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Phosphatidylserine is polarized and required for proper. Cdc42 localization and for development of cell polarity. Gregory D. Fairn1, Martin Hermansson2, Pentti ...

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Phosphatidylserine is polarized and required for proper Cdc42 localization and for development of cell polarity Gregory D. Fairn1 , Martin Hermansson2 , Pentti Somerharju2 and Sergio Grinstein1,3 Polarity is key to the function of eukaryotic cells. On the establishment of a polarity axis, cells can vectorially target secretion, generating an asymmetric distribution of plasma membrane proteins. From Saccharomyces cerevisiae to mammals, the small GTPase Cdc42 is a pivotal regulator of polarity. We used a fluorescent probe to visualize the distribution of phosphatidylserine in live S. cerevisiae. Remarkably, phosphatidylserine was polarized in the plasma membrane, accumulating in bud necks, the bud cortex and the tips of mating projections. Polarization required vectorial delivery of phosphatidylserine-containing secretory vesicles, and phosphatidylserine was largely excluded from endocytic vesicles, contributing to its polarized retention. Mutants lacking phosphatidylserine synthase had impaired polarization of the Cdc42 complex, leading to a delay in bud emergence, and defective mating. The addition of lysophosphatidylserine resulted in resynthesis and polarization of phosphatidylserine, as well as repolarization of Cdc42. The results indicate that phosphatidylserine—and presumably its polarization—are required for optimal Cdc42 targeting and activation during cell division and mating. Phospholipids, including phosphatidylserine, are asymmetrically distributed between and across membranes. Phosphatidylserine comprises only 3–10% of the phospholipids in eukaryotic cells and is enriched in the plasma membrane, where it is found almost exclusively in the inner leaflet1 . In mammalian cells the plasmalemmal cytosolic leaflet is composed of ≈20% phosphatidylserine, and the percentage is even higher in yeast2 . Furthermore, in yeast, phosphatidylserine levels show peak concentrations during bud emergence, followed by a decrease through the remainder of the cell cycle. This contrasts with the behaviour of phosphatidylcholine and phosphatidylethanolamine, which increase linearly in abundance as the cycle progresses. This differential behaviour is reflected in the cellular phosphatidylcholine/phosphatidylserine ratio, which is ≈2 : 1 at the time of bud emergence, whereas at later stages of the cycle it approaches 6:1 (ref. 3).

We generated a fusion of GFP with the discoidin-like C2 domain of lactadherin4 (GFP–Lact-C2) to visualize the intracellular distribution of phosphatidylserine in S. cerevisiae. At all stages of the cell cycle, the fluorescence signal of GFP–Lact-C2 was largely confined to the plasma membrane. However, the probe showed differential accumulation in distinct regions of the membrane depending on the stage of the cycle (Fig. 1a,d). Phosphatidylserine was concentrated at incipient bud sites (stage i in Fig. 1) and in small buds (stage ii). As the cells progress through the cycle and the bud becomes larger, increased signal is seen at the bud neck and the bud itself is enriched in phosphatidylserine when compared with the mother cell (stages iii and iv). In contrast, the distribution of phosphatidylinositol-4,5-bisphosphate (PtdIns(4,5)P2 ), monitored using a tandem PH (pleckstrin homology) domain of PLCδ fused to GFP, showed only modest polarization during the cycle (Fig. 1b,d). PtdIns(4,5)P2 was enriched ≈30% in the bud cortex and neck when compared with the remainder of the mother cell. In comparison, GFP–Lact-C2 was increased by >200% in the bud cortex and >300% at the bud neck. Hereafter, phosphatidylserine polarization was deemed to occur only when the GFP–Lact-C2 fluorescence intensity at the bud was at least twofold greater than that of the mother. We also used the plasmalemmal protein GFP–Ras2 as a marker to ensure that the observed differences in fluorescence intensity were not caused by increased membrane density5 . As shown in Fig. 1c, Ras2 is rather evenly distributed throughout the membrane at all stages of the cycle. These results demonstrate that phosphatidylserine enrichment at sites of bud formation is not a general property of plasma membrane lipids or lipid-modified plasmalemmal proteins. S. cerevisiae also undergo polarization during the formation of projections in response to mating factors. Stimulation of the receptor Ste2 by mating factor α leads to recruitment of Far1, which in turn recruits Cdc24, promoting the activation of Cdc42 (refs 6,7). As during bud formation, Cdc42 activation generates a polarity axis leading to the directed secretion of vesicles to the tips of mating projections. Figure 1e shows wild-type cells expressing GFP–Lact-C2, exposed to mating factor α for 3 h. The tips of mating projections were clearly enriched in phosphatidylserine. As in vegetative yeast, GFP–Ras2 distributed

