Article |
Address correspondence to Aaron Neiman, 332 Life Sciences, Dept. of Biochemistry and Cell Biology, SUNY Stony Brook, Stony Brook, NY 11794-5215. Tel.: (631) 632-1543. Fax: (631) 632-8575. E-mail: Aaron.Neiman{at}sunysb.edu
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Abstract |
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Key Words: sporulation; GIP1; GLC7; septin; spore wall
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Introduction |
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The spore wall consists of four layers. The two inner layers are composed of glucan and mannan, components of the vegetative cell wall (Klis, 1994). The third layer is composed largely of chitosan, a glucosamine polymer, produced by the combined action of the chitin synthetase, Chs3p, and chitin deacetylases (Briza et al., 1988; Pammer et al., 1992; Christodoulidou et al., 1996; Mishra et al., 1997). The outermost layer consists largely of cross-linked dityrosine molecules (Briza et al., 1986). Dityrosine precursors are synthesized in the spore cytosol by the enzymes encoded by DIT1 and DIT2 (Briza et al., 1990, 1994). These genes are induced specifically around the time of prospore membrane closure. The chitosan and dityrosine layers are specific to spores, and provide much of the spore's resistance to environmental stress. Several mutants and genes related to spore wall formation have been isolated and characterized. Among these are genes coding for protein kinases, SPS1, SMK1, CAK1, and MPS1, and a gene encoding a nuclear protein, SWM1, raising the possibility that signal transduction pathways might regulate the spore wall formation (Friesen et al., 1994; Krisak et al., 1994; Wagner et al., 1997; Ufano et al., 1999; Straight et al., 2000).
Septins are a conserved family of proteins characterized by a central core domain and P-loop nucleotide-binding motif (Longtine et al., 1996; Kartmann and Roth, 2001). Most of them also contain a coiled-coil domain that could be involved in their assembly into filaments. In vegetatively growing yeast cells, septins localize to the bud neck and function as a scaffold interacting with a variety of proteins (DeMarini et al., 1997; Longtine et al., 2000). There are seven septin genes in S. cerevisiae, four of which are transcriptionally upregulated during sporulation: SPR3 and SPR28 are sporulation specific, and CDC3 and CDC10 are induced more than tenfold (Kaback and Feldberg, 1985; Chu et al., 1998; De Virgilio et al., 1996; Fares et al., 1996; Primig et al., 2000). During spore development, septin localization is different than that seen in vegetative cells (Fares et al., 1996). No septin localization to the cell periphery is seen. Rather, the septins appear in early meiosis II as four ring-like structures around each SPB. As the prospore membrane extends, septins disappear from the SPB region and display an extended band-like pattern underlying the prospore membrane. After closure of the prospore membrane, the septins become diffusely localized in the spore periphery (Fares et al., 1996). Surprisingly, disruption studies have revealed only modest sporulation defects in septin mutants; spr3 spr28
homozygous diploids and cdc10
diploids sporulate well (De Virgilio et al., 1996; Fares et al., 1996). Although no strong sporulation phenotype has been observed in septin mutants, their expression profile and specific localization suggest that they have some function in sporulation.
Protein phosphatase type 1 (PP1) is a highly conserved phosphoserine-/phosphothreonine-specific protein phosphatase that plays important roles in a variety of cellular processes including cell cycle progression, glycogen metabolism, glucose repression, and sporulation (Mumby and Walter, 1993; Depaoli-Roach et al., 1994; Shenolikar, 1994). In S. cerevisiae, PP1 is encoded by the essential gene GLC7 (Feng et al., 1991). Consistent with the multiple functions of Glc7p, the localization of this protein is dynamic throughout the cell cycle. In addition to a predominant nuclear localization, it is also observed at the SPB, bud neck, and actomyosin ring at distinct times in the cell cycle (Bloecher and Tatchell, 2000). The localization and substrate specificity of Glc7p are thought to be regulated by interaction with different targeting or regulatory subunits. For instance, bud neck localization of Glc7p requires the septin-binding protein Bni4p (unpublished data), and Gac1p is required to localize Glc7p to glycogen particles (Francois et al., 1992; Stuart et al., 1994). As in vegetative cells, there are multiple points at which GLC7 is required during sporulation, including premeiotic DNA synthesis and passage through meiosis I (Ramaswamy et al., 1998; Bailis and Roeder, 2000).
