• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of eukcellPermissionsJournals.ASM.orgJournalEC ArticleJournal InfoAuthorsReviewers
Eukaryot Cell. Nov 2010; 9(11): 1650–1660.
PMCID: PMC2976300

Characterization of Glycoside Hydrolase Family 5 Proteins in Schizosaccharomyces pombe [down-pointing small open triangle]

Abstract

In yeast, enzymes with β-glucanase activity are thought to be necessary in morphogenetic events that require controlled hydrolysis of the cell wall. Comparison of the sequence of the Saccharomyces cerevisiae exo-β(1,3)-glucanase Exg1 with the Schizosaccharomyces pombe genome allowed the identification of three genes that were named exg1+ (locus SPBC1105.05), exg2+ (SPAC12B10.11), and exg3+ (SPBC2D10.05). The three proteins have different localizations: Exg1 is secreted to the periplasmic space, Exg2 is a membrane protein, and Exg3 is a cytoplasmic protein. Characterization of the biochemical activity of the proteins indicated that Exg1 and Exg3 are active only against β(1,6)-glucans while no activity was detected for Exg2. Interestingly, Exg1 cleaves the glucans with an endohydrolytic mode of action. exg1+ showed periodic expression during the cell cycle, with a maximum coinciding with the septation process, and its expression was dependent on the transcription factor Sep1. The Exg1 protein localizes to the septum region in a pattern that was different from that of the endo-β(1,3)-glucanase Eng1. Overexpression of Exg2 resulted in an increase in cell wall material at the poles and in the septum, but the putative catalytic activity of the protein was not required for this effect.

Schizosaccharomyces pombe cells, like other yeasts, are surrounded by a rigid cell wall that provides mechanical strength and protection from environmental stresses. The cell wall determines cellular morphology during the different stages of the life cycle, and it is continuously remodeled during the cell cycle to allow cellular growth. Extracellular cues that trigger shape changes, such as nutrient deprivation or exposure to mating factors, also result in cell wall remodeling. During vegetative growth, S. pombe cells are rod shaped and grow by tip elongation, first at the old end and then at both ends. Subsequently, cell wall deposition occurs during septation, and the two new ends are thus sealed off (39).

The S. pombe cell wall is composed of mannoproteins, α-glucan and β-glucan (7, 32). Recent studies have shown that S. pombe cell walls also contain up to 15% of a branched β(1,3)-β(1,6)-glucan, which has been termed diglucan (31). β(1,3)-Glucan is a major structural component of the fungal cell wall, and it forms a fibrillary network that is thought to be responsible for the mechanical strength of the cell wall. These components are assembled in different layers that can be visualized by electron microscopy (26, 45). There is an outer layer enriched in glycoproteins and an inner layer of carbohydrates. In situ localization studies have indicated that β(1,6)-branched β(1,3)-glucan is localized all over the cell wall and throughout the septum; β(1,6)-glucan appears in the same layer and in the secondary septum, whereas linear β(1,3)-glucan is present only at the primary septum (23).

Synthesis of the cell wall is a complex process that requires the participation of different enzymes, some of which have been identified. The biosynthesis of β(1,3)-glucan is carried out by the β(1,3)-glucan synthase complex, whose catalytic subunit in S. pombe is encoded by the bgs genes. S. pombe contains four proteins of this family; Cps1/Bgs1, Bgs3, and Bgs4 are essential for cell viability during vegetative growth (14, 15, 29, 33), while Bgs2 performs an essential role during spore wall formation (27, 34). In yeast and filamentous fungi, the β(1,3)-glucan chains synthesized by the glucan synthase complex are extruded to the periplasmic space in a vectorial process (6). It has been postulated that the nascent β(1,3)-glucan chains should be cross-linked to other components of the cell wall by the action of glycoside hydrolases (GH) and transglycosidases. However, in vivo evidence of such a mechanism has been shown only for the Saccharomyces cerevisiae Crh1 and Crh2 proteins, which are involved in the cross-linking of β(1,3)-glucan to chitin (8, 9). Other proteins involved in cell wall assembly are the β(1,3)-glucanosyl-transferases of glycoside hydrolase family 72 (GH72), which in vitro are able to elongate β(1,3)-glucan oligosaccharides (42, 47). Four genes encoding proteins of this family (gas1+, gas2+, gas4+, and gas5+) are present in the S. pombe genome, performing specific functions at different moments of the life cycle (21).

Other proteins that may be involved in the modification of the cell wall β(1,3)-glucans are the exo-β(1,3)-glucanases from the GH5 family. Three proteins of this family have been characterized in S. cerevisiae. Exg1 of S. cerevisiae (ScExg1) is a polypeptide whose differential glycosylation accounts for the two main extracellular exo-β(1,3)-glucanases detected in culture supernatants, while Exg2 is a highly glycosylated minor exo-β(1,3)-glucanase with a C-terminal glycosylphosphatidylinositol (GPI) anchor site (12, 13, 43, 57). The third protein, Ssg1/Spr1, is a sporulation-specific glucanase (52). These enzymes are exo-β(1,3)-glucanases, but they also usually act on β(1,6)-linkages, although with less efficiency. It has been proposed that within the cell wall, these proteins would catalyze a transglycosylation rather than a hydrolytic reaction since transferase activity has also been demonstrated for the S. cerevisiae and Candida albicans Exg1 proteins (54, 55). Here, we report the characterization of the three GH5 family proteins present in S. pombe, which were named Exg1, Exg2, and Exg3. They were present in different compartments: the cell wall, membrane, and cytoplasm, respectively. Two of the proteins were active against β(1,6)-glucans but not against β(1,3)-glucans. Unexpectedly, enzymatic assays with purified Exg1 indicated that the protein has an endohydrolytic mode of action.

MATERIALS AND METHODS

Strains, growth conditions, and genetic manipulations.

The S. pombe strains used in this study are listed in Table 1. Yeast cells were grown on YES (yeast extract with supplements) medium or Edinburgh minimal medium (EMM) with appropriate supplements (41). For overexpression experiments using the nmt1+ promoter, cells were grown in EMM containing 20 μg/ml thiamine up to logarithmic phase. Then, the cells were harvested, washed five times with EMM, and inoculated in fresh medium (with or without thiamine) at an optical density at 595 nm (OD595) of 0.05 to 0.1. Synchronization of strains carrying the thermosensitive cdc25-22 mutation was achieved by growing the cells at the permissive temperature (25°C) to early log phase (OD595 of 0.5) and then shifting the cultures to 37°C for 4 h. Cells were released from arrest by transfer to 25°C, and samples were taken every 20 min.

Table 1.
Yeast strains used in this study

Plasmids and DNA manipulations.

The oligonucleotides used for different DNA manipulations are shown in Table 2. Construction of plasmid pED138 carrying the exg1+ coding sequence under the control of the nmt1+ promoter was achieved by PCR amplification of the coding sequence using oligonucleotides 424 and 425B (which introduced XhoI and BamHI sites) and cloning of the resulting fragment between the XhoI and BamHI sites of plasmid pJCR-3XL (40). A similar approach was used to construct plasmids pML2 (pJCR-3XL carrying exg2+) and pED139 (pJCR-3XL carrying exg3+) by using oligonucleotide pairs 426-427B and 428-429B, respectively.

