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© The Rockefeller University Press,
0021-9525/1998//1217 $5.00
The Journal of Cell Biology, Volume 141, Number 5,
, 1998 1217-1228
Articles |
FH3, A Domain Found in Formins, Targets the Fission Yeast Formin Fus1 to the Projection Tip During Conjugation
School of Biological Sciences, University of Manchester, Manchester M13 9PT, United Kingdom
Formins are involved in diverse aspects of morphogenesis, and share two regions of homology: FH1 and FH2. We describe a new formin homology region, FH3. FH3 is an amino-terminal domain that differs from the Rho binding site identified in Bni1p and p140mDia. The Schizosaccharomyces pombe formin Fus1 is required for conjugation, and is localized to the projection tip in cells of mating pairs. We replaced genomic fus1+ with green fluorescent protein (GFP)- tagged versions that lacked either the FH1, FH2, or FH3 domain. Deletion of any FH domain essentially abolished mating. FH3, but neither FH1 nor FH2, was required for Fus1 localization. An FH3 domain–GFP fusion protein localized to the projection tips of mating pairs. Thus, the FH3 domain alone can direct protein localization. The FH3 domains of both Fus1 and the S. pombe cytokinesis formin Cdc12 were able to localize GFP to the spindle pole body in half of the late G2 cells in a vegetatively growing population. Expression of both FH3-GFP fusions also affected cytokinesis. Overexpression of the spindle pole body component Sad1 altered the distribution of both Sad1 and the FH3-GFP domain. Together these data suggest that proteins at multiple sites can interact with FH3 domains.
Abbreviations used in this paper: DAPI, 4',6-diamidino-2-phenylindole; FH, formin homology; GFP, green fluorescent protein; GST, glutathione S transferase; MSL, minimal sporulation media liquid; SPB, spindle pole body.
EXECUTION of the correct morphogenic program is essential for the growth fidelity of eukaryotes, be it during complex developmental processes such as limb formation, or in linear cell extension in fungi. Polarization of individual cells in response to diverse signals, e.g. internal programs, external factors, or cell-cell contact, can simply be defined as the generation of asymmetric distribution of specific molecules or factors that direct global alterations in the cytoskeleton. In simple eukaryotes such as the yeasts Saccharomyces cerevisiae and Schizosaccharomyces pombe, the actin cytoskeleton plays a key role in establishing and maintaining polarized growth, and in executing cell division (reviewed in Bretscher et al., 1994; Robinow and Hyams, 1989). Structural studies of yeast actin show that there are two types of actin filaments: cytoplasmic cables and cortical dots (Kilmartin and Adams, 1984; Marks and Hyams, 1985). Cortical dots cluster at the growing tip and the cytokinetic ring, while cables extend from the tip towards the main body of the cell. In Saccharomyces cerevisiae, it has been demonstrated that F-actin cortical dots are motile, responding rapidly to external stimuli (Waddle et al., 1996; Doyle and Botstein, 1996).
Several recent observations suggest that the members of the formin protein family are important for actin-related processes during polarization in diverse systems. In vertebrates, the founder member, formin, plays a key role in limb development, and p140mDia is involved in regulating actin polymerization. The Drosophila formins diaphanous and cappuccino execute roles in cytokinesis and polarity establishment. In fungi, formins play key roles in polarized growth and cytokinesis. Budding yeast Bni1p and Bnr1p are required for bud site selection and cytokinesis, while the S. pombe formin Cdc12 and the Aspergillus nidulans FIGA/SEPA are both required for cytokinesis (Marhoul and Adams, 1995; Woychik et al., 1990; Jackson-Grusby et al., 1992; Castrillon and Wasserman, 1994; Emmons et al., 1995; Evangelista et al., 1997; Imamura et al., 1997; Chang et al., 1997; Harris et al., 1997; Watanabe et al., 1997). Two regions of sequence homology—formin homology regions 1 and 2 (FH1 and FH2, respectively)—are found in all formins. FH1 is a proline-rich sequence that has been postulated to interact with profilin (Evangelista et al., 1997; Jansen et al., 1996; Chang et al., 1997; Imamura et al., 1997), and FH2 is defined by a consensus sequence (Emmons et al., 1995).