GIP1 encodes a potential developmentally regulated targeting subunit of Glc7p. GIP1 was isolated as a GLC7 interacting gene in a two-hybrid screen (Tu et al., 1996). This interaction was confirmed by coimmunoprecipitation (Tu et al., 1996). GIP1 is a sporulation-specific gene and diploid strains homozygous for a deletion of this gene are blocked in sporulation and fail to induce SPS100, one of the very late sporulation genes (Tu et al., 1996). Additionally, several glc7 mutant alleles defective in sporulation have been isolated, and in most instances sporulation efficiency correlates with the strength of the two-hybrid interaction between the glc7 mutant protein and Gip1p (Baker et al., 1997; Ramaswamy et al., 1998). These previous reports raise the possibility that GIP1 dependent regulation of Glc7p might be important for sporulation.
In this study, we examine the roles of GIP1 and GLC7 in sporulation. Cytological analyses of mutant strains revealed that GIP1 and GLC7 are required for spore wall formation. Both proteins are found to colocalize with septins and are required for septin assembly during sporulation. These results suggest that GIP1 and GLC7, and perhaps the septins, participate in a signal transduction pathway necessary to monitor prospore membrane growth and initiate the synthesis of the spore wall.
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Results |
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The DIT1 gene is required for the synthesis of dityrosine and expression of DIT1 is induced in sporulation before SPS100, but after middle genes such as GIP1 or SPR3 (Briza et al., 1990; Chu et al., 1998; Primig et al., 2000). GIP1 is required for expression of the late gene SPS100 (Tu et al., 1996). This previously described effect of gip1 mutation on SPS100 expression, as well as the lack of dityrosine fluorescence, led us to examine DIT1 expression in the gip1
mutant. A DIT1lacZ fusion gene was introduced into wild-type and gip1
mutant strains. As a control, a lacZ fusion to SPR3 was used. Whereas SPR3lacZ was induced with the same kinetics in the isogenic wild-type and gip1
diploids, no induction of DIT1lacZ was observed when GIP1 was absent (Fig. 3). Thus, the absence of dityrosine staining in the gip1
mutant can be explained by the failure to transcribe DIT1.
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Gip1p colocalizes with Spr28p during spore formation
To examine where in the cell Gip1p functions, the Gip1 protein was localized by immunofluorescence analysis. COOH-terminaltagged versions of Gip1p were nonfunctional (unpublished data). Therefore, targeted integration was used to fuse the hemagglutinin (HA) epitope to the 5' end of the GIP1 gene. To preserve the sporulation-specific expression of GIP1, the fusion gene was under the control of SPO20 promoter. SPO20 is a sporulation specific gene (Neiman, 1998) with a similar expression profile to GIP1 (Chu et al., 1998). When NY509 (in which both copies of the genomic GIP1 genes are replaced with the fusion gene) was sporulated, the efficiency of sporulation was similar to wild-type cells (82% asci). However, only 26% of the asci were tetrads, with the remainder predominantly dyads and triads, suggesting that the fusion gene is not completely functional. When the fusion gene was cloned into a multicopy vector and introduced into the homozygous gip1 deletion strain, NY501, NY501 produced tetrads at a comparable level (>60%) to the wild-type strain. This strain carrying multicopy HA-GIP1 was therefore used for examination of Gip1p localization.
Gip1p localization during sporulation was examined by immunofluorescence analysis using anti-HA antibodies. HAGip1p appears initially in sporulating cells as four rings during meiosis II, and then localizes to the region of extending prospore membranes following the leading edge (Fig. 5). At this stage, parallel bar-like structures were observed. After completion of meiosis II, round structures surrounding the four nuclei were seen. Identical results were obtained when the integrated HAGIP1 was examined, indicating that the localization is not a consequence of overexpression (unpublished data). This localization pattern of Gip1p resembles the reported localization of septins during sporulation. Therefore, we examined whether Gip1p colocalizes with septins. Gip1pHA and Spr28pgreen fluorescent protein (GFP) were expressed in wild type cells and their localization during sporulation was compared. The distribution of the two proteins overlapped extensively with each other (Fig. 6, AD). These results indicate that Gip1p colocalizes with septins during spore formation.
