Research Article

Developmental control of plant Rho GTPase nano-organization by the lipid phosphatidylserine

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Science  05 Apr 2019:
Vol. 364, Issue 6435, pp. 57-62
DOI: 10.1126/science.aav9959

GTPase clustering in response to a hormone

Some lipid variants that are rare in plasma membranes function as signaling components. Studying root tip cells of the model plant Arabidopsis, Platre et al. found that phosphatidylserine, which is relatively abundant in plasma membranes, also modulates signaling pathways. Phosphatidylserine is required for the clustering of ROP6, a small guanosine triphosphatase (GTPase), in membranes in response to signals from the plant hormone auxin. Changes in phosphatidylserine concentration altered the clustering of ROP6 and thus the auxin signaling response.

Science, this issue p. 57


Rho guanosine triphosphatases (GTPases) are master regulators of cell signaling, but how they are regulated depending on the cellular context is unclear. We found that the phospholipid phosphatidylserine acts as a developmentally controlled lipid rheostat that tunes Rho GTPase signaling in Arabidopsis. Live superresolution single-molecule imaging revealed that the protein Rho of Plants 6 (ROP6) is stabilized by phosphatidylserine into plasma membrane nanodomains, which are required for auxin signaling. Our experiments also revealed that the plasma membrane phosphatidylserine content varies during plant root development and that the level of phosphatidylserine modulates the quantity of ROP6 nanoclusters induced by auxin and hence downstream signaling, including regulation of endocytosis and gravitropism. Our work shows that variations in phosphatidylserine levels are a physiological process that may be leveraged to regulate small GTPase signaling during development.

Proteins from the Rho/Ras superfamily are small guanosine triphosphatases (GTPases) that regulate fundamental eukaryotic functions, including cell signaling, cell polarity, intracellular trafficking, and cytoskeleton dynamics (1, 2). Furthermore, they control the morphology and behavior of cells and organisms by integrating signaling pathways at the cell surface into various cellular outputs. GTPases are considered to be in an “inactive” form when bound to guanosine diphosphate (GDP) and in an “active” form when bound to GTP. However, emerging evidence suggests that this view is likely oversimplified, because the membrane environment of GTPases also dictates their signaling capacity (2). For example, Ras/Rho signaling is intimately linked with membrane lipids in all eukaryotes. Interaction with anionic lipids is important for their plasma membrane targeting (3, 4) but also mediates the clustering of these small GTPases at the cell surface into nanometer-scale membrane domains (58). In particular, the phospholipid phosphatidylserine is involved in the nanoclustering and signaling of some GTPases, such as K-Ras in humans and Cdc42 in yeast (2, 7, 8). In contrast to other anionic phospholipids, phosphatidylserine is relatively abundant, representing as much as 10 to 20% of the total phospholipids at the plasma membrane inner leaflet (9). In addition, phosphatidylserine is not constantly modified by specialized metabolizing enzymes, and its subcellular repartition is thought to be relatively stable across cell types (9, 10). Therefore, phosphatidylserine appears to be a structural component of the membrane that is required for K-Ras/Cdc42 nanoclustering. It is unknown, however, whether phosphatidylserine also has a regulatory role in vivo in modulating nanocluster formation and subsequent signaling. In other words, is phosphatidylserine function rate-limiting in GTPase nanoclustering? If so, are phosphatidylserine levels regulated during development, and what are the consequences of such changes on small GTPase signaling capacity? Here, we addressed these questions using the Arabidopsis thaliana root as a model system because it is a genetically tractable multicellular organ, with a variety of cell types and cell differentiation states, and is amenable to live imaging.

In plants, there is a single protein family in the Ras/Rho GTPase superclade, called ROP (Rho of Plants) (11). ROPs are master regulators of cell polarity and cell morphogenesis, but they also sit at the nexus of plant hormone signaling (including auxin and abscisic acid), cell wall sensing pathways, and receptor-like kinase signaling (involved in development, reproduction, and immunity) (1118). We found that auxin triggers ROP6 nanoclustering within minutes in a phosphatidylserine-dependent manner. Furthermore, we found that phosphatidylserine is required for ROP6 signaling, and that variations in the cellular phosphatidylserine content directly affect the quantity of ROP6 nanoclusters and thereby subsequent downstream auxin signaling, including the regulation of endocytosis and root gravitropism. Therefore, phosphatidylserine is not a mere structural component of the membrane; it is a bona fide signaling lipid that acts as a developmentally controlled lipid rheostat to regulate small GTPases in a cell context–dependent manner.

