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Spontaneous Cell Polarization Through Actomyosin-Based Delivery of the Cdc42 GTPase

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Science  21 Feb 2003:
Vol. 299, Issue 5610, pp. 1231-1235
DOI: 10.1126/science.1080944

Abstract

Cell polarization can occur in the absence of any spatial cues. To investigate the mechanism of spontaneous cell polarization, we used an assay in yeast where expression of an activated form of Cdc42, a Rho-type guanosine triphosphatase (GTPase) required for cell polarization, could generate cell polarity without any recourse to a preestablished physical cue. The polar distribution of Cdc42 in this assay required targeted secretion directed by the actin cytoskeleton. A mathematical simulation showed that a stable polarity axis could be generated through a positive feedback loop in which a stochastic increase in the local concentration of activated Cdc42 on the plasma membrane enhanced the probability of actin polymerization and increased the probability of further Cdc42 accumulation to that site.

The ability to self-organize is a fundamental property of living systems (1, 2). Cell polarity, or the generation of a vectorial axis controlling cell organization and behavior, is an important example. Although cell polarization is often directed by asymmetric cues from the environment or the cell's history (3), several cell types polarize randomly in response to global temporal signals, presumably through a self-organization mechanism. One view is that directed polarization in response to spatial cues could occur by biasing an intrinsic self-polarization system (4).

A simple and genetically tractable system such as yeast is useful for studying spontaneous cell polarization at the molecular level. However, in normal haploid yeast, cell polarization is guided by spatial cues such as the bud scar or a pheromone gradient (5, 6). Cue-dependent signaling generally involves localization of Cdc24, a guanine nucleotide exchange factor (GEF) for Cdc42 (7, 8). To render cell polarization in yeast independent of any specific physiological signals, we took advantage of the observation that expression of a constitutively activated form of Cdc42 is sufficient to cause polarization in otherwise nonpolarized cells, arrested in the G1 phase of the cell cycle, where Cdc24 is inactive (9, 10). This polarization event presumably bypasses the GEF and the spatial signals from the bud scar.

The original assays (11, 12) were modified (13) to improve the efficiency of polarization and to enable us to view live cells. Cells were arrested in G1 with the use of a strain with a methionine-repressible Cln2 as the sole source of G1cyclins (14). Expression of a constitutively active form of Cdc42, Cdc42Q61L (15), tagged with (myc)6-GFP (green fluorescent protein) at the N-terminus (referred to as MG-Cdc42Q61L), was induced under the control of the Gal1/10 promoter. The level of MG-Cdc42Q61Lrapidly increased during the initial 3 hours of induction and then reached a plateau estimated to be higher than the endogenous level of Cdc42 by a factor of 5 to 6 (fig. S1A) (16). Shortly after induction, the constitutively active MG-Cdc42Q61L exhibited a uniform distribution in the plasma membrane (Fig. 1A). With increasing time of induction, cells formed one or two polar caps of MG-Cdc42Q61L on the plasma membrane (Fig. 1, B and C, and Table 1). MG-Cdc42Q61L was found on the plasma membrane and on various internal membranes including vacuolar, nuclear, and endocytic membranes (17) (Fig. 1B) (fig. S1B), as well as on a large number of highly motile, dot-like structures that accumulated below the MG-Cdc42Q61L caps (arrows in fig. S1C).

Figure 1

MG-Cdc42Q61L in G1 cells leads to cell polarization. (A) Distribution of MG-Cdc42Q61L after 1 hour of induction. Note that the image is overexposed compared to (B). (B) Formation of polar MG-Cdc42Q61L caps after 3 hours of induction. V and N indicate vacuoles and the nucleus. (C) Kinetics of MG-Cdc42Q61L polarization. Percentages of cells with one or two caps were plotted against time of induction. (D) Time series showing the development of a polar cap (left cell, large arrow) and stability of a polar cap (right cell, small arrow). Time in minutes is given on each frame (time 0 = 90 min of induction). Linescans around the plasma membrane of the cell on the left are shown on the right of each frame. (E) Chymographs around the periphery of the cells represented in (D) in left-to-right order. Time is given in minutes. (F) MG-Cdc42Q61L localization and rhodamine-phalloidin staining of F-actin (arrow denotes an actin cable) in a cell after induction of MG-Cdc42Q61L for 3 hours. (G) Sec4 staining in cells expressing MG-Cdc42Q61L. Scale bars, 3 μm.