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Program in Cell Biology, The Hospital for Sick Children, 555 University Avenue, Toronto, Ontario M5G 1X8, Canada. 2 Institute of Biomedicine, Department of Biochemistry and Developmental Biology, Haartmaninkatu 8, 00014 University of Helsinki, Finland. 3 Correspondence should be addressed to S.G. (e-mail: [email protected]) Received 3 August 2011; accepted 24 August 2011; published online 2 October 2011; DOI: 10.1038/ncb2351

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Figure 1 phosphatidylserine distribution in budding yeast. (a–c) Distribution of phosphatidylserine (a), monitored using the GFP–Lact-C2 probe, of PtdIns(4,5)P2 (b), monitored using the GFP–2×PH-PLCδ probe, and of the plasma-membrane marker GFP–Ras2 (c) during the cell cycle of BY4741 wild-type yeast cells grown to early log phase. Cells were examined at different points during bud growth; (i) bud emergence, (ii) tiny bud, (iii) small bud, (iv) large bud. In this and subsequent figures, images were acquired by spinning-disc confocal microscopy. The images in a–c are representative of three experiments, with >50 cells analysed per condition. (d) Quantification of the subcellular distribution of GFP–Lact-C2 and 2×PH-PLCδ–GFP in cells with small buds (similar to cells in column iii); data are means ± s.e.m. of 30 individual determinations pooled from three individual experiments. (e) Wild-type cells expressing GFP–Lact-C2, GFP–2×PH-PLCδ or GFP–Ras2 were grown to early log phase at 25 ◦ C and treated with mating factor α for 3 h before imaging. The most frequent phenotype and its prevalence as a percentage of total cells is shown. Scale bars, 4 µm.

evenly throughout the membrane, without concentrating in the mating projections (Fig. 1e). Moreover, unlike a recent report8 , we found that PtdIns(4,5)P2 did not accumulate measurably in the projections. Phosphatidylserine biosynthesis takes place in the endoplasmic reticulum, followed by delivery to other organelles, including the plasma membrane. It was conceivable that the phosphatidylserine found to accumulate at bud sites originated from the secretory pathway. We used temperature-sensitive mutants of Sec1 and Sec6 to investigate the role of secretion9 in generating phosphatidylserine polarization. For reference, we also examined the yeast synaptobrevin homologue, Snc1 (ref. 10). In wild-type yeast at steady state, Snc1 is found primarily

