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Address correspondence to Fred Chang, Department of Microbiology, Columbia University College of Physicians and Surgeons, 701 W. 168th St., New York, NY 10032. Tel.: (212) 305-0252. Fax: (212) 305-1468. email: fc99{at}columbia.edu
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Abstract |
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Key Words: actin; microtubules; cell polarity; fission yeast; formin
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Introduction |
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The fission yeast Schizosaccharomyces pombe serves as a model cell in which to study microtubuleactin interactions, as microtubules and actin regulate cell polarity and cell shape in these cells (Chang, 2001). Fission yeast are simple rod-shaped cells that grow at cell tips in a regulated manner. After cell division, they initially grow only at the previous cell tips (the old ends), and then later in G2 phase, initiate cell growth at the new ends (Mitchison and Nurse, 1985). Microtubules are organized in linear arrays of anti-parallel bundles so that microtubule plus ends repeatedly touch and shrink at both cell tips (Brunner and Nurse, 2000; Drummond and Cross, 2000; Tran et al., 2001). Disruption of these microtubules causes aberrant cell shapes such as bent or branched (T-shaped) cells (Toda et al., 1983; Sawin and Nurse, 1998).
The analysis of the kelch repeat protein tea1p has begun to elucidate how microtubules may regulate fission yeast cell polarity. tea1 mutants exhibit abnormal cell shapes much like cells with disrupted microtubules, and grow only from one cell tip (Snell and Nurse, 1994; Verde et al., 1995; Mata and Nurse, 1997). Tea1p is located on the growing plus ends of microtubules and in dots at the cell tip (Mata and Nurse, 1997; Behrens and Nurse, 2002). The localization of tea1p at the microtubule plus end is dependent on the CLIP-170 tip1p and the Kip2-like kinesin tea2p (Browning et al., 2000; Brunner and Nurse, 2000). Indirect observations and time-lapse images of cells with abnormal tea1p dynamics suggest that microtubule plus ends deliver tea1p to the cell tip; when the microtubule shrinks away, tea1p may be released from the microtubule and "docks" at the cell cortex (Mata and Nurse, 1997; Behrens and Nurse, 2002; unpublished data). For instance, in a mod5 mutant, tea1p localizes on the microtubule but does not dock at the cell tip (Snaith and Sawin, 2003). However, direct observation of tea1p deposition at the cell tips in wild-type cells has not been definitively shown. As tea1p has strong effects on cell polarity but only subtle effects on microtubule dynamics, tea1p may directly regulate cell polarity and possible actin distribution at the cell tip.
Formins are a conserved family of proteins with roles in cell polarization and cytokinesis (Wallar and Alberts, 2003). Recent reports show that they directly nucleate actin filament assembly in vitro and regulate actin filament elongation while bound to the growing barbed end of actin filaments (Evangelista et al., 2002; Pruyne et al., 2002; Sagot et al., 2002b; Li and Higgs, 2003; Zigmond et al., 2003; Moseley et al., 2004). Formins are responsible for the formation of diverse actin structures including actin cables, contractile rings, filopodia, endosome actin tails, and adherens junctions (Evangelista et al., 2002; Sagot et al., 2002a; Gasman et al., 2003; Peng et al., 2003; Kobielak et al., 2004). The S. pombe formin for3p, which is located at cell tips, is required specifically for assembly of actin cables in interphase cells (Feierbach and Chang, 2001; Nakano et al., 2002). These actin cables may contribute to polarized growth by functioning as tracks to guide polarized targeting of secretory vesicles to the growing cell tip (Schott et al., 1999). One likely regulator of for3p is the actin-binding protein bud6p/aip3p (Glynn et al., 2001; Jin and Amberg, 2001). Its budding yeast homologue (Bud6p/Aip3p) is an actin monomerbinding protein that interacts with the formins Bni1p and Bnr1p (Evangelista et al., 1997; Kikyo et al., 1999; Jin and Amberg, 2000, 2001) and acts as a cofactor with profilin to increase actin assembly by Bni1p in vitro (Moseley et al., 2004). S. pombe mutants lacking tea1p, for3p, or bud6p have varying defects in cell shape and cell polarity establishment at one or both cell tips (Snell and Nurse, 1994; Feierbach and Chang, 2001; Glynn et al., 2001). We have shown previously that S. pombe bud6p interacts with tea1p (Glynn et al., 2001).