Table 1

Polarization of GFP-Cdc42 in wild-type and mutant strains.

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The process of MG-Cdc42Q61L polar cap formation was followed by time-lapse imaging using confocal microscopy. In all six cells that developed a polar cap in the course of the imaging experiments, the GFP signal started to accumulate at a single site on the plasma membrane and increased in intensity and width in a time-dependent fashion, until a steady state was reached (Fig. 1D). The position of the caps did not drift during or after their formation (Fig. 1, D and E) (movie S1), which suggests that stable polarity axes were established. Besides MG-Cdc42Q61L, fluorescence staining showed that two other polarity markers—F-actin and Sec4, a component of secretory vesicles (18)—were also concentrated at the caps, which often became protrusions (Fig. 1, F and G). Additionally, when the polarized Cdc42Q61L-expressing cells were released from G1 arrest, buds were seen to develop from polar caps (fig. S1E). Thus, the sites of Cdc42Q61Laccumulation were true sites of polarized growth.

Next, we investigated whether polarization of MG-Cdc42Q61L occurred independently of any preexisting structural or chemical asymmetry. First, staining with calcofluor showed no correlation between the localization of the MG-Cdc42Q61L polar caps and the bud scars, which normally dictate the positions of the new buds (5) (Fig. 2, A and B). Additionally, deletion of the BUD1 gene, encoding a factor essential for bud site selection (5), resulted in only a small reduction in the fraction of cells that formed MG-Cdc42Q61L polar caps (Table 1) (fig. S1F). Another potential spatial cue might come from an asymmetric distribution of cytoplasmic microtubules. However, treatment with the microtubule-destabilizing drug nocodazole had no effect on the formation of MG-Cdc42Q61L caps (Table 1) (fig. S1G). A third possible spatial cue could come from the geometry of the cell. To test this possibility, we induced MG-Cdc42Q61L cap formation in shmoo-shaped cells that were the result of a prior incubation with α-factor (a mating pheromone). In 77.0 ± 2.9% of the cells (n = 3 experiments with ≥100 cells each), MG-Cdc42Q61L caps did not overlap with, and were distributed randomly relative to, the original shmoo tip (Fig. 2C). Finally, we found that phosphatidylinositol 4,5-bisphosphate [PI(4,5)P2] and sterol-rich lipids—implicated in actin assembly (19) and cell polarization (20), respectively—were uniformly distributed in the plasma membrane of G1-arrested cells (Fig. 2, D and E). Thus, the MG-Cdc42Q61L–induced cell polarization was likely to reflect an underlying self-organizing process that was not directed by preexisting spatial cues.

Figure 2

Role of spatial cues and membrane transport in MG-Cdc42Q61L–induced polarization. (A) Localization of actin and bud scars (stained with calcofluor, CF) in a G1-arrested cell expressing MG-Cdc42Q61L. The arrow shows the site of polarization. (B) Quantification of the position of the MG-Cdc42Q61L cap relative to the closest bud scar. Bud scars were categorized as “adjacent” if the closest scar was less than 45° away from the polar cap on either side, as “opposite” if less than 45° away from the pole opposite of the polar cap, or as “side” if not in one of the above categories. (C) MG-Cdc42Q61L cap formation in a cell with a shmoo shape due to prior αfactor treatment. Arrow, shmoo tip; arrowhead, MG-Cdc42Q61L cap. (D and E) G1-arrested cell (before MG-Cdc42Q61Lexpression), showing a uniform distribution of PI(4,5)P2[(D), visualized with GFP-2xPH-PLCδ (31)] and sterol-rich lipids [(E), stained with filipin (20)], respectively. (F) G1-arrested cells were allowed to express MG-Cdc42Q61L for 3 hours and were then treated with 100 μM LatA for 15 min. (G to J) MG-Cdc42Q61L cap formation in various temperature-sensitive yeast strains. Cells were induced for 3 hours at 23°C and then incubated for 1 hour at 23° or 30°C (restrictive or semi-permissive temperature). (G) myo2-66; (H) Δtpm2 tpm1-2; (I) sec3-2; (J)sec6-4. Scale bars, 3 μm. (K) Subcellular fractionation of MG-Cdc42Q61L and Sec4. Low-speed supernatant (S2) and pellet (P2) and high-speed supernatant (S3) and pellet (P3) from wild-type and sec6-4 cells were obtained as described (13, 26). The P3 fraction from sec6-4cells was further extracted with vesicle buffer containing no salt (LS), 0.5 M KCl (HS), or 1% Triton X-100 (TX).