Table 2.
Oligonucleotides used in this study

To construct the plasmids carrying the different mutated versions of exg2+, first the exg2+ coding sequence and flanking regions were amplified with oligonucleotides 305 and 306. Chromosomal HindIII and SacI sites were used to clone the gene with its promoter and terminator (from −473 to +278) at the same sites of vector pUC19, generating plasmid pEX22. Then, an NdeI site was introduced before the stop codon by recombinant PCR using oligonucleotides 643, 644, 645, and 646, yielding plasmid pED162. The NdeI site was used to clone the c-myc epitope obtained from plasmid pGEMMH (16), producing plasmid pED168′. Finally, a SalI-SacI fragment containing the exg2-myc region was cloned in plasmid pML2, generating plasmid pED172. The different mutations were generated by amplifying specific DNA fragments carrying the desired mutations by recombinant PCR and cloning the fragments in plasmid pED168′ or pED172. Thus, the following constructs were generated with the indicated oligonucleotides: exg2-A1-myc (where exg2 harbors the mutation E338A, designated A1) with oligonucleotides 646, 653, 654, and 655; exg2-A2-myc (where exg2 harbors the mutation E349Q, designated A2) with oligonucleotides 646, 655, 640, and 641; exg2-ΔN-myc (where is exg2-ΔN is a mutant lacking the cytoplasmic tail) with oligonucleotides 647, 648, 649, and 650; exg2-ΔTM-myc (where exg2-ΔTM is an exg2 mutant lacking the transmembrane [TM] domain) with oligonucleotides 647, 650, 651, and 652; exg2-Δout-myc (where exg2-Δout lacks the spacer domain that separates the transmembrane and catalytic domains) with oligonucleotides 647, 657, 658, and 650; and exg2-ΔGH5-myc (where exg2-ΔGH5 lacks the catalytic domain) with oligonucleotides 1010 and 1011. The amplified fragments carrying the exg2-A1, exg2-A2 and exg2-ΔGH5 mutations were first cloned in plasmid pED168′, and then the complete exg2 coding sequence fused to c-myc was cloned in plasmid pML2 using SalI-SacI sites, generating plasmids pED188 (exg2-A1-myc), pED193 (exg2-A2-myc), pED195 (exg2-A1A2-myc, where exg2 harbors both the A1 and A2 mutations), and pED215 (exg2-ΔGH5-myc). For exg2-Δout, exg2-ΔN and exg2-ΔTM, the amplified fragments carrying the mutations were cloned in plasmid pED172 using different restriction sites, yielding plasmids pED177 (exg2-Δout-myc), pED178 (exg2-ΔN-myc), and pED179 (exg2-ΔTM-myc). In all cases, the presence of the desired mutations was confirmed by sequencing the amplified fragments.

To construct plasmid pED173, carrying exg2-green fluorescent protein (GFP) under the control of the nmt1+ promoter (41X), the NdeI site of plasmid pED162 was used to clone the GFP coding sequence obtained from plasmid pGEM-EGFP (16), producing plasmid pED166. The exg2+ coding region obtained from plasmid pML2 as a XhoI-BamHI fragment was cloned under the control of the nmt1+ promoter present in plasmid pJCR-41XU (40) to create plasmid pED169. Then, a SalI-SacI fragment from plasmid pED166 containing the C-terminal region of exg2+ and the GFP was used to replace the same region of plasmid pED169, generating plasmid pED173.

To construct null mutants lacking the exg1+ (locus SPBC1105.05), exg2+ (SPAC12B10.11), or exg3+ (SPBC2D10.05) genes, the entire coding sequences were replaced by the ura4+ or kanMX4 cassette. The deletion cassettes were constructed using recombinant PCR. For this purpose, DNA fragments of 300 to 500 bp corresponding to the 5′ and 3′ flanking regions of each gene were PCR amplified using specific oligonucleotide pairs, and the resulting fragments were then fused, by recombinant PCR, to the kanMX4 cassette, which confers resistance to the G418 antibiotic (4) or to the ura4+ gene. The C-terminally tagged strains carrying exg1-HA (where HA is hemagglutinin), exg2-HA, exg3-HA, exg1-GFP, exg2-GFP, or exg3-GFP were constructed by direct chromosome integration of PCR fragments generated using plasmid pFA6a-3HA-kanMX6 or pFA6a-GFP-kanMX6 as a template and specific oligonucleotides (4). The amplified fragments contained the HA or GFP coding regions fused in frame to the last codon of the gene and the kanMX6 cassette to select for transformants. Correct integration of the DNA fragment was verified by PCR or Southern blotting.

RNA isolation and Northern blot analyses.

Cells (1.3 × 109) were collected at different time intervals after release from the restrictive temperature (37°C) or from different mutant strains, and total RNA was prepared using the method described by Percival-Smith and Segall (46). For Northern blot analyses, 12.5 μg of RNA was used. The DNA probes used to detect the different transcripts were DNA fragments (400 to 500 nucleotides [nt]) obtained by PCR amplification with specific oligonucleotides. For act1+, a 1.1-kb fragment containing the whole coding region obtained by PCR was used.

Microscopy techniques.

For light microscopy, cells were fixed in 3.7% formaldehyde and stained with aniline blue or calcofluor white. Samples were viewed using a Leica DMRXA microscope equipped for Nomarski optics and epifluorescence and photographed with a Photometrics Sensys charge-coupled-device (CCD) camera. For transmission electron microscopy (TEM), cells were stained with potassium permanganate and examined on a Zeiss EM 902 transmission electron microscope.

Immunoblotting and protein methods.

Total cell extracts of S. pombe were prepared by breaking the cells with glass beads in radioimmunoprecipitation assay (RIPA) buffer (10 mM sodium phosphate, pH 7, 1% Triton X-100, 0.1% SDS, 2 mM EDTA, 150 mM NaCl). Cell extracts were centrifuged at 10,000 × g for 10 min to separate a pellet from the supernatant. Culture supernatants were concentrated 10 times using Amicon units. The conditions used to examine whether Exg2 is a membrane protein have been described previously (28). The lysates were spun at maximal speed in a microcentrifuge for 30 min at 4°C. Supernatants were recovered, and the pellets were resuspended in SDS-PAGE loading buffer. For immunoblotting, 50 μg of protein extract was resolved by SDS-PAGE on 8% gels. Protein transfer, blotting, and enhanced chemiluminescence (ECL) detection were performed using standard procedures. Mouse monoclonal anti-myc antibodies (clone 9E10; Roche) were used.

Expression in Pichia pastoris.

To purify Exg1, the catalytic domain lacking the signal sequence (amino acids 28 to 407) was cloned into the EcoRI-XbaI sites of plasmid pPICZαA (Invitrogen). The resulting plasmid (pED204), containing the exg1 gene under the control of the AOX1 promoter, and a C-terminal fusion to the c-myc epitope and His6 was introduced into strain KM71H (arg4 aox1::ARG4). Production of the fusion protein was accomplished according to the protocol described by the supplier (Invitrogen). Exg1-His6 was purified using His-Trap columns (Amersham Pharmacia).

Assay for β-glucanase activity.