Here, we address two questions about tea1p: is tea1p directly deposited by microtubules at the cell tip, and how does tea1p regulate cell polarity and actin cable distribution? We observed directly that tea1p is deposited by plus ends of microtubules. Biochemical analyses show that tea1p associates with for3p, bud6p, and the CLIP-170 tip1p in distinct high molecular weight complexes. Localization experiments suggest that tea1p acts to regulate the localization of formin and actin cables at specific cell tips. These experiments contribute key insights into the molecular mechanisms of tea1p trafficking and function and suggest a model for how microtubule plus ends regulate actin assembly through regulation of a formin.
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Results |
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Tea1p associates with for3p and bud6p in high molecular weight complexes
After being deposited at the cell tips, tea1p may regulate cell polarity by interacting with other polarity factors. We tested whether tea1p associates with the cell polarity factors bud6p and for3p (formin) using four different approaches. Epitope-tagged tea1p, for3p, and bud6p protein fusions were expressed from the endogenous promoters at their chromosomal locus. These tagged proteins were functional, as they supported normal cell polarity in the absence of untagged protein.
First, using these epitope-tagged strains, we tested for coimmunoprecipitation from yeast extracts. For3p coimmunoprecipitated with bud6p-HA (Fig. 2 a). For3p and tea1p also coimmunoprecipitated (Fig. 2, b and c). A coimmunoprecipitation association between tea1p and bud6p was shown previously (Glynn et al., 2001).
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Fourth, we examined these complexes in yeast extracts derived from different mutant backgrounds (Fig. 3). Using sucrose gradients, we found that in for3 extracts, bud6p and tea1p were still in complexes, but they sedimented with smaller S values (Fig. 3 a); bud6p and tea1p migrated together 23 fractions smaller in complex D (the 75S complex), and possibly one fraction smaller in complex C (the 45S complex). In tea1
extracts, for3p in complex D similarly shifted two fractions smaller, although there was no detectable shift in complex C (Fig. 3 a). In bud6
extracts, tea1p in complex D shifted together to a smaller complex by 12 fractions, and may shift one fraction smaller in complex C. Interestingly, although for3p co-migrated with tea1p in the smaller complex D, it was absent or greatly reduced from complex C (Fig. 3 b). This finding suggests that bud6p may be required to attach for3p to complex C. Because of the large size of these complexes, a lack of changes in sedimentation behavior in mutant extracts does not rule out that these proteins interact in that particular complex. However, the observed changes provide further evidence, especially in complex D, that these proteins reside together in common protein complexes.
It is not clear whether for3p and bud6p actually interact in the 20S complex B. In contrast to the shifts in the large complexes, there were little or no effects on the migration of complex B in either for3 or bud6
extracts. In addition, the lower molecular weight form of bud6p that co-migrated with for3p at 20S was not the dominant form of bud6p that coimmunoprecipitated with for3p. However, we found that the 75S complexes disassembled over time into 20S complexes containing for3p and bud6p, with bud6p primarily in the higher molecular weight forms (unpublished data). Together, the coimmunoprecipitation, two-hybrid, and sucrose gradient data provide strong evidence that tea1p, bud6p, and for3p physically associate.
Tea1p and bud6p regulate for3p localization
To determine how tea1p and bud6p may affect for3p in vivo, we tested first whether tea1p and bud6p regulate for3p localization. These three gene products localize during interphase to multiple dots at both cell tips, even at nongrowing (preNETO) cell tips. These proteins also reside at the cell division site during cell division. During mitosis, tea1p persists at the cell tips, whereas for3p and bud6p leave the cell tip and accumulate more at the cell division plane. Thus, in just-divided cells, tea1p localization precedes for3p and bud6p at the old cell tips. At cell tips in interphase cells, a subset of for3p dots colocalized with tea1p dots, and all the for3p dots colocalized precisely with bud6p dots (Fig. 4, a and b). In addition, tea1p (but not for3p or bud6p) localizes to the plus ends of growing cytoplasmic microtubules (Mata and Nurse, 1997; Feierbach and Chang, 2001; Glynn et al., 2001; Behrens and Nurse, 2002; Nakano et al., 2002).
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In contrast, tea1p and bud6p localization patterns were largely independent of for3p. In for3 cells, tea1p-YFP was localized normally at cell tips in the large majority of cells (Fig. 4 g, right; 92% cells, n = 100). Bud6p-CFP localized normally in most for3
cells (65% cells, n = 35 cells). Bud6p-CFP was reduced from one or both ends in other cells (Fig. 4 h, right), showing some interdependency between bud6p and for3p. In contrast to the cell tip localization patterns, the localization of for3p, tea1p, and bud6p at the cell division site were all independent of each other (unpublished data). Thus, these data suggest a dependency pathway in which at certain cell tips, tea1p acts to position bud6p, and bud6p positions for3p.