in the plasmalemma and is enriched in the bud, whether the cells are grown at 25 ◦ C or 37 ◦ C (Fig. 2a)11 . A similar distribution of Snc1 was noted in sec1ts or sec6 ts cells grown at 25 ◦ C, the permissive temperature (Fig. 2a). When secretion was impaired by incubation at 37 ◦ C, Snc1 accumulated in vesicular structures at or near budding sites (Fig. 2b). The sec1ts and sec6 ts cells had a normal distribution of phosphatidylserine at 25 ◦ C (Fig. 2a). The fraction of polarized cells and the degree of phosphatidylserine accumulation were quantified and the collated data are presented in Fig. 2c,d. In wild-type, sec1ts and sec6 ts cells at the permissive temperature, >95% of the small-budded cells have polarized GFP–Lact-C2 (Fig. 2c). In contrast, vesicles labelled with GFP–Lact-C2 accumulated in the cytosol as early as 10 min after shifting the temperature to 37 ◦ C; most cells exhibited multiple phosphatidylserine-enriched vesicles, as well as net depletion and loss of polarity of plasmalemmal phosphatidylserine after 60 min (Fig. 2b); only 4.8% and 3.2% of the sec1ts and sec6 ts mutants, respectively, showed polarization (Fig. 2c). As delivery to the surface membrane is arrested, phosphatidylserine that accumulated at the bud before the temperature shift diffuses along the plasma membrane, dissipating the concentration gradient (Fig. 2b–d). Dissipation of the gradient takes 20–30 min, indicating that phosphatidylserine may not diffuse freely from the bud to the mother. Indeed, in agreement with previous findings12 , we found that the lipid surrogate GFP–Ras2-tail has a diffusion coefficient of 0.046 µm2 s−1 , implying that lipids diffuse ×10–20 more slowly in yeast than in mammalian cells. Furthermore, a barrier at the mother–bud neck could further restrict lipid flow13 . We also studied a temperature-sensitive mutation that blocks exit from the trans-Golgi. Sec7, a nucleotide-exchange factor for Arf1/2, is required for vesicular exit from the Golgi14 . In Sec7 ts cells incubated at the non-permissive temperature, Snc1 and GFP–Lact-C2 accumulated in internal structures, identified earlier as an enlarged trans-Golgi9,15 (Fig. 2e). More importantly, by blocking secretion, the sec7 mutation prevented accumulation of phosphatidylserine at budding sites (Fig. 2e,f). Jointly, these findings indicate that targeted secretion of phosphatidylserine-containing vesicles underlies the polarization of the phospholipid in budding yeast. Growing yeast cells produce two main populations of secretory vesicles: high-density (HDSVs) and low-density secretory vesicles16,17 (LDSVs). To determine whether one type of vesicle is more important in phosphatidylserine polarity, we used two different mutants of Exo70 described to have impaired LDSV secretion18 ; the exo70-38 mutant is defective at 37 ◦ C, whereas the exo70-35 is defective at 25 ◦ C. A strain with plasmid-borne wild-type EXO70 was used as a control. Both mutants exhibited a near-normal phenotype at their respective permissive temperature, but at the restrictive temperature underwent a marked accumulation of phosphatidylserine- and Snc1-positive vesicles in the bud cytosol, while failing to accumulate these markers in the bud membrane (Supplementary Fig. S1a–d). These observations imply that LDSVs are at least partly responsible for phosphatidylserine polarization. HDSVs are less abundant than LDSVs and are a product of biosynthetic cargo trafficking through endosomes en route to the membrane. A mutant lacking the yeast dynamin homologue Vps1 impairs the generation of HDSVs (refs 16,17). Expression of GFP–LactC2 in Vps1-deficient cells (vps1∆) showed that, as in wild-type cells, most of the fluorescence signal emanates from the plasmalemma (Supplementary Fig. S1e). Moreover, ≈80% of the mutant cells with

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Figure 2 Secretion is required for the polarized distribution of phosphatidylserine. (a,b) Wild-type, sec1ts and sec6 ts cells co-expressing GFP–Lact-C2 and mRFP–Snc1 were grown to early log phase at 25 ◦ C (a), an aliquot shifted to 37 ◦ C for 60 min (b) and images of live cells were acquired. (c) Classification of GFP–Lact-C2 distribution in early mitotic cells. GFP–Lact-C2 was visualized by spinning–disc confocal microscopy as in a and b; small-budded cells were catalogued as having a polarized (yellow), non-polarized (black) or accumulated (blocked; grey) distribution of phosphatidylserine; data are means of 150 determinations pooled from three individual experiments. (d) Quantification of the fluorescence intensity of the GFP–Lact-C2 probe at the bud in wild-type, sec1ts and sec6 ts cells at both permissive and non-permissive temperatures (as in a and b). The fluorescence intensity of the bud membrane (PMbud )

is compared with that of the bud cytoplasm (Cytobud ) or that of the membrane of the corresponding mother (PMmother ). Data are means ± s.e.m. of 30 determinations pooled from three individual experiments. (e) sec7 ts cells co-expressing GFP–Lact-C2 and mRFP–Snc1 were grown to early log phase at 25 ◦ C, an aliquot shifted to 37 ◦ C for 60 min and images of live cells acquired. Spinning-disc confocal images are representative of three experiments. (f) Localization of GFP–Lact-C2 in early mitotic sec7 ts cells at 25 ◦ C or 37 ◦ C. GFP–Lact-C2 was visualized by spinning-disc confocal microscopy as in e; small-budded cells were catalogued as having a polarized (yellow), non-polarized (grey), blocked (black) or partially blocked appearance (blue) of GFP–Lact-C2; data are means of 150 individual determinations pooled from three individual experiments. Scale bars, 3 µm.