β-Glucanase activity was assayed as previously described (5). The substrates used to test activity were laminarin [β(1,3)-glucan], pustulan [β(1,6)-glucan], scleroglucan [β(1,3)-glucan with one β(1,6)-glucose side chain every three residues], or lichenan [mixed linkage β(1,3)-β(1,4) glucan]. Determination of the reducing sugars released in the reactions was performed by the methods of Somogyi (53) and Nelson (44). One unit of activity was defined as the amount of enzyme that catalyzed the release of reducing sugar groups equivalent to 1 μmol of glucose per hour, and specific activity was expressed as units per milligram of protein. For activity against p-nitrophenyl-β-d-galactopyranoside (pNPG), the amount of p-nitrophenol released was determined by measuring optical density at 420 nm. One unit of enzyme catalyzed the release of 1 μmol of p-nitrophenol per h under the reaction conditions used. The degree of polymerization (DP) of oligosaccharides released from laminarin or pustulan by Exg1 was determined by high-performance anion-exchange (HPAE) chromatography using a pulsed electrochemical detector (PED) and an anion-exchange column (Carbo-PAC PA1; 4.6 mm by 250 mm; Dionex) under the following conditions: flow rate of 1 ml/min; buffer A, 50 mM NaOH; buffer B, 500 mM sodium acetate in 50 mM NaOH; gradient, 0 to 2 min with 98% buffer A and 2% buffer B (isocratic); 2 to 15 min with 75% buffer A and 25% buffer B (linear); 15 to 45 min with 60% buffer A and 40% buffer B (linear).

Labeling and fractionation of cell wall polysaccharides.

Labeling and fractionation of cell wall polysaccharides were performed as described previously (3). Cultures of S. pombe cells that had been incubated in the presence or absence of thiamine for 18 h were diluted in medium supplemented with d-[U-14C]glucose (3 μCi/ml) and incubated for 4 additional hours. Total glucose incorporation was monitored by measuring radioactivity in trichloroacetic acid-insoluble material. Cells were harvested and broken with glass beads. Cell walls were purified by repeated washing and differential centrifugation (once with 1 mM EDTA, twice with 2 M NaCl, and twice with 1 mM EDTA) at 1,000 × g for 5 min. Finally, they were heated at 100°C for 5 min. Aliquots of the cell walls were incubated with 100 μg of Zymolyase 100T (Seikagaku Kogyo Co.) in 50 mM citrate-phosphate buffer (pH 5.6) or with 100 U of Quantazyme (Qbio) in 50 mM Tris, pH 7.5, for 36 h at 30°C. After incubation, samples were centrifuged and washed with the same buffer. One milliliter of 10% trichloroacetic acid was added to the pellets, and their radioactivity levels were measured. The supernatants from the Zymolyase 100T reaction were considered β-glucan plus galactomannan, and the pellet was considered α-glucan. The supernatants from the Quantazyme reactions were considered β(1,3)-glucan, and the pellet was considered α-glucan plus galactomannan. All determinations were carried out in duplicate.

RESULTS

S. pombe contains three proteins belonging to glycoside hydrolase family 5.

Family 5 glycoside hydrolases (GH5) comprise a group of enzymes that cleave different glucan polymers [β(1,3)- and β(1,6)-glucans, cellulose, lichenan, or xylan] that are widespread in fungi and bacteria. S. cerevisiae contains four different polypeptides belonging to this family (Exg1, Exg2, Ssg1, and YBR056w). Comparison of the ScExg1 sequence with the S. pombe genome database revealed the presence of three proteins belonging to this family, exg1+ (locus SPBC1105.05), exg2+ (SPAC12B10.11), and exg3+ (SPBC2D10.05). The S. pombe Exg1, Exg2, and Exg3 proteins show 40.5%, 25%, and 21% overall identity to ScExg1, respectively.

exg1+ expression peaks during the septation process.

Northern blot analysis in asynchronous cultures indicated that the three open reading frames (ORFs) were expressed during the vegetative cycle (data not shown). When RNAs obtained from a synchronized cdc25-22 mutant strain were analyzed, a periodic cell cycle variation was found for exg1+ (Fig. 1A), with the maximum accumulation of mRNA occurring during the septation process, suggesting that the product of this gene might exert its function during the last stages of the cell cycle, namely, septum assembly or cell separation. No significant variations that correlated with cell cycle progression were observed for the other two genes, exg2+ and exg3+. These results are in good agreement with the results obtained in a large-scale analysis (50). To study whether the periodic expression of exg1+ was dependent on the transcription factors Ace2 or Sep1, Northern analyses were performed to compare the expression in wild-type, ace2Δ, and sep1Δ mutants (2, 50, 58). The results revealed that the expression of exg1+ was clearly reduced in the sep1Δ mutant (Fig. 1B), suggesting that this transcription factor is required for its expression. As expected for genes that do not show fluctuations during the cell cycle, no significant differences were seen in the expression of exg2+ and exg3+.

Fig. 1.
Transcription pattern of exg genes. (A) Expression during the cell cycle. Synchrony was induced by arrest-release of a cdc25-22 mutant, and samples were taken at the indicated time points after the release for RNA extraction. RNA was hybridized with specific ...

The three β-glucanases have different cellular fates.

Analysis of the sequence of the three Exg proteins revealed the presence of different features in addition to the GH5 domain. Thus, Exg1 contained an N-terminal signal peptide for secretion, with the likely cleavage site between positions 22 and 23 (AFS-YV). In contrast, Exg2 contained a predicted transmembrane region (amino acids 39 to 60), while Exg3 did not contain any sequence for extracellular localization (Fig. 2A). These observations suggested that the three proteins might have different cellular localizations: the cell wall, the plasma membrane, and the cytoplasm, respectively. To test these predictions, they were tagged with the HA epitope at their C termini, and the distribution of the proteins in culture supernatants, the membrane/cell wall fraction, and the cytoplasm was analyzed. The results confirmed that only Exg1 was present in culture supernatants, indicating that the protein was secreted to the exterior of the cell and then released to the surrounding medium (Fig. 2B). In contrast, Exg3 was found mainly in the cytoplasm (supernatant) of the cell extracts, with a minor fraction in the pellet. Although Exg2 was difficult to detect, it was present mainly in the pellet fraction. To analyze the localization of Exg2 with more detail, the gene was placed under the control of the nmt1 promoter and tagged with the c-myc epitope. Overexpression of the gene confirmed that Exg2 was present in the membrane/cell wall fraction (Fig. 2B, Exg2 hc). Thus, these results confirm the predictions of the sequence analysis and indicate that Exg1 is a secreted protein, Exg2 is a possible membrane-associated protein, and Exg3 is a cytoplasmic protein.

Fig. 2.
Cellular fractionation of the Exg proteins. (A) Schematic representation of the characteristics of the Exg1, Exg2, and Exg3 proteins. The GH5 domain, common to the three proteins, is indicated. The black box represents a hydrophobic region with the characteristics ...

To ascertain whether Exg2 was indeed an integral membrane protein, protein extracts were prepared under different conditions. Cells were extracted with buffer alone, buffer containing 0.6 M NaCl or 1.6 M urea (to solubilize peripheral membrane proteins), 0.1 M Na2CO3 (to solubilize intracellular vesicles), 4% Triton X-100 (a nonionic detergent that solubilizes most membrane proteins), or 2% SDS (an ionic detergent that solubilizes all membrane proteins), and extracts were separated into the pellet and supernatant fractions. With the exception of SDS treatment, Exg2 was found to be insoluble under all conditions tested and was detected in the pellet fraction (Fig. 2C). Thus, Exg2 is a novel integral membrane protein that is solubilized only by strong ionic detergents but not by nonionic detergents, as has been previously described for the Cps1/Bgs1 subunit of the β-glucan synthase (28).

Cellular localization of Exg proteins.