Tea1p and bud6p regulate actin cable distribution
As for3p is thought to nucleate actin cable assembly, we then examined if tea1p and bud6p influence the formation or distribution of actin cables. We fixed and stained cells for F-actin using Alexa Fluor® phalloidin and imaged them using confocal microscopy. In bipolar wild-type cells, actin was localized in patches that were concentrated at two growing ends, and in a network of actin cables that traversed the long axis of the cell (Fig. 5 a, first panel; 95% cells, n = 35). bud6 mutants exhibited normal concentration of actin patches at the growing end, but only faint actin cables. We measured the fluorescence intensity of Alexa Fluor® phalloidinstained actin cables (normalized to the intensity of actin patches, which were similar in the bud6
and wild-type strains) and found that individual actin cables in bud6
mutants were
50% less bright than those in wild-type cells (Fig. 5 b). These findings are consistent with the fact that for3p is partially delocalized in a bud6
mutant and with a recent finding that budding yeast Bud6p stimulates formin activity in vitro (Moseley et al., 2004). As actin cables consist of bundles of actin filaments, there may be fewer actin filaments present in bud6
actin cables.
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Tea1p, bud6p, and for3p contribute to general cell polarity
To determine if tea1p and bud6p function solely to regulate for3p or whether they have additional functions, we analyzed double- and triple-mutant combinations (Fig. 6). If these proteins function in a strict linear pathway, we predicted that multiple mutants would have similar phenotypes as single mutants. However, for3tea1
cells grew slower (Fig. 6 a) and had more aberrant morphology than either single mutant, as most cells were ovoid in shape (Fig. 6, b and c). These double mutants did not form T-shaped cells seen in tea1
cells (Fig. 6 b, compare tea1
panel to for3
tea1
panel), suggesting that for3p is required for cell growth from the sides of cells, as is bud6p (Jin and Amberg, 2001). In addition, 30% of these for3
tea1
cells exhibited a different morphology that was not apparent in either single mutant: long cells with multiple septa (Fig. 6 b; for3
tea1
, right cell). This phenotype is indicative of a cellcell separation defect similar to those of septin mutants (Longtine et al., 1996) or exocyst mutants (Wang et al., 2002), and suggests a defect in initiating cell growth at the new ends (Fig. 6 b). for3
bud6
double mutants were similar to for3
single mutants, but were slightly rounder (Fig. 6, b and c), whereas bud6
tea1
mutants exhibited no synthetic effects and resembled tea1
mutants (Glynn et al., 2001). Strikingly, the for3
bud6
tea1
triple-mutant cells were extremely slow growing and formed round or oval cell shapes (Fig. 6). The severe polarization defect in the triple mutant indicates that these proteins do not simply regulate the transitions from monopolar to bipolar growth as previously thought, but work together to organize general polarized growth. These synthetic genetic interactions demonstrate that these genes do not operate in a linear pathway, but may function in parallel pathways or in the context of a common complex (the "polarisome;" see Discussion) that is required for polarized cell growth.
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Previous papers have shown that tip1p (but not for3p or bud6p) colocalizes with tea1p on the microtubule plus end (Brunner and Nurse, 2000; Niccoli and Nurse, 2002). Tip1p is required for proper localization of tea1p to the microtubule plus ends and cell tips, and tip1 mutants exhibit similar morphological defects as tea1
cells (Brunner and Nurse, 2000). We examined the abnormal distribution of tea1p in tip1
cells using time-lapse microscopy. Dual imaging of tea1p and microtubules showed that tea1p dots were in multiple dots or dashes all along the microtubule, with increased concentration around medial regions of microtubule overlap (Video 4, available at http://www.jcb.org/cgi/content/full/jcb.200403090/DC1). Many tea1p dots on the microtubules were not motile. Other tea1p dots moved in either a minus end or plus enddirected manner and were present on growing or shrinking microtubule ends and sites along the microtubule bundles. The large number of tea1p dots and their occasional distribution in lines suggested that they are localized not only at microtubule plus ends, but also along the length of microtubules. A previous report showed that in a tea1
mutant, tip1p localizes normally at the microtubule plus end, but does not accumulate at the cell end (Brunner and Nurse, 2000). Thus, the interaction between tea1p and tip1p may help tip1p attach tea1p to the plus end of the microtubule, and allow tea1p to retain tip1p at the cell end.
