Arabidopsis RhoGDIs Are Critical for Cellular Homeostasis of Pollen Tubes1[OPEN] Qiang-Nan Feng, Hui Kang, Shi-Jian Song, Fu-Rong Ge, Yu-Ling Zhang, En Li, Sha Li, and Yan Zhang* State Key Laboratory of Crop Biology, College of Life Sciences, Shandong Agricultural University, Tai’an, 271018, China ORCID ID: 0000-0002-3501-5857 (Y.Z.).

Rhos of plants (ROPs) play a key role in plant cell morphogenesis, especially in tip-growing pollen tubes and root hairs, by regulating an array of intracellular activities such as dynamic polymerization of actin microfilaments. ROPs are regulated by guanine nucleotide exchange factors (RopGEFs), GTPase activating proteins (RopGAPs), and guanine nucleotide dissociation inhibitors (RhoGDIs). RopGEFs and RopGAPs play evolutionarily conserved function in ROP signaling. By contrast, although plant RhoGDIs regulate the membrane extraction and cytoplasmic sequestration of ROPs, less clear are their positive roles in ROP signaling as do their yeast and metazoan counterparts. We report here that functional loss of all three Arabidopsis (Arabidopsis thaliana) GDIs (tri-gdi) significantly reduced male transmission due to impaired pollen tube growth in vitro and in vivo. We demonstrate that ROPs were ectopically activated at the lateral plasma membrane of the tri-gdi pollen tubes. However, total ROPs were reduced posttranslationally in the tri-gdi mutant, resulting in overall dampened ROP signaling. Indeed, a ROP5 mutant that was unable to interact with GDIs failed to induce growth, indicating the importance of the ROP-GDI interaction for ROP signaling. Functional loss of GDIs impaired cellular homeostasis, resulting in excess apical accumulation of wall components in pollen tubes, similar to that resulting from ectopic phosphatidylinositol 4,5-bisphosphate signaling. GDIs and phosphatidylinositol 4,5-bisphosphate may antagonistically coordinate to maintain cellular homeostasis during pollen tube growth. Our results thus demonstrate a more complex role of GDIs in ROP-mediated pollen tube growth.

The plant-specific Rho GTPases, ROP/Rac, play a central role in plant cell morphogenesis, especially in tip-growing cells such as pollen tubes and root hairs (Yang, 2002; Kost, 2008; Bloch and Yalovsky, 2013). ROP activation is translated into an array of intracellular events, such as dynamic polymerization of actin microfilaments (MFs) and dynamic maintenance of a Ca 2+ gradient during tip growth (Fu et al., 2001; Foreman et al., 2003; Gu et al., 2005) through effector binding (Wu et al., 2001; Lavy et al., 2007). In addition to actin MF and Ca2+ gradients, other key intracellular activities including regulated membrane trafficking (Lavy et al., 2007; Lee et al., 2008; Szumlanski and 1

This work was supported by Major Research Plan (2013CB945102) from the Ministry of Science, Technology of China, Natural Science Foundation of China (31271578), and by Shandong Provincial Funds for Outstanding Young Scientists (to Y.Z.). Y.Z.’s laboratory is partially supported by Tai-Shan Scholar Program by Shandong Provincial Government. * Address correspondence to [email protected]. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Yan Zhang ([email protected]). Y.Z. conceived and supervised the project; Q.F. performed most of the experiments; H.K. and S.S. provided technical assistance to Q.F.; F.G., Y.-L.Z., and E.L. provided materials for this project; Y.Z., S.L, and Q.F. designed the experiments and analyzed the data; Y.Z. wrote the article with contributions of all the authors. [OPEN] Articles can be viewed without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.15.01600

Nielsen, 2009; Hazak et al., 2010) and the asymmetric distribution of phosphatidylinositol 4,5-bisphosphate [PI(4,5)P2; Dowd et al., 2006; Helling et al., 2006; Ischebeck et al., 2008; Sousa et al., 2008; Ischebeck et al., 2011] may also be regulated by ROPs based on their association in tip-growing cells (Kost et al., 1999; Lee et al., 2008; Ischebeck et al., 2011) and their interactions in other plant cells (Bloch et al., 2005; Hála et al., 2008; Hazak et al., 2010; Ischebeck et al., 2013). Interdependency and cross-talk between these intracellular activities are crucial for tip growth (Ischebeck et al., 2008; Sousa et al., 2008; Zhang et al., 2010; Ischebeck et al., 2011). As key molecular switches, ROPs alternate between a GTP-bound active form and a GDP-bound inactive form, whose distinct spatiotemporal distribution is critical for tip growth (Bloch et al., 2005; Nibau et al., 2006; Kost, 2008). Guanine nucleotide exchange factors (RopGEFs) or GTPase activating proteins (RopGAPs) turn “on” or “off” the ROP switches, respectively (Yang, 2002; Nibau et al., 2006; Berken and Wittinghofer, 2008; Kost, 2008; Bloch and Yalovsky, 2013). Despite being a plant-specific family distinct in domain organization from their counterparts in other phyla, RopGEFs positively regulate ROP activation by catalyzing the GTPGDP exchange (Berken et al., 2005; Gu et al., 2006; Zhang and McCormick, 2007). In contrast, the catalytic domain of plant RopGAPs is evolutionarily conserved except for their distinct regulatory domains (Wu et al., 2000; Klahre and Kost, 2006; Hwang et al., 2008). Of the three classes of RopGAPs found in plant genomes, two have been demonstrated to regulate ROP signaling

Plant PhysiologyÒ, February 2016, Vol. 170, pp. 841–856, www.plantphysiol.org Ó 2016 American Society of Plant Biologists. All Rights Reserved.

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(Wu et al., 2000; Klahre and Kost, 2006; Hwang et al., 2008). Guanine nucleotide dissociation inhibitors (GDIs) are another group of evolutionarily conserved regulators for Rho GTPases. In yeast and metazoans, GDIs play crucial roles in Rho signaling through membrane extraction, cytoplasmic sequestration, cytoplasmic stabilization, and recycling of Rho GTPases (Boulter et al., 2010; Garcia-Mata et al., 2011). Recent studies suggested that GDI-dependent recycling of Cdc42 is critical for maintaining a single and unique polarization site during yeast budding and mating (Slaughter et al., 2009; Freisinger et al., 2013). Studies in pollen tubes have clearly shown that plant GDIs retain the ability to extract ROPs from membranes and to sequester ROPs in the cytoplasm (Klahre et al., 2006; Hwang et al., 2010). Overexpression of GDIs suppressed depolarized growth induced by ROP overexpression in pollen tubes, suggesting a negative role of GDIs in ROP-induced growth (Klahre et al., 2006; Hwang et al., 2010). In root hairs, the normally polar distribution of ROP2 (Molendijk et al., 2001) became apolar in the GDI1 loss-of-function mutant supercentipede1 (scn1), in which multiple initiation sites appeared in a single trichoblast, indicating impaired polarity control (Carol et al., 2005). Results in both types of tip-growing plant cells indicate that GDIs act as inhibitors for ROP-mediated signaling (Hwang et al., 2010). However, it is unclear whether plant GDIs are involved in the dynamic recycling of ROPs as are their counterparts in other phyla. The Arabidopsis (Arabidopsis thaliana) genome encodes three RhoGDIs, designated GDI1, GDI2a, and GDI2b (Bischoff et al., 2000). GDI1 (SCN1) expression is constitutive, but the other two GDIs are enriched in pollen by reverse transcription PCRs (RT-PCRs; Hwang et al., 2010). Due to the lack of null mutants for these GDIs, attempts were made to understand their function in pollen tubes with a RNA interference approach (Hwang et al., 2010). However, RNA interference often causes off-target down-regulation, thus preventing the functionality of GDIs to be revealed unequivocally. We carried functional characterization of Arabidopsis GDIs by a reverse genetic approach. Functional loss of all three GDIs (tri-gdi) significantly reduced male transmission due to impaired growth of pollen tubes in vitro and in vivo. Active ROPs were ectopically distributed to the lateral plasma membrane (PM) of the trigdi pollen tubes, in which altered polymerization of actin MF rendered it more resistant to latrunculin B (LatB). Despite that, total ROPs were reduced in the trigdi mutant, indicating overall dampening of ROP signaling. We discovered that functional loss of GDIs resulted in impaired cellular homeostasis, i.e. excess accumulation of pectins and cellulose at the apex of pollen tubes, in a way similar to that induced by ectopic PI(4,5)P2 signaling. We further show that GDIs and PI(4,5)P2 may antagonistically coordinate to regulate pollen tube growth, likely by coupling exocytosis and endocytosis. Our results thus demonstrate a more complex role of GDIs in ROP-mediated pollen tube growth. 842