To determine the in vivo localization of the three Exg proteins, the GFP was inserted in frame before the stop codon of each gene. The localization of the three proteins was monitored in live cells, but the fluorescence from only Exg1-GFP and Exg3-GFP was detectable. In good agreement with the peak of transcription during cytokinesis observed for exg1+, the protein was found mainly at the center of the cells by the time the septum was starting to be visible by differential interference contrast (DIC). Exg1 first started to accumulate as a ring surrounding the septa during the initial steps of their assembly (Fig. 3A, asterisks). At later stages, when the septum was clearly visible by DIC, Exg1 was found spanning the whole septum, and it had disappeared by the time the cells were separating (Fig. 3A, arrowhead). Staining of the cells with calcofluor white or aniline blue to visualize the dynamics of septum assembly more clearly resulted in a loss of the GFP fluorescence.

Fig. 3.
Localization of Exg proteins. (A) Wild-type cells expressing exg1-GFP were grown to early log phase. Photographs of differential interference contrast microscopy (DIC) or Exg1-GFP are shown. (B) Exponentially growing wild-type cells expressing Exg1-GFP ...

We have previously shown that the S. pombe endo-β(1,3)-glucanase Eng1 has a localization similar to that of Exg1 (35). To determine whether the two hydrolases colocalized, we studied the localization of both proteins in the same cell using a strain that simultaneously expressed Exg1-GFP and Eng1-red fluorescent protein (RFP). Exg1-GFP was seen in the septum before the Eng1-RFP fluorescence could be detected (Fig. 3B, asterisks), consistent with the fact that exg1+ expression is dependent on Sep1 while that of eng1+ requires Ace2. Only when the septum was clearly apparent by DIC did Eng1-RFP start to accumulate in the region, indicating that Eng1 secretion occurs after septum assembly has been completed. Even though Eng1 and Exg1 were present in the septum region, the distributions were different, and there was no clear colocalization of either protein. The fluorescence of Exg1-GFP was generally wider and less uniform than that of Eng1-RFP, suggesting that Exg1 localization might not be restricted to the primary septum but also spread to the secondary septum. This is in contrast with the perfect colocalization found for the two endoglucanases involved in cell separation, Eng1 and Agn1 (38).

In the case of Exg3, we found that this was a cytoplasmic protein, consistent with the fractionation experiments, since the fluorescence was detected as a diffuse cytoplasmic staining excluded from the nucleus although no specific pattern or accumulations could be seen (Fig. 3C). Exg2 could not be visualized when the Exg2-GFP protein was expressed from its own promoter, suggesting that the gene is transcribed at low levels. When Exg2-GFP was mildly overexpressed using the weak P41X-nmt1+ promoter, the localization of the protein could be determined. Exg2 was localized at the sites of cell growth, that is, the poles of the cell and the septum (Fig. 3D, arrowheads). Thus, these results are in good agreement with the fractionation experiments and indicate that the three GH5 proteins have different cellular localizations.

Exg1 and Exg3 are β(1,6)-glucanases.

The observed amino acid sequence similarity between S. pombe Exg proteins was an indication that they might also be β(1,3)-glucanases. However, since S. pombe lacks any detectable exo-β(1,3)-glucanase activity (48), two possibilities can be envisioned. First, it is possible that the proteins might be related only in sequence but lack any enzymatic activity. Alternatively, they might have different substrate specificities since it has been reported that the S. cerevisiae and C. albicans Exg1 proteins are active against not only β(1,3)-glucan but also β(1,6)-glucans (54, 55). To check whether the S. pombe Exg proteins indeed contained glucanase activity, Exg1 was purified from P. pastoris cells expressing an Exg1-His6-myc construct. The enzymatic activity of the purified protein was assayed using laminarin [a β(1,3) polymer], pustulan [a β(1,6) polymer], scleroglucan [a β(1,3)-glucan with β(1,6) branches], lichenan [a mixed β(1,3)-β(1,4)-glucan], nigeran [an insoluble α(1,3) polymer], or the synthetic compound p-nitrophenyl-β-d-glucopyranoside (pNPG) as a substrate. As shown in Fig. 4A, activity was detected only when pustulan was used as a substrate, indicating that Exg1 was a glucanase specific for β(1,6) linkages, in contrast to the S. cerevisiae and C. albicans proteins, which are able to cleave both β(1,3) and β(1,6) polymers. In addition, no activity was detected against pNPG, suggesting that the mechanism of action was not exohydrolytic.

Fig. 4.
Exg proteins have activity against β(1,6)-glucans. (A) β-Glucanase activity of purified Exg1 against laminarin [β(1,3)-glucan], pustulan [β(1,6)-glucan], schleroglucan [β(1,3)-glucan with β(1,6) ramifications], ...

To further confirm these results and to investigate the kinetics and degradation pattern of different substrates, high-performance liquid chromatography (HPLC) was used to analyze the reaction products. Soluble pustulan was used as substrate for the assay. After 2 h of incubation, Exg1 had released small oligosaccharides (DP of 2 to 30). The progressive degradation resulted in the formation of numerous reducing β(1,6)-oligosaccharides of various size (Fig. 4B). Gentobiose to gentohexose (Fig. 4B, G2 to G6) were the main reaction products after 4 h of incubation. The production of small amounts of glucose and large amounts of oligosaccharides of different sizes proved that Exg1 degraded linear β(1,6)-glucan substrates with an endolytic mode of action. No degradation of laminari-oligosaccharides [β(1,3)-glucans] such as laminarin, laminari-hexaose, or octaose was observed (data not shown), confirming that β(1,3)-glucans are not a substrate for Exg1.

Having determined that Exg1 was a β(1,6)-glucanase, we tested whether Exg2 and Exg3 possessed similar activities. To this end, extracts from cells overexpressing each of the three genes under the control of the thiamine-repressible nmt1+ promoter were prepared and assayed using pustulan as a substrate. The results indicated that overexpression of exg1+ and exg3+ produced a 3-fold and 6-fold increase in activity, respectively, in comparison with the activity found in cells grown in the presence of thiamine (promoter repressed) or in wild-type cells, confirming that the two proteins are β(1,6)-glucanases (Fig. 4C). However, no activity could be detected in cells overexpressing exg2+.

Deletion of exg genes presents no apparent phenotype.

To determine the biological role of the three Exg proteins during the life cycle of the fission yeast, mutant strains lacking each of the three genes were constructed. The strains were viable, and no growth defects were observed in different media (YES medium or minimal medium) or at different temperatures (25 to 37°C). Growth on plates supplemented with cell wall-disturbing agents, such as calcofluor white or Congo red was also tested, and no significant differences with the wild-type strain were detected. This could be due to a redundant function of the three Exg proteins. However, similar results were obtained when the triple mutant exg1Δ exg2Δ exg3Δ was tested, indicating that these proteins do not perform an essential role in cell wall construction or that the cells have additional mechanisms to compensate for the absence of Exg proteins. Since Exg1 is expressed during the cell cycle slightly before the endo-β(1,3)-glucanase Eng1 and since it also localizes to the septum region during cell division, the double mutant exg1Δ eng1Δ was constructed to analyze whether Exg1 plays a minor role during cell separation. The phenotype of exg1Δ eng1Δ cells was almost indistinguishable from that of eng1Δ cells.

Overexpression of exg2+ causes alterations in the cell wall.