Phosphatidylserine localization varies during root cell differentiation

Phosphatidylserine is an anionic phospholipid that accumulates in the cytosolic leaflets of the plasma membrane and endosomes (10). Bulk phosphatidylserine measurements in Arabidopsis thaliana suggested that the relative phosphatidylserine concentration can vary in vivo, depending on the organ (19). To observe phosphatidylserine distribution at tissue and cellular resolution, we recently validated the use of two phosphatidylserine reporters in Arabidopsis: the C2 domain of lactadherin (C2LACT) and the pleckstrin homology (PH) domain of EVECTIN2 (2xPHEVCT2) (10, 20). In both cases, the proportion of phosphatidylserine sensors was markedly more pronounced at the plasma membrane than at endosomes in the root basal meristem compared to cells in the elongation zone (Fig. 1A and fig. S1). This developmental gradient appeared to be in part regulated by the plant hormone auxin, because relatively short treatment (60 min) with the synthetic auxin naphthalene-1-acetic acid (NAA) increased the level of both phosphatidylserine sensors at the plasma membrane at the expense of their endosomal localization in the elongation zone (Fig. 1B and fig. S1C). Therefore, not only does the overall phosphatidylserine level vary depending on the organ, but also there are local variations of the phosphatidylserine content at the plasma membrane within an organ during cell differentiation and in response to hormonal cues.

Fig. 1 Plasma membrane phosphatidylserine levels vary during root cell differentiation.

(A and B) Confocal images and quantification of the plasma membrane (PM)/cytosol ratio of mCIT-C2LACT (phosphatidylserine sensor) root epidermis in the basal meristem and elongation zone (A) and in the absence or presence of 10 μM NAA (60 min, n = 150 cells) (B). Wilcoxon-Mann-Whitney/two-tailed test, significance level 5%. Scale bars, 10 μm. For this and all figures, red crosses and red horizontal lines denote mean and median values. (For this and all figures, see supplementary materials for details of statistical tests.)

Graded phosphatidylserine levels tune ROP6 signaling

To test the potential impact of phosphatidylserine variations during development, we experimentally manipulated the plant phosphatidylserine content. We produced three variants: one with no phosphatidylserine biosynthesis in the phosphatidylserine synthase1 (pss1) mutant (10), another with mild phosphatidylserine levels in transgenic lines expressing artificial microRNAs against PSS1 (amiPSS1), and a third with high phosphatidylserine levels in transgenic lines overexpressing PSS1 (PSS1-OX) (fig. S2). The changes in phosphatidylserine content measured in amiPSS1 and PSS1-OX lines (by a factor of ~ ±2) were in the physiological range, because phosphatidylserine levels in Arabidopsis vary by a factor of ~5 between root and leaf tissues (19). The pss1 mutant showed defects in root gravitropism (fig. S3, A and B). Quantitative analyses of root bending after gravistimulation (fig. S3C) revealed that the pss1-3 mutant had no gravitropic response (Fig. 2A) and that amiPSS1 lines had an attenuated response, whereas PSS1-OX lines were hypergravitropic (Fig. 2B). These opposite gravitropic phenotypes of PSS1 loss and gain of function resembled those of ROP6, a ROP GTPase that is activated by auxin and regulates root gravitropism (15, 16). Like PSS1-OX lines, lines overexpressing either ROP6 (ROP6-OX) or constitutively active GTP-lock ROP6 (ROP6CA) showed a hypergravitropic phenotype, which was abolished in a pss1-3 background (Fig. 2A). During root gravitropism, ROP6 acts downstream of auxin to inhibit endocytosis and regulate microtubule orientation (15, 16, 21). Similar to rop6 (15, 16, 21), we observed that in pss1-3 mutants, (i) uptake of the fluorescent endocytic tracer FM4-64 and PIN2-GFP (a fusion of the auxin efflux carrier PIN-FORMED 2 and green fluorescent protein) in the presence of brefeldin A (BFA) was increased (Fig. 2C and fig. S4, A to D), (ii) auxin failed to inhibit FM4-64 and PIN2-GFP endocytosis (Fig. 2C and fig. S4, A to D), (iii) clathrin light-chain plasma membrane association was insensitive to auxin treatment (fig. S4E), and (iv) auxin-triggered microtubule reorientation was abolished (fig. S4F). FM4-64 uptake in pss1-3xROP6CA plants was identical to that of the pss1-3 single mutant and opposite to ROP6CA (Fig. 2C), which shows that PSS1 is required for ROP6-mediated inhibition of endocytosis. Furthermore, transgenic lines with low phosphatidylserine content (amiPSS1) had decreased auxin-mediated inhibition of endocytosis, whereas lines with heightened phosphatidylserine content (PSS1-OX) mimicked ROP6CA phenotypes with pronounced inhibition of endocytosis upon auxin treatment (Fig. 2D and fig. S4, G to I). Together, our analyses suggest that (i) phosphatidylserine is required for auxin-mediated ROP6 signaling during root gravitropism and (ii) phosphatidylserine levels affect the strength of ROP6 signaling output in a dose-dependent manner.