RESULTS The tri-gdi Mutant Was Defective in Male Transmission

To test the expression pattern of GDI2 (GDI2a) and GDI3 (GDI2b), we first generated the promoter: NLS-YFP transgenic lines. By fluorescence microscopy, we determined that both GDI2 and GDI3 were highly expressed in mature pollen and pollen tubes (Supplemental Fig. S1), as suggested previously by RTPCRs (Hwang et al., 2010), implying their function mainly in male gametophytes. However, quantitative real-time PCRs (qRT-PCRs) showed that GDI3 was also detectable in other tissues, such as seedlings, pistils, and roots (Supplemental Fig. S1), suggesting its potential roles in other developmental or cellular processes. To understand the function of Arabidopsis GDIs, we took a reverse genetic approach by generating a mutant defective in all three GDIs (Fig. 1A). A null mutant of GDI1/SCN1, scn1-1, was described previously (Carol et al., 2005). We obtained T-DNA insertion mutants for GDI2 (FLAG_184A02, gdi2) and GDI3 (FLAG_259E06, gdi3). Transcriptional analyses showed that gdi2 was a null mutant for GDI2 (Fig. 1B). Although no full-length GDI3 was expressed in gdi3 (Fig. 1B), a partial transcript including the first four exons of GDI3 was detected in gdi3. To exclude the possibility that the partial GDI3 transcript encoded a functional protein, we verified the interaction of its potential protein product with ROPs by yeast two-hybrid assay (Y2H) and bimolecular fluorescence complementation (BiFC) because GDIs function through interacting with ROPs. Both Y2H and BiFC showed that the partial GDI3 was unable to interact with ROPs (Supplemental Fig. S2), as suggested by structural studies demonstrating the importance of the C-terminal GDI in Rho interaction (Dovas and Couchman, 2005). In addition, GDI3 driven by the pollen-specific promoter ProLAT52 was able to restore growth of tri-gdi pollen tubes, in the same way as ProLAT52: GDI1/SCN1 (Supplemental Fig. S3), confirming the null identity of the triple GDI mutant. Single mutants for GDIs did not show any growth defects both in tube width and length (Supplemental Fig. S3), implying functional redundancy. Thus, hierarchy mutants were analyzed for gametophytic transmission. Among different double mutant combinations, reduced male transmission was only detected for pollen mutated at both SCN1 and GDI2 (Table I), which is to a lesser extent than pollen mutated at all three GDIs (Table I), indicating that GDIs play redundant roles in male gametophytic function.

The tri-gdi Mutant Was Impaired in Pollen Tube Growth in Vitro and in Vivo

The tri-gdi mutant was comparable to wild type with regards to vegetative and reproductive growth under greenhouse condition (Fig. 1, C and D), except for its shorter siliques that contained only a few seeds at the top (Fig. 1, E–H). To determine the stage when GDI loss Plant Physiol. Vol. 170, 2016

GDIs Coordinate Pollen Tube Homeostasis

Figure 1. The tri-gdi mutant is defective in male transmission. A, Schematic illustration of the genomic regions and the T-DNA insertions of GDI2 and GDI3. Arrows indicate the binding sites for RTPCR primers. B, Transcript analysis of GDI2 and GDI3 in the triple GDI mutant (tri-gdi ). ACTIN2 (ACT2) was used as the internal control. C and D, Representative images of the wild type (left) and tri-gdi (right) at the vegetative (D) and reproductive (C) stages. E, A representative main inflorescence of the wild type (left) and tri-gdi (right). F, Quantitative analysis of fertility in the wild type (WT) and tri-gdi (gdi). Results are means 6 SD (n = 20). Asterisk indicates significant difference (t test, P , 0.01). G and H, Representative open siliques of the wild type (G) and tri-gdi (H). Arrows point at the developing seeds in tri-gdi. Bars = 1 cm for C–E; 1 mm for G and H.

of function resulted in male gametophytic defects, we analyzed pollen development by scanning electron microscopy to assess pollen coat structure, by Alexander staining for cytoplasmic viability, and by 49,6-diamidino2-phenylindole staining for nuclear organization (JohnsonBrousseau and McCormick, 2004; Li et al., 2013a). By all approaches, the tri-gdi mutant was comparable to wild type (Supplemental Fig. S4). Thus, the significantly reduced male transmission in the tri-gdi mutant was due to defects in pollen germination or tube growth. Mature pollen germinate on the stigma. The tubes grow inside pistils in a polar and guided way, penetrate the embryo sac, and finally rupture to deliver sperm for double fertilization (McCormick, 1993). To determine at which stage male transmission was compromised due to GDI functional loss, we examined pollen germination and tube growth in vitro. Pollen of the tri-gdi plants germinated at a comparable rate to that of the wild type at earlier hours (Supplemental Fig. S4). However, tri-gdi pollen showed a higher germination potential than that of the wild type: extended incubation resulted in around 95% germination in the tri-gdi mutant but only around 85% in the wild type (Supplemental Fig. S2). In addition, GDI loss of function significantly affected pollen tube growth such that tri-gdi pollen tubes were stubby, with tube diameters 3 times that of wild-type tubes and rarely Plant Physiol. Vol. 170, 2016

longer than 100 mm (Fig. 2, A–D). Unlike the case in root hairs (Carol et al., 2005), no branches were formed in trigdi pollen tubes (Fig. 2B), suggesting distinct mechanisms underlying these similar tip-growing cells. To determine the growth dynamics of tri-gdi pollen tubes in vivo, we pollinated wild-type pistils with either wild-type or tri-gdi pollen. At 9 h after pollination (HAP), wild-type pollen tubes mostly grew to the bottom of pistils (Fig. 2E). By contrast, tri-gdi pollen tubes had just exited the style (Fig. 2F). At 48 HAP, when wild-type pollen tubes turned toward ovules and entered the micropyle (Fig. 2, G and I), the majority of tri-gdi pollen tubes were still in the upper part of the transmitting tract (Fig. 2H). A few tri-gdi pollen tubes managed to grow deep down in the transmitting tract but were later defective in funicular guidance (Fig. 2J). Only a few seeds could be obtained in self-fertilized tri-gdi plants (Fig. 1F). These results demonstrated that GDI loss of function impaired pollen tube growth both in vitro and in vivo. Ectopic ROP Activation and Dynamic Actin MF in tri-gdi Pollen Tubes