Bni1p interacts with a number of molecules that are important for polarized growth in budding yeast (reviewed in Chant, 1996; Roemer et al., 1996). Bni1p has been shown to interact directly with Rho1p, Cdc42p, actin, and the two actin-binding proteins profilin and Bud6p (Kohno et al., 1996; Evangelista et al., 1997). Cdc42p is a Rho-family GTPase that is required for establishing cell polarity during the mitotic cell cycle, and for mating (Adams et al., 1990; Simon et al., 1995). Cdc42p localizes to the projection tip during mating in an actin-independent manner (Ayscough et al., 1997). Therefore, the interactions between Cdc42p and Bni1p and between Bni1p and actin suggest that this S. cerevisiae formin homolog serves as a link between the actin cytoskeleton and actin-independent polarization, and thus probably plays a key role in directing markers to the cell tip. Recent studies provide further evidence for such a role for formins, as the other S. cerevisiae formin, Bnr1p, binds to the GTPase Rho4p and the actin-binding protein profilin (Imamura et al., 1997).
The S. pombe formin homolog Fus1 is required for cell fusion during mating (Bresch et al., 1968; Petersen et al., 1995). Upon nitrogen starvation, diffusible mating pheromones induce polarized cell growth in cells of the opposite mating types, P and M, towards one another. Upon contact and agglutination, the cells grow towards one another, and localized cell wall degradation between the partner cells at the projection tips results in cell fusion, enabling karyogamy. After karyogamy, the resulting diploid zygote enters meiosis and sporulates (reviewed in Nielsen and Davey, 1995). Conjugation is blocked after agglutination and formation of the projection tip in the fus1.B20 mutant, and the cell walls separating the mating partners are not degraded (Petersen et al., 1995). Thus, fus1 mutants are blocked at the prezygote stage with a characteristic fus– phenotype (two touching cells attempt to mate, but the cell wall between them remains intact).
The ability to study a formin homolog that is required for an inducible process, and hence is nonessential for normal mitotic growth, has enabled us to ask a number of key questions about the domain structure of formins. We have documented interactions of Fus1 with the actin cytoskeleton, and propose potential functions for different portions of the molecule. We have identified a new formin homology domain, which we call the FH3 domain. We use fusions of the FH3 domains of both Fus1 and Cdc12 to green fluorescent protein (GFP) to confirm the prediction, arising from deletion analyses, that the FH3 domain targets formins to their site of action.
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To do a complete deletion of fus1 from the genome, the Saccharomyces cerevisiae LEU2 gene from pSL19 (kindly provided by Antony Carr, Medical Research Council Cell Mutation Unit, Sussex University, United Kingdom) was cloned into Sph1-digested pDW375 (Petersen et al., 1995), creating pJP98. The HindIII fragment of pDW234 (Petersen et al., 1995) was cloned into HindIII-digested pJP98 (pJP101). pJP101 was used to delete the fus1+ gene by homologous recombination in EG640 (Kjærulff et al., 1994) using LEU2 as a selective marker (EG999). The integration was confirmed by PCR.
Generation of Anti-fus1 Antibodies
The amino-terminal portion of the fus1+ gene (1–1712 bp; Petersen et al., 1995) was cloned by PCR into pGEX2T (Pharmacia Biotech, Inc., Piscataway, NJ) by using a fus1-specific primer (JPP36) that generated a BamHI site at the initiating ATG codon. The glutathione S transferase (GST)- Fus1 fusion protein was incorporated into inclusion bodies upon induction in E. coli BL21, enabling the GST-Fus1 fusion protein to be purified as described in Studier et al. (1990) and used for production of polyclonal antibodies. Antibodies were affinity-purified from sera of two rabbits—1446 and 1447—using nitrocellulose-immobilized GST-Fus1 (Harlow and Lane, 1988). Both sets of antibodies stained immunoblots identically. Antibodies from 1447 were used for immunofluorescence microscopy.