Because actin filaments accumulated at polar MG-Cdc42Q61Lcaps, we tested whether F-actin was required for polarization of MG-Cdc42Q61L. When the actin polymerization inhibitor latrunculin A (LatA) was added during induction of MG-Cdc42Q61L expression, cells completely failed to form MG-Cdc42Q61L caps (16). Furthermore, when LatA was added after the polar caps had formed, polarized distribution of MG-Cdc42Q61L was abolished in less than 15 min (Fig. 2F). This effect was completely reversible after LatA washout (fig. S1H). Thus, F-actin was required for both the establishment and maintenance of the Cdc42Q61L polar cap in G1-arrested cells.

Actin cables in yeast are known to have a crucial role in membrane transport toward the sites of polarized growth (21,22) and might be required for MG-Cdc42Q61L polar cap formation. To test this hypothesis, we examined the effects of mutations in the secretory pathway. myo2-66, a temperature-sensitive mutation affecting a type V myosin known to be responsible for the transport of secretory vesicles along actin cables (22–24), prevented the formation and maintenance of the MG-Cdc42Q61L polar cap at the restrictive but not the permissive temperature (Fig. 2G and Table 1). A similar defect was also observed in a temperature-sensitive tropomyosin mutant defective specifically in actin cable formation (Fig. 2H and Table 1). Additionally, mutations (sec3-2 and sec6-4) in components of the exocyst complex required for fusion of secretory vesicles at the plasma membrane (25) also blocked MG-Cdc42Q61L polar cap formation (Fig. 2, I and J, andTable 1). Thus, actomyosin-directed vesicle transport and fusion were essential for MG-Cdc42Q61L polarization.

To investigate whether Cdc42Q61L localized to secretory vesicles, we fractionated MG-Cdc42Q61L–expressing cells by differential centrifugation (26). We found that MG-Cdc42Q61Lbehaved similarly to the secretory vesicle marker Sec4 (Fig. 2K). In wild-type extracts, both proteins were present in the P2 fraction (containing plasma membrane) and to a lesser extent in P3 (containing secretory vesicles). In extracts from a sec6-4 strain at 37°C, both proteins were enriched in P3 (relative to the wild type, amounts of P3/P2 in sec6-4 extracts were increased by a factor of 3.33 in Sec4 and by a factor of 3.18 in MG-Cdc42Q61L). Both MG-Cdc42Q61L and Sec4 could be extracted from P3 by Triton X-100 but not by 0.5 M KCl (Fig. 2K), which suggests that the association was mediated through lipid interactions.

Thus, Cdc42Q61L polarization involved transport of GTP-bound Cdc42 along actin cables. The assembly of the actin cables is in turn dependent on the formin-like proteins, which can be activated by GTP-bound Cdc42 present on the plasma membrane (27–29). Newly synthesized Cdc42Q61L was originally (shortly after induction) deposited randomly in the plasma membrane (Fig. 3A, time 1). If, at a certain location, actin nucleation happened to occur, leading to formation of actin cables toward this site (Fig. 3A, time 2), new Cdc42Q61L would be selectively recruited to this site through membrane transport along actin cables (Fig. 3A, times 2 and 3). Deposition of Cdc42 would further stimulate actin nucleation at this site, leading to the formation of a Cdc42Q61L cap.