Because GDIs are regulators of ROPs, pollen tube defects detected in GDI loss of function presumably resulted from defective ROP signaling. Therefore, we 843

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Table I. Segregation ratios of trans-heterozygous plants for GDIs WT, Wild type. Parents (Female 3 Male)

Progenies (Genotype)

Expected

Observed

scn1 gdi2 +/2 3 WT WT 3 scn1 gdi2 +/2 gdi2 +/2 3 gdi2 +/2 (scn1) scn1 gdi3 +/2 3 WT WT 3 scn1 gdi3 +/2 gdi3 +/2 3 gdi3 +/2 (scn1) gdi3 gdi2 +/2 3 WT WT 3 gdi3 gdi2 +/2 gdi2 +/2 3 gdi2 +/2 (gdi3) gdi2 gdi3 +/2 3 WT WT 3 gdi2 gdi3 +/2 gdi3 +/2 3 gdi3 +/2 (gdi2) scn1 +/2 (gdi2 gdi3) 3 scn1 +/2 (gdi2 gdi3) gdi2 +/2 (scn1 gdi3) 3 gdi2 +/2 (scn1 gdi3) gdi3 +/2 (scn1 gdi2) 3 gdi3 +/2 (scn1 gdi2)

gdi2 +/+: gdi2 +/2 (scn1 +/2) gdi2 +/+: gdi2 +/2 (scn1 +/2) gdi2 +/+ & +/2: gdi2 (scn1) gdi3 +/+: gdi3 +/2 (scn1 +/2) gdi3 +/+: gdi3 +/2 (scn1 +/2) gdi3 +/+ & +/2: gdi3 (scn1) gdi2 +/+: gdi2 +/2 (gdi3 +/2) gdi2 +/+: gdi2 +/2 (gdi3 +/2) gdi2 +/+: gdi2 +/2: gdi2 (gdi3) gdi3 +/+: gdi3 +/2 (gdi2 +/2) gdi3 +/+: gdi3 +/2 (gdi2 +/2) gdi3 +/+: gdi3 +/2: gdi3 2/2 (gdi2) scn1 +/+ or scn1 +/2: scn1 (gdi2 gdi3) gdi2 +/+: gdi2 +/2: gdi2 (scn1 gdi3) gdi3 +/+: gdi3 +/2: gdi3 (scn1 gdi2)

1:1 1:1 3:1 1:1 1:1 3:1 1:1 1:1 1:2:1 1:1 1:1 1:2:1 3:1 1:2:1 1:2:1

84:80 131:24a 463:182b 119:105 84:73 122:43 66:63 50:48 32:61:28 89:91 101:97 25:60:30 920:2b 50:46:0c 37:27:2c

a Significantly different from 1:1 (x 2, P , 0.01). 0.01).

b

Significantly different from 3:1 (x 2, P , 0.01).

introduced ProLAT52:CRIBRIC1-mRFP (Li et al., 2013b; Zhao et al., 2013), a pollen-specific fluorescent probe for GTP-bound ROPs (Wu et al., 2001), into the wild type by transformation and into tri-gdi by crosses so that comparable expression from the same transgene was reached in both backgrounds. As reported previously (Hwang et al., 2005; Hwang et al., 2010; Li et al., 2013b; Zhao et al., 2013), active ROPs are restricted to the apical PM surrounding the clear zone in wild-type pollen tubes (Fig. 3, A and C; Supplemental Movie S1). By contrast, GDI loss of function resulted in expansion of the CRIB signal (Fig. 3G) to the shank PM, far away from the apex and with no obvious asymmetry (Fig. 3, B and D; Supplemental Movie S2), indicating ectopic ROP activation in tri-gdi pollen tubes. The compromised polar distribution of active ROPs in tri-gdi pollen tubes indicated impaired intracellular activities mediated by ROP signaling, among which the dynamic polymerization of actin MF is the most extensively studied (Kost et al., 1999; Li et al., 1999; Fu et al., 2001). We therefore analyzed the effect of GDI loss of function on actin MF by introducing ProLAT52:LifeactmEGFP, whose expression faithfully reflected actin MF dynamics in wild-type pollen tubes (Vidali et al., 2009; Qu et al., 2013), i.e. an apical fringe or collar composed of short actin bundles at the cortical apex and longitudinal actin cables in the shank (Fig. 3E). In contrast to the distribution pattern in wild-type pollen tubes, actin MFs were less organized in tri-gdi pollen tubes (Fig. 3F). Extensive short actin bundles were present at the cortical region from the apical PM extending to the shank region away from the apex (Fig. 3F, midsection), suggesting ectopic dynamics of actin MF. Interestingly, tri-gdi pollen tubes were hyposensitive to the actin MFdisrupting drug LatB both in germination percentage (Fig. 3H) and in tube growth (Fig. 3I), further supporting ectopic actin MF dynamics in tri-gdi pollen tubes. However, helical or transverse actin cables in tri844

c

Significantly different from 1:2:1 (x 2, P ,

gdi pollen tubes, although irregularly organized, were distributed only in the shank or base rather than penetrating to the apical region (Fig. 3F, projection). GDIs Were Essential for ROP-Induced Growth in a GTPase-Dependent Way

Metazoan RhoGDIs play a key role in Rho signaling through stabilizing Rho GTPases by protecting their C-terminal hydrophobic regions (Boulter et al., 2010; Garcia-Mata et al., 2011). To determine the influence of GDI loss of function on the stability of ROP GTPases, we analyzed the level of total ROPs using an antibody against ROP2 for its demonstrated specificity (Tao et al., 2002; Xu et al., 2010; Chen et al., 2013; Huang et al., 2013). We found that the level of total ROP2 was reduced by GDI loss of function (Fig. 4A). The reduced level of ROP2 in tri-gdi was not due to transcriptional changes because no significant differences were detected for the transcript levels of either ROP2 (Fig. 4B) or other tip-growth-related ROPs (Supplemental Fig. S5). To determine whether GDI gain of function had an opposite effect on ROP2 level, we generated transgenic plants overexpressing GDI1 in seedlings because GDI1 is constitutively expressed while GDI2 and GDI3 are mostly restricted in pollen (Supplemental Fig. S1). Indeed, overexpressing GDI1 resulted in increased total ROP2 (Fig. 4A). Much reduced signals were detected in rop2-1 (Fig. 4A), a knockout mutant of ROP2 (Jeon et al., 2008), suggesting relatively good specificity of the antibody. Next, we generated stable transgenic plants expressing ProLAT52-driven Arabidopsis ROP5, which is highly expressed in pollen (Kost et al., 1999), or its mutant variants, for which its GDI affinity was abolished (ROP5R69A; Supplemental Figs. S6 and S7), its RopGEF affinity was abolished (ROP5G15V), or both (Supplemental Fig. S8). As expected, overexpression of Plant Physiol. Vol. 170, 2016

GDIs Coordinate Pollen Tube Homeostasis

Figure 2. Pollen tube growth is compromised in the tri-gdi mutant. A and B, Growth of wild-type (A) or tri-gdi pollen tubes (B) in vitro. C and D, Quantitative analysis of pollen tube length (C) and width (D). Results are means 6 SE. Four independent experiments were analyzed. In total, 480 to 500 pollen tubes were used for pollen tube length measurements, while 50 pollen tubes were measured for tube width. Asterisks indicate significant difference (t test, P , 0.01). WT, Wild type. E and F, Growth of wild-type (E) or tri-gdi pollen tubes (F) inside wildtype pistils at 9 HAP, visualized by aniline blue staining. G and H, Growth of wild-type (G) or tri-gdi pollen tubes (H) inside wild-type pistils at 48 HAP by aniline blue staining. Arrowheads point at the front of pollen tube growth. I and J, Close-up of wild-type (I) or tri-gdi pollen tubes (J) inside wild-type pistils at 48 HAP by aniline blue staining. Arrows point at the micropyle where incoming pollen tubes are seen in pistils pollinated with wild-type pollen (I) but not in those pollinated with tri-gdi pollen (J). Bars = 50 mm for A, B, I, and J; 200 mm for E to H.