Immunolocalization
For fluorescence microscopy of conjugating cells, cells were grown in MSL to a density of 5 x 106, washed in MSL -N, and starved in MSL -N for 5 h. Actin was localized by fluorescence microscopy using rhodamine-conjugated phalloidin (Marks and Hyams, 1985) after fixation in 3% formaldehyde (Hagan and Hyams, 1988) in PM buffer (Marks and Hyams, 1985). Affinity-purified anti-Cdc3 antibodies (Balasubramanian et al., 1994) were used to visualize Cdc3 by combined formaldehyde and glutaraldehyde fixation (Hagan and Hyams, 1988) in PM buffer (Marks and Hyams, 1985). Fus1, Sad1, and 
tubulin were visualized after fixation in formaldehyde (Hagan and Hyams, 1988) and using affinity-purified anti-Fus1 antibodies (1:20), affinity-purified anti-Sad1 antibodies (1:25; Hagan and Yanagida, 1995), or anti- tubulin (1:5; gift from M.J. Heitz) respectively. To enhance the GFP signal, anti-GFP antibodies were used (kindly provided by K. Sawin, Imperial Cancer Research Fund, London, United Kingdom), and the cells were scraped from MSA plates with a toothpick and smeared onto a coverslip, which was rapidly placed into methanol at –80°C. After 10 min at –80°C, cells were washed off in PEM buffer and processed as described (Hagan and Hyams, 1988). The advantages of fixing cells grown on solid MSA mating medium was that all stages of mating and sporulation were present in one sample. FITC-conjugated anti–rabbit, Cy3-conjugated anti–rabbit, or FITC-conjugated anti–mouse secondary antibodies were all from Sigma Chemical Co. (St. Louis, MO). To observe autonomous fluorescence of Fus1-GFP signals, cells were starved in MSL-N or on MSA, and were dried onto coverslips and inverted onto drops of glycerol containing 1 µg ml–1 diamidinophenylindole (DAPI).1 Calcofluor was used for septum staining at 10 µg ml–1. Color images were produced using a SIT CameraTM and a C2400 processor (Hamamatsu Phototonics, Bridgewater, NJ) for capturing the fluorescent signal into National Institutes of Health image software package on a MacIntosh Quadra 8500/ 100AV. A Pixel pipeline framegrabber was used to integrate 250 images to produce each single channel image, which were then merged in Adobe Photoshop.
Immunoblot Analysis
Crude protein extracts were prepared from 1 x 108 cells that had been starved for nitrogen in MSL-N for 5 h. Cells were broken using glass beads in lysis buffer (50mM Tris-HCl, pH 7.5, 5mM EDTA, 100 mM NaCl, 1% Triton X-100, 0.1 mM PMSF, and 3.4 g/ml aprotinin) in a FASTprepTM FP120 machine (BIO 101 SAVANT; Savant Instrument Inc., Holbrook, NY) at max power for 15 s. Immunoblot analysis was performed according to Meloche et al. (1992) with two exceptions: transfer buffer 1 described in Harlow and Lane (1988) was used for electrotransfer of the proteins to the membranes, and proteins were detected with the Super Signal UltraTM Western blotting detection reagent (Pierce Chemical Co., Rockford, IL).
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Rabbit antibodies were raised against and affinity-purified with a bacterially produced fusion of the amino-terminal portion of Fus1 (residues 1–572) and GST. These antibodies recognized a single 160-kD band on an immunoblot (Fig. 2 A). The 160-kD band was only seen when the Fus1 molecule was overexpressed from a plasmid (pJP54: Petersen et al., 1995), suggesting that Fus1 is present below the detection threshold of these antibodies in wild-type conjugating cells. By immunofluorescence, these antibodies stained a single dot at the very tip of each cell in wild-type pre-zygotes (Fig. 2, C and D). In addition, cytoplasmic dots were detectable before and after fusion, during karyogamy, and during horsetail movement (data not shown; horsetail movement is a stage that follows karyogamy, but precedes the first meiotic division during which the nucleus moves from one end of the cell to the other (Chikashige et al., 1994). The staining with the anti-Fus1 antibodies was not due to cross-reaction with an antigenically related molecule, because these antibodies failed to stain prezygotes that were attempting mating in the complete absence of a fus1+ gene (Fig. 2 B).
