Fission Yeast α-Glucan Synthase Mok1 Requires the Actin Cytoskeleton to Localize the Sites of Growth and Plays an Essential Role in Cell Morphogenesis Downstream of Protein Kinase C Function

Strains used in this study are listed in Table I. Complete medium, YPD (1% yeast extract, 2% polypeptone, 2% dextrose) which contains 10 μg/ml Phloxine B (called YPDP), YES5 (0.5% yeast extract, 3% dextrose, and 75 μg/ml of adenine, histidine, leucine, and uracil), and modified synthetic EMM2 have been described (Moreno et al., 1991).

Standard procedures for S. pombe genetics were followed according to Moreno et al. (1991). Gene disruptions are abbreviated by putting the symbol Δ before the gene name, e.g., Δmok1. Proteins are designated by an uppercase first letter, e.g., Mok1.

As an initial screen collection of temperature-sensitive (ts) or cold-sensitive (cs) mutants (2,822 or 1,961, the restrictive temperatures were set at 36 and 20°C, respectively) were visually selected by calcofluor staining and morphological mutants with altered cell shape at the restrictive temperature were isolated. Four types of morphological mutants were observed, namely round or pear-like, elongated, bent or branched, and septation defective. Round or pear-shaped mutants were further screened for the supersensitivity to a protein kinase inhibitor staurosporine (STS, 1.5 μg/ml) for which Pck1 and Pck2 are two of the major targets (Toda et al., 1993). To exclude mutants that were nonspecifically supersensitive to a broad range of drugs, sensitivity to K-252a (3 μg/ml), which structurally related but has distinct biological targets to STS (Tamaoki and Nakano, 1990), was also examined.

After mating of each strain, free spores were directly plated on four YPDP plates, three of which were incubated at 20, 29, or 36°C and the fourth was incubated on a YPDP plate containing 1.5 μg/ml STS at 29°C. If the difference in colony number between the plates growing under these conditions was >10⁴-fold, the two mutants were assumed to be allelic.

Standard molecular biology techniques were followed as described (Sambrook et al., 1989). Enzymes were used according to the recommendation of the suppliers (New England Biolabs Inc.). Nucleotide sequence was determined by use of dideoxy-method (Sanger et al., 1977).

An S. pombe genomic library constructed in the vector pDB248 (Moreno et al., 1991) was used for the isolation of plasmids that complemented the ts mok1 mutant (DH664). 2 out of 20,000 colonies were capable of growing at 36°C and the segregation analysis showed that the Ts⁺ phenotype was plasmid-dependent. Two different plasmids (pMK100 and pMK1100) containing different, but overlapping, inserts were isolated. Subcloning analysis indicated that the internal SmaI site was essential for the complementing activity of pMK1100. When the 5.5-kb SmaI/BamHI fragment was subcloned from pMK1100, the resulting plasmid (pMK1101) lost the complementing activity. However, Ts⁺ colonies appeared from transformants containing pMK1101 at a frequency of ∼10⁻³. It was found that pMK1101 directed integration of the LEU2 marker into the genome via homologous recombination in these Ts⁺ transformants. Subsequent analysis showed that no ts recombinants arose from 10⁴ recombinants between this integrant and a wild-type strain. This result demonstrated that pMK1101 contains the mok1⁺ gene itself.

Homology searching using Mok1 as query against the fission yeast genome database (Sanger Centre, Cambridge, UK) showed that there are four additional mok1⁺ homologues [designated mok11⁺ (SPAC23D3), mok12⁺ (SPAB4538), mok13⁺ (c16D10S), and mok14⁺ (c63), respectively]. The nucleotide sequence data reported in this paper will appear in the DDBJ/EMBL/Genbank nucleotide sequence databases under the accession numbers: AB019183 (mok1⁺), AB018380 (mok11⁺), AB018381 (mok12⁺), AB018382 (mok13⁺), and AB018383 (mok14⁺).

Three different strategies were taken to disrupt the mok1⁺ gene by use of a PCR-generated fragment containing the ura4⁺ marker (Bähler et al., 1998): one resulted in the complete deletion of the ORF (SKDP1), the second in deletion of a 1,764 bp NH₂-terminal fragment (corresponding to amino acids 66–654) and the third in fusing the DNA encoding HA or GFP tags to the COOH terminus, which was accidentally found during the attempt to construct the tagged mok1⁺ gene. Correct disruption was verified by PCR. In both cases, tetrad analysis of the heterozygous diploid showed two viable (Ura⁻) and two inviable spores, indicating that the mok1⁺ gene is essential for cell viability. The mok11⁺, mok12⁺, mok13⁺, and mok14⁺ genes were also disrupted. Tetrad analysis of a heterozygous diploid yielded four viable spores, in which Ura⁺ and Ura⁻ segregates 2:2 in each case. This indicated that neither mok11⁺, mok12⁺, nor mok13⁺ gene is essential for cell viability. It was also found that double mok11mok12 and triple mok11mok12mok14 disruptions were viable.