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Figure 3. ROP activation and ROP-mediated actin MF polymerization in tri-gdi pollen tubes. A and B, Representative in vitro growing pollen tubes from the wild type (A) and the tri-gdi mutant (B) expressing CRIBRIC1-mRFP from the ProLAT52:CRIBRIC1-mRFP transgene. C and D, Schematic illustration of CRIBRIC1 distribution in wild-type (C) or tri-gdi (D) pollen tubes. E and F, Representative confocal fluorescence images of in vitro growing pollen tubes from the wild type (WT; E) and tri-gdi (F) expressing Lifeact-mEGFP from the same transgenic line. In total, 30 pollen tubes were documented for each genotype in four independent experiments. G, Fluorescence intensity of CRIBRIC1 in wild-type and tri-gdi pollen tubes expressing the ProLAT52:CRIBRIC1-mRFP transgene. AU, Arbitrary units of fluorescence intensity. Results are means 6 SE. Four independent experiments and measurements of 60 to 66 pollen grains were performed. Asterisk indicates significant difference (t test, P , 0.01). H and I, Germination percentage (H) and tube width (I) upon LatB treatment. Germination in the presence of DMSO or 3 nM LatB was documented 6 h after incubation. Three independent experiments including 500 to 600 pollen grains were performed. Results shown are means 6 SE. Different letters indicate significant difference (t test, P , 0.01). Bars = 10 mm for A, B, E, and F.

wild-type ROP5 caused tube widening in a dosedependent way (Fig. 5E) due to depolarized growth (Fig. 5A). Fluorescence signals were detected mostly at the apical PM and in the cytoplasm (Fig. 5A). By contrast, overexpression of ROP5R69A did not induce tube depolarization (Fig. 5B) over a wide range of expression levels (Fig. 5E). Fluorescence signals were uniformly distributed along the PM of pollen tubes rather than restricted to the apex (Fig. 5B). Overexpression of ROP5-CA induced isotropic growth of pollen tubes (Fig. 5, C and E), as reported for other ROPs (Kost et al., 1999; Li et al., 1999; Fu et al., 2001). In comparison, a R69A mutation did not abolish the isotropic growth induced by overexpressing 846

the GTP-locked, GEF-independent ROP5G15V (Fig. 5, D and E), indicating that GDI-regulated ROP signaling requires cycles of activation/inactivation, either through intact GTPase activity or through GEF interaction. GDI Loss of Function Impaired the Distribution of Cell Wall Components

The dynamic organization of cell wall contributes to plant cell morphogenesis. While analyzing pollen tube growth in vitro, we noticed that a substantial portion of tri-gdi pollen tubes displayed abnormal accumulation of cell wall components and, in extreme cases, membrane Plant Physiol. Vol. 170, 2016

GDIs Coordinate Pollen Tube Homeostasis

tri-gdi than wild-type pollen tubes, as deduced by the intensity of the JIM7 signal (Fig. 6, D and H). On the other hand, immunofluorescence labeling with JIM5 showed uniform signals along the PM of pollen tubes in both the wild type (Fig. 6E) and tri-gdi (Fig. 6F). However, JIM5-positive signals were substantially elevated at the base of tri-gdi pollen tubes (Fig. 6F). Callose deposition was comparable in wild-type and tri-gdi pollen tubes, i.e. signals were hardly detectable at the apex but strong at the shank and base of pollen tubes (Supplemental Fig. S10). These results showed that GDI loss of function impaired the organized asymmetry of cellulose and pectins in pollen tubes. GDI Loss of Function Interfered With Cellular Homeostasis Balanced by Exocytosis and Endocytosis

Figure 4. GDI loss of function and gain of function resulted in decreased or increased ROP level, respectively. A, Proteins were isolated from seedlings of the wild type (WT), tri-gdi, two independent lines of Pro35S:GDI1, and rop2-1. Protein blots were incubated with an antiROP2 antibody or an anti-ACTIN (ACT) antibody. Levels of total ROP2 (means 6 SE, n = 3) were calculated against the level of total ACTIN in each sample. B, qRT-PCRs detecting the expression of ROP2 in 4 DAG seedlings of the wild type, tri-gdi, two independent lines of Pro35S:GDI1, or rop2-1. GAPDH and TUBULIN2 were used as internal controls. Results shown are means 6 SE (n = 3). Means with the same letters are not significantly different (one-way ANOVA, Tukey-Kramer method, P . 0.05).

invagination at the apex (Supplemental Fig. S9), a phenomenon resulting from impaired cell homeostasis and defined as protoplast trapping (Ischebeck et al., 2008; Sousa et al., 2008; Boisson-Dernier et al., 2013). That observation prompted us to determine whether cell wall deposition was compromised in tri-gdi pollen tubes. Specifically, we used the JIM7 and JIM5 antibodies in immunofluorescence assays to detect highly esterified and de-esterified pectins, respectively, used calcofluor white staining to detect cellulose or polysaccharides, and used decolorized aniline blue staining to detect the distribution of callose (Li et al., 2013a). In wild-type pollen tubes growing in vitro, staining with calcofluor white resulted in distinct signals at the shank region but hardly any at the apex (Fig. 6A), correlating with the role of cellulose in wall rigidity. By contrast, tri-gdi pollen tubes displayed strong signals at the apex and the shank (Fig. 6B), indicating impaired distribution of cellulose. Immunofluorescence with JIM7 resulted in strong signals restricted to the very apical region in wild-type pollen tubes (Fig. 6C), which indicated highly esterified pectins for wall plasticity and extensibility. In comparison, the JIM7 signal was much more expanded in tri-gdi pollen tubes (Fig. 6, D and H). In addition, substantially larger amounts of highly esterified pectins were secreted to the apex of Plant Physiol. Vol. 170, 2016

Dynamic cell wall deposition during pollen tube growth depends on cellular homeostasis maintained by targeted exocytosis and endocytosis. Protoplast trapping occurs in pollen tubes when membrane trafficking is compromised (Szumlanski and Nielsen, 2009) or when PI(4,5)P2 biosynthetic genes are overexpressed (Ischebeck et al., 2008; Sousa et al., 2008), which may have interfered with clathrin-mediated endocytosis (Ischebeck et al., 2013). Therefore, we were curious as to whether the abnormal cell wall deposition in tri-gdi pollen tubes resulted from uncoupled exocytosis and endocytosis. To test that hypothesis, we analyzed the dynamic uptake of the lipophilic fluorescence dye FM4-64 and the distribution of RabA4b in growing pollen tubes. FM4-64 uptake was used to mark endocytic vesicles (Ischebeck et al., 2008; Sousa et al., 2008), whereas the fluorescencefused RabA4s are faithful markers for post-Golgi secretory vesicles in pollen tubes (Szumlanski and Nielsen, 2009; Zhang et al., 2010). As previously documented (Szumlanski and Nielsen, 2009; Zhang et al., 2010), RabA4b formed an inverted cone at the apical clear zone whose central axis overlaid with the growth axis of wildtype pollen tubes (Fig. 7A; Supplemental Movie S3). In growing tri-gdi pollen tubes, the inverted cone labeled by RabA4b was slightly distorted (Fig. 7). Instead, RabA4b labeled a much more expanded apical region without forming an extended trail (Fig. 7B). In tri-gdi pollen tubes that randomly grew out a new axis after a period of growth arrest, RabA4b-positive signals regrouped into an inverted cone whose central region correlated with the new growth axis (Fig. 7B; Supplemental Movie S4). We then pulse-labeled pollen tubes expressing YFPRabA4b with FM4-64 and examined FM4-64 uptake within a short period of time (5–30 min). Internalization of FM4-64 in the wild type was indicated by a gradual increase of apical signals at the apex, leading to an inverted cone-shaped distribution pattern, largely overlapping that of RabA4b (Fig. 7C). In comparison, tri-gdi pollen tubes internalized FM4-64 slightly more slowly. In contrast to an inverted cone with a long substantial tail in wild-type pollen tubes (Fig. 7C), FM4-64 signals were mostly focused at the expanded apex in a portion of 847