small–medium-sized buds have polarized phosphatidylserine, only slightly less than the >95% found for wild-type cells (Supplementary Fig. S1f). The degree of polarization is also slightly decreased (Supplementary Fig. S3g). Together, these findings indicate that LDSVs are the primary carriers of phosphatidylserine, whereas HDSVs make only a minor contribution. There is a precipitous decrease in the mole percentage (mol%) of phosphatidylserine in membranes during endocytic progression

from the plasma membrane to the vacuole19 . This indicates that mechanism(s) exist that segregate phosphatidylserine, retaining it in the plasmalemma. We therefore investigated whether phosphatidylserine is excluded from endocytic vesicles, which would contribute to its enrichment in the plasma membrane. Furthermore, as endocytosis rates are very high at sites of polarized growth, endocytosis may also contribute to phosphatidylserine polarization20 . We analysed several markers of endocytic traffic. First, Abp1–mCherry, a protein that

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Figure 3 phosphatidylserine is excluded from newly formed endocytic vesicles and endocytic organelles. (a) Wild-type cells expressing Abp1–mCherry and GFP–Lact-C2 were grown to early log phase, collected, resuspended in fresh medium and transferred to an agar slab for imaging. (b) Wild-type cells expressing GFP–Lact-C2 were grown to early log phase, washed and immobilized on a coverslip using concanavalin A. FM4-64 was added at a final concentration of 3.2 µM and incubated with cells for 5 min. The solution was then replaced with medium lacking

FM4-64 and images were acquired immediately. (c,d) Wild-type cells co-expressing RFP–Lact-C2 and either GFP–FYVE (c) or GFP–Vps27 (d). (e) Quantification of phosphatidylserine in newly formed endocytic vesicles and endocytic organelles. The corrected (cor.) fluorescence intensity of GFP–Lact-C2 in endosomes (endo.; identified by the indicated marker) and the plasma membrane (PM) is compared. Data are means ± s.e.m. of 20 determinations pooled from three experiments. Scale bars, 3 µm.

marks the internalization phase of endocytosis21 , was visualized along with GFP–Lact-C2. As shown in Fig. 3a and Supplementary Movies S1 and S2, whereas the Abp1–mCherry cluster forms initially at the membrane, overlapping with GFP–Lact-C2, once internalized the endocytic structures identified by Abp1–mCherry are largely (≈75%) devoid of GFP–Lact-C2 (Fig. 3e). Similar dynamics are seen using the dye FM4-64. When added for 5 min to wild-type cells expressing GFP–Lact-C2, FM4-64 labels early endocytic structures22 (Fig. 3b). The internal structures also label with GFP–Lact-C2, but the GFP intensity is much decreased when compared with the plasmalemma (Fig. 3e). These observations indicate that phosphatidylserine is excluded from the membrane of forming endosomes, a conclusion supported by earlier determinations of lipid content19 . We also found similarly low levels of GFP–Lact-C2 on endosomes and vacuolar membranes, as indicated by GFP–FYVE (a probe for phosphatidylinositol-3-phosphate23 ; Fig. 3c), GFP–Vps27 (the Hrs homologue24 ; Fig. 3d) and GFP–Atg18 (ref 25; not depicted) labelling. Thus, directed delivery to sites of bud formation, together with selective depletion from endocytic vesicles, contributes to the accumulation and polarization of phosphatidylserine at the bud. We next examined whether phosphatidylserine is required for optimal cell growth. First, the growth rate of yeast lacking phosphatidylserine was compared with wild-type cells. In S. cerevisiae, a single phosphatidylserine synthase is responsible for the biosynthesis of phosphatidylserine. Cells lacking the phosphatidylserine-synthase gene, referred to as cho1∆ (or pss1∆), grew more slowly than the parental strain, implying that phosphatidylserine is important for optimal growth (Fig. 4a). However, most functions examined in cho1∆ cells proceed normally: FM4-64 uptake, α-factor maturation, CPY trafficking and localization of the septin ring are unaltered26 (data not shown). The initial delivery of phosphatidylserine to the incipient bud site through polarized exocytosis may be required to support optimal bud growth. This hypothesis predicts that cho1∆ cells may have a delay