Fig. 2 Variation in phosphatidylserine concentration tunes ROP6-mediated auxin response.

(A and B) Quantification of root bending after gravistimulation [means ± SEM, from top: n = 40, 28, 39, 33, 70, 64, 51 roots analyzed in (A); Kruskal-Wallis bilateral test combined with Conover-Iman procedure/two-tailed test, significance level 15%; n = 40, 52, 65, 77, 57 in (B), two-way analysis of variance (ANOVA), confidence index 95%, combined with Fisher test]. (C and D) Confocal images of FM4-64 staining in root epidermis (25 μM BFA, 5 μM NAA) and related quantification [from left: n = 26, 37, 14, 21, 28, 21 roots analyzed in (C), two-way ANOVA, confidence index 95%, combined with Fisher test; n = 33, 32, 36, 25, 24 in (D), two-way ANOVA, confidence index 95%, combined with Tukey test]. Scale bars, 10 μm. Dotted blue line is the tendency curve; letters indicate statistical differences. WT, wild type.

Auxin triggers ROP6 nanoclustering

Phosphatidylserine and ROP6 both accumulate at the plasma membrane, which suggests that phosphatidylserine may contribute to ROP6 localization. However, GFP-ROP6 localization, as seen by confocal microscopy, was almost identical in pss1-3 and wild-type plants, being mainly at the plasma membrane and only faintly delocalized in intracellular compartments in pss1-3 plants (fig. S5). In leaves, ROP6CA was previously shown to be confined in membrane domains (22), raising the possibility that phosphatidylserine could contribute to ROP6 signaling by regulating its lateral segregation at the plasma membrane. To analyze ROP6 plasma membrane partitioning in root cells and in the context of auxin response, we used several microscopy-based assays, including fluorescence recovery after photobleaching (FRAP), total internal reflection fluorescence microscopy (TIRFM), and photoactivated localization microscopy (PALM) (fig. S6). As shown for ROP6CA in leaf (22), activation of ROP6 (here using auxin treatment) delayed GFP-ROP6 FRAP in roots and increased the proportion of immobile GFP-ROP6 (Fig. 3A and fig. S7). TIRFM on root tip epidermal cells allowed us to focus only on the plane of the plasma membrane with an axial resolution of ~100 nm (fig. S6B) and revealed that GFP-ROP6 mostly localized uniformly at the plasma membrane (Fig. 3B). By contrast, in plants treated with two different auxins [NAA or the naturally occurring auxin indole-3-acetic acid (IAA)], GFP-ROP6 additionally resided in diffraction-limited spots present in the plane of the plasma membrane (Fig. 3B and fig. S8A), which suggests that auxin treatment triggers the clustering of ROP6 in membrane domains. By using stochastic photoswitching on live roots, single-particle tracking PALM (sptPALM) experiments provided tracks of single-molecule localization through time, and therefore allowed us to analyze the diffusion behavior of single ROP6 molecules in response to auxin (fig. S6D). Whereas mEos-ROP6 molecules in the untreated condition were almost exclusively diffusing, mEos-ROP6 molecules in plants treated for 5 min with NAA or IAA (or mEos-ROP6CA molecules) existed in two states at the plasma membrane of epidermal cells: immobile or diffusing (Fig. 3, C and D, fig. S9, and movie S1). Clustering analyses on live PALM images (23) showed that auxin triggered the clustering of mEos-ROP6 in plasma membrane nanodomains ~50 to 70 nm wide (Fig. 3, E and F, and fig. S10). Together, our data indicate that activation, either genetically (i.e., ROP6CA) or by an endogenous activator (i.e., auxin), triggers ROP6 recruitment, immobilization, and stabilization into nanodomains and that these events happen minutes after auxin treatment.