Three different fragments of the mok1⁺ gene were amplified with PCR and inserted in pREP1 or pREP2 (the nmt1 promoter-containing vector; Maundrell, 1990); a 7.2-kb fragment containing the whole ORF (5′-TTTGGATCCTATGCATGGTCTTCAAGGGTTATGTTTTAGA-3′ and 5′-TTTGGATCCCTAAGGACGACTAAGGTTTTCACGACGGAA-3′ were used for PCR, yielding pREP1-mok1), an NH₂-terminal 3.4 kb containing amino acid residues 1–1,236 (5′-TTTAGGCCTCTACGAGACTACAGGAGTGACCTT-3′ and 5′-TTTGGATCCTATGCATGGTCTTCAAGGGTTATGTTTTAGA-3′ were used for PCR, yielding pREP2-mok1N) and a COOH-terminal 3.9 kb containing amino acid residues 1,017 to 2,410 (5′-TTTGGATCCTATGCATGGTCTTCAAGGGTTATGTTTTAGA-3′ and 5′-TTTGGATCCCTAAGGACGACTAAGGTTTTCACGACGGAA-3′ were used for PCR, yielding pREP1-mok1C). The nmt1 promoter was integrated in the genome in front of the initiator ATG of the mok1⁺ gene by a PCR-based gene targeting method (SKP103; Bähler et al., 1998).

Fission yeast cells, in which the nmt1 promoter was integrated in front of the ORF of mok1⁺ (SKP103), were grown in liquid minimal medium at 29°C for 12 h and placed on a slide glass embedded in a slice (1 mm) of agar made of minimal medium supplemented with leucine. This slice was then overlaid with a coverslip and sealed with liquid sealer. The slide glass was set under the phase microscope (Zeiss Co.) at room temperature (22°C) and the cells were viewed with a chilled video-rated CCD camera (model C5985; Hamamatsu) connected to a computer (Apple Power Macintosh 8600/200). Photographs of live cells in a fixed field were taken every 30 min. Images were processed by use of Adobe^® Photoshop (version 4).

Cultures of S. pombe cells were supplemented with U-[¹⁴C]glucose (1 μCi/ml) and incubated for additional 4 h. To label cells overproducing mok1⁺ (SKP103 and SKP170), cultures were induced for 14 h in the absence of thiamine before addition of U-[¹⁴C]glucose. Total glucose incorporation was monitored by measuring the radioactivity in trichloroacetic acid insoluble material. Mechanical breakage of cells was done as described (Arellano et al., 1996, 1997) and cell walls were pelleted at 1,000 g for 5 min. 100 μl aliquots of the total wall were incubated with 100 units of zymolyase 100T (Seikagaku Kogyo Co. Ltd.) or Quantazyme (Quantum Biotechnologies Inc.) for 36 h at 30°C. The samples were centrifuged and the supernatant and washed pellet were counted separately. The polysaccharides in the supernatant from the zymolyase 100T reaction were considered to be β-glucan plus galactomannan, and the pellet α-glucan, whereas that from the Quantazyme reactions was considered to be β-glucan and the pellet α-glucan plus galactomannan.

Rabbit polyclonal anti-Mok1 antibody was prepared as follows. To express the fused Mok1 protein, the 0.9-kb fragment corresponding amino acid residues 1,583 to 1,904 (5′-TTTCATATGTCTCAACGTACCCGTGCTCGACTT-3′ and 5′-TTTGGATCCATCTCTGATTTCATTAAGTATTGT-3′ were used for PCR) was inserted into pET10c (Invitrogen Co.). Insoluble proteins were purified using Ni-NTA column. Two rabbits were injected with the Mok1 fusion protein (200 μg per injection). Immunoblotting was performed with crude sera or affinity-purified anti-Mok1 antibodies. Bacterially made Mok1 fusion protein was bound to nitrocellulose filters in order to purify anti-Mok1 antibodies from crude sera. Monoclonal anti–α-tubulin antibody (Sigma Chemical Co.) was also used. Horseradish peroxidase-conjugated goat anti–rabbit IgG, goat anti– mouse IgG (Bio-Rad Laboratories Ltd.) and a chemiluminescence system (ECL; Amersham Corp.) were used to detect bound antibody.