in bud emergence/formation. Cell-cycle synchronization was used to compare the time required for bud emergence in wild-type cells and in an isogenic cho1::kanMX strain. Large unbudded cells accumulated when growth was arrested in late G1 phase by depletion of G1 cyclins27 . At 60 min after release from G1, ≈30% of the wild-type cells had buds, and this number increased to ≈80% after 120 min. In contrast, only ≈2% of cho1∆ cells had buds after 60 min, and ≈10% did so after 120 min (Fig. 4b). We concluded that phosphatidylserine—and possibly its polarization at sites of bud formation—are important to support cell polarity and bud emergence. Cdc42 clusters at sites of bud emergence and resides on vesicles in close proximity to the bud28,29 , resembling the pattern of phosphatidylserine polarization described above. We therefore investigated whether clustering of Cdc42 requires phosphatidylserine. In wild-type yeast, GFP–Cdc42 was enriched in the plasma membrane at sites of bud emergence and in the bud cortex of small-budded cells (Fig. 4c). In cho1∆ cells, in contrast, GFP–Cdc42 was not visibly enriched at sites of bud emergence or in small buds, and there was an overall decrease in plasmalemmal binding. Instead, the amount of GFP–Cdc42 associated with internal organelles increased (Fig. 4c). These results demonstrate that phosphatidylserine is required for proper localization of Cdc42 to the plasma membrane and for its clustering during bud formation. Bem1 is a scaffold protein required for bud emergence30,31 . To further examine the role of phosphatidylserine in Cdc42-complex signalling, wild-type and cho1∆ cells were transformed with a plasmid expressing Bem1–GFP. In wild-type cells, Bem1–GFP localized to the site of bud emergence and to the tips of buds (Fig. 4d). However, as was the case for GFP–Cdc42, the polarization of Bem1–GFP to sites of bud emergence or bud tips was greatly decreased in cho1∆ cells (Fig. 4d). As Cdc42 is also required for the formation of mating projections, we investigated whether cells lacking phosphatidylserine may have impaired ability to mate. Indeed, cho1∆ cells failed to form proper

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Figure 4 phosphatidylserine is required for optimal Cdc42 location, activity and cell polarity. (a) Wild-type and cho1∆ cells were grown to early log phase at 25 ◦ C; subsequently, cells were plated in 1:10 serial dilutions onto YPD agar and incubated at 25 ◦ C for 3 days. (b) Quantification of bud emergence in wild-type and the cho1∆ cells. Cells were grown to early log phase in medium containing galactose, collected and arrested in G1 by incubating in medium containing glucose for 3 h. Next, cells were shifted back to galactose-containing medium. At the indicated times, an aliquot of the culture was removed, fixed, imaged and the percentage of budded cells counted. The histogram shows the percentage of cells with new buds ± s.e.m. (n = 3 with >100 cells counted per condition). ∗ indicates P < 0.025 and ∗∗ indicates P < 0.005. (c) Wild-type and cho1∆ cells were transformed with p415MET–GFP–Cdc42. Arrows indicate the buds in cho1∆ cells. (d) Wild-type and cho1∆ cells were transformed with a plasmid expressing Bem1–GFP. The arrow indicates the buds in the cho1∆ cells. Owing to the low fluorescence signal of the Bem1–GFP 2x 2 binning was used during image acquisition. (e) Quantification of

the degree of polarization of Cdc42 (black bars) and Bem1 (open bars) in wild-type and cho1∆ cells. Data are means ± s.e.m. of three experiments, with >50 cells counted for each condition per experiment. (f) Bright-field microscopy images of wild-type and cho1∆ cells grown to early log phase at 25 ◦ C and treated with mating factor α for 3 h. The most frequent phenotypes and their prevalence as a percentage of total cells are shown. (g) BY4741 or cho1∆ cells (106 ) were allowed to mate with SEY6210, Matα cells (107 ). After 4 h the cells were resuspended in medium and serial dilutions were plated on medium selective for diploids. Mating efficiency was calculated as the percentage of input Mata cells that formed diploids. Data are means ± s.d. (n = 3; with >1,000 cells counted for each condition/experiment). (h,i) Confocal microscopy images of wild-type and cho1∆ cells expressing GFP–Cdc42 (h) or Bem1–GFP (i) following exposure to mating factor for 3 h. The arrows in h indicate the tips of the mating projections. Spinning-disc confocal microscopy images in c,d,h,i are representative of 3 experiments, with >50 cells analysed per condition. Scale bars, 4 µm.