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Glc7p shows a dynamic localization pattern during sporulation and colocalizes with Gip1p in meiosis II
Because GIP1 encodes a potential Glc7p targeting protein, we examined Glc7p localization in sporulating cells. GFPGLC7 was introduced into the wild-type strain and the localization of the fusion protein was observed by GFP fluorescence (Fig. 7). Early in sporulation, cells displayed a predominant nuclear staining with a bright nucleolar dot as observed in vegetative cells (Bloecher and Tatchell, 2000). In early meiosis II, in addition to the nuclear staining, four dots were observed near the nuclear periphery suggesting some Glc7p is localized to the SPB. At later stages of meiosis II, Glc7p clearly displayed a bar-like pattern that resembles those of Spr28p and Gip1p. Finally, in postmeiotic cells, Glc7p was again found predominantly in the spore nucleus. These observations indicate that the localization of Glc7p is dynamic during sporulation, and that Glc7p displays a similar localization pattern to Gip1p and Spr28p during meiosis II.
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Glc7p and Septins are not properly localized in the gip1 mutant
By analogy to other Glc7p targeting proteins, Gip1p might be a targeting subunit that recruits Glc7p to the septins at the prospore membrane. To test this hypothesis, we examined the localization of Glc7p in the gip1 mutant. In this strain, Glc7p did not show a septin-like staining pattern in meiosis II. Instead, it often showed cytosolic staining with four discrete dots, which may indicate SPB localization (Fig. 8 C). Thus, localization of Glc7p to the prospore membrane is dependent on GIP1.
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The Don1p ring is not affected by gip1
DON1 encodes a coiled-coil protein that localizes to a ring-like structure at the leading edge of the prospore membrane, and this structure is distinct from the septin bars that underlie the prospore membrane (Knop and Strasser, 2000). To examine whether the effect of gip1 deletion is specific to septins or is perhaps a more general effect on prospore membraneassociated structures, we examined the localization of Don1p in the gip1 mutant. DON1-GFP was introduced into wild type and gip1
cells, and the localization of the fusion protein was observed using anti-GFP antibodies. In both wild-type and gip1
cells, ring-like structures were observed at the lip of the prospore membranes (Fig. 9). Thus, assembly of the Don1p ring is not affected by gip1 deletion, suggesting that the effects of gip1
are specific to assembly of the septin complex.
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Discussion |
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CHS3 encodes the chitin synthase responsible for the synthesis of the chitosan layer. In contrast to DIT1, the CHS3 gene is not transcriptionally induced during sporulation (Pammer et al., 1992). Therefore, the failure of gip1 mutants to produce chitosan suggests that GIP1 may be required for posttranslational regulation of this enzyme. In vegetative cells, Chs3p activity is controlled in part by regulated recycling through an endosomal compartment (Ziman et al., 1998). It will be of interest to examine the localization of Chs3p in sporulating wild-type and gip1
cells.
Spore wall formation is an ordered process
Previous studies have shown that Calcofluor white transiently stains the chitosan layer of the developing spore (Briza et al., 1990). The loss of Calcoflour white staining in mature asci is due to maturation of the overlying dityrosine layer blocking access of the dye to its ligand (Briza et al., 1988, 1990). We interpret the transient staining of spores with the antiß-glucan antibody in the same way. That is, spores are positive for staining until the overlying chitosan layer forms and obscures access of the antibody to the glucan layer. Surprisingly, staining with FITC-ConA is not lost as spores mature, even though the inner mannan layer should also be obscured as the outer layers form. However, at the same time that ß-glucan staining is lost, the appearance of the FITC-ConA staining changes markedly. This change probably reflects a loss of access to the mannan layer of the inner spore wall, but residual staining of mannoproteins present in the outer prospore membrane or ascal cytoplasm condensed around the spores. Thus, our immunofluorescence assay of spore wall formation suggests a definitive order of spore wall assembly. Mannoproteins are deposited as the prospore membrane forms. After prospore membrane closure, the glucan layer forms, followed by the chitosan layer and finally the dityrosine layer.
Surprisingly, the layers are deposited from innermost to outermost, with respect to the spore cytoplasm. This suggests that precursors to the chitosan and dityrosine layer must be able to pass through the glucan and mannan layers before assembling. Alternatively, some of the spore wall layers may be constructed from precursors introduced from the ascal cytoplasm rather than the spore cytosol. This possibility has been suggested previously based on specific localization of some sporulation-induced transcripts to the ascal cytoplasm (Kurtz and Lindquist, 1986), and is also consistent with the accumulation of darkly staining vesicles in the region of the ascal cytoplasm surrounding the prospores that is seen in gip1 (Fig. 2) and other spore walldefective mutants (Christodoulidou et al., 1999; Wagner et al., 1999).