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The fluorescent signal emitted by GFP depends upon cyclization of the molecule to generate the active fluorophore (Cubitt et al., 1995). Because cyclization can take some time, it was possible that we were not detecting all of the Fus1 GFP fusion protein. To this end we used anti-GFP antibodies to localize the fusion by indirect immunofluorescence. This approach has the added advantage that the sensitivity of detection should be considerably enhanced. The staining was identical to that seen with the autonomous fluorescence of GFP, with the sole exception that the dots persisted until sporulation (Fig. 3), when they were excluded from the forming spores.
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The FH3 Domain: Fus1 and Other Formins Share a Previously Unidentified Region of Homology
Two regions of homology have been described in the carboxy termini of formins: FH1 and FH2 (formin homology regions 1 and 2; Castrillon and Wasserman, 1994). Motivated by small but consistent homology islands in dot-plot comparisons of Fus1 with other formins, the sequences that lie between the initiator ATG codon and the start of the FH1 domain of all formins were aligned individually to Fus1 using the GCG program Eclustalw (Genetics Computer Group, Madison, WI). Based on the information from the dot plot analyses and these secondary alignments, three potential homology regions were identified. These regions were then finally aligned using the LasergeneTM multiple alignment function (DNASTAR Inc., Madison, WI) with final minor adjustments by eye. From these analyses, it was apparent that there is a third FH region near the amino termini of formins. This region consists of three blocks of similarity in the same relative order in each formin (Fig. 5). The first part of the first FH3 block and the whole of the third block shows the highest level of conservation. The sequence identity, spacing of the sequence blocks, and the similarity at the amino acid level suggest that the fungal sequences form a distinct subgroup with higher similarity than the other members. In line with preceding nomenclature, we call this region FH3. The FH3 domain of formins is the most variant of the homology regions, which probably explains why it has eluded identification to date. The FH3 domain in Bni1p is distinct from the region that interacts with Rho1. The algorithm of Lupas et al. (1991) predicted an additional feature in the Fus1 sequence: a coiled-coil motif between residues 1145 and 1240 (Fig. 6 A). Similar regions of coiled coil have been described in the carboxy terminus of Cdc12 and SEPA (Chang et al., 1997; Harris et al., 1997). Furthermore, we have found a stretch predicted to form a coiled-coil structure in the comparable region of S. cerevisiae Bni1p (data not shown).
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FH1 and Fus1 FH2 to the tips, deletion of FH3 abolished Fus1-GFP localization (Fig. 6 B, Table III). The requirement for FH3 for tip localization of Fus1 is consistent with the stronger effect on the mating efficiency seen in the FH3 deleted strain (Fig. 6 A). Similar to fus1.B20 (Egel, 1973a), the two cells attempting conjugation in strains lacking the FH1 and FH2 domains often had flat ends rather than the rounded points seen in FH3 and fus1 deletants attempting conjugation. This observation may suggest that cell wall expansion, but not breakdown, can occur in the former class of mutants. Alternatively, it may reflect the leakier nature of the block in these mutants. These data strongly suggest that the FH3 domain binds to the cortex, while FH1 and FH2 are required to mediate other essential interactions of Fus1.
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-tubulin (M. Heitz and I.M. Hagan, unpublished data) showed that, in more than half of the late G2 cells with a GFP signal above background, Fus1 FH3 colocalized with the -tubulin at the SPB (Fig. 11 B). In septated binucleate cells and cells in early G2 phase, colocalization was seen in up to a fifth of the cells with a GFP signal. Staining was notably different with anti-Sad1 antibodies. Cells with a strong nucleus-associated GFP dot failed to stain with anti-Sad1 antibodies, while cells with no GFP signal had a normal Sad1 signal (Fig. 11 A). This result raised the intriguing possibility that the overexpressed Fus1 FH3-GFP was blocking anti-Sad1 antibody binding either by directly binding to Sad1, or to a Sad1-containing complex.