The effect of overexpression of the three exg+ genes was also assessed. Strains containing the genes under the control of the strong version of the nmt1+ promoter were constructed and grown in the absence of thiamine. Overexpression of exg1+ and exg3+ produced a moderate defect in the growth of the strains, but the microscopic appearance of the cells was similar to that of wild-type cells (data not shown). In contrast, cells overexpressing exg2+ had a severe growth defect, and they stopped growing at 10 to 12 h after induction (Fig. 5A). To determine the nature of the defect, the morphology of cells was analyzed. By 12 h, the cells had an abnormal morphology and had become rounded and irregularly shaped. Also, large amounts of abnormal material had accumulated at the poles of the cell. In an effort to determine the identity of the accumulated material, cells overexpressing exg2+ were stained with aniline blue, a dye that preferentially binds to β(1,3)-glucans and is used to stain the cell wall and septum in S. pombe (25). The material stained well with aniline blue (Fig. 5B), indicating that it was cell wall and that at least a portion of it was comprised of β-glucans. The excess of cell wall accumulated specifically at the poles and septum of the cells.

Fig. 5.
Overexpression of exg2+ produces severe defects in cell growth. (A) Growth of wild-type (WT) strains carrying P3Xnmt1-exg2 (3X-exg2) or vector alone grown in the presence (+) or absence (−) of thiamine (T). Cells overexpressing exg2+ cease growth ...

The previous observations were confirmed when the cells were observed by electron microscopy (Fig. 6 A to F). In comparison with the normal appearance of the wall of wild-type cells (Fig. 6A and E), the wall of cells overexpressing exg2+ clearly contained extra material that accumulated at the poles of the cell and the separation septum (Fig. 6B to D and F). Interestingly, the excess material was incorporated into the cell wall and had a similar appearance to the rest of the cell wall.

Fig. 6.
Effects of exg2+ overexpression. Wild-type (WT) cells containing vector or plasmid pML2 (overexpressing [OE] exg2+), as indicated, were grown for 22 h in the absence of thiamine and prepared for transmission electron microscopy. Images of panels A to ...

Cells overexpressing exg2+ accumulate α- and β-glucans.

To identify the nature of the material that accumulated in cells overexpressing exg2+ more precisely, cell wall constituents were isolated and characterized after growing the cells in the presence of [U-14C]glucose. Incorporation of radioactive glucose into the wall of cells grown in the presence of thiamine was similar to that of wild-type cells, but it increased considerably in the absence of thiamine (from 34 to 47% of total glucose incorporated) (Fig. 6G). A dramatic increase in the amount of α(1,3)- and β(1,3)-glucan was detected under conditions of exg2+ overexpression (Fig. 6H), but the β/α-glucan ratios were similar in the three strains, indicating a simultaneous increase in both glucan polymers. Additionally, the amount of galactomannan was not significantly affected, leading to an alteration in the ratio of glucan to galactomannan (around two times more glucan than galactomannan). These results suggest that strains overexpressing exg2+ have lost the ability to properly coordinate glucan polymer synthesis with cell growth and that glucan synthases produce an excess of cell wall material.

The N-terminal region and the catalytic domain of Exg2 are required for cell wall accumulation.

To investigate the reason for increased cell wall accumulation in cells overexpressing exg2+, versions of the Exg2 protein in which different domains were deleted were constructed. When the exg2 sequence was analyzed using topology prediction programs such as TopPred (http://bioweb.pasteur.fr/seqanal/interfaces/toppred.html) or TMHMM (http://www.cbs.dtu.dk/services/TMHMM-2.0/), the N terminus was predicted to be cytoplasmic while the catalytic domain would be extracellular. Thus, mutant proteins lacking the putative cytoplasmic tail of the protein (Exg2-ΔN), the transmembrane domain (Exg2-ΔTM), the spacer domain that separates the transmembrane and catalytic domains (Exg2-Δout), or the catalytic domain (Exg2-ΔGH5) were generated (Fig. 7A). One possibility for explaining these results is that an excess of enzymatic activity could weaken the cell wall, and as a consequence the synthesis of glucans would be induced. To test this possibility, we also created proteins containing single amino acid changes in the two glutamic acid residues that form part of the catalytic center of GH5 proteins, which are also conserved in exg2+. These mutations involved replacing E338 with Ala (E338A, yielding Exg2-A1), the replacement of E439 with Gln (E349Q, yielding Exg2-A2), and the double mutant (E338A-E349Q, yielding Exg2-A1A2). As described for other members of this family (11, 30), these mutations should completely eliminate the catalytic activity of the enzyme, if present. All the constructs were cloned under the control of the strong version of the nmt1+ promoter and introduced into the wild-type strain to test their effects.

Fig. 7.
Overexpression of mutant forms of exg2+. (A) Schematic representation of Exg2 and different mutant versions generated. The gray box represents the GH5 domain, and the hatched box represents the putative transmembrane domain. The different constructs were ...

Overexpression of Exg2-ΔN resulted in cells that had an abnormal morphology (the cells were rounder than wild-type cells) but did not exhibit a large accumulation of cell wall material (Fig. 7B). Overexpression of Exg2-ΔTM resulted in almost wild-type cells, but since the protein levels were significantly lower than those found in WT cells (Fig. 7C), the absence of phenotype could be due to the small amount of protein. The same reason—reduced protein levels—could account for the fact that the overexpression of Exg2-Δout resulted in a modest phenotype, with a minor accumulation of glucans in the septum region. When the construct lacking the catalytic domain (Exg2-ΔGH5) was overexpressed, the cells showed a wild-type morphology, indicating that this region of the protein is required for the activation of glucan synthesis. However, the putative catalytic activity of the enzyme was not required for this effect since cells carrying the double mutant Exg2-A1A2 (and also the two single mutants Exg2-A1 and Exg2-A2) were almost identical to those overexpressing the full-length protein (Fig. 7B). Thus, these results indicate that the N-terminal region and the GH5 domains are required to induce abnormal glucan synthesis but that the catalytic activity, if present, is not essential.

DISCUSSION

Cell wall growth and extension represent a delicate balance between the hydrolysis of the existing cell wall and the synthesis of new wall. A considerable body of evidence suggests that fungal β(1,3)-glucanases play key roles in morphogenetic processes during development and differentiation. Since β-glucans are major components of fungal and yeast cell walls, it seems likely that β-glucanases would play a crucial role in this process, partially hydrolyzing localized areas and enabling the insertion of new cell wall material, without disturbing the overall integrity of the cell (1). The complement of glucan-modifying enzymes present in S. cerevisiae is very complex, and a large number of proteins have been characterized, but much less is known about the role of hydrolases and glucan-remodeling enzymes during the life cycle of fission yeasts. Only a few glycoside hydrolases have been studied in fission yeast, such as the endo-α(1,3)-glucanases Agn1 and Agn2, the endo-β(1,3)-glucanases Eng1 and Eng2, and the glycanosyl-transferases from family GH72 (18, 19, 21, 22, 35,38). In this study, we have characterized the three genes belonging to family GH5 that are present in the S. pombe genome. They were identified by comparison of the S. cerevisiae Exg1 protein with the fission yeast genomic sequences and were named exg1+ (SPBC1105.05), exg2+ (SPAC12B10.11), and exg3+ (SPBC2D10.05).