Relationship of GIP1 to other spore wall synthesis genes.
Our data indicate that not only is spore wall deposition coordinated with respect to prospore membrane closure, but also a temporal order of deposition, glucan before chitosan before dityrosine, must be maintained. Previously reported spore walldefective mutants fall broadly into one of two categories: those such as dit1 or chs3 that remove specific spore wall layers (Briza et al., 1990; Pammer et al., 1992), or those such as smk1 and swm1 in which individual spores within an ascus display heterogeneous wall defects (Krisak et al., 1994; Ufano et al., 1999). The heterogeneous spore wall defects seen in these latter mutants could result from an inability to coordinate the timing of deposition within individual spores.
SMK1 is particularly interesting in this regard. Though deletion of SMK1 causes a heterogeneous spore wall phenotype, point mutations produce more homogeneous arrests (Wagner et al., 1999). In the most severe allele, smk14, this arrest appears similar to gip1. Further, it has been reported that formation of the glucan, chitosan, and dityrosine layers requires increasing levels of Smk1p activity, respectively (Wagner et al., 1999). As this matches the temporal order of deposition of these layers, SMK1 activity might provide a mechanism to control the timing of synthesis of the different layers. Gip1pGlc7p phosphatase could control the initiation of spore wall synthesis, at least in part, by regulating the activation of Smk1p.
Gip1pGlc7p phosphatase regulates septin organization
The Gip1pGlc7p complex colocalizes with the septins during prospore membrane growth and the first evident phenotype of the gip1 mutation is not the spore wall defect, but a failure to organize the septins onto the prospore membrane. However, at times after prospore closure, in a fraction of the gip1
mutant cells the septins can be found in the spore periphery as in wild-type cells. These results indicate a specific requirement for the Gip1pGlc7p phosphatase to organize the septins into structures associated with the growing prospore membrane. Whether this effect is mediated by direct dephosphorylation of the septin proteins or some other modifier of septin organization is not yet known. In vegetative cells, mutation of several different protein kinases has effects on organization of the septin ring (Bouquin et al., 2000; Longtine et al., 2000). For instance, mutation of GIN4 leads to the arrangement of the septins as a series of parallel bars (Longtine et al., 1998a). It has not yet been reported whether any of the septin proteins are direct substrates of any of these kinases. Nonetheless, taken together with our results, it seems likely that phosphorylation and dephosphorylation regulate septin architecture.
The fact that septins are lost from the prospore membrane in the gip1 mutant but that the prospore membrane nonetheless forms, indicates that the septins are not required for membrane assembly or growth. One possibility is that the septins function to keep Gip1pGlc7p associated with the prospore membrane. In this instance, mutation of the septin genes might cause a similar spore wall phenotype to the gip1
mutation. However, no significant sporulation phenotype has been reported in septin mutants (De Virgilio et al., 1996; Fares et al., 1996). Therefore, it remains to be determined whether the spore wall and septin phenotypes of the gip1
mutant are linked, or whether they represent two independent aspects of Gip1pGlc7p function.
In higher cells, as in yeast, septins are found at sites of cytokinesis (Neufeld and Rubin, 1994; Kinoshita et al., 1997). However, in contrast to mitotic yeast cells, the septins of higher cells exhibit dynamic localization and are often found in association with regions of active membrane growth (Fares et al., 1995; Kinoshita et al., 1997; Hsu et al., 1998). Thus, the behavior of septins during sporulation is similar to that in higher cells. Our results indicate that the Gip1pGlc7p complex is required to organize the septins onto the prospore membrane. Given the high degree of conservation of PP1, it will be of interest to learn if septin organization is also regulated by this enzyme in higher cells.
Bailis and Roeder (2000) have reported that GLC7-dependent dephosphorylation of the synaptonemal complex component Red1p is required for progression through the meiotic pachytene checkpoint. In contrast to our findings, this study reported that gip1 mutants arrest in pachytene. Further, GIP1 was required for the localization of Glc7p to meiotic chromosomes and overexpression of GLC7 suppressed the gip1
sporulation defect. From these results it was suggested that the function of Gip1p is to target Glc7p to meiotic chromosomes. We found no evidence of a meiotic arrest in gip1
in two different strain backgrounds, SK1 (Fig. 1) and JC482. Additionally, overexpression of GLC7 does not relieve the spore formation defect of gip1
mutants in our strains (unpublished data), as was observed in the BR strain background (Bailis and Roeder, 2000). The reason for the differing phenotypes of gip1
in different strain backgrounds remains unclear; however, our results clearly demonstrate roles for GIP1 and GLC7 in postmeiotic events.