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To determine whether the localization to these structures was specific to the FH3 domain of Fus1, or whether other fission yeast FH3 domains would direct GFP to similar structures, the FH3 domain of Cdc12 was fused to GFP and localized in logarithmically growing cells, either with or without a sad1+-bearing plasmid. Overexpression of Cdc12 FH3-GFP from the nmt1+ promoter on AA media without thiamine resulted in similar defects to those seen with expression of Fus1 FH3. In many cases the septum or the nucleus was misplaced from the center of the cell, and cytokinesis was not always completed (Fig 11 F; Table IV). In wild-type cells, the Cdc12 FH3-GFP fusion protein colocalized with tubulin in 46% of late G2 cells having a GFP signal (Fig. 11 C), and in 15% of binucleate cells and cells in early G2. In Sad1-overexpressing cells, the fusion protein went to the nuclear periphery in 25% of cells that had a GFP signal (Fig. 11 E).
These data show that FH3 domains are capable of directing the localization of chimeric fusion proteins to discrete structures within the cell, and are consistent with the presence of FH3 domain interacting protein(s) at multiple locations.
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The Mechanism of Fus1 Tip Localization in S. pombe
Once polarized cell growth has brought two mating partners into contact, the cell walls are degraded at the contact point in a highly localized and regulated fashion to enable the two genomes to fuse after karyogamy. Fus1 is required for this cell wall degradation process (Petersen et al., 1995). We have consistently localized Fus1 to the fusion point by three different approaches: tagging at the amino terminus with the HA-tag, carboxy-terminal tagging with GFP, and immunofluorescence with anti-Fus1 antibodies.
F-actin associates with the cell tip during growth towards the pheromone source that is produced by prospective mating partners (Petersen et al., 1998). This association with the projection tip is before and independent of Fus1 localization to the tip (Fig. 4 A); however, the phenotype of the fus1+ deletion and disruption strains indicated that Fus1 is required to stabilize or mediate F-actin association with the fusion zone after cell–cell contact. F-actin was rarely seen at the tip in cells in which the fus1+ gene had been disrupted by the insertion of the ura4+ gene at the nucleotide corresponding to amino acid residue 270. Instead, F-actin was seen slightly back along the projection tubes of the majority of prezygotes in this strain. When the orientation of the specimen was appropriate, the F-actin dots at this stage were often arranged in a circular fashion reminiscent of a ring. While the lack of conjugation showed that the gene disruption had blocked the production of full-length Fus1 protein, it is highly likely that some portion of the protein was produced, as complete deletion of fus1+ resulted in random cytoplasmic distribution of F-actin dots in the prezygotes after an initial association with the tip. It would therefore seem likely that after cell–cell contact, the role of the actin cytoskeleton is to expand the cell wall at the point of fusion, and that this expansion is regulated by dilation of an F-actin ring between the two cells in the newly formed zygote. In the disruption strain the ring remains intact, associated with the cortex, still expands, and so moves back along the projection tip.
Multiple Commitment Points During Conjugation
The two different roles played by the actin cytoskeleton during mating—polarized cell growth and coordinated cell wall degradation—underline the importance of correctly coordinating fusion events. Cell–cell contact is required before Fus1 is localized to the projection tip. Thus, two decision points are defined: commitment to polarized growth and the attachment to a mating partner, which stimulates the recruitment of Fus1 to the tip and chromatin rearrangements within the nucleus (Chikashige et al., 1997). Recruitment of Fus1 may be involved in a feedback loop to signal the cessation of tip growth, as cells that harbor the fus1.B20 mutant continue to elongate when fusion is defective (Petersen et al., 1995). Alternatively, this feedback loop may be activated by cytoplasmic mixing after cell fusion. In this case, fus1 mutants continue to elongate because the cytoplasms do not mix.
How Does the Cell Sense the Binding of a Partner?
It is possible that cell–cell contact is registered by localized pheromone gradients through the established signal transduction pathway (reviewed by Nielsen and Davey, 1995). If a pheromone gradient is important, the high levels required to generate a signal have to be localized at the point of cell–cell contact. In this case, adding a pheromone to a heterothallic strain fails to induce Fus1 localization and heterochromatin rearrangements (Chikashiga et al., 1994) because the same level of pheromone is registered all around the cell (Fig. 2). Alternatively, there may be a distinct receptor-mediated signal transduction pathway that is activated after cell–cell contact-specific agglutination at the projection tips. A potential candidate for such a pathway may involve the MAP kinase Spm1 (Zaitsevskaya-Carter and Cooper, 1997). Disruption of spm1 interferes with cytokinesis and morphogenesis, and greatly perturbs conjugation.