Fig. 3 Auxin triggers ROP6 nanoclustering at the plasma membrane of root epidermal cells.

(A) Kymograph images of GFP-ROP6 during FRAP experiments (100 nM NAA, 30 min) and quantification of GFP-ROP6 immobile fraction determined by FRAP analyses [n = 48, 34 regions of interest (ROIs), Student t test for two independent samples/two-tailed test, significance level 5%]. Red arrowhead indicates bleaching; white arrow is a 9-s time scale. (B) TIRFM micrograph of GFP-ROP6 (10 μM NAA, 20 min) and related quantification (n = 151, 277 ROIs; Wilcoxon-Mann-Whitney/two-tailed test, significance level 5%). (C) Representative mEos-ROP6 trajectories obtained by sptPALM analyses. Immobile molecules are in blue, mobile molecules in orange. (D) Distribution of mEos-ROP6 molecules according to their apparent diffusion coefficient D obtained by analyzing sptPALM trajectories (10 μM NAA, 5 min) and related quantification [n = 14, 14, 14 independent acquisitions (different cells), two-way ANOVA, confidence index 95%, combined with Fisher test]. (E) Live PALM analysis of mEos-ROP6 molecules density by tessellation-based automatic segmentation of superresolution images (10 μM NAA, 5 min). (F) Distribution of mEos-ROP6 molecules according to their local density and related quantification (n = 24, 24, 34 acquisitions; Kruskal-Wallis bilateral test combined with Steel-Dwass-Critchlow-Fligner multiple pairwise comparison, significance level 5%). Scale bars, 5 μm [(A) and (B)], 1 μm (E). Letters indicate statistical differences.

Dose-dependent regulation of ROP6 nanoclustering by phosphatidylserine

Next, we tested the impact of phosphatidylserine on ROP6 plasma membrane dynamics. In FRAP experiments, GFP-ROP6 sensitivity to auxin was reduced in pss1-3 plants (Fig. 4A and fig. S7), which implies that phosphatidylserine is critical for the immobilization of ROP6 by auxin. In wild-type plants, the presence of NAA-induced GFP-ROP6 in nanodomains was more pronounced in the basal meristem than in the elongation zone in TIRFM experiments (Fig. 4B), which correlated with the observed differential presence of phosphatidylserine content at the plasma membrane in these regions (Fig. 1A). To analyze whether this differential auxin sensitivity was dependent on the amount of phosphatidylserine present in these cells, we performed phosphatidylserine loss- and gain-of-function experiments. First, auxin failed to induce GFP-ROP6 nanodomains in both regions of the root in pss1-3 plants (Fig. 4C), which suggests that phosphatidylserine is indeed required for auxin-triggered ROP6 nanoclustering. Second, exogenous treatment with lysophosphatidylserine, a more soluble lipid than phosphatidylserine but with an identical head group (10), boosted the number of auxin-induced GFP-ROP6 nanodomains observed in TIRFM in wild-type plants (fig. S8B). Third, sptPALM analyses showed that the fraction of immobile mEos-ROP6CA (for which all ROP6 molecules are loaded with GTP) was significantly enhanced in PSS1-OX lines relative to the wild type (Fig. 4D and fig. S11). Together, these data suggest that the quantity of phosphatidylserine at the plasma membrane affects ROP6 nanoclustering.