mating projections when stimulated by mating factor (Fig. 4f). Moreover, when crossed with a tenfold excess of CHO1 cells, the mating efficiency of cho1∆ cells was only 5%, compared with nearly 100% for their wild-type counterparts (Fig. 4g). As with bud emergence, in wild-type cells, GFP–Cdc42 was enriched at the tip of the mating projection (Fig. 4d). The cho1∆ cells expressing GFP–Cdc42 had atypical morphology, often having either a peanut shape or forming multiple projections (Fig. 4h). Regardless of their shape, however, little enrichment of GFP–Cdc42 was seen at the tips of projections in cho1∆ cells. Cdc24, similarly to Cdc42, localizes to the tips of mating projections in wild-type cells32 . In cho1∆ cells, GFP–Cdc24 showed a very modest association with the plasma membrane (not shown). As Bem1 not

only stabilizes the Cdc42–Cdc24 complex, but is known to localize to the tip of the mating projections (Fig. 4e), this adaptor may itself require phosphatidylserine for normal targeting. Consistent with this, Bem1–GFP was not recruited to the plasma membrane of phosphatidylserine-deficient cells (Fig. 4i). Jointly, these findings indicate that phosphatidylserine, and probably its polarization, are required for proper formation of projections and mating. The absence of phosphatidylserine in cho1∆ cells can cause secondary alterations in phospholipid content19 . We verified this by mass spectrometry: whereas in the parental strain, phosphatidylethanolamine comprised 15.9 mol% of the total phospholipids, cho1∆ cells contained only 2.7 mol% of phosphatidylethanolamine. Phosphatidylinositol and phosphatidylcholine showed a modest increase in cho1∆ cells, compen-

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Figure 5 Effects of phosphatidylserine repletion and phosphatidylethanolamine depletion on Cdc42 localization. (a) Cho1∆ and psd 1∆ cells expressing the GFP–Lact-C2 construct were grown to early log phase. An aliquot of the cho1∆ cells was incubated with NP40/lysophosphatidylserine (lyso-PS) mixed micelles for 1 h. As indicated, 85% of the cho1∆ cells expressing the GFP–Lact-C2 had the probe translocate to the plasma membrane following the addition of lysophosphatidylserine. (b) Quantification of the localization of GFP–Lact-C2 in early mitotic cells, from experiments similar to those in a. (c) Mass spectrometric determination of the mol% of phosphatidylethanolamine, phosphatidylserine and lysophosphatidylserine in wild-type and in

cho1∆ cells before and after the addition of lysophosphatidylserine. See Supplementary Tables S1 and S2 for a detailed breakdown of lipid species. (d) Psd1∆ and cho1∆ cells were transformed with p415MET–GFP–Cdc42 and grown in SC-leu medium containing 400 µM methionine to allow for limited expression of the GFP–Cdc42 protein. Where indicated the cho1∆ cells were incubated with lysophosphatidylserine/NP-40 mixed micelles for 1 h. Arrowheads indicate the location of buds in cho1∆ cells. Spinning-disc confocal images in d are representative of three experiments, with >50 cells analysed per condition. (e) Quantification of the fraction of cells with polarized GFP–Cdc42. Data are means ± s.e.m. of three experiments, with >50 cells counted for each condition per experiment. Scale bars, 4 µm.