Family GH5 is a large and diverse group of hydrolases with different substrate specificities, such as exo-β(1,3)-glucanases, endo-β(1,4)-glucanases (cellulases), endo-β(1,6)-glucanases, endo-β(1,4)-xylanases, or β(1,3)-mannanases (Carbohydrate Active Enzymes database [http://www.cazy.org/]). In many cases, hydrolysis of glycoside bonds takes place via a general acid catalysis mechanism, which requires two acidic residues, one acting as an acid/base catalyst (proton donor) and the other acting as a nucleophile (17). There is a low degree of conservation of the primary sequence of these proteins, but all of them contain a conserved fold consisting of a (β/α)8 barrel. Despite the considerable sequence divergence, all of the proteins share the signature [LIV]-[LIVMFYWGA](2)-[DNEQG]-[LIVMGST]-{SENR}-N-E-[PV]-[RHDNSTLIVFY] as well as eight invariant residues that are involved in the catalysis and the recognition of the glycosyl group attacked during cleavage. Comparison of the three S. pombe proteins to GH5 proteins revealed that the signature and the eight invariant residues were conserved, suggesting that they are new members of this family.

The S. pombe proteins share sequence homology to S. cerevisiae Exg1 and C. albicans Xog1 (CaXog1), which are exo-β(1,3)-glucanases that also act on β(1,6) linkages (10, 43, 57). However, since it has been reported that S. pombe lacks any detectable exo-β(1,3)-glucanase activity (48), the biochemical activity and substrate specificity of the three S. pombe proteins were analyzed in strains overexpressing each of them. In contrast to ScExg1 and CaXog1, the S. pombe Exg1 and Exg3 were highly specific for β(1,6)-glucans (pustulan), being unable to degrade linear or branched β(1,3)-glucans. However, no enzymatic activity was detected for Exg2 in different assays or using different substrates. Therefore, it is possible that Exg2 may have diverged from other GH5 members, losing its catalytic activity during evolution. Alternatively, Exg2 might act on a substrate different from the substrates used in the assay, or it could catalyze a transglycosidase reaction that cannot be detected with the assay used. Indeed, transglycosidase activity has been reported for ScExg1 and CaXog1 (54, 55). Analysis of the reaction products released by Exg1 on β(1,6)-glucans by HPLC revealed that the enzyme had endolytic activity since the main products were gento-oligosaccharides (DP of 2 to 6). Therefore, the mode of action of S. pombe Exg1 is similar to that previously suggested for the BGN16.2 glucanase from the filamentous fungus Trichoderma harzianum (20), and it is different from that found in ScExg1 or CaXog1, which have nonspecific β(1,3)-glucanase, β(1,6)-glucanase, and β-glucosidase activities, with an exolytic mode of action (10, 43, 51). Multiple sequence alignment of yeast and fungal GH5 proteins indicated that the S. pombe proteins cluster in three different branches, but none of them associated with the branch that contains β(1,6)-glucanases such as T. harzianum BGN16.2 (see B9VQ16_TRIHA in Fig. S1 in the supplemental material).

Although no phenotypes have been detected for exg1Δ mutants, based on the fact that Exg1 was secreted and that it localized to the cell wall, it should be involved in the metabolism of cell wall β(1,6)-glucans in vivo. Similarly, the physiological role of ScExg1 has not been clearly established, but it has been proposed that in vivo it would be involved in the metabolism of β(1,6)-glucans since its deletion results in an increase in killer toxin sensitivity, while its overexpression produces resistance (24). Furthermore, overproduction or deletion of the gene leads to detectable in vivo alterations in the cell wall β(1,6)-glucan content, suggesting that its function in the cell wall could be related to the metabolism of β(1,6)-glucan. This protein localized to the septum region in a pattern that was different from that found for Eng1, the endo-β(1,3)-glucanase responsible for primary septum degradation (35). Interestingly, the β(1,6)-branched β(1,3)-glucan spans the whole thickness of the septum, with a tendency to become more concentrated in the primary septum, while β(1,6)-glucan is close to the cell membrane, labeling only the secondary septum (23). Thus, it is possible that Exg1 could play a minor role during cell separation, acting as an endo-β(1,6)-glucanase required for the hydrolysis the β(1,6)-glucans of the secondary septum.

Exg2 protein is different from the other S. pombe GH5 proteins in that it contains a putative transmembrane region before the catalytic domain. Sequence alignment indicates that Exg2 clusters in a branch containing a group of proteins of fungal origin (Aspergillus fumigatus, Aspergillus nidulans, Aspergillus terreus, Aspergillus clavatus, Neurospora crassa, or Magnaporthe grisea), all of which contain a putative transmembrane domain before the catalytic domain (see Fig. S1 in the supplemental material). However, the extent of the putative cytoplasmic region is longer in the fungal proteins than in S. pombe Exg2 although no function has been described for any of them. We have demonstrated that Exg2 is indeed an integral membrane protein that fractionates in the detergent-resistant membrane fraction, a biochemical test for lipid-raft association (56), and that its overexpression produces abnormal cell wall deposition. Interestingly, it has recently been reported that overexpression of a catalytically inactive form of Gas3 is toxic for gas1Δ mutants in S. cerevisiae (49), and it was proposed that hyperaccumulation of Gas3 might produce a physical disturbance of the cell wall structure. Thus, it is possible that the defect in exg2+ overexpression in S. pombe might be due to a similar cause. Alternatively, it is possible that Exg2 might activate cell wall synthesis through an unknown mechanism. Unfortunately, attempts to identify proteins that might interact with the Exg2 cytoplasmic tail by two-hybrid screenings were negative (data not shown). Further analysis will be necessary to analyze this effect in more detail.

Finally, Exg3 is a β(1,6)-glucanase that localizes to the cytoplasm. The function of cytoplasmic glucanases is not currently known although they are present in different yeast organisms. Indeed, Exg3 clusters in a tree branch in which all the members lack a putative signal secretion sequence (see Fig. S1 in the supplemental material). Also, S. pombe contains cytoplasmic glucanases from other families, such as the endo-α(1,3)-glucanase Agn2 (family GH71) and the endo-β(1,3)-glucanase Eng2 (family GH81). It has been shown that these two proteins are required to hydrolyze the cell wall of asci, allowing the dehiscence of spores and their dispersal (19, 22). Thus, cytoplasmic glucanases might perform their function at specific moments of the life cycle.

Supplementary Material

[Supplemental material]

ACKNOWLEDGMENTS

We thank members of the lab for helpful comments and discussions on the manuscript and Nick Skinner for revision of the manuscript.

This research was supported by grants from the Comisión Interministerial de Ciencia y Tecnología (BFU2007-60390/BMC) and Junta de Castilla y Leon (GR231). A. B. Martín-Cuadrado was the recipient of a fellowship from Ministerio de Educación y Ciencia (Spain).

Footnotes

Supplemental material for this article may be found at http://ec.asm.org/.

[down-pointing small open triangle]Published ahead of print on 17 September 2010.