In sum, our data suggest a model in which Gip1p is induced during sporulation, binds to Glc7p, and the Gip1pGlc7p complex then organizes the septins onto the prospore membrane. The Gip1pGlc7p complex remains associated with the septins during meiosis II, and this places the phosphatase in a position to act as a monitor of prospore membrane growth. Upon closure of the prospore membrane, we propose that Gip1p and Glc7p initiate a signaling cascade that triggers events leading to the synthesis of the spore wall. This includes the induction of DIT1 as well as the activation of chitin and ß-glucan synthases. Further work will be necessary to elucidate the nature of this intracellular signaling pathway.
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Materials and methods |
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Plasmids
To create pFA6a-His3MX6-prSPO20-HA, the promoter sequence of the SPO20 gene was amplified directly from yeast genome using ANO195 (5'-CCTTGAGATCTAAGTCTAGGCGCTTTCAAC-3') and ANO201 (5'-CCTTGTTAATTAAAGACATTATATATCTAAAAATGGC-3'). The PCR product was then digested with PacI and BglII and used to replace GAL1 promoter region of pFA6a-His3MX6-PGAL1-3HA (Longtine et al., 1998b). To clone the prSPO20-HA-GIP1 fusion gene, genomic DNA from NY13 was subjected to PCR with ANO195 and HT6 (5'-CAGTTGCCTACCAATGTTTC-3'). The product was cloned into SmaI sites of pRS424 and pRS426 (Christianson et al., 1992) to create pSB5 and pSB6, respectively. pNC160-GFP-GLC7 (Bloecher and Tatchell, 2000) was used to express GFP-tagged version of Glc7p. pSB7 was created by excising the SPR28GFP fusion gene from YEplac181-SPR28-GFP (De Virgilio et al., 1996) with EcoRI and BamHI and cloning it into pRS314 (Sikorski and Hieter, 1989). To clone DON1GFP into plasmid, genomic DNA prepared from NY17 was subjected to PCR using HT10 (5'-AAACAGATCTATATTACCCTG-3') and HT19 (5'-GAAGAATTCGATATAGCTCTGAACAATTC-3'). The product was digested with EcoRI and BglII and ligated into similarly digested pRS314 to create pSB8. To integrate glc7136 at the ura3 locus, the glc7136 gene was cut out from pSB56 (Baker et al., 1997) with HindIII and SacI, and cloned into pRS306, creating pSB14. pSB14 was linearized with PstI before transformation to target integration to the ura3 locus. The plasmids pHindIII-DIT1-lacZ, a gift of J. Segall (University of Toronto, Toronto, Canada), and pGK16 (Holaway et al., 1987) carrying the DIT1lacZ and SPR3lacZ reporters, respectively, were used to monitor gene induction.
Microscopy
For analysis of meiotic progression, cells were sporulated as described (Neiman, 1998). At intervals, aliquots were removed and fixed with 3.7% formaldehyde at 4°C overnight. Cells were collected and mounted in medium containing DAPI (Molecular Probes). EM was performed as described (Neiman, 1998). Samples were analyzed using a JEOL 1200EX transmission electron microscope at the Stony Brook University Microscopy Imaging Center (Stony Brook, NY).
For immunofluorescence analysis, cells were sporulated and fixed with 3.7% formaldehyde for 2 h, collected, resuspended in SHA buffer (1 M Sorbitol, 0.1 M Hepes, pH 7.5, 5 mM NaN3) and stored at 4°C. Immunofluorescence analysis of proteins was performed as described previously (Gao and Dean, 2000). Monoclonal anti-HA antibodies (12CA5) were provided by N. Dean (SUNY Stony Brook, Stony Brook, NY), and affinity-purified anti-GFP antibodies, a gift of N. Davis (Louisiana State University Medical Center, Shreveport, LA), were used at a 1:10 and 1:200 dilution, respectively. Anti-Spr3p antibodies (Fares et al., 1996) were used at a 1:10 dilution. Alexa Fluor 488 goat antimouse IgG conjugate, Alexa Fluor 488 goat antirabbit IgG conjugate, Alexa Fluor 546 goat antirabbit IgG conjugate, and Texas red goat antirabbit IgG conjugate (Molecular Probes) were used as secondary antibodies at a 1:400 dilution.