Fission yeast whole cell extracts were prepared using glass beads to disrupt yeast cells in TEG buffer (50 mM Tris-HCl, pH 7.5, 1 mM EDTA, 10% glycerol, 30 mM NaCl, 1 mM DTT, 60 mM β-glycerophosphate, 15 mM p-nitrophenylphosphate, 0.1 mM NaF, 10 μg/ml soybean trypsin inhibitor, 20 μg/ml leupeptin, 50 μg/ml aprotinin, 2 μg/ml pepstatin, 1 mM PMSF, and 0.1 mM Na-orthovanadate). For the subcellular fractionation, cell extracts were microcentrifuged at 13,000 rpm for 15 min at 4°C in various conditions in order to separate soluble from insoluble fractions.

Rhodamine-conjugated phalloidin (Molecular Probes Inc.) or monoclonal anti-chicken gizzard actin antibody (N-350; Amersham Corp.) followed by Cy3-conjugated sheep anti–mouse IgG (Sigma Chemical Co.) was used to visualize actin as described previously (Toda et al., 1996b). Rabbit polyclonal anti-Mok1 serum (1:10 dilution) or undiluted affinity-purified antibodies and Cy3-conjugated goat anti–rabbit IgG (Sigma Chemical Co.) or FITC-conjugated swine anti–rabbit IgG (DAKO Ltd.) was used to localize the Mok1 protein.

A stock solution of latrunculin A (lat A; Molecular Probes Inc.) was made in DMSO (50 mM) and used at a concentration of 100 μM. DMSO alone did not affect the F-actin cytoskeleton or Mok1 distribution.

Cells containing nmt1-mok1⁺ (SP103) grown at 29°C in minimal medium in the presence of 5 μM thiamine were filtered, resuspended in minimal medium without thiamine, and incubated for further 12 h to induce the nmt1 promoter. Then thiamine (5 μM) and cycloheximide (100 μg/ml) were added, cell extracts prepared every 15 min and immunoblotted with anti-Mok1 antibody.

A genetic screen was performed in order to identify genes that regulate cell morphogenesis and function in the protein kinase C (PKC) pathway. Mutants that showed round or pear-shaped morphology at the restrictive temperature were visually isolated from a collection of ts or cs mutants. We have previously shown that two of the major targets of a protein kinase inhibitor staurosporine (STS) in fission yeast are PKC-related molecules, Pck1 and Pck2 (Toda et al., 1993), and that mutants defective in Pck1 and Pck2 function are supersensitive to this drug (Toda et al., 1996b). Accordingly, the sensitivity of each round or pear-like mutant to STS was examined. In total 25 ts and 6 cs mutants were identified that satisfied both criteria. These loci were designated mok (morphological and kinase-inhibitor supersensitive).

Complementation analysis of these mok mutants and known morphological mutants showed that they defined 10 different loci (mok1-10, Table II). mok2 (eight alleles) and mok3 (one allele) were allelic to sts5/orb4 and pck2/ sts6, respectively, which have previously been shown to produce STS-supersensitive round mutants (Toda et al., 1993, 1996b). The most frequently isolated locus was mok1 (twelve ts and two cs alleles, Table II).

The localization of F-actin was examined to characterize the defective phenotypes of mok1 mutants. In fission yeast F-actin localizes to either the growing tips or the medial regions of dividing cells where the septum forms (Marks and Hyams, 1985; Marks et al., 1986, see Fig. 1 a, left). In contrast to wild-type patterns, cortical actin did not show the specific localization in the mok1-664 mutant incubated at 35.5°C for 6 h. Instead, it localized in the cell cortex in a random punctate (right). Percentage of septated cells slightly increased at the restrictive temperature (19% at 26°C and 22% at 36°C for 6 h, Fig. 1 b), suggesting that mok1 mutant is also defective in cell separation. The mok1⁺ gene is, therefore, required for the maintenance of rod-shape, cytokinesis and the cellular localization of F-actin.

pck2 mutants are defective in cell wall integrity and, as a result, these cells are supersensitive to treatment with cell wall digesting enzymes (Shiozaki and Russell, 1995; Toda et al., 1996a,b). To determine the morphology defect of mok1 mutants was also due to an inability to maintain cell wall integrity, mutant cells were treated with β-glucanase. Like pck2, the mok1 mutants were hypersensitive to β-glucanase even at the temperature permissive of 28°C (Fig. 1 c). Defects in cell wall integrity can be sometimes compensated for by increases in the osmolarity of the growth media (Levin and Bartrett-Heubusch, 1992). Consistently mok1 mutant cells were capable of forming colonies at 35.5°C on rich YPD plates containing 1.2 M sorbitol (Fig. 1 d). Also the mok1 mutants were hypersensitive to calcofluor, which disrupts cell wall architecture (data not shown). Therefore, the ts and morphological defective phenotypes of mok1 mutants were, at least in part, ascribable to defects in cell wall integrity.