REFERENCES

1. Adams D. J. 2004. Fungal cell wall chitinases and glucanases. Microbiology 150:2029–2035. [PubMed]
2. Alonso-Núñez M., An H., Martín-Cuadrado A. B., Mehta S., Petit C., Sipiczki M., del Rey F., Gould K., Vázquez de Aldana C. R. 2005. Ace2p controls the expression of genes required for cell separation in Schizosaccharomyces pombe. Mol. Biol. Cell 16:2003–2017. [PMC free article] [PubMed]
3. Arellano M., Duran A., Perez P. 1997. Localisation of the Schizosaccharomyces pombe rho1p GTPase and its involvement in the organisation of the actin cytoskeleton. J. Cell Sci. 110:2547–2555. [PubMed]
4. Bähler J., Wu J. Q., Longtine M. S., Shah N. G., McKenzie A., Steever A. B., Wach A., Philippsen P., Pringle J. R. 1998. Heterologous modules for efficient and versatile PCR-based gene targeting in Schizosaccharomyces pombe. Yeast 14:943–951. [PubMed]
5. Baladrón V., Ufano S., Dueñas E., Martín-Cuadrado A. B., del Rey F., Vázquez de Aldana C. R. 2002. Eng1p, an endo-1,3-β-glucanase localized at the daughter side of the septum, is involved in cell separation in Saccharomyces cerevisiae. Eukaryot. Cell 1:774–786. [PMC free article] [PubMed]
6. Beauvais A., Drake R., Ng K., Diaquin M., Latgé J. P. 1993. Characterization of the 1,3-β-glucan synthase of Aspergillus fumigatus. J. Gen. Microbiol. 139:3071–3078. [PubMed]
7. Bush D. A., Horisberger M., Horman I., Wursch P. 1974. The wall structure of Schizosaccharomyces pombe. J. Gen. Microbiol. 81:199–206. [PubMed]
8. Cabib E., Blanco N., Grau C., Rodríguez-Peña J. M., Arroyo J. 2007. Crh1p and Crh2p are required for the cross-linking of chitin to β(1-6)glucan in the Saccharomyces cerevisiae cell wall. Mol. Microbiol. 63:921–935. [PubMed]
9. Cabib E., Farkas V., Kosik O., Blanco N., Arroyo J., McPhie P. 2008. Assembly of the yeast cell wall. Crh1p and Crh2p act as transglycosylases in vivo and in vitro. J. Biol. Chem. 283:29859–29872. [PMC free article] [PubMed]
10. Chambers R. S., Broughton M. J., Cannon R. D., Carne A., Emerson G. W., Sullivan P. A. 1993. An exo-β-(1,3)-glucanase of Candida albicans: purification of the enzyme and molecular cloning of the gene. J. Gen. Microbiol. 139:325–334. [PubMed]
11. Chambers R. S., Walden A. R., Brooke G. S., Cutfield J. F., Sullivan P. A. 1993. Identification of a putative active site residue in the exo-β-(1,3)-glucanase of Candida albicans. FEBS Lett. 327:366–369. [PubMed]
12. Cid V. J., Durán A., del Rey F., Snyder M., Nombela C., Sánchez M. 1995. Molecular basis of cell integrity and morphogenesis in Saccharomyces cerevisiae. Microbiol. Rev. 59:345–386. [PMC free article] [PubMed]
13. Correa J., Vázquez de Aldana C. R., San Segundo P., del Rey F. 1992. Genetic mapping of 1,3-β-glucanase-encoding genes in Saccharomyces cerevisiae. Curr. Genet. 22:283–288. [PubMed]
14. Cortés J. C., Konomi M., Martins I. M., Muñoz J., Moreno M. B., Osumi M., Durán A., Ribas J. C. 2007. The (1,3)β-d-glucan synthase subunit Bgs1p is responsible for the fission yeast primary septum formation. Mol. Microbiol. 65:201–217. [PubMed]
15. Cortés J. C. G., Carnero E., Ishiguro J., Sánchez Y., Durán A., Ribas J. C. 2005. The novel fission yeast (1,3)β-d-glucan synthase catalytic subunit Bgs4p is essential during both cytokinesis and polarized growth. J. Cell Sci. 118:157–174. [PubMed]
16. Craven R. A., Griffiths D. J., Sheldrick K. S., Randall R. E., Hagan I. M., Carr A. M. 1998. Vectors for the expression of tagged proteins in Schizosaccharomyces pombe. Gene 221:59–68. [PubMed]
17. Davies G., Henrissat B. 1995. Structures and mechanisms of glycosyl hydrolases. Structure 3:853–859. [PubMed]
18. Dekker N., Speijer D., Grün C. H., van den Berg M., de Haan A., Hochstenbach F. 2004. Role of the α-glucanase Agn1p in fission-yeast cell separation. Mol. Biol. Cell 15:3903–3914. [PMC free article] [PubMed]
19. Dekker N., van Rijssel J., Distel B., Hochstenbach F. 2007. Role of the α-glucanase Agn2p in ascus-wall endolysis following sporulation in fission yeast. Yeast 24:279–288. [PubMed]
20. de la Cruz J., Pintor-Toro J. A., Benitez T., Llobell A. 1995. Purification and characterization of an endo-β-1,6-glucanase from Trichoderma harzianum that is related to its mycoparasitism. J. Bacteriol. 177:1864–1871. [PMC free article] [PubMed]
21. de Medina-Redondo M., Arnáiz-Pita Y., Fontaine T., del Rey F., Latgé J. P., Vázquez de Aldana C. R. 2008. The β-1,3-glucanosyltransferase gas4p is essential for ascospore wall maturation and spore viability in Schizosaccharomyces pombe. Mol. Microbiol. 68:1283–1299. [PubMed]
22. Encinar del Dedo J., Dueñas E., Arnáiz Y., del Rey F., Vázquez de Aldana C. R. 2009. β-Glucanase Eng2 is required for ascus wall endolysis after sporulation in the fission yeast Schizosaccharomyces pombe. Eukaryot. Cell 8:1278–1286. [PMC free article] [PubMed]
23. Humbel B. M., Konomi M., Takagi T., Kamasawa N., Ishijima S. A., Osumi M. 2001. In situ localization of β-glucans in the cell wall of Schizosaccharomyces pombe. Yeast 18:433–444. [PubMed]
24. Jiang B., Ram A. F., Sheraton J., Klis F. M., Bussey H. 1995. Regulation of cell wall β-glucan assembly: PTC1 negatively affects PBS2 action in a pathway that includes modulation of EXG1 transcription. Mol. Gen. Genet. 248:260–269. [PubMed]
25. Kippert F., Lloyd D. 1995. The aniline blue fluorochrome specifically stains the septum of both live and fixed Schizosaccharomyces pombe cells. FEMS Microbiol. Lett. 132:215–219. [PubMed]
26. Kopecka M., Fleet G. H., Phaff H. J. 1995. Ultrastructure of the cell wall of Schizosaccharomyces pombe following treatment with various glucanases. J. Struct. Biol. 114:140–152. [PubMed]
27. Liu J., Tang X., Wang H., Balasubramanian M. 2000. Bgs2p, a 1,3-β-glucan synthase subunit, is essential for maturation of ascospore wall in Schizosaccharomyces pombe. FEBS Lett. 478:105–108. [PubMed]
28. Liu J., Tang X., Wang H., Oliferenko S., Balasubramanian M. K. 2002. The localization of the integral membrane protein Cps1p to the cell division site is dependent on the actomyosin ring and the septation-inducing network in Schizosaccharomyces pombe. Mol. Biol. Cell 13:989–1000. [PMC free article] [PubMed]
29. Liu J., Wang H., McCollum D., Balasubramanian M. K. 1999. Drc1p/Cps1p, a 1,3-β-glucan synthase subunit, is essential for division septum assembly in Schizosaccharomyces pombe. Genetics 153:1193–1203. [PMC free article] [PubMed]
30. Mackenzie L. F., Brooke G. S., Cutfield J. F., Sullivan P. A., Withers S. G. 1997. Identification of Glu-330 as the catalytic nucleophile of Candida albicans exo-β-(1,3)-glucanase. J. Biol. Chem. 272:3161–3167. [PubMed]