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Discussion |
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Tea1p may somehow regulate the localization of for3p and bud6p at cell tips. In tea1 mutants, for3p, bud6p, and actin cables are concentrated at only one cell tip, indicating that tea1p is needed for for3p and bud6p localization to the second cell tip. The physical interactions among these proteins suggest that tea1p may directly recruit or stabilize for3p and bud6p to the second cell tip. However, it is also possible that tea1p affects these factors in a more indirect manner. Because of photobleaching problems for for3p-YFP, we have been unable use time-lapse microscopy to observe if tea1p dots stimulate the formation of for3p dots at cell tips. We should also stress that for3p and bud6p still localize to one of the cell tips and to the septum in the absence of tea1p, suggesting that additional mechanisms (such as cortical landmarks) contribute to for3p localization and activity.
As for3p, bud6p, and tea1p function in actin regulation and cell polarity, we speculate that large complexes represent polarisome complexes. An analogous polarisome complex in budding yeast has been proposed (Sheu et al., 1998; Pruyne and Bretscher, 2000), but large complexes containing formins have not yet been directly demonstrated in any other organism. The round cell phenotype of the for3 bud6
tea1
triple mutant may reflect the phenotype of a nonfunctional polarisome complex. Biochemical analyses of the S. pombe complexes suggest that complexes are still mostly intact in single mutants, but may have defects in their regulation or activity. The synthetic genetic effects indicate that these polarisome factors collectively not only regulate actin cable formation, but additional aspects of polarized cell growth, such as secretion. Future identification of additional polarisome complex components will provide further molecular insights into polarisome functions.
CLIP-170s and microtubule plus ends
CLIP-170 is a conserved microtubule plus endbinding protein that regulates microtubule stability (Carvalho et al., 2003). In budding yeast and mammalian cells, CLIP-170s have been found to attach the dynactin complex to the microtubule plus ends (Goodson et al., 2003; Sheeman et al., 2003). However, dynein appears to have only meiotic-specific functions in fission yeast (Yamamoto et al., 1999). In fission yeast, the CLIP-170 tip1p functions to stabilize microtubules when they contact the sides of cells (Brunner and Nurse, 2000). In addition, tip1p also functions to attach tea1p to the microtubule plus end. In the absence of tip1p, it is interesting that tea1p still associates with microtubules and moves along microtubules in both minus end and plus end directions, suggesting that tea1p also associates with other microtubule-associated proteins, and possibly one or more motor proteins. One candidate motor protein is tea2p, a Kip2-like kinesin that appears to move factors such as tea1p and CLIP-170 to the microtubule plus end (Browning et al., 2000, 2003). Interactions among other microtubule plus endbinding proteins including tea2p and the EB1 homologue mal3p have been found (Browning et al., 2003). Further analyses will test whether the tea1ptip1p associate complex represents a microtubule plus end complex.
Interactions between microtubules and actin
It is becoming increasingly clear that many cytoskeletal processes, including cell polarization, organelle transport, and cytokinesis, depend on interactions between the microtubule and actin cytoskeletons. Our analyses suggest a simple model for how microtubules may instruct the actin cytoskeleton. Conversely, in some cell types the actin cytoskeleton and/or formins may also function to move or stabilize microtubules (Palazzo et al., 2001; Gundersen, 2002). For instance, in cytokinesis, microtubule plus ends are stabilized at the cortex specifically in the region of the future cell division site (Canman et al., 2003). However, there is little evidence that actin or for3p directly control microtubule dynamics in interphase fission yeast cells (Feierbach and Chang, 2001; unpublished data). Mammalian CLIP-170 has been implicated in microtubuleactin interactions, for instance through interaction with IQGAP protein (Fukata et al., 2002). The animal equivalents to tea1p and bud6p are not yet clear. The formin-interacting regions of yeast Bud6 proteins are similar to a region in Rho kinase (Glynn et al., 2001; Moseley et al., 2004), but the functional significance of this Rho kinase region has not been investigated in animal cells. Of kelch repeat proteins similar to tea1p (Adams et al., 2000), budding yeast Kel1 and Kel2 are required for polarized cell growth (Philips and Herskowitz, 1998), mammalian Keap protein is localized on focal adhesions and zipper junctions (Velichkova et al., 2002), p97 is a Rab effector (Diaz et al., 1997), and Drosophila kelch is a component of ring canals (Robinson and Cooley, 1997). Future work will be needed to test how these conserved sets of proteins may function together in the integration of the microtubule and actin cytoskeletal networks in different cell types.