sating for the loss in phosphatidylserine and phosphatidylethanolamine (Supplementary Table S1). It was important to determine whether the defective Cdc42 localization in cho1∆ cells was a direct consequence of the lack of phosphatidylserine, or was instead caused by the decreased phosphatidylethanolamine content. Two independent approaches were used. The first involved the addition of exogenous lysophosphatidylserine to the phosphatidylserine-deficient mutants. Lysophosphatidylserine is rapidly incorporated into the membrane and flipped to its inner monolayer. As documented in Fig. 5c and Supplementary Table S1, phosphatidylserine can then be generated de novo, probably by membrane-bound acyltransferases33 . The effectiveness of this approach was demonstrated using GFP–Lact-C2. The phosphatidylserine probe, which is largely cytoplasmic in the phosphatidylserine-deficient cells, redistributed rapidly (95%. The assay was carried out three times with >2,000 cells examined per condition.

area were determined at various time points and corrected for photobleaching incurred during image acquisition by dividing by the total cell fluorescence intensity at the corresponding time point.

Image analysis. Images were captured and analysed using Volocity (Perkin Elmer) or NIH ImageJ v1.38x to quantify fluorescence intensities from un-manipulated raw images. Regions of interest on the plasma membrane, endomembranes, in the cytosol and in an area outside the cell (background) were traced on highly magnified (typically 1,600%) images using the free-hand tool and their mean fluorescence intensities were measured. After background subtraction, the desired functions were carried out. For Fig. 1d, the polarization of signal in the plasma membrane of the bud was compared with the mother after subtraction of the cytosolic contribution (plasma membranebud − cytosolbud )/(plasma membranemother − cytosolmother ). For Fig. 3d, the indicated markers are used only to identify and trace the organelle of interest. The ratios are for the background-corrected GFP– or RFP–Lact-C2 signal for the organelle, compared with the average corrected signal of the plasma membrane.

Analysis of phospholipids by mass spectrometry. The yeast cells were disrupted by glass beads (0.5 mm; Sigma) at 4 ◦ C and phospholipds were extracted according to a previously published method38 . For quantification of the lipid species, a mixture of internal standards was added at the one-phase stage of extraction. The extract was evaporated and redissolved in chloroform/methanol (1:2, v/v). After the addition of 1% aqueous NH3 the sample was infused (6 µl min−1 ) into a Micromass Quattro Micro triple-quadrupole mass spectrometer operated as described previously39 . The phospholipid species were detected using specific precursor ion and constant neutral-loss scanning40 and then quantified using LIMSA software41 .

Statistical analysis. Results are expressed as the mean ± s.e.m. or ± s.d. from the specified number of experiments, as indicated in the figure legends. Student’s t -test was used to analyse statistical significance.

FRAP analysis. To determine the rate of recovery, the fluorescence intensity of the bleached area was compared with the intensity of an unbleached area of the plasma membrane. For each time point, the intensity of the bleached area was normalized to that of the corresponding unbleached area to correct for photobleaching incurred during the sampling. Data were fitted to a simple diffusion, zero-flow model using the formula: F(t ) = (F(t =0) + F(t =∞) × (t /t1/2 ))/1 + (t /t1/2 ) where the fluorescence intensity (F ) at a given time (t ) is related to the maximal fluorescence (F(t =∞) ) and the half-time of maximal recovery (t1/2 ). Using this equation, recovery curves were fitted by least-squares using Prism 4 (GraphPad Software). Diffusion coefficients were calculated from the t1/2 of the recovery curves as previously described using

Fluorescence microscopy. All fluorescence micrographs were acquired using spinning-disc confocal microscopy. The systems used (Quorum) are based on a Zeiss Axiovert 200M microscope with a ×100 objective. The units are equipped with diode-pumped solid-state laser lines (440 nm, 491 nm, 561 nm, 638 nm, 655 nm; Spectral Applied Research), a motorized x–y stage (ASI) and a piezo focus drive. Images were acquired using either back-thinned, electron-multiplied or conventional cooled CCD (charge-coupled device) cameras (Hamamatsu), driven by Volocity 4.1.1 software (Improvision).