31. Magnelli P. E., Cipollo J. F., Robbins P. W. 2005. A glucanase-driven fractionation allows redefinition of Schizosaccharomyces pombe cell wall composition and structure: assignment of diglucan. Anal. Biochem. 336:202–212. [PubMed]
32. Manners D. J., Meyer M. T. 1977. The molecular structures of some glucans from the cell wall of Schizosaccharomyces pombe. Carbohydr. Res. 57:189–203.
33. Martín V., García B., Carnero E., Durán A., Sánchez Y. 2003. Bgs3p, a putative 1,3-β-glucan synthase subunit, is required for cell wall assembly in Schizosaccharomyces pombe. Eukaryot. Cell 2:159–169. [PMC free article] [PubMed]
34. Martín V., Ribas J. C., Carnero E., Durán A., Sánchez Y. 2000. bgs2+, a sporulation-specific glucan synthase homologue is required for proper ascospore wall maturation in fission yeast. Mol. Microbiol. 38:308–321. [PubMed]
35. Martín-Cuadrado A. B., Dueñas E., Sipiczki M., Vázquez de Aldana C. R., del Rey F. 2003. The endo-β-1,3-glucanase Eng1p is required for dissolution of the primary septum during cell separation in Schizosaccharomyces pombe. J. Cell Sci. 116:1689–1698. [PubMed]
36. Martín-Cuadrado A. B., Encinar del Dedo J., de Medina-Redondo M., Fontaine T., del Rey F., Latgé J. P., Vázquez de Aldana C. R. 2008. The Schizosaccharomyces pombe endo-1,3-β-glucanase Eng1 contains a novel carbohydrate binding module required for septum localization. Mol. Microbiol. 69:188–200. [PubMed]
37. Martín-Cuadrado A. B., Fontaine T., Esteban P. F., Encinar del Dedo J., de Medina-Redondo M., del Rey F., Latgé J. P., Vázquez de Aldana C. R. 2008. Characterization of the endo-β-1,3-glucanase activity of S. cerevisiae Eng2 and other members of the GH81 family. Fungal Genet. Biol. 45:542–553. [PubMed]
38. Martín-Cuadrado A. B., Morrell J. L., Konomi M., An H., Petit C., Osumi M., Balasubramanian M., Gould K. L., del Rey F., Vázquez de Aldana C. R. 2005. Role of septins and the exocyst complex in the function of hydrolytic enzymes responsible for fission yeast cell separation. Mol. Biol. Cell 16:4867–4881. [PMC free article] [PubMed]
39. Mitchison J. M., Nurse P. 1985. Growth in cell length in the fission yeast Schizosaccharomyces pombe. J. Cell Sci. 75:357–376. [PubMed]
40. Moreno M. B., Durán A., Ribas J. C. 2000. A family of multifunctional thiamine-repressible expression vectors for fission yeast. Yeast 16:861–872. [PubMed]
41. Moreno S., Klar A., Nurse P. 1991. Molecular genetics analysis of fission yeast Schizosaccharomyces pombe. Methods Enzymol. 194:795–823. [PubMed]
42. Mouyna I., Fontaine T., Vai M., Monod M., Fonzi W. A., Diaquin M., Popolo L., Latgé H. R. P. J. P. 2000. Glycosylphosphatidylinositol-anchored glucanosyltransferases play an active role in the biosynthesis of the fungal cell wall. J. Biol. Chem. 275:14882–14889. [PubMed]
43. Nebreda A. R., Vázquez C. R., Villa T. G., Villanueva J. R., del Rey F. 1987. Heterogeneous glycosylation of the EXG1 gene product accounts for the two extracellular exo-β-glucanases of Saccharomyces cerevisiae. FEBS Lett. 220:27–30. [PubMed]
44. Nelson M. J. 1957. Colorimetric analysis of sugars. Methods Enzymol. 3:85–86.
45. Osumi M., Sato M., Ishijima S. A., Konomi M., Takagi T., Yaguchi H. 1998. Dynamics of cell wall formation in fission yeast, Schizosaccharomyces pombe. Fungal Genet. Biol. 24:178–206. [PubMed]
46. Percival-Smith A., Segall J. 1984. Isolation of DNA sequences preferentially expressed during sporulation in Saccharomyces cerevisiae. Mol. Cell. Biol. 4:142–150. [PMC free article] [PubMed]
47. Ragni E., Fontaine T., Gissi C., Latgé J. P., Popolo L. 2007. The Gas family of proteins of Saccharomyces cerevisiae: characterization and evolutionary analysis. Yeast 24:297–308. [PubMed]
48. Reichelt B. Y., Fleet G. H. 1981. Isolation, properties, function, and regulation of endo-(1-3)-β-glucanases in Schizosaccharomyces pombe. J. Bacteriol. 147:1085–1094. [PMC free article] [PubMed]
49. Rolli E., Ragni E., Rodriguez-Peña J. M., Arroyo J., Popolo L. 2010. GAS3, a developmentally regulated gene, encodes a highly mannosylated and inactive protein of the Gas family of Saccharomyces cerevisiae. Yeast 27:597–610. [PubMed]
50. Rustici G., Mata J., Kivinen K., Lió P., Penkett C. J., Burns G., Hayles J., Brazma A., Nurse P., Bähler J. 2004. Periodic gene expression program of the fission yeast cell cycle. Nat. Genet. 36:809–817. [PubMed]
51. Sánchez M., Villanueva J. R., Villa T. G. 1982. Saccharomyces cerevisiae secretes two exoglucanases. FEBS Lett. 138:209–212. [PubMed]
52. San Segundo P., Correa J., Vázquez de Aldana C. R., del Rey F. 1993. SSG1, a gene encoding a sporulation-specific 1,3-β-glucanase in Saccharomyces cerevisiae. J. Bacteriol. 175:3823–3837. [PMC free article] [PubMed]
53. Smogyi M. 1952. Notes on sugar determination. J. Biol. Chem. 195:19–23. [PubMed]
54. Stubbs H. J., Brasch D. J., Emerson G. W., Sullivan P. A. 1999. Hydrolase and transferase activities of the β-1,3-exoglucanase of Candida albicans. Eur. J. Biochem. 263:889–895. [PubMed]
55. Suzuki K., Yabe T., Maruyama Y., Abe K., Nakajima T. 2001. Characterization of recombinant yeast exo-β-1,3-glucanase (Exg1p) expressed in Escherichia coli cells. Biosci. Biotechnol. Biochem. 65:1310–1314. [PubMed]
56. Takeda T., Kawate T., Chang F. 2004. Organization of a sterol-rich membrane domain by cdc15p during cytokinesis in fission yeast. Nat. Cell Biol. 6:1142–1144. [PubMed]
57. Vázquez de Aldana C. R., Correa J., San Segundo P., Bueno A., Nebreda A. R., Méndez E., del Rey F. 1991. Nucleotide sequence of the exo-1,3-β-glucanase gene, EXG1, of the yeast Saccharomyces cerevisiae. Gene 97:173–182. [PubMed]
58. Zilahi E., Salimova E., Simanis V., Sipiczki M. 2000. The S. pombe sep1 gene encodes a nuclear protein that is required for periodic expression of the cdc15 gene. FEBS Lett. 481:105–108. [PubMed]

Articles from Eukaryotic Cell are provided here courtesy of American Society for Microbiology (ASM)

Formats:

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...

Links

  • MedGen
    MedGen
    Related information in MedGen
  • Pathways + GO
    Pathways + GO
    Pathways, annotations and biological systems (BioSystems) that cite the current article.
  • PubMed
    PubMed
    PubMed citations for these articles
  • Taxonomy
    Taxonomy
    Related taxonomy entry
  • Taxonomy Tree
    Taxonomy Tree

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...