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Materials and methods |
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Microscopy
Wide-field and spinning disc confocal microscopy were performed as described previously (Pelham and Chang, 2001). For live-cell imaging, 12 µm cell slurry in media was placed under the coverslip, with no agarose pad or sealant. Actin staining was performed as described previously (Pelham and Chang, 2001) using Alexa Fluor® 488 phalloidin (Molecular Probes, Inc.). Acquisition, three-dimensional reconstruction, and restrained iterative deconvolution were performed using Openlab software (Improvision).
Immunoprecipitations and sucrose gradients
For extract preparation, yeast cells were grown in Edinburgh minimal media with appropriate amino acid supplements, harvested, washed, and resuspended in an equal volume of CXS buffer (50 mM Hepes, pH 7.5, 20 mM KCl, 1 mM MgCl2, 2 mM EDTA, and protease inhibitor cocktail). The cell slurry was quick-frozen as pellets in liquid nitrogen and ground while frozen into a powder using a mortar and pestle. The resulting powder was thawed and protease inhibitors and 0.1% Triton X-100 were added. The difference in Triton X-100 concentration accounts for the difference in 1220S complex mobilities seen in Glynn et al. (2001), which used 1% Triton X-100. Velocity sucrose gradients were performed as described previously (Glynn et al., 2001). For Western blotting, we used monoclonal anti-HA antibody HA.11 (Covance), polyclonal anti-myc antibody A-14 (Santa Cruz Biotechnology, Inc.), polyclonal rabbit anti-tea1p antibody (a gift from P. Nurse, Imperial Cancer Research Fund, London, UK), and rabbit polyclonal anti-tip1p antibody (a gift from D. Brunner, EMBL, Heidelberg, Germany). For estimation of complex size, we probed sucrose gradient fractions for ribosomal subunits using anti-ribosomal antibodies (gifts from J. Warner, Einstein College of Medicine, Bronx, NY and L. Pon, Columbia University, New York, NY) and ran gel filtration markers (Bio-Rad Laboratories) in parallel sucrose gradients.
For immunoprecipitations, 50 µl soluble yeast extract was added to 25 µl protein ASepharose bead slurry (Sigma-Aldrich; Fig. 2 a), 25 µl Dynal mouse antirabbit magnetic bead slurry (Dynal Corp.; Fig. 2 c), or 25 µl Dynal sheep antimouse magnetic bead slurry (Dynal Corp.; Fig. 2, b and c). The protein ASepharose beads were washed twice in 1x PBS, pH 7.4, and preabsorbed for 2 h with either monoclonal anti-HA antibody HA.11 (Covance) or monoclonal anti-myc antibody 9E10 (Santa Cruz Biotechnology, Inc.). Dynal mouse antirabbit beads were washed twice in 1x PBS, pH 7.4, and complexed with 2 µg of 10 mg/ml IgG antibody (Sigma-Aldrich). Dynal sheep antimouse beads were washed twice in 1x PBS, pH 7.4, and complexed with 2 µg of 5 mg/ml anti-HA antibody. After a 90-min incubation, all reactions were washed three times with CXS buffer (Fig. 2, a and b) or CXS buffer with 150 or 250 mM NaCl (Fig. 2 c). Dynal beads were collected using a Magnetic Particle Concentrator (Dynal Corp.). Immunoprecipitations were then boiled in 45 µl sample buffer and loaded onto SDS-PAGE gels.
Two-hybrid protein interaction analyses
A fragment of the bud6+ gene (nt 33614801) was inserted into pGAD GH vector (CLONTECH Laboratories, Inc.). The for3 two-hybrid constructs were gifts from K. Nakano and I. Mabuchi (University of Tokyo, Tokyo, Japan; Nakano et al., 2002). Two-hybrid constructs were transformed into TAT7 (Mat a his3-200 leu2-3,112 trp1-
90 ade2-101 gal80 lys::LYS2::lexAopHIS3 ura3::URA3::lexAop-lacZ). Lac Z expression and/or histidine auxotrophy were scored.
Online supplemental material
Time-lapse movies of tea1p dynamics in wild-type and rsp1-1 mutants are available at http://www.jcb.org/cgi/content/full/jcb.200403090/DC1.
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Acknowledgments |
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This research was supported by National Institutes of Health grant GM R01-GM56836, a research project grant from the American Cancer Society, a Nikon summer fellowship at the Marine Biological Laboratory (Woods Hole, MA) to F. Chang, and a National Institutes of Health postdoctoral fellowship (GM20283) to B. Feierbach.