Imaging of cells. Cells were grown to early log phase, collected by centrifugation, washed with PBS and resuspended in fresh medium. Mating projections were formed by incubating cells with 5 µM mating factor α for ≈3 h. Cells were attached to coverslips treated with concanavalin A or mounted on agarose pads. For FRAP (fluorescence recovery after photobleaching) experiments, cells were resuspended in growth medium, spotted on medium containing agarose pads, and covered with glass coverslips. Cells were imaged with a ×100 objective and a bleaching area was defined, as indicated. The fluorescence intensities of the bleached

D = (ω2 /4τ1/2 )γD where ω is the width of the bleach area, τ1/2 is the time of half-maximal recovery and γD is a constant equal to 0.88. 38. Folch, J., Lees, M. & Sloane Stanley, G. H. A simple method for the isolation and purification of total lipides from animal tissues. J. Biol. Chem. 226, 497–509 (1957). 39. Hermansson, M., Uphoff, A., Käkelä, R. & Somerharju, P. Automated quantitative analysis of complex lipidomes by liquid chromatography/mass spectrometry. Anal. Chem. 77, 2166–2175 (2005). 40. Koivusalo, M., Haimi, P., Heikinheimo, L., Kostiainen, R. & Somerharju, P. Quantitative determination of phospholipid compositions by ESI-MS: effects of acyl chain length, unsaturation, and lipid concentration on instrument response. J. Lipid Res. 42, 663–672 (2001). 41. Haimi, P., Uphoff, A., Hermansson, M. & Somerharju, P. Software tools for analysis of mass spectrometric lipidome data. Anal. Chem. 78, 8324–8331 (2006).

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DOI: 10.1038/ncb2351

o 25 C

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Figure S1 Delivery of low-density secretory vesicles is required for PS polarization. (a,b) EXO70, exo70-35ts and exo70-38ts cells co-expressing GFP-Lact-C2 and mRFP-Snc1 were grown to early log phase at 25°C, an aliquot shifted to 37°C for 60 min and images of live cells were acquired as in Fig. 1. Note: the non-permissive temperature for the exo70-35 mutant is 25°C. (c) Classification of the distribution of GFP-Lact-C2 in early mitotic cells. GFP-Lact-C2 was visualized by spinning-disc confocal microscopy as in a and b, small budded cells were catalogued as having a polarized (yellow), non-polarized (black) or accumulated (blocked; gray) appearance (n>50 cells; performed in triplicate). (d) Quantification of the fluorescence intensity of the GFP-Lact-C2 probe in the plasma membrane of the bud (PMbud) relative

37°C

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to the plasma membrane of the mother (PMmother) in the EXO70, exo70-35ts and exo70-38ts cells at both permissive and non-permissive temperatures (as in panels a and b). Data are means +/-± S.E.M. of 30 individual observations pooled from three individual experiments (n=30). (e) Cells lacking the yeast dynamin protein, Vps1, expressing the GFP-Lact-C2 fusion construct were grown to early log phase at 25°C and live cells imaged as above. (f) Localization of GFP-Lact-C2 in early mitotic cells. (g) Quantification of the fluorescence intensity of the GFP-Lact-C2 probe in the plasma membrane of the bud (PMbud) compared to the plasma membrane of the mother (PMmother) in the vps1D and wild type cells; data are means +/- ± S.E..M. of 30 determinations pooled from three individual experiments (n=30). Size bars = 3 mm.

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Supplemental Table 1. Summary of lipid analysis. Summary showing the mole % and % of wild-type (in bold) of each of the major classes of phospholipids and lyso-PS in wild-type, and cho1D grown with or without lyso-PS. Analysis of the molecular species and corresponding fatty acid composition referred to the accompanying excel file - “Supplemental Spreadsheet 1”. Supplemental Table 2. Strain List. A table containing a description of the yeast strains used in this study. Supplemental Table 3. Plasmid List. A table containing a description of the plasmids used in this study. Supplemental Spreadsheet 1. Lipidomics analysis. Analysis of the molecular species and corresponding fatty acid composition of the relevant phospholipids in yeast. Supplemental movie 1. ABP1 was genomically tagged at the C-terminus with mCherry:kanR. Cells were transformed with the plasmid encoding the GFPLact-C2 and maintained on synthetic complete medium lacking uracil. ABP1 is shown in red and Lact-C2 in green. Images were acquired at 3 sec intervals and are played at 2 frames/s. Supplemental movie 2: Cells co-expressing ABP1-mCherry and GFP-Lact-C2 were grown at 26 C. ABP1 is shown in red and Lact-C2 in green. Images were acquired at 3 sec intervals and are played at 10 frames/s.

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