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Figure 5. GDIs are essential for ROP5-induced pollen tube growth in a GTPase-dependent way. A to D, Representative in vitro growing pollen tubes expressing RFP-ROP5 (A), RFP-ROP5R69A (B), RFP-ROP5CA (C), or RFP-ROP5CAR69A (D). Right images are merges of fluorescence and brightfield images. E and F, Scatter plots showing the correlation of transgene expression levels and pollen tube width. The signal intensity of pollen tubes was defined as the fluorescence intensity of apical to subapical PM. a.u., Arbitrary units of fluorescence intensity. Bar = 25 mm.

tri-gdi pollen tubes, in a way overlapping with the pattern of RabA4b (Fig. 7E). In another substantial portion of trigdi pollen tubes, internalization of FM4-64 was substantially reduced despite the spread-out signals of RabA4b at the apical region (Fig. 7D). These results suggested that although tri-gdi pollen tubes retained the activities of exocytosis and endocytosis, they were defective in balancing exocytosis with endocytosis. The Polar Distribution of PI(4,5)P2 in tri-gdi Pollen Tubes

The impaired cell wall deposition and altered vesicle trafficking patterns suggested that GDIs were important for cellular homeostasis through ROP signaling. We next explored the possibility that GDI loss of function might have affected the distribution of PI(4,5)P2 for 848

a few reasons. First, the dynamic localization of PI(4,5) P2 at the apical PM of pollen tubes (Kost et al., 1999; Dowd et al., 2006; Helling et al., 2006; Ischebeck et al., 2008; Sousa et al., 2008; Ischebeck et al., 2011) was proposed to associate with ROP activity (Kost et al., 1999; Ischebeck et al., 2011). Second, overexpressing PI(4,5)P2 biosynthetic genes caused massive apical pectin deposition and PM invagination (Ischebeck et al., 2008; Sousa et al., 2008), similar to that caused by GDI loss of function. Third, PI(4,5)P 2 regulates clathrin-mediated endocytosis in plant cells (Ischebeck et al., 2013), the mechanism by which FM4-64 internalizes. Finally, as in yeast and metazoans, PI(4,5)P2 may act as a displacement factor for GDI-mediated ROP recycling in pollen tubes (Ischebeck et al., 2011). Plant Physiol. Vol. 170, 2016

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Figure 6. Differential deposition of cell wall components in wild-type and tri-gdi pollen tubes. A and B, Confocal fluorescence images showing representative wild-type (A) and tri-gdi (B) pollen tubes stained with calcofluor white, indicating the deposition of cellulose. Arrowheads point at the apex. C and D, Immunofluorescence labeling of esterified pectins in wild-type (C) and tri-gdi (D) pollen tubes using the JIM7 antibody. C and D show images taken at the same gain value. Arrowheads point at the region flanking the fluorescence signals. E and F, Immunofluorescence labeling of de-esterified pectins in wild-type (E) and tri-gdi (F) pollen tubes using the JIM5 antibody. E and F show images taken at the same gain value. G and H, Quantitative analysis of cellulose distribution from calcofluor white staining (G) or esterified pectins with JIM7 immunofluorescence staining (H) in wild-type (WT) and tri-gdi pollen tubes. Both fluorescence and immunofluorescence labeling were performed in three independent experiments involving 20 to 30 pollen tubes. Results shown are means 6 SE. Asterisks indicate significant difference (t test, P , 0.01). Fluorescence images are merged with their corresponding bright-field images. Bars = 10 mm for A to F.

To determine whether the polar distribution of PI (4,5)P2 was impaired in tri-gdi pollen tubes, we needed a valid biosensor for PI(4,5)P2 whose expression did not affect tube growth on its own. Previous studies often used the PH domain of PLCd1 (PHPLCd1) as the PI(4,5)P2 sensor (Kost et al., 1999; Dowd et al., 2006; Helling et al., 2006). However, strong expression of PHPLCd1 disturbed pollen tube growth likely by sequestering endogenous PI(4,5)P2 (Kost et al., 1999; Dowd et al., 2006; Helling et al., 2006). We thus decided to use a recently reported Plant Physiol. Vol. 170, 2016

Pro UBQ10:CITRINE-TUBBY-C transgenic line (P15Y) whose mild expression and specificity make it a good biosensor for PI(4,5)P2 in planta (Simon et al., 2014). In wild-type pollen tubes growing in vitro, fluorescence signals were exclusively associated with the apical PM and slightly weak at the very apex (Fig. 8A), as reported previously (Dowd et al., 2006; Helling et al., 2006; Ischebeck et al., 2008; Sousa et al., 2008). In comparison, PI(4,5)P2 was still restricted to the apical PM of tri-gdi pollen tubes (Fig. 8B). However, its distribution was 849

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Figure 7. RabA4b-positive post-Golgi secretion and FM4-64 uptake in wild-type or tri-gdi pollen tubes. A and B, Distribution of RabA4b-positive post-Golgi vesicles in growing pollen tubes of the wild type (A) and tri-gdi (B). Images were captured using time-lapse confocal fluorescence microscopy. Left-most is the corresponding brightfield image. In total, 24 pollen tubes were documented for each in three independent experiments. C to E, Uptake of FM4-64 (red) in wild-type (C) and tri-gdi (D and E) pollen tubes expressing YFP-RabA4b (green). Bright-field and fluorescence merged images are next to their corresponding fluorescence images. The incidence of fluorescence distribution patterns is listed at the bottom of the corresponding images. Experiments were performed in three independent replicates involving 35 to 67 pollen tubes. Bars = 10 mm.

significantly expanded (Fig. 8E). Instead of showing less intense signals at the very apex as in wild-type pollen tubes (Fig. 8A), PI(4,5)P2 was distributed at the very apex as the signal was as strong as the apical flank (Fig. 8B), likely resulting from the slower growth of trigdi pollen tubes. Because PI(4,5)P2 regulates clathrin-mediated endocytosis (Ischebeck et al., 2013), we analyzed the distribution of clathrin-coated vesicles by immunofluorescence 850

labeling with an antibody against clathrin light chain (CLC; Wang et al., 2013a). CLC signals were detected along the PM of pollen tubes as puncta, in addition to cytoplasmic signals, and were concentrated more at the apical than lateral regions in wild-type pollen tubes (Fig. 8C). Similar to the pattern for PI(4,5)P2, CLC signals were less intense at the very apex in growing wild-type pollen tubes (Fig. 8A). In comparison, tri-gdi pollen tubes also showed enhanced accumulation of CLC signals at the Plant Physiol. Vol. 170, 2016