Submitted: 16 March 2004
Accepted: 28 April 2004
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References |
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Adams, J., R. Kelso, and L. Cooley. 2000. The kelch repeat superfamily of proteins: propellers of cell function. Trends Cell Biol. 10:1724.
Behrens, R., and P. Nurse. 2002. Roles of fission yeast tea1p in the localization of polarity factors and in organizing the microtubular cytoskeleton. J. Cell Biol. 157:783793.
Browning, H., J. Hayles, J. Mata, L. Aveline, P. Nurse, and J.R. McIntosh. 2000. Tea2p is a kinesin-like protein required to generate polarized growth in fission yeast. J. Cell Biol. 151:1528.
Browning, H., D.D. Hackney, and P. Nurse. 2003. Targeted movement of cell end factors in fission yeast. Nat. Cell Biol. 5:812818.
Brunner, D., and P. Nurse. 2000. CLIP170-like tip1p spatially organizes microtubular dynamics in fission yeast. Cell. 102:695704.
Canman, J.C., L.A. Cameron, P.S. Maddox, A. Straight, J.S. Tirnauer, T.J. Mitchison, G. Fang, T.M. Kapoor, and E.D. Salmon. 2003. Determining the position of the cell division plane. Nature. 424:10741078.
Carvalho, P., J.S. Tirnauer, and D. Pellman. 2003. Surfing on microtubule ends. Trends Cell Biol. 13:229237.
Chang, F. 2001. Establishment of a cellular axis in fission yeast. Trends Genet. 17:273278.
Dent, E.W., and F.B. Gertler. 2003. Cytoskeletal dynamics and transport in growth cone motility and axon guidance. Neuron. 40:209227.
Diaz, E., F. Schimmoller, and S.R. Pfeffer. 1997. A novel Rab9 effector required for endosome-to-TGN transport. J. Cell Biol. 138:283290.
Drummond, D.R., and R.A. Cross. 2000. Dynamics of interphase microtubules in Schizosaccharomyces pombe. Curr. Biol. 10:766775.
Evangelista, M., K. Blundell, M.S. Longtine, C.J. Chow, N. Adames, J.R. Pringle, M. Peter, and C. Boone. 1997. Bni1p, a yeast formin linking cdc42p and the actin cytoskeleton during polarized morphogenesis. Science. 276:118122.
Evangelista, M., D. Pruyne, D.C. Amberg, C. Boone, and A. Bretscher. 2002. Formins direct Arp2/3-independent actin filament assembly to polarize cell growth in yeast. Nat. Cell Biol. 4:3241.
Feierbach, B., and F. Chang. 2001. Roles of the fission yeast formin for3p in cell polarity, actin cable formation and symmetric cell division. Curr. Biol. 11:16561665.
Fukata, M., T. Watanabe, J. Noritake, M. Nakagawa, M. Yamaga, S. Kuroda, Y. Matsuura, A. Iwamatsu, F. Perez, and K. Kaibuchi. 2002. Rac1 and Cdc42 capture microtubules through IQGAP1 and CLIP-170. Cell. 109:873885.
Gasman, S., Y. Kalaidzidis, and M. Zerial. 2003. RhoD regulates endosome dynamics through Diaphanous-related Formin and Src tyrosine kinase. Nat. Cell Biol. 5:195204.
Glynn, J.M., R.J. Lustig, A. Berlin, and F. Chang. 2001. Role of bud6p and tea1p in the interaction between actin and microtubules for the establishment of cell polarity in fission yeast. Curr. Biol. 11:836845.
Goode, B.L., D.G. Drubin, and G. Barnes. 2000. Functional cooperation between the microtubule and actin cytoskeletons. Curr. Opin. Cell Biol. 12:6371.
Goodson, H.V., S.B. Skube, R. Stalder, C. Valetti, T.E. Kreis, E.E. Morrison, and T.A. Schroer. 2003. CLIP-170 interacts with dynactin complex and the APC-binding protein EB1 by different mechanisms. Cell Motil. Cytoskeleton. 55:156173.
Gundersen, G.G. 2002. Evolutionary conservation of microtubule-capture mechanisms. Nat. Rev. Mol. Cell Biol. 3:296304.
Jin, H., and D.C. Amberg. 2000. The secretory pathway mediates localization of the cell polarity regulator Aip3p/Bud6p. Mol. Biol. Cell. 11:647661.