The mok1⁺ gene was cloned by complementation of the ts defect. Nucleotide sequencing of the cloned DNA fragment identified an uninterrupted ORF consisting of 7,230 bp which encoded a protein of 2,410 amino acid residues (the predicted molecular mass is 272 kD). Homology searching and structural analysis showed that the Mok1 protein comprises five structural domains (Fig. 2, a and b). The first is the NH₂-terminal 30 amino acid residues, which are highly hydrophobic and probably act as a signal peptide. The second domain follows the first 1,000 amino acid residues and has three transmembrane domains. A significant homology to a group of glycoside hydrolases, including bacterial α-amylase (Henrissat and Davies, 1997) was found within this region (Fig. 2 b). The third domain is a central, putative transmembrane, domain (amino acid residues 1,070– 1,090). The fourth comprises the next 1,000 amino acid residues which show significant homology to both bacterial glycogen synthase (Kumar et al., 1986) and plant starch synthase (van der Leij et al., 1991; Fig. 2 b). Consistent with the homology between Mok1 and glucan synthases, two consensus sequences for UDP-glucose binding motifs (Furukawa et al., 1993) were found in Mok1 (Fig. 2 c). Mok1 and plant starch synthase further share a similarity in the N-terminal signal peptide and the region after the amylase-homologous region (Fig. 2 b). The fifth domain is the COOH-terminal 400 amino acid residues that predict 12 membrane spanning domains (Fig. 2 a). Thus the predicted structure suggests that Mok1 is an integral membrane protein that plays a role in degradation or synthesis of cell wall components.

Homology searching against the fission yeast genome database (Sanger Centre) indicated that mok1⁺ constitutes a gene family; four putative homologues (designated mok11⁺, mok12⁺, mok13⁺, and mok14⁺) are found in the genome sequences that are currently available. The predicted mok11⁺ (COOH terminus is unsequenced), mok12⁺ and mok13⁺ gene products also encode larger proteins (1,204, 2,352, and 2,371 amino acid residues, respectively), whereas mok14⁺ encodes a smaller protein (1,369 amino acid residues) which lacks the NH₂-terminal amylase-homologous region. The overall homology among these proteins is 40–50% identity (50–60% similarity; Table III) and all of them share virtually identical hydrophobicity profiles. It is of note that budding yeast does not contain any proteins that share significant homology to Mok1.

Different mok1 mutants alleles (mok1-559, -664, -905, -936, -1071, -1172, -1221, -1224, and -1963) were transformed with the noncomplementing plasmid pMK1101 (containing amino acid residues 1–876) and transformants were streaked and incubated at 35.5°C. In all nine ts mok1 mutant alleles tested, Ts⁺ integrants appeared at a frequency of ∼10⁻³. Thus, the mutation sites in these mok1 mutants are in the region of the NH₂-terminal 876 amino acid residues.

The mok1⁺ gene was disrupted by three different strategies (Materials and Methods). In each case, tetrad analysis indicated that two viable and two inviable spores were obtained (Fig. 2 d) and that viable spores produced Ura⁻ colonies. Microscopic observation of inviable spores showed that most of the Δmok1 spores (18 out of 20 spores) failed to germinate. Therefore, mok1⁺ is essential for cell viability and also is required for germination.

To examine phenotypes arising from overexpression of mok1⁺, three different plasmids, which contained the whole ORF, the NH₂-terminal 1,236 amino acid residues, and the COOH-terminal 1,393 amino acid residues respectively, were constructed in a vector carrying the thiamine-repressible nmt1 promoter (Fig. 3 a). In addition, the nmt1 promoter was integrated into the genome in front of the mok1⁺ gene initiator methionine (ATG). It was found that overexpression of the entire mok1⁺ gene (nmt-mok1⁺), either episomal or integrated, was toxic (Fig. 3 b). Viability dropped sharply upon induction of the nmt1 promoter (25 and 15% after 14 and 18 h induction, respectively, Fig. 3 c). In contrast, overexpression of the NH₂-terminal or COOH-terminal half did not affect viability (Fig. 3 a).

Cell morphology after induction of nmt1-mok1⁺ was examined. Two types of cell shape defects were found. One showed an asymmetrical shape in which one end of the cell swelled abnormally to produce a tadpole (Fig. 3 d, 62% after 20 h induction). The other was observed as pairs of divided cells associated side-by-side (12%), the cell wall of which appeared fragile and often lysed upon division.