FH3-mediated Fus1 Binding to the Tip
Two formin homology regions have been described previously (FH1 and FH2), and a domain that can interact with members of the rho family of small GTP binding proteins has been described in two of the formins: Bni1p and p140mDia. We describe an additional amino-terminal tripartite formin homology region that we call FH3 (Fig. 5). The similarity between FH3 domains is strongest in comparisons of different fungal members. Different parts of the sequence are conserved to varying degrees in the metazoan family members. The FH3, but not the FH1 or FH2 domain of Fus1, was required to direct Fus1 localization to the tip. Since deletion of all three domains drastically reduces mating efficiency, FH1 and FH2 are presumably required for some other aspects of Fus1 function.
The ability of the FH3 domain to direct a GFP fusion protein to the projection tip and compete with native Fus1 at this site, thus generating a fus– phenotype with cells accumulating as prezygote pairs, suggests that it alone is sufficient to locate Fus1 to the tip. However, other parts of the protein are likely to stabilize or enhance the binding to the tip (see below). The Fus1 FH3 domain can also compete with additional proteins required for other events in the life cycle when Fus1 is not normally present. Thus, Fus1 FH3 targets the fusion protein to the SPB and the equatorial ring. This targeting to new locations leads to defects in nuclear positioning and cytokinesis (Fig. 10). Considering the similarity between this localization pattern and that reported for Cdc12 (Chang et al., 1997) and the concomitant cytokinesis defects, it is possible that the Fus1 FH3 domain is affecting cytokinesis by binding to the normal partner of the FH3 domain of Cdc12. Consistently, we found that the FH3 domain of Cdc12 behaved like that of Fus1; it colocalized with the SPB marker tubulin and followed Sad1 to the nuclear periphery when it was driven there by moderate overexpression (Hagan and Yanagida, 1995). The block to anti-Sad1 antibody binding to the Sad1 protein that results from expression of the Fus1 FH3-GFP fusion suggests that epitope masking of Sad1 is occurring for some reason. Whether or not the epitope masking is due to a direct interaction with Sad1 is not clear, but this finer point does not detract from the fact that the FH3 domain is never seen around the nuclear periphery in wild-type cells, but is in strains overexpressing Sad1. Thus, the FH3 domain is capable of directing location to more than one site within the cell.
The Fus1 FH1 Domain
It has been suggested that the proline-rich formin homology domain, FH1, is responsible for the association of formins with the actin-binding protein profilin in vivo. This suggestion was stimulated by the demonstration that profilin binds to polyproline regions in in vitro assays (Tanaka and Shibata, 1985). Several reports describe an in vitro interaction between profilin and the proline-rich FH1 domain of formins (Chang et al., 1997; Evangelista et al., 1997; Imamura et al., 1997), and some use two hybrid and synthetic lethality data to argue for the same interaction in vivo. While the fission yeast profilin homolog Cdc3 is absolutely required for conjugation (Petersen et al., 1998), it localizes to the tip independently of the FH1 domain of Fus1. We have also determined that Fus1-GFP localizes to the tip in a cdc3.124 mutant at a temperature that is restrictive for both conjugation and mitotic growth, suggesting that the converse may be true, and that Fus1 may localize independently of Cdc3 function. One potential problem with this conclusion, however, is that we cannot rule out the possibility that a Fus1-interacting function of Cdc3 is unaffected at this restrictive temperature, and that it is some other function of this multifunctional protein that is temperature-sensitive in cdc3.124. However, it is clear that Fus1 localizes to the tip when its FH1 domain has been completely removed. Finally, attempts to detect any interaction between Fus1 and Cdc3 by immunoprecipitation have failed, and we have detected only weak interactions between the FH1 domain of Fus1 and Cdc3 in the budding yeast two-hybrid assay (as would be expected for a proline-rich sequence; J. Petersen, unpublished data). It is perhaps important in assessing this body of evidence that argues against an interaction between the Fus1 FH1 domain and Cdc3 to note that the FH1 domains of the other formins contain far more prolines than does the FH1 domain of Fus1, and that profilin binding requires runs of 8–10 prolines (Sohn and Goldschmidt-Clermont, 1994). If the FH1 domain of Fus1 does not bind to profilin, it may confer the ability to interact with SH3 or WW domains in target molecules (Sudol, 1996). Profilin may therefore execute its role in conjugation independently of the Fus1 FH1 domain.