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apical region (Fig. 8D), forming a crescent at the apical PM (Fig. 8D) and likely correlating with the expanded distribution of PI(4,5)P2. The polar distribution of PI(4,5) P2 and CLC in tri-gdi pollen tubes indicated that GDI loss of function affected but did not impair clathrin-mediated endocytosis. GDI and PI(4,5)P2 Coordinate Cellular Homeostasis During Pollen Tube Growth

GDI loss of function resulted in enhanced exocytosis but did not compromise PI(4,5)P2-dependent clathrinmediated endocytosis, suggesting that ROP-mediated cellular homeostasis depends on antagonistic interactions between GDIs and PI(4,5)P2, as proposed earlier (Ischebeck et al., 2011). We thus took several approaches to provide further evidence for this hypothesis. First, we applied the pharmacological drug U-73122 to wild-type (Fig. 9, A–C) and tri-gdi pollen tubes (Fig. 9, D–F) and analyzed their growth dynamics. U-73122

specifically inhibits the activity of phospholipase C (PLC) and thus induces ectopic distribution of PI(4,5)P2 at the PM of pollen tubes (Dowd et al., 2006; Helling et al., 2006). Application of U-73122 at low concentrations inhibited the germination of wild-type pollen (Fig. 9G) and induced significant tube bulging (Fig. 9H), indicating impaired polarity. By contrast, the germination of tri-gdi pollen was hyposensitive to U-73122; U-73122 at 0.5 mM completely suppressed wild-type germination, whereas it had only a slight effect on tri-gdi (Fig. 9G). Also, U-73122 did not substantially induce ectopic distribution of PI(4,5)P2 in tri-gdi pollen tubes (Fig. 9, E and F), whereas it did in the wild type (Fig. 9, B and C). Second, we reasoned that if GDIs and PI(4,5)P2 antagonistically regulate cellular homeostasis, GDI gain-offunction pollen tubes would resemble tubes containing reduced PI(4,5)P2. We thus generated stable transgenic Arabidopsis plants expressing ProLAT52:RFP-GDI1 or RFP-GDI3. Overexpression of GDIs dramatically reduced pollen germination percentage (Supplemental Figure 8. The distribution of PI(4,5)P2 and clathrincoated vesicles was altered but not impaired by GDI loss of function. A and B, Representative in vitro growing pollen tubes from the wild type (A) and tri-gdi (B) expressing CITRINE-TUBBY-C (CLC) from the ProUBQ10:CITRINE-TUBBY-C transgene. C and D, Immunofluorescence labeling of CLC of representative in vitro growing pollen tubes from the wild type (C) and tri-gdi (D). Merges of brightfield and fluorescence images are below the corresponding fluorescence image. E, Distribution of PI(4,5)P2 as indicated by fluorescence intensity of CITRINE-TUBBY-C. a.u., Arbitrary units of fluorescence intensity. Results shown are means 6 SE from three independent replicates involving 32 to 36 pollen tubes. Asterisk indicates significant difference (t test, P , 0.05). WT, Wild type. Bars = 10 mm.

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Fig. S11), the opposite to tri-gdi (Fig. 2; Supplemental Fig. S4) but similar to mutants defective in production of PI (4,5)P2 (Ischebeck et al., 2008; Sousa et al., 2008). Third, we transiently expressed GDI1 and Arabidopsis PIP5K11 to gain further proof using the tobacco pollen in vitro system, which has been widely adopted in Arabidopsis pollen biology (Cheung et al., 2003; Zhang and McCormick, 2007; Ischebeck et al., 2008; Hwang et al., 2010; Ischebeck et al., 2011). Similar to what was reported (Ischebeck et al., 2011), the tube bulging phenotype induced by PIP5K11 overexpression was suppressed by coexpressed GDI1 (Supplemental Fig. S12). GDI1 also suppressed the disturbed post-Golgi secretion seen in PIP5K11 overexpression, as indicated by the distribution of RabA4b (Supplemental Fig. S12). DISCUSSION GDIs Are Critical for Male Gametophytic Function

Compared to root hairs, which also grow by tip growth, pollen tubes seem to have high demand for GDI-mediated ROP signaling since GDI2 and GDI3 are highly enriched in pollen tubes (Supplemental Fig. S1). A likely explanation is that the tip growth of pollen tubes is more demanding than root hair growth, possibly due to the necessity of pollen tubes to perceive female cues and change growth axis rapidly. Ectopic expression of Arabidopsis ROPs (Cheung et al., 2003) or functional loss of maize ROP2 (Arthur et al., 2003) compromised male transmission, suggesting fine regulation of ROP activity. However, despite extensive studies on ROPs, their roles during in vivo growth were scarce due to high redundancy (Yang, 2002). Thus, the stable GDI loss-of-function line analyzed in this study provides the opportunity to dissect the contribution of ROP signaling during directional pollen tube growth not only in vitro but also in vivo (Fig. 2). We found that trigdi pollen tubes were stubby when germinated in vitro, exhibited severely reduced growth inside the transmitting tract, and failed to respond to funicular guidance cues (Fig. 2). Because its funicular guidance was already defective, it is difficult to know whether micropylar guidance of tri-gdi pollen tubes was also impaired. These growth defects resulted in significantly reduced male transmission (Table I) and fertility (Fig. 2). Because ROPs are involved in multiple developmental processes such as hormone signaling, root growth, and cell differentiation (Yang, 2002; Nibau et al., 2006), it was somehow surprising that plants with all GDIs mutated still managed to complete growth without noticeable developmental defects (Fig. 1). One likely scenario is that there are other factors playing similar roles to GDIs in ROP signaling, as is the case in other phyla in which caveolin-1, a protein critical for lipid raft-mediated membrane dynamics, functions as a GDI for Rac GTPases (Boulter et al., 2010; Garcia-Mata et al., 2011). There are other possibilities. The tri-gdi mutant may have other developmental or cellular defects that could be detected only upon closer scrutiny. In 852

Figure 9. Functional loss of GDIs reduced sensitivity to U-73122 in pollen germination and tube growth. A to C, A representative P15Y pollen tube expressing ProUBQ10:CITRINE-TUBBY-C treated with DMSO (A) or with 0.5 mM U-73122 for 1 h (B) or 1.5 h (C). D to F, A representative P15Y;tri-gdi pollen tube expressing Pro UBQ10 :CITRINETUBBY-C treated with DMSO (D) or with 0.5 mM U-73122 for 1 h (E) or 1.5 h (F). G, Pollen germination percentage of the wild type (WT) and trigdi upon treatment with U-73122 at different concentrations for 4 h. H, Pollen tube width of the wild type and tri-gdi upon 0.5 mM U-73122 treatment for 1 h. Results shown in G and H are means 6 SE from three independent replicates involving 400 to 500 pollen grains (G) or 32 to 36 pollen tubes (H). Different letters in H indicate significant difference (t test, P , 0.05). Bars = 10 mm.