Fig. 4 Phosphatidylserine is necessary for auxin-induced stabilization of ROP6 into nanodomains.

(A) Quantification of FRAP experiments in wild-type and pss1-3 root epidermal cells (1 nM or 100 nM NAA, 30 min; n = 48, 54, 34, 36, 54, 64 ROIs; two-way ANOVA, confidence index 95%, combined with Fisher test). (B and C) Quantification of TIRFM experiment in wild-type (n = 65, 78, 149, 116 ROIs) (B) and pss1-3 (n = 68, 170, 73, 168 ROIs) (C) root epidermal cells (10 μM NAA, 20 min) in basal meristem (BM) and elongation zone (EZ) (data are the same as in Fig. 3B but split into the respective zones); Kruskal-Wallis bilateral test combined with Steel-Dwass-Critchlow-Fligner multiple pairwise comparison, significance level 5%. (D) Distribution of mEos-ROP6CA molecules according to their apparent diffusion coefficient obtained by analyzing sptPALM trajectories in the wild-type and PSS1-OX background (n = 18, 18 acquisitions; Student t test for two independent samples/two-tailed test, significance level 5%). (E) Kymograph of GFP-ROP6 localization obtained by TIRFM, with images of a single GFP-ROP6 nanocluster (9-s interval) and related quantification (n = 282, 121, 195, 91 GFP-ROP6 nanodomains; Kruskal-Wallis bilateral test combined with Steel-Dwass-Critchlow-Fligner multiple pairwise comparison, significance level 5%). The black arrow is a 12-s time scale; the red arrowhead indicates bleaching; asterisks indicate GFP-ROP6–containing nanodomains. Scale bars, 5 μm. Letters indicate statistical differences.

Although phosphatidylserine was required for auxin-triggered ROP6 nanoclustering, a certain amount of ROP6 was still found in plasma membrane domains in pss1 plants, independent of the presence of auxin (Fig. 4C). Kymograph analyses revealed that ROP6-containing nanodomains observed by TIRFM in the presence of auxin were immobile in both wild-type and pss1-3 backgrounds (Fig. 4E). Photobleaching experiments showed that GFP-ROP6 was highly stable in these nanodomains in the wild type (i.e., no fluorescence recovery of GFP-ROP6 in nanodomains, and fast recovery of fluorescence outside of these domains) (Fig. 4E and movie S2). By contrast, GFP-ROP6 fluorescence in nanodomains was rapidly recovered in pss1-3 plants, which suggests that ROP6 was not stabilized into nanodomains in the absence of phosphatidylserine (Fig. 4E and movie S3). Together, our results show that phosphatidylserine is necessary for both ROP6 stabilization into nanodomains and downstream ROP6 signaling, including regulation of endocytosis and root gravitropism.

Immobile phosphatidylserine molecules accumulate in nanodomains

Next, we addressed whether phosphatidylserine, like ROP6, also localizes in nanodomains. Using sptPALM and clustering analyses, we found that in both the absence and presence of exogenous auxin, (i) the phosphatidylserine reporter mEos-2xPHEVCT2 segregated into nanodomains at the plasma membrane of root epidermal cells (Fig. 5A), and (ii) about 50% of mEos-2xPHEVCT2 molecules were present as a slow-diffusible population (Fig. 5B and fig. S12). The apparent diffusion coefficient of mEos-2xPHEVCT2 in the absence and presence of auxin was similar to that of immobile mEos-ROP6 in the presence of auxin (Fig. 3D), which suggests that ROP6 may cluster in preexisting phosphatidylserine nanodomains. Accordingly, co-visualization of GFP-ROP6 and the phosphatidylserine sensor 2xmCHERRY-C2LACT in TIRFM confirmed that they at least partially reside in the same nanodomains in response to auxin, whereas only 2xmCHERRY-C2LACT was present in nanoclusters in the absence of auxin (Fig. 5C and fig. S12D).