Figure 3

Mathematical simulation of MG-Cdc42Q61L polarization. (A) Schematic representation of the model for the establishment of Cdc42Q61L polar cap. A horizontal black bar represents an activated section of plasma membrane. Light gray circles, Cdc42Q61L; dark gray vertical bars, actin cables. Direction of transport is indicated by arrows. GW indicates the Gaussian distribution. Time is in arbitrary units. (B) Representative examples of simulations showing initial distribution of Cdc42Q61L and final distribution in cells that formed one or two caps (initial Cdc42Q61L on the plasma membrane = 1%, c = 5). (C) Kinetics of cap formation with low initial Cdc42Q61L concentration on the membrane. The evolution of 100 cells was simulated (c = 5; initial percentage of Cdc42Q61L on the plasma membrane = 0.1%). (D) Histograms showing the distribution of cells with 0, 1, and ≥2 caps simulated under different conditions (c = 1, 5, and 10; fractions of Cdc42Q61Linitially on the plasma membrane = 0.001, 0.01, and 0.1). (E) Formation of Cdc42Q61L caps with various amounts of initial Cdc42Q61L on the plasma membrane. MG-Cdc42Q61L expression was induced in arrested cells for 0, 30, and 180 min. Cells were then treated with LatA for 15 min, washed, and allowed to repolarize for 3 hours. (F) Examples of cells that formed multiple Cdc42Q61L caps (arrowheads) after LatA washout. Scale bar, 3 μm. (G) Quantification of Cdc42Q61L distribution in the cells shown in (F) using linescans around the cell periphery. Dashed line denotes background.

To assess whether the above positive feedback circuit is sufficient to induce cell polarization, we performed mathematical simulations (13) that demonstrate the generation of Cdc42Q61L peaks, similar in shape and number to those observed in living cells, from either a random initial distribution (Fig. 3B) or a completely uniform distribution of Cdc42Q61L(fig. S2A). The observed evolution (Fig. 3C) (fig. S2A) strongly resembled the observed kinetics of polarization after induction of MG-Cdc42Q61L (Fig. 1C). Variation of the initial concentration of plasma membrane–bound Cdc42Q61L or the transport rate (c) affected the number of caps formed per cell: The smaller the value of c, or the more Cdc42Q61L initially deposited on the plasma membrane, the more caps were predicted to form (Fig. 3D). Furthermore, the amplitude of individual peaks became smaller as more peaks formed per cell (Fig. 3B).

We experimentally tested one prediction of the simulation by varying the initial amount of Cdc42Q61L on the plasma membrane. Expression of MG-Cdc42Q61L was induced for various amounts of time in G1-arrested cells. The cells were then depolarized by brief treatment with LatA and allowed to repolarize for 3 hours after LatA washout. In this way, cells could begin polarization with different initial concentrations of MG-Cdc42Q61L on the plasma membrane. Because the level of MG-Cdc42Q61L plateaus after 3 hours of induction by galactose (fig. S1A), the final levels of MG-Cdc42Q61L in each of the treatments should be the same. Consistent with the prediction, we observed an increase in the fraction of cells with more than two MG-Cdc42Q61L caps with increasing initial concentrations of MG-Cdc42Q61L on the plasma membrane (Fig. 3, E and F). Furthermore, in cells with multiple polar caps, the size of the caps was smaller relative to those in cells with fewer caps (Fig. 3G).

We have demonstrated a cytoskeleton- dependent mechanism that could account for the intrinsic ability of cells to polarize in response to Cdc42 activation. This mechanism involves a positive feedback loop between Cdc42-dependent actin polymerization and F-actin–dependent delivery of Cdc42 to the plasma membrane. It remains to be determined to what extent such an intrinsic polarization mechanism contributes under physiological conditions where cell polarization is controlled by spatial signals. In neutrophils, the actin cytoskeleton plays an important role in the amplification of the spatial signal provided by gradients of chemoattractants (30). Thus, the cytoskeleton-dependent positive feedback loop could also be used as a powerful signal amplification mechanism that amplifies a small initial asymmetry in the distribution of polarity inducers, thereby establishing the polarity axis toward a physiologically relevant orientation.

Supporting Online Material

www.sciencemag.org/cgi/content/full/1080944/DC1

Materials and Methods

References

Figs. S1 and S2

Movie S1

  • * To whom correspondence should be addressed. E-mail: rong_li{at}hms.harvard.edu

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