Jin, H., and D.C. Amberg. 2001. Fission yeast Aip3p (spAip3p) is required for an alternative actin-directed polarity program. Mol. Biol. Cell. 12:12751291.
Kikyo, M., K. Tanaka, T. Kamei, K. Ozaki, T. Fujiwara, E. Inoue, Y. Takita, Y. Ohya, and Y. Takai. 1999. An FH domain-containing Bnr1p is a multifunctional protein interacting with a variety of cytoskeletal proteins in Saccharomyces cerevisiae. Oncogene. 18:70467054.
Kim, H., P. Yang, P. Catanuto, F. Verde, H. Lai, H. Du, F. Chang, and S. Marcus. 2003. The kelch repeat protein, Tea1, is a potential substrate target of the p21-activated kinase, Shk1, in the fission yeast, Schizosaccharomyces pombe. J. Biol. Chem. 278:3007430082.
Kobielak, A., H.A. Pasolli, and E. Fuchs. 2004. Mammalian formin-1 participates in adherens junctions and polymerization of linear actin cables. Nat. Cell Biol. 6:2130.
Li, F., and H.N. Higgs. 2003. The mouse Formin mDia1 is a potent actin nucleation factor regulated by autoinhibition. Curr. Biol. 13:13351340.
Longtine, M.S., D.J. DeMarini, M.L. Valencik, O.S. Al-Alwar, H. Fares, C. De Virgilio, and J.R. Pringle. 1996. The septins: roles in cytokinesis and other processes. Curr. Opin. Cell Biol. 8:106119.
Maddox, A.S., and K. Oegema. 2003. Deconstructing cytokinesis. Nat. Cell Biol. 5:773776.
Mata, J., and P. Nurse. 1997. tea1 and the microtubular cytoskeleton are important for generating global spatial order within the fission yeast cell. Cell. 89:939949.
Mitchison, J.M., and P. Nurse. 1985. Growth in cell length in the fission yeast Schizosaccharomyces pombe. J. Cell Sci. 75:357376.
Moseley, J.B., I. Sagot, A.L. Manning, Y. Xu, M.J. Eck, D. Pellman, and B.L. Goode. 2004. A conserved mechanism for Bni1- and mDia1-induced actin assembly and dual regulation of Bni1 by Bud6 and profilin. Mol. Biol. Cell.15:896907.
Nakano, K., J. Imai, R. Arai, E.A. Toh, Y. Matsui, and I. Mabuchi. 2002. The small GTPase Rho3 and the diaphanous/formin For3 function in polarized cell growth in fission yeast. J. Cell Sci. 115:46294639.
Niccoli, T., and P. Nurse. 2002. Different mechanisms of cell polarisation in vegetative and shmooing growth in fission yeast. J. Cell Sci. 115:16511662.
Palazzo, A.F., T.A. Cook, A.S. Alberts, and G.G. Gundersen. 2001. mDia mediates Rho-regulated formation and orientation of stable microtubules. Nat. Cell Biol. 3:723729.
Pelham, R.J., and F. Chang. 2001. Role of actin polymerization and actin cables in actin-patch movement in Schizosaccharomyces pombe. Nat. Cell Biol. 3:235244.
Peng, J., B.J. Wallar, A. Flanders, P.J. Swiatek, and A.S. Alberts. 2003. Disruption of the Diaphanous-related formin Drf1 gene encoding mDia1 reveals a role for Drf3 as an effector for Cdc42. Curr. Biol. 13:534545.
Philips, J., and I. Herskowitz. 1998. Identification of Kel1p, a kelch domain-containing protein involved in cell fusion and morphology in Saccharomyces cerevisiae. J. Cell Biol. 143:375389.
Pruyne, D., and A. Bretscher. 2000. Polarization of cell growth in yeast. I. Establishment and maintenance of polarity states. J. Cell Sci. 113:365375.
Pruyne, D., M. Evangelista, C. Yang, E. Bi, S. Zigmond, A. Bretscher, and C. Boone. 2002. Role of formins in actin assembly: nucleation and barbed-end association. Science. 297:612615.
Rappaport, R. 1996. Cytokinesis in Animal Cells. Cambridge University Press, Cambridge, UK. 386 pp.
Robinson, D.N., and L. Cooley. 1997. Drosophila kelch is an oligomeric ring canal actin organizer. J. Cell Biol. 138:799810.
Sagot, I., S.K. Klee, and D. Pellman. 2002a. Yeast formins regulate cell polarity by controlling the assembly of actin cables. Nat. Cell Biol. 4:4250.