To examine the deposition of cell wall material and the localization of actin in mok1⁺-overexpressing cells, cells were stained with DAPI, calcofluor and rhodamine-conjugated phalloidin. In tadpole cells, cortical F-actin localized exclusively in the swollen tips (Fig. 4 a, middle). Calcofluor-staining showed a huge accumulation of cell wall material in the swollen parts of the tadpole cells (Fig. 4 b, arrow). Time-lapse microscopy was performed after gene induction in order to follow morphological changes leading to the tadpole phenotype. Either the new or old end became swollen (the new end is the end that is produced after cytokinesis; the old end is the one that has already existed in the previous cycle). In Fig. 4 c, asymmetrical cell swelling that took place in the new end (marked with arrows) is shown. This analysis suggested that overproduction of Mok1 inhibited the translocation of F-actin (and cell wall materials) from one end to the other.

In contrast to the asymmetrical accumulation of actin in tadpole cells, side-by-side cells showed neither actin staining (Fig. 4 a, bottom) nor the accumulation of cell wall material (Fig. 4 b, arrowhead). These phenotypes suggest that these cells died after lysis. Nuclear staining by DAPI showed that each cell had a nucleus, indicating that chromosome segregation had been completed. In summary, the loss of Mok1 protein resulted in randomization of F-actin, whereas its overproduction abrogated translocation of F-actin from one end of the cell to the other as well as an abnormal accumulation of cell wall material and the lysis of some cells after cell division.

As Mok1 contains two separate regions that display homology to amylase and glucan synthase respectively, we were interested in measuring the level of cell wall components in mutants in which the activity of Mok1 was either down- or upregulated. For this purpose, the level of glucan in cell walls was measured through the incorporation of radioactive glucose in either ts mok1 mutants or cells in which mok1⁺ was overexpressed. As shown in Fig. 5 and Table IV, the level of glucan was significantly altered by manipulation of Mok1. Most remarkably, α-glucan levels in Mok1-overproducing cells were elevated more than threefold over compared with wild-type cells (344%). The level of galactomannan also showed a modest increase (202%), whereas the level of β-glucan decreased slightly (75%).

On the other hand, incubation of ts mok1 mutants at the restrictive temperature resulted in a decrease in α-glucan level by 69%. In contrast, galactomannan and β-glucan levels were virtually unchanged. These results strongly suggested that Mok1 is an α-glucan synthase.

Polyclonal anti-Mok1 antibodies were prepared in order to characterize the Mok1 protein (Materials and Methods). These antibodies detected a band ∼280 kD after immunoblotting of total cell extracts from wild-type cells (Fig. 6 a, lane 1). The intensity of this band increased in cells containing the mok1⁺ gene on a multicopy plasmid (lane 2) and further dramatically increased in a strain in which mok1⁺ was overexpressed by induction of the nmt1 promoter (lane 3). These data showed that the antibody specifically recognized the mok1⁺ gene product.

Total cell extracts were fractionated and immunoblotted using anti-Mok1 antibody. Mok1 was insoluble when a low salt buffer was used for the preparation of whole cell extracts (30 mM NaCl, Fig. 6 b, lanes 1–3) or when the buffer contained high salt (0.5 M, lanes 4 and 5). Treatment with a nonionic detergent solubilized ∼50% of Mok1 (lanes 6 and 7) and ionic detergent resulted in complete solubilization (lanes 8 and 9). Protein denaturation with 8 M urea also solubilized Mok1 (lanes 10 and 11). Next the half-life of the Mok1 protein was examined. After a short induction of the mok1⁺ gene from the nmt1 promoter in minimal medium without thiamine (derepressed), p280^mok1 levels were followed by immunoblotting after the addition of thiamine and cycloheximide to repress promoter and protein synthesis respectively. It was found that Mok1 is a stable protein, a half-life is >120 min (Fig. 6 c). These results suggested that Mok1 is an stable integral membrane protein, which is consistent with the presence of several transmembrane domains (Fig. 2 a).

Immunofluorescence microscopy was performed using anti-Mok1 in order to determine the subcellular localization of Mok1. As shown in Fig. 7 a, Mok1 mainly localizes in two regions, cell tips and the medial region. This localization of Mok1 is very similar to that for F-actin (Marks et al., 1986; Jochová et al., 1991).