Multiprotein Complex at the Tip
Formins interact with molecules such as actin and actin-binding proteins (Imamura et al., 1997; Evangalista et al., 1997), suggesting that further work may well identify large complexes containing Fus1. Two pieces of data suggest that Fus1 may interact with multiple partners. The first is that while deletion of the FH1 or FH2 domains severely reduces conjugation efficiency, it does not reduce it to the zero level seen with deletion of either FH3 or the entire molecule. This fact suggests that although the functions of FH1 and FH2 are virtually essential, the molecules with which they interact must be able to bind to at least one other partner or other regions of Fus1. Thus, in the absence of interaction of the domain with Fus1, some activity of the overall complex is achieved. Clearly, if the complex were unable to localize at all, as is the case for Fus1 which lacks the FH3 domain, its function would be completely lost, as we have seen. Precedents for such a situation include the interaction between the budding yeast SPB components Cdc31p and Kar1p. Both are essential. Kar1p is required to localize Cdc31p to the SPB, but in extragenic suppressors the requirement for Kar1p binding is bypassed, suggesting that the complex contains more than just Kar1p and Cdc31p (Biggins and Rose, 1984; Vallen et al., 1994). Thus, extragenic suppressors of FH1 or FH2 deletions could be expected to identify other components of the complex. Complexes are also suggested by the large aggregates of Fus1 seen before and after conjugation.
The identification of the FH3 domain and its ability to localize in a similar way to, and compete with, the full-length molecule suggest that this is a functionally relevant motif. The ability of Cdc12 FH3 domain to target GFP to discrete locations suggests that the FH3 motif in other formins will similarly target them to discrete locations. We have shown that the ability to manipulate formins that are absolutely required for an inducible event offers a key to unravelling the functional complexities of this expanding family of complex molecules.
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Acknowledgments |
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-tubulin antibodies, Tony Carr and Jim Haselhof for plasmids, and to Viesturs Simanis for stimulating discussions. This work was supported by a European Molecular Biology Organization short-term fellowship to J. Petersen, as well as by grants from the Danish Natural Science Research Council (to R. Egel), the Novo-Nordisk Foundation (to O. Nielsen), and the Cancer Research Campaign (I. Hagan). Imaging technology in Manchester was supported by the Cancer Research Campaign and a Welcome Trust Equipment grant.
Submitted: 21 November 1997
Revised: 3 April 1998
Address all correspondence to Janni Petersen, Department of Genetics, Institute of Molecular Biology, Østerfarimagsgade 2A, University of Copenhagen, DK-1353 Copenhagen K, Denmark. Tel: 45-35-32-21-03; Fax: 45-35-32-21-13; E-mail: jan0302{at}biobase.dk
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160 kD in cells overexpressing Fus1 (A, lane 3). No signal was seen in lanes containing extracts from fus1-deleted strains or wild-type cells (A, lanes 1 and 2); the protein extract was made from conjugation cells. Localization of Fus1 with these antibodies in wild-type cells showed an association to the tips of conjugating cell pairs. C and D illustrate Fus1 localization (top) and DAPI/ phase contrast (bottom) of the same cells. (E) Autonomous fluorescence of an integrated version of Fus1 bearing a carboxy-terminal GFP tag showed an identical localization pattern to that obtained with the antibodies; Fus1-GFP was associated with the tip at the point of contact of the conjugating pair, and after fusion cytoplasmic dots were seen. This localization of Fus1 to the tip was identical to that which we have reported before for an amino-terminal HA epitope-tagged molecule (