Sagot, I., A.A. Rodal, J. Moseley, B.L. Goode, and D. Pellman. 2002b. An actin nucleation mechanism mediated by Bni1 and profilin. Nat. Cell Biol. 4:626631.
Sawin, K.E., and P. Nurse. 1998. Regulation of cell polarity by microtubules in fission yeast. J. Cell Biol. 142:457471.
Schott, D., J. Ho, D. Pruyne, and A. Bretscher. 1999. The COOH-terminal domain of Myo2p, a yeast myosin V, has a direct role in secretory vesicle targeting. J. Cell Biol. 147:791808.
Sheeman, B., P. Carvalho, I. Sagot, J. Geiser, D. Kho, M.A. Hoyt, and D. Pellman. 2003. Determinants of S. cerevisiae dynein localization and activation: implications for the mechanism of spindle positioning. Curr. Biol. 13:364372.
Sheu, Y.J., B. Santos, N. Fortin, C. Costigan, and M. Snyder. 1998. Spa2p interacts with cell polarity proteins and signaling components involved in yeast cell morphogenesis. Mol. Cell. Biol. 18:40534069.
Small, J.V., B. Geiger, I. Kaverina, and A. Bershadsky. 2002. How do microtubules guide migrating cells? Nat. Rev. Mol. Cell Biol. 3:957964.
Snaith, H.A., and K.E. Sawin. 2003. Fission yeast mod5p regulates polarized growth through anchoring of tea1p at cell tips. Nature. 423:647651.
Snell, V., and P. Nurse. 1994. Genetic analysis of cell morphogenesis in fission yeasta role for casein kinase II in the establishment of polarized growth. EMBO J. 13:20662074.
Tanaka, E., and M.W. Kirschner. 1995. The role of microtubules in growth cone turning at substrate boundaries. J. Cell Biol. 128:127137.
Tasto, J.J., R.H. Carnahan, W.H. McDonald, and K.L. Gould. 2001. Vectors and gene targeting modules for tandem affinity purification in Schizosaccharomyces pombe. Yeast. 18:657662.
Toda, T., K. Umesono, A. Hirata, and M. Yanagida. 1983. Cold-sensitive nuclear division arrest mutants of the fission yeast Schizosaccharomyces pombe. J. Mol. Biol. 168:251270.
Tran, P.T., L. Marsh, V. Doye, S. Inoue, and F. Chang. 2001. A mechanism for nuclear positioning in fission yeast based on microtubule pushing. J. Cell Biol. 153:397411.
Velichkova, M., J. Guttman, C. Warren, L. Eng, K. Kline, A.W. Vogl, and T. Hasson. 2002. A human homologue of Drosophila kelch associates with myosin-VIIa in specialized adhesion junctions. Cell Motil. Cytoskeleton. 51:147164.
Verde, F., J. Mata, and P. Nurse. 1995. Fission yeast cell morphogenesis: Identification of new genes and analysis of their role during the cell cycle. J. Cell Biol. 131:15291538.
Wallar, B.J., and A.S. Alberts. 2003. The formins: active scaffolds that remodel the cytoskeleton. Trends Cell Biol. 13:435446.
Wang, H., X. Tang, J. Liu, S. Trautmann, D. Balasundaram, D. McCollum, and M.K. Balasubramanian. 2002. The multiprotein exocyst complex is essential for cell separation in Schizosaccharomyces pombe. Mol. Biol. Cell. 13:515529.
Waterman-Storer, C.M., and E. Salmon. 1999. Positive feedback interactions between microtubule and actin dynamics during cell motility. Curr. Opin. Cell Biol. 11:6167.
Yamamoto, A., R.R. West, J.R. McIntosh, and Y. Hiraoka. 1999. A cytoplasmic dynein heavy chain is required for oscillatory nuclear movement of meiotic prophase and efficient meiotic recombination in fission yeast. J. Cell Biol. 145:12331249.
Zigmond, S.H., M. Evangelista, C. Boone, C. Yang, A.C. Dar, F. Sicheri, J. Forkey, and M. Pring. 2003. Formin leaky cap allows elongation in the presence of tight capping proteins. Curr. Biol. 13:18201823.
Zimmerman, S., P.T. Tran, R. Daga, O. Niwa, and F. Chang. 2004. Rsp1p, a J-domain protein required for disassembly and assembly of microtubule organizing centers during the fission yeast cell cycle. Dev. Cell. 6:497509.