outer layer with a high density of mannoproteins, and that the outer boundary of the cell wall is irregular. Therefore, in these mutants, the mannoproteins do not localize in the outer layer at a high density, and, instead, may be spread around the whole cell wall.
Rot1p appears to be a membrane protein required for normal levels of the cell wall 1,6--glucan, and has homologues with high identity in Sz. pombe and C. albicans. Although in both SzRot1p and CaRot1p the presence of a transmembrane domain is predicted, the functional role of those proteins has not yet been defined. Analysis of Rot1p function should contribute to an understanding of fungal 1,6--glucan biosynthesis. Rot1p is also potentially interesting as a target for antifungal drugs because the protein is essential for growth and has a homologue in the pathogenic yeast C. albicans.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Azuma, M., Levinson, J. N., Page, N. & Bussey, H. (2002). Saccharomyces cerevisiae Big1p, a putative endoplasmic reticulum membrane protein required for normal levels of cell wall -1,6-glucan. Yeast 19, 783–793.[CrossRef][Medline]
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Received 30 April 2004;
revised 24 July 2004;
accepted 29 July 2004.
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), cwh41
), big1
) and rot1
) cells carrying pMUF-GFP were centrifuged, and fresh medium (0·6 ml) was added to the supernatant (0·9 ml). Fluorescence intensities of the solutions (1·5 ml) were measured. Values were expressed as percentages relative to the intensity of the solution from the culture of rot1