Next, cells were simultaneously double stained with anti-Mok1 and anti-actin antibodies. Anti-Mok1 antibodies stained the region of the cell that overlapped with, but not identical to, actin staining (Fig. 7 b). In a newly born small cell (Fig. 7 a, <10 μm), just after cytokinesis, Mok1 and actin localized still in the new end in which the septum had previously been formed. Then these two proteins moved to the old end (Fig. 7 b). Upon NETO (Fig. 7 c, new end take off; Mitchison and Nurse, 1985), both actin and Mok1 were seen at the new end and these two proteins existed hereafter at both ends. It is of note that, while actin mostly localized as a patched pattern, Mok1 localization at the tips was much more homogeneous and produced a uniform staining pattern (Fig. 7, b and c). At mitosis, actin and Mok1 disappeared from cell tips, and upon entry into anaphase (Fig. 7, d and e), both proteins localized in the medial region of the cell as ring-like structures, where septa would subsequently form. Finally in septating cells (Fig. 7 f), actin and Mok1 colocalized with septa and during septation (g), Mok1 staining broadened and looked as if it was doubled in width. In contrast, actin also now looked duplicated but the staining was less uniform and patched structures were more obvious. Finally, just before cell separation (Fig. 7 h), Mok1 localized as dense dot that was bounded by two sets of actin patches in the center of the two dividing cells. It should be noted that localization to dense dot upon cytokinesis has been observed in several actin binding proteins such as Cdc8 (tropomyosin; Balasuramanian et al., 1992) and Myo2 (Type II myosin; Kitayama et al., 1997). In summary Mok1 closely localizes with actin, at the growing cell tips in interphase, the medial regions in mitosis and a central dense dot during the final stage of septation.

Given the localization of actin and Mok1 to the similar regions of the cell, we were keen to determine whether the actin cytoskeleton was required for the localization of Mok1. To address this question, F-actin was depolymerized with latrunculin A (lat A; Spector et al., 1989; Ayscough et al., 1997), and the consequence for the cellular localization of Mok1 determined. As shown in Fig. 8 a, after 10 min upon lat A addition (middle), Mok1 localization to both the growing ends and the medial regions was abolished. Immunoblotting showed that the total level of Mok1 unchanged during lat A treatment (Fig. 8 b). These results suggested that cortical actin is essential for the correct localization of Mok1.

To disrupt the actin cytoskeleton by another approach, the ts profilin mutant cdc3 and the ts tropomyosin mutant cdc8 (Balasubramanian et al., 1992, 1994) were incubated at the restrictive temperature to disassociate F-actin patches from the cell tips and medial regions. Under these conditions, the F-actin patches associated with the cell periphery and the actomyosin ring structure failed to form, resulting in mitosis without cytokinesis and the accumulation of multi-nucleated cells (Fig. 8 c, top and bottom). Mok1, as in lat A–treated cells, dislocalized completely in these cells (middle). We conclude that the cellular localization of Mok1 is dependent on the integrity of the F-actin cytoskeleton.

The localization of Mok1 was examined in the ts mok1 mutants. As shown in Fig. 9 a, the Mok1 protein was no longer localized to discrete regions of the cell, instead the whole cytoplasm was stained. It should be noted that the presence of patch-like actin dots indicated that actin polymers attached with the plasma membrane as in wild-type cells in mok1 mutant cells (Mulholland et al., 1994). This mislocalized pattern of Mok1 was reminiscent of that in cells in which the actin cytoskeleton had been disrupted (Fig. 8). The level of the Mok1 protein was indistinguishable between wild-type and ts mok1 mutants grown at the restrictive temperature (Fig. 9 b), indicating that the apparent lack of the specific localization of Mok1 was not due to reduced protein levels. These results suggested that Mok1 localization requires intact Mok1 activity and/or structure and that the ts Mok1 proteins fail to be transported to the plasma membrane, which leads to randomization of F-actin and cell shape defects.

We sought to examine the functional relationship between Mok1 and Pck1/2. To this end, the cellular localization of Mok1 was examined in pck2 mutant. It was found that Mok1 became mislocalized in this mutant (Fig. 10 a, upper left); neither the cell tips nor the medial regions were stained with anti-Mok1 antibody, instead a dispersed pattern was observed. Immunoblotting showed that the amount of Mok1 did not alter between wild-type and pck2 mutant (lower). In contrast the localization of F-actin appeared not to be severely impaired as cortical actin existed at either the cell tips or the medial regions (upper right).

Next the double mutants between ts mok1 and Δpck2 were constructed. mok1-664Δpck2 was synthetically lethal at 30°C, a temperature at which either single mutant could grew (Fig. 10 b). Synthetic lethality was not observed between mok1-664 and Δpck1, consistent with our previous result showing that pck2⁺ plays the major role (Toda et al., 1993). Although mok1-664Δpck2 managed to form colonies at 27°C, double mutant cells were round and spontaneously lysed (Fig. 10 c), suggesting that cell wall integrity was substantially impaired. This was confirmed by glucanase treatment of the double mutants grown at 27°C, as this strain was lysed much faster than either wild-type or each single mutant alone (Fig. 10 d). These data indicate that Pck2 is required for the specific localization of Mok1 and that Mok1 regulates cell wall integrity either in parallel with or downstream of Pck1/2.