Fig. 5 ROP6 nanoclustering in response to auxin and downstream signaling requires interaction with anionic phospholipids.

(A) Live PALM analysis of mEos-2xPHEVCT2 molecule density by tessellation-based automatic segmentation of superresolution images (10 μM NAA, 5 min) in root epidermal cells. (B) Distribution of mEos-2xPHEVCT2 molecules according to their apparent diffusion coefficient obtained by analyzing sptPALM trajectories (n = 24, 34 independent acquisitions; Student t test for two independent samples/two-tailed test, significance level 5%); n.s., not significant. (C) TIRFM micrograph of root cells coexpressing GFP-ROP6 with the phosphatidylserine marker 2xmCH-C2LACT (10 μM NAA, 20 min). Arrows indicate phosphatidylserine-containing nanodomains in the absence of auxin; arrowheads indicate phosphatidylserine- and ROP6-containing nanodomains in the presence of auxin. (D and E) Quantification of root bending after gravistimulation (n = 49, 43, 42 roots; two-way ANOVA, confidence index 95%, combined with Tukey test) (D) and the size of FM4-64–stained BFA bodies (data for wild type are the same as in Fig. 2C; n = 26, 29, 22 roots; two-way ANOVA, confidence index 95%, combined with Fisher test) (E). (F) Distribution of mEos-ROP6 molecules in pss1-3 roots (left) and mEos-ROP67Q molecules in wild-type roots (right) according to their apparent diffusion coefficient, obtained by analyzing sptPALM trajectories (10 μM NAA, 5 min), and related quantification (n = 22, 22, 13, 15 acquisitions; two-way ANOVA, confidence index 95%, combined with Fisher test). Scale bars, 1 μm (A), 5 μm (C), 10 μm (E). Letters indicate statistical differences.

Direct ROP6-lipid interactions control nanoclustering and signaling

ROP6 possesses in its C terminus a polybasic region adjacent to a prenylation site (fig. S13A). Such a polybasic region is anticipated to bind to anionic phospholipids, including phosphatidylserine (3, 4, 6), which we confirmed in protein-lipid overlay experiments (fig. S13B). Substitution of seven lysines into neutral glutamines in ROP6 C terminus (ROP67Q) abolished in vitro interactions with all anionic lipids (fig. S13B). In planta, diminishing the net positive charges of ROP6 C terminus or the net negative charge of the plasma membrane gradually induced ROP6 mislocalization into intracellular compartments (fig. S13, C and D). To test the functionality of ROP67Q at the plasma membrane, we selected transgenic lines that had strong expression levels to compensate for their intracellular localization and therefore had comparable levels of ROP67Q and ROP6 at the plasma membrane (fig. S14, A, D, E, and F). ROP67Q mutants were not functional in planta (Fig. 5, D and E, and fig. S14, B and C), even though the 7Q mutations had no impact on ROP6 intrinsic GTPase activity in vitro and ROP6-GTP conformation in vivo (fig. S15). We next analyzed the dynamics of mEos-ROP67Q at the plasma membrane of wild-type roots by sptPALM experiments and found that it had the same proportion of immobile molecules as mEos-ROP6 in pss1-3 plants, and that in both cases they were insensitive to auxin (Fig. 5F and fig. S16). Therefore, impairing phosphatidylserine-ROP6 interaction by either removing phosphatidylserine from the membranes (pss1 mutant) or by mutating the anionic lipid-binding site on ROP6 (ROP67Q) had a similar impact on ROP6 signaling and its auxin-induced nanoclustering.