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addition, although the GDI triple mutant behaved normally under optimal growth conditions, it might show defects when environmental conditions are not favorable or when competition for resources is intense, a plausible scenario worthy of further exploration. Dual Role of GDIs in ROP-Mediated Pollen Tube Growth

Being evolutionarily conserved components, GDIs play inhibitory roles in Rho signaling by membrane extraction and cytoplasmic sequestration (Boulter et al., 2010; Garcia-Mata et al., 2011). As reported in tobacco (Klahre et al., 2006), overexpression of GDIs in Arabidopsis pollen tubes reduced pollen germination and tube growth (Supplemental Fig. S11). Functional loss of GDIs caused expansion of ROP-GTP to a uniform rather than a restricted distribution pattern at the PM of pollen tubes (Fig. 3). In addition, expression of a mutant ROP that is unable to interact with GDIs (Supplemental Figs. S6 and S7) showed ectopic distribution along the pollen tube PM rather than being restricted to the apical PM (Fig. 5), supporting an inhibitory role of GDIs in ROP signaling by extracting ROPs from the lateral PM of pollen tubes. On the other hand, several lines of evidence support a positive role of GDIs in ROP signaling during pollen tube growth. First, the ability of ROP5 to induce depolarized growth was abolished by a R69A mutation (Fig. 5) that interfered with its binding to GDIs (Supplemental Figs. S6 and S7). However, GDI dependency of ROP signaling relies on active GTP-GDP cycle or GEF interaction (Fig. 5). It was previously reported that overexpression of GDIs suppresses tube depolarization caused by wild-type ROPs but not that caused by constitutive active ROPs (Klahre et al., 2006; Hwang et al., 2010). Indeed, the ability of ROPs to bind to GDIs is not necessary for induced isotropic growth of pollen tubes as long as ROPs are in GTP-locked, GEF-independent forms (Fig. 5; Supplemental Fig. S8). Interestingly, similar mechanisms have been demonstrated in yeast and metazoans (Lin et al., 2003; Slaughter et al., 2009; Freisinger et al., 2013), suggesting an evolutionarily recurring paradigm. Second, the level of total ROP2 was reduced without transcriptional changes (Fig. 4) that may result in dampened ROP signaling. Third, dynamic actin MF in tri-gdi was affected in a way similar to that caused by dominant negative ROP expression, i.e. actin cables rarely penetrated to the apex and were less organized (Fig. 3). GDI and PI(4,5)P2 May Antagonistically Mediate Cellular Homeostasis During Pollen Tube Growth

Dynamic cell wall organization is critical for plant tip growth in which plasticity and extensibility ensure growth at the apex while mechanical strength maintains the shape of the distal region (Zonia and Munnik, 2009). Targeted exocytosis constitutively deposits building materials for tip growth while active endocytosis not only retrieves excess materials deposited from exocytosis but also controls the dynamic spatial distribution of signaling components. Although direct evidence is scarce as to Plant Physiol. Vol. 170, 2016

whether and how ROPs regulate post-Golgi secretion in pollen tubes (Lee et al., 2008), studies on vegetative tissues supported a key role of ROPs in post-Golgi secretion (Bloch et al., 2005; Lavy et al., 2007). For example, the ROP effector ICR1 interacts with SEC3 (Lavy et al., 2007), a component of the exocyst critical for polarized secretion (Hála et al., 2008; Hazak et al., 2010). In addition, ROPmediated dynamic polymerization of actin MF (Fu et al., 2001; Gu et al., 2005) coordinates vesicle targeting during pollen tube growth (Lee et al., 2008; Zhang et al., 2010). As a key polarity factor, PI(4,5)P2 is restricted to the apical PM of growing pollen tubes (Kost et al., 1999; Dowd et al., 2006; Helling et al., 2006; Ischebeck et al., 2008; Sousa et al., 2008; Ischebeck et al., 2011). The dynamic asymmetry of PI(4,5)P2 distribution depends on antagonistic activities of PI4P5Ks (Ischebeck et al., 2008; Sousa et al., 2008; Ischebeck et al., 2011) and PLCs (Dowd et al., 2006; Helling et al., 2006). The delivery of PI4P via the trans-Golgi network/early endosome to the PM may also contribute to PI(4,5)P2 accumulation by providing substrates for PI4P5Ks (van Leeuwen et al., 2004; Boss and Im, 2012). Overexpression of a dominant negative PLC (PLC-DN; Dowd et al., 2006; Helling et al., 2006) or overexpression of PI4P5Ks (Ischebeck et al., 2008; Sousa et al., 2008; Ischebeck et al., 2011) resulted in depolarized pollen tube growth, accompanied by ectopic PI(4,5)P2 distribution along the PM. Overexpression of PLC-DN, which caused ectopic distribution of PI(4,5)P2, resulted in the formation of transverse actin cables at the pollen tube apex (Dowd et al., 2006), similar to that seen with constitutive active ROPs (Kost et al., 1999; Fu et al., 2001). It was recently reported that PI(4,5)P2 is critical for clathrin-mediated endocytosis (Ischebeck et al., 2013), suggesting its role in maintaining cellular homeostasis. We report here that GDI loss of function resulted in ectopic ROP activation (Fig. 3) but did not impair PI(4,5)P2 or clathrin-mediated endocytosis (Figs. 7 and 8). GDI loss of function enhanced the potential of pollen germination (Supplemental Fig. S4), during which the proportion of GTP-ROP increases (Chen et al., 2013). By contrast, mutants defective in PI4P5Ks (Ischebeck et al., 2008; Sousa et al., 2008) and overexpressing GDIs (Supplemental Fig. S11) reduced pollen germination. In addition, germination and tube growth of tri-gdi pollen are hyposensitive to LatB (Fig. 3), whereas PI4P5K loss-of-function pollen tubes were hypersensitive to LatB (Ischebeck et al., 2011). Furthermore, GDIs restored the RabA4b-positive postGolgi secretion in pollen tubes overexpressing PIP5Ks (Supplemental Fig. S12). These results provide more evidence supporting the hypothesis that GDIs and PI(4,5)P2 antagonistically act in ROP signaling to maintain cellular homeostasis (Ischebeck et al., 2008; Sousa et al., 2008; Szumlanski and Nielsen, 2009; Ischebeck et al., 2011).

MATERIALS AND METHODS Plant Materials and Growth Conditions The T-DNA insertion lines for GDI2 (FLAG_184A02) and GDI3 (FLAG_259E06) were obtained from INRA FLAG collections (www.ijpb-versailles/inra/flag). The 853

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T-DNA insertion line rop2-1 (SALK_055328C) as described previously (Jeon et al., 2008) was obtained from the Arabidopsis Biological Resource Center (ABRC). Other materials, i.e. scn1-1 (Carol et al., 2005), P15Y (Simon et al., 2014), ProLAT52: CRIBRIC1-RFP (Zhao et al., 2013), ProLAT52:YFP-RabA4b (Zhang et al., 2010), and ProLAT52:Lifeact-mEGFP (Qu et al., 2013), were described previously. The Wassilewskija ecotype was used as the wild type. Plant growth, transformation, and selection were as described (Zhou et al., 2013).

RNA Extraction, RT-PCR, and qRT-PCR Total RNAs were isolated from either open flower (for RT-PCR analysis) or 4 d after germination (DAG) seedlings (for qRT-PCRs) using a Qiagen RNeasy plant miniprep kit according to the manufacturer’s instructions. Oligo(dT)primed cDNAs were synthesized using Superscript III reverse transcriptase with on-column DNase digestion (Invitrogen). Primers used in RT-PCRs were ZP337/ZP338 for gdi2, ZP483/ZP2377 for gdi3, and ZP16/ZP17 for ACTIN2. The qRT-PCRs of ROPs were performed with the Bio-Rad CFX96 real-time system using SYBR Green real-time PCR master mix (Toyobo) as described (Zhou et al., 2013). Primers used in qRT-PCRs were specifically designed to nondiscriminatively amplify ROP1/ROP3/ROP5 (ZP2713/ZP2714) or ROP2/ ROP4/ROP6 (ZP2715/ZP2716). Primers for GAPDH and TUBULIN2 in qRTPCRs were as described (Zhou et al., 2013). All primers are listed in Supplemental Table S1. Genotypic verification of scn1-1 in the double and triple GDI mutant backgrounds was performed by examining root hair defect (Carol et al., 2005) together with sequencing.