To distinguish between the possibilities that Mok1 acts in parallel with or downstream of the Pck1 and Pck2 pathway, the nmt1-mok1⁺ gene was overexpressed in Δpck2 cells (chromosomally integrated, SKP170). If Mok1 is somehow regulated by Pck1 and Pck2, Δpck2 cells should become resistant to simple overexpression of mok1⁺ as Mok1 activity is expected to be compromised in the absence of Pck2 function. On the other hand, if Mok1 and Pck2 (and Pck1) regulate cell wall integrity in a parallel manner, nmt1-mok1⁺ should be as toxic in Δpck2 cells as in wild-type cells or it might even be more toxic. As shown in Fig. 11 a, Δpck2 cells became insensitive to nmt1-mok1⁺, they were capable of forming colonies in the derepressed conditions, whereas wild-type cells were not. Consistent with this, calcofluor staining showed that morphology of cells in which mok1⁺ was overexpressed was less abnormal in Δpck2 than in wild-type; neither tadpole cells nor asymmetrical accumulation of cell wall materials were observed (Fig. 11 b). The lack of toxicity of nmt1-mok1⁺ was not due to a reduction in the levels of Mok1 in the Δpck2 background (Fig. 11 c).

The level of cell wall glucan was measured in Δpck2 mutant or that containing nmt1-mok1⁺. α-glucan level was decreased in Δpck2 mutant (24% reduction, Fig. 11 d and Table IV). The level of α-glucan in Δpck2 cells containing nmt1-mok1⁺ was elevated, but the increase was greatly compromised compared with a wild-type strain containing nmt1-mok1 (46% lower). The reduced level of α-glucan together with mislocalization of Mok1 might explain why Δpck2 mutant is tolerant to Mok1 overproduction. Taken together, the data suggested that Mok1 acts downstream, either directly or indirectly, of Pck1 and Pck2.

Three lines of evidence indicate that Mok1 is a fission yeast α-glucan synthase. The amino acid sequence of Mok1 shows high homology to glycogen or starch synthase, and contains the consensus sequence for UDP-glucose binding. UDP-glucose is used as a substrate for glucan synthase. Second, overexpression of mok1⁺ results in 3.5-fold increase, while mutation of the gene results in a 30% reduction in the amount of α-glucan. Third, no homologous proteins exist in budding yeast, whose cell wall lacks the α-glucan (Cabib et al., 1997). During the preparation of this manuscript, we learned that Hochstenbach et al. (1998) have also isolated mok1⁺, and concluded that it encodes an α-glucan synthase (the gene was designated ags1⁺).

The cell cycle–dependent targeting of Mok1 to the growing tips, the medial region and contractile ring-like structures is consistent with the identification of Mok1 as α-glucan synthase. These regions correspond to the sites at which cell surface structures are being actively remodeled, and both synthesis and degradation of the cell wall glucan take place to ensure tip growth, septum formation and cell separation respectively. We propose that in fission yeast 1,3-β-d-glucan synthase and glucanases also colocalize at these sites. Previous and ongoing work has suggested that Rho1 (Arellano et al., 1997; Nakano et al., 1997) and Pck1 and Pck2 (Sayers, L., K. Nakano, I. Mabuchi, S. Katayama, T. Toda, and P. Parker, unpublished results) are intimately associated with the actin cytoskeleton. It is intriguing that all the proteins that constitute a sequential effector pathway (Rho1-Pck1/2-Mok1) appear to colocalize. The molecular mechanisms underlying the integration of these multiple enzymatic systems at these sites remain to be addressed.

What is the molecular basis of Mok1 association with the plasma membrane? The presence of a putative signal peptide in the NH₂ terminus suggests that Mok1 is first translocated into the endoplasmic reticulum via the secretory pathway. It would be logical to suppose that this transport is followed by translocation into the plasma membrane through the Golgi and vesicle fusion (Kaiser et al., 1997). The dependency of Mok1 localization upon the integrity of the F-actin cytoskeleton is consistent with this idea. Recent analysis shows that polarized delivery of secretory vesicles requires actin cables rather than cortical actin (Pruyne et al., 1998) and Mok1 localization may be also dependent on actin cables. Then why would F-actin be required to maintain Mok1 localization? The total level of Mok1 remained unchanged upon F-actin disruption, suggesting that it is unlikely that protein instability leads to the loss of specific localization of Mok1. It is possible that Mok1 turns over rapidly at the cell tips or medial regions and that, in the absence of F-actin integrity, it accumulates at some stage of the secretory pathway. This situation is analogous to actin, which turns over rapidly and disassembles instantly upon lat A treatment (Ayscough et al., 1997).