Our results show that in root tip epidermal cells, (i) ROP6 is immobilized in plasma membrane–localized nanodomains upon activation by auxin; (ii) phosphatidylserine is necessary for both ROP6 stabilization into nanodomains and signaling; (iii) ROP6 directly interacts with anionic lipids, including phosphatidylserine; and (iv) phosphatidylserine itself is present and immobile in these nanodomains, which suggests that stabilized ROP6 in phosphatidylserine-containing nanoclusters constitutes the functional signaling unit of this GTPase. Our imaging pipeline revealed that ROP6 nano-organization is rapidly remodeled by auxin, and as such it provides a quantitative in vivo readout to reevaluate how auxin may be perceived upstream of ROP6 activation. Plants have 11 ROPs that can respond to a wide range of signals (11), which raises the question of whether nanoclustering is specific to auxin response or common to other signals and to various ROPs, and to what extent it may contribute to signal integration by plant Rho GTPases. All ROP proteins have polybasic clusters at their C terminus (fig. S17A), and phosphatidylserine could therefore potentially regulate additional members of this family. In addition to root gravitropic defects, pss1 mutants had many developmental phenotypes that may be linked to altered ROP function (e.g., pavement cell and root hair morphology, planar polarity defects; see fig. S17, B to F) but that are not found in rop6 single mutants and could therefore involve additional ROP proteins. Furthermore, we found that ROP6 interacts with phosphoinositides and phosphatidic acid, which are required for its proper targeting to the plasma membrane (fig. S13). Additional studies are required to determine whether these anionic phospholipids may participate in ROP nanoclustering, perhaps in synergy with phosphatidylserine. Nanoclustering is a shared feature of several yeast and animal small GTPases, including K-Ras, Rac1, and Cdc42 (58), and both K-Ras and Cdc42 require phosphatidylserine for nanoclustering (2, 68). Our finding that ROP6 function depends on phosphatidylserine level opens the possibility that variations of phosphatidylserine concentration at the plasma membrane of both animal and plant systems may have specific regulatory functions and could control the signaling capacity of small GTPases, either during normal or pathological development.

Supplementary Materials

Materials and Methods

Figs. S1 to S17

Movies S1 to S3

References (2457)

References and Notes

Acknowledgments: We thank M. Dreux, E. Bayer, G. Ingram, O. Hamant, S. Mongrand, Y. Boutté, J. Gronnier, J. Reed, T. Vernoux, and the SiCE group for discussions and comments; T. Stanislas for help with root hair phenotyping; C. Burny for help with statistics; B. Peret for sharing his root template; S. Bednarek, S. Yalovsky, B. Scheres, and the NASC collection for providing transgenic Arabidopsis lines; A. Lacroix, J. Berger, and P. Bolland for plant care; and J. C. Mulatier for help in preparing lipids. We acknowledge the contribution of SFR Biosciences (UMS3444/CNRS, US8/Inserm, ENS de Lyon, UCBL) facilities: C. Lionnet, E. Chattre, and C. Chamot at the LBI-PLATIM-MICROSCOPY for assistance with imaging, and V. Guegen-Chaignon at the Protein Science Facility for assistance with protein purification. We thank the PHIV and MRI platform for access to microscopes. Funding: Y.J. is funded by ERC no. 3363360-APPL under FP/2007-2013; Y.J. and A.M. by the innovative project iRhobot from the department of “Biologie et Amélioration des Plantes” (BAP) of INRA. We acknowledge support by France-BioImaging (ANR-10-INBS-04, “Investments for the future”). Author contributions: M.P.P. generated all transgenic material and was responsible for all experiments and for performing statistics; V.B., M.P.P., and A.M. conceived, performed, and analyzed superresolution imaging; V.B. performed and analyzed TIRFM and FRAP imaging; L.M.-P. and P.M. performed lipid measurements; A.M. imaged Raichu-ROP6 sensors; J.B. produced recombinant ROP6 and performed GTPase assays; M.P.P. and L.A. performed lipid overlay experiments; M.d.M.M.-B. and C.M. assisted with phenotyping and cloning; A.C. performed PSS1 gene expression analyses; J.-B.F. and M.N. designed the sptPALM analyses pipeline; M.P.P., V.B., and Y.J. conceived the study, designed experiments, and wrote the manuscript; and all the authors discussed the results and commented on the manuscript. Competing interests: Authors declare no competing interests. Data and materials availability: All data are available in the main text or the supplementary materials.
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