Immunofluorescence analysis of spore walls was performed as follows. Fixed sporulating cells were suspended in Zymolyase solution (27 µg/ml Zymolyase and 0.2% ß-mercaptoethanol in SHA buffer) and incubated at 30° for 1 h. Cells were then treated 5 min with SHA buffer containing 0.1% Triton X-100, and 10 min with SHA buffer containing 0.2 mg/ml Calcofluor white (Sigma-Aldrich), washed twice with SHA buffer, and adhered to wells of a polylysine-coated slide. Cells on the slide were treated with methanol for 6 min, followed by acetone for 30 s, and then dried. After blocking with WT buffer (50 mM Hepes, pH 7.5, 150 mM NaCl, 1% Milk, 5 mg/ml bovine serum albumin), cells were incubated with a monoclonal antiß-1,3 glucan antibody (Biosupplies) (Humbel et al., 2001) at a 1:300 dilution in WT buffer overnight. Wells were washed with PBS-BSA (7 mM Na2HPO4, 3 mM NaH2PO4, 130 mM NaCl, 5 mg/ml bovine serum albumin) and incubated with PBS-BSA containing 1:400 diluted Alexa Fluor 546 goat antimouse IgG and 0.1 mg/ml FITC-ConA (Sigma-Aldrich) for 2 h. Finally, wells were washed with PBS-BSA and covered with mounting media. For detection of native GFP fluorescence, cells were fixed with 3.7% formaldehyde for 10 min, washed once with PBS, and mounted with mounting media containing DAPI or processed for immunofluorescence.
Immunofluorescence images were acquired using a Zeiss Axioskop with attached SPOT camera (Diagnostic Instruments) and Adobe Photoshop 5.0, or on a Zeiss Axioplan 2 microscope with a Zeiss Axiocam and Zeiss Axiovision 3.0.6 software. Figures were prepared using Adobe Photoshop 6.0 and Canvas 5.0 (Deneba Software).
ß-Galactosidase and dityrosine assays
Natural fluorescence of the dityrosine molecules was observed as described previously (Briza et al., 1990). DIT1lacZ and SPR3lacZ were separately introduced into both AN120 and NY501. The resulting cells were sporulated and aliquots were collected at each time point and subjected to ß-galactosidase assay as described (Stern et al., 1984).
Online supplemental material
To create three-dimensional projections of Gip1 immunofluorescence, cells carrying HAGIP1 were prepared after 7 h in sporulation medium, and stacks of images were collected with a Bio-Rad Radiance 2000 confocal microscroscope (25mW Arlaser, Nikon TE300, 100 X Plan Apo objective 1.4 NA). Images were collected at 0.2-µm intervals in the Z-axis at 1.5% laser power, pinhole aperture at 2.6, Kalman filtration of four images. Three-dimensional projections were created using Bio-Rad Lasersharp2000 software. Projected images were tilted at angles of -45°45° at 1°-intervals.
To create three-dimensional projects of HA-Gip1p and DAPI fluorescence, stacks of images were collected through an Olympus UPlanFl 100 x 1.3 NA objective with a Roper Coolmax HQ CCD Camera using Scanalytics IPlab Spectrum software. Images were collected at 0.2-µm intervals in the Z-axis, and three-dimensional projections were created using Iplab software. Projected images were tilted at angles of -45°45° at 5°-intervals. Both projections were saved as Apple QuickTime movies, available at http://www.jcb.org/cgi/content/full/jcb.200107008/DC1
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Footnotes |
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A. Bloecher's present address is Division of Basic Sciences, Fred Hutchinson Cancer Research Center, 1100 Fairview Ave. North, Seattle, WA 98109.
* Abbreviations used in this paper: GFP, green fluorescent protein; HA, hemagglutinin; PP1, protein phosphatase type 1; SPB, spindle pole body.
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Acknowledgments |
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This work was supported by National Institutes of Health grants GM62184 to A.M. Neiman and GM47789 to K. Tatchell.
Submitted: 2 July 2001
Revised: 24 September 2001
Accepted: 15 October 2001
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References |
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