pck2 mutant shows the reduced level of cell wall α-glucan and is resistant to toxic overproduction of Mok1. This suggests that Pck2 positively regulates Mok1 activity. In addition, Pck2 is required for Mok1 localization at the specific sites. Mok1 and Pck1/2 do not interact in yeast two-hybrid methods (Katayama, S., and T. Toda, unpublished results). It is possible that Pck2 regulates Mok1 indirectly via some regulators. Alternatively Mok1 may be a direct substrate of Pck2 such that Pck2-dependent phosphorylation is essential for the cellular localization and enzymatic activities of Mok1. The predicted amino acid sequence of Mok1 contains thirty consensus phosphorylation sites for PKC. It might be significant that all these sites are found in the domain that shows a homology to glycogen synthase.

Although we have shown that Mok1 could be a downstream target of Pck1 and Pck2, it may not be the sole molecule by which Pck1 and Pck2 regulate cell morphology. In sharp contrast to the resistance of pck2 mutants to Mok1 overproduction, overproduction of Pck2 is as toxic in the ts mok1 mutants as in wild-type (Toda et al., 1993; Katayama, S., and T. Toda, unpublished results). This suggests that Pck1 and Pck2 regulate cell wall integrity via some effectors in addition to Mok1. Candidates include β-glucan metabolic enzymes, synthases or glucanases. Thus, in cells in which pck2⁺ is overexpressed, the level of β- as well as α-glucan increases (Arellano, M., and P. Pérez, unpublished results). It is highly possible that Pck1 and Pck2 regulate both α-glucan and β-glucan levels to ensure a coordinated regulation of the levels of these two major cell wall components.

The Mok1 protein consists of several putative transmembrane domains, three of which lie in the NH₂-terminal amylase-homologous domains, one in the central region of the protein and twelve at the COOH terminus. Consistent with this, the protein behaved as a membrane-bound protein. COOH-terminal tagging by fusion with DNA sequences encoding either HA-peptide or GFP completely abolishes Mok1 function. We suspect that tagging blocks the correct localization of Mok1 by disrupting the function of the membrane-spanning domains.

In this context, it is of interest that the Mok1 protein derived from a ts mok1-664 strain also failed to localize correctly. As with the other eight ts mok1 mutant alleles examined, the mutation site in mok1-664 is not in the COOH terminus, but rather in the NH₂-terminal amylase-homologous domain. The precise role of this domain is unclear at present, but it may be required not only for reorganization of α-glucan via a transglycosylase activity as suggested (Hochstenbach et al., 1998) but also for the specific localization of Mok1. Multiple domains, therefore, appear to be responsible for the plasma membrane-bound localization of Mok1, and this association appears to be essential for Mok1 function.

Generally genes encoding glucan synthases appear to comprise multi-gene families. In addition to five members of the Mok1 family of α-glucan synthase, the fission yeast genome contains at least three 1,3-β-d-glucan synthase homologues (Katayama, S., and T. Toda, unpublished results). In budding yeast, three 1,3-β-d-glucan synthase-encoding genes and two for 1,6-β-d-glucan synthase-encoding genes have been identified (Orlean, 1997). Furthermore there are three chitin synthase-encoding genes in budding yeast and six in Aspergillus nidulans (Orlean, 1997). The biological significance of five Mok1-related proteins is currently unclear as mok1⁺ is by itself essential for cell viability. Simultaneous triple deletions of the other four homologues (Δmok11Δmok12Δmok14) do not affect viability in normal media. It is possible that these homologues become important in certain growth conditions such as stress and different growth media. It would be of significant to know how these multiple α-glucan synthase homologues are functionally distinct and differentially regulated during mitotic and meiotic cycles.

We thank Drs. Hirofumi Nakano for staurosporine and K-252a, Iain Hagan for FITC-conjugated anti–rabbit IgG secondary antibody, Paul Nurse for strains, John Sgouros for help for bioinformatics and structural predictions, and Mohan K. Balasubramanian and Yasuhisa Adachi for discussion. We are grateful to Dr. Frans Hochstenbach for exchanging data. We thank Drs. Iain Hagan, Paul Nurse, and Peter J. Parker for careful reading of the manuscript and useful suggestions.

cs

cold-sensitive

GFP

green fluorescent protein

HA

hemagglutinin

lat A

latrunculin A

PKC

protein kinase C

STS

staurosporine

ts

temperature-sensitive