Plasmid Construction All constructs were generated using the Gateway technology (Invitrogen). All entry vectors were generated in the pENTR/D/TOPO vector (Invitrogen). Promoters for GDI2 and GDI3 were cloned with primers ZP425/ZP426 and ZP427/ZP428 from Columbia-0 genomic DNA. ProGDI2 or ProGDI3, containing 1415 bp or 1196 bp sequence upstream of their corresponding start codon, was introduced into a destination vector (GW:NLS-YFP) described earlier (Wang et al., 2013b). Wild-type coding sequences were cloned with the following primer pairs: ZP945/ZP946 for ROP5, ZP1990/ZP1992 for PRONE1RopGEF1, ZP784/ZP785 for GDI1, and ZP2499/ZP2500 for REN1. The entry vector for ROP2 (Huang et al., 2013) and for GDI3 (Li et al., 2013b) was described. Mutations of ROP2 or ROP5 were generated using the corresponding entry vectors as the templates using Phusion site-directed mutagenesis kit. The destination vector for pollen-specific expression was described earlier (Zhang and McCormick, 2007). The destination vectors for Y2H, pDEST32 and pDEST22 for BD- and AD-fusions, respectively, were purchased from Invitrogen. The destination vectors for BiFC, pSITE-nEYFP-C1 and pSITE-cEYFP-C1 (Citovsky et al., 2008), were obtained from the ABRC. Expression vectors were generated by LR reactions using LR Clonase II (Invitrogen). Expression vectors, including ProLAT52:CRIBRIC1-mRFP, ProLAT52:YFP-RabA4b, and ProLAT52:RFP-mTalin, were described earlier (Zhang et al., 2010; Zhao et al., 2013). All PCR amplifications were performed using Phusion hot start high-fidelity DNA polymerase with the annealing temperature and extension times recommended by the manufacturer (Finnzyme). All entry vectors were sequenced, and sequences were analyzed using Vector NTI (Invitrogen). The Bioneer PCR purification kit and the Bioneer Spin miniprep kit were used for PCR product recovery and plasmid DNA extraction, respectively. All primers are listed in Supplemental Table S1, and related vectors are listed in Supplemental Table S2.

Protein Interaction Assays Y2H was performed with the ProQuest system (Invitrogen). YSD media supplemented with all amino acids except Trp and Leu (2WL) were used to select diploids. YSD media, supplemented with 10 mM 3-amino-1,2,4-triazole and all amino acids except Trp, Leu, and His (2WLH + 10 mM 3AT), were used to select colonies showing bait-prey interactions. BiFC was performed by tobacco infiltration as described (Huang et al., 2013). Each infiltration was performed in eight duplicates. Confocal imaging was performed 48 h after infiltration.

Pollen Development, Germination, Growth, Transformation, and Pharmacological Treatments Pollen analyses, including scanning electron microscopy, Alexander staining, and 49,6-diamidino-2-phenylindole staining of mature pollen, pollen germination, and growth in vitro, as well as aniline blue staining, were performed as 854

described (Johnson-Brousseau and McCormick, 2004; Li et al., 2013a). For the sensitivity of pollen germination to LatB, LatB at a final concentration of 3 nM was added in pollen germination medium. Dimethyl sulfoxide (DMSO) at the same dilution ratio was added in pollen germination medium as controls. Germination percentage was measured at different time points after incubation. For the sensitivity of pollen tube growth to LatB, LatB at a final concentration of 3 nM in liquid germination medium was dropped on in vitro growing pollen tubes 2.5 h after incubation. Images were captured after 1 h incubation with either LatB or DMSO at the same dilution. Transient expression of constructs in tobacco pollen by particle bombardment was performed as described (Zhang and McCormick, 2007). For the sensitivity of pollen germination to U-73122, U-73122 at different concentrations (0.25 mM, 0.5 mM, 1 mM, and 5 mM) was added in pollen germination medium. DMSO at the same dilution was added as controls. For the sensitivity of pollen tube growth to U-73122, 0.5 mM U-73122 in liquid germination medium was dropped on in vitro growing pollen tubes 2.5 h after incubation. Images were captured after 1 h incubation with either U-73122 or DMSO at the same dilution.

Fluorescence, Immunofluorescence, and Confocal Microscopy For double labeling experiments, FM4-64 at a final concentration of 4 mM in liquid germination medium was dropped on in vitro growing pollen tubes 2.5 h after incubation. Image was collected after 15 min incubation at 28°C. Cellulose staining by calcofluor white, callose staining by aniline blue, and immunolabeling of different pectin species with JIM5 and JIM7 antibodies were as described (Li et al., 2013a). Microscopic imaging was performed using either an Axio Observer D1 microscope (Zeiss; www.zeiss.com) with epifluorescence optics equipped with a CCD camera or by confocal imaging using a Leica TCS SP5 confocal laser-scanning microscope (Leica) with a 488-nm argon laser/BP 505-550 filter for GFP and a 561-nm laser/BP 600-650 filter for RFP. Images were exported and processed using Adobe Photoshop CS3.

Biochemical Assays of Total ROP2 Analyses of total ROP2 were performed as described (Tao et al., 2002; Xu et al., 2010; Chen et al., 2013; Huang et al., 2013). Briefly, about 500 mg seedlings of 4 DAG were pulverized in liquid nitrogen, suspended in 500 mL of pull-down buffer (40 mM Tris-HCl, pH 7.5, 100 mM NaCl, 1 mM Na2-EDTA, 5% glycerol, 0.75% Triton X-100, 1 mM phenylmethylsulfonyl fluoride, and 1% proteinase inhibitor cocktail; Sigma), centrifuging three times for 15 min at 13,000 rpm at 4°C. About 40 mL of supernatant was used to check protein concentration. Ten microliters of total proteins and eluted protein/beads mix were applied for SDS-PAGE for immunodetection of ROP2 with anti-ROP2 antibody (Abiocode Uniprot ID Q38919; Catalog No. R2165-2). The mouse monoclonal antibody for plant ACTIN was purchased from Abmart (M20009).

Accession Numbers Arabidopsis Genome Initiative locus identifiers for the genes mentioned in this article are as follows: SCN1/GDI1, At3g07880; GDI2, At1g62450; GDI3, At1g12070; RabA4d, At3g12160; RIC1, At2g33460; ROP2, At1g20090; ROP5, At4g35950; RopGEF1, At4g38430; REN1, At4g24580; and PIP5K11, At1g01460.

Supplemental Data The following supplemental materials are available. Supplemental Figure S1. GDI2 and GDI3 are enriched in mature pollen and pollen tubes. Supplemental Figure S2. The partial GDI3 transcript produced in gdi3 did not encode a functional GDI3. Supplemental Figure S3. Length and width of pollen tubes growing in vitro. Supplemental Figure S4. Pollen development and pollen germination percentage are comparable between the wild type and tri-gdi mutant. Supplemental Figure S5. The collective transcript level of type I ROPs is comparable between the wild type and tri-gdi mutant. Supplemental Figure S6. Interaction of SCN1 with ROPs by Y2H analysis. Plant Physiol. Vol. 170, 2016

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Supplemental Figure S7. Interaction of SCN1 with ROPs by BiFC analysis. Supplemental Figure S8. ROP GTPases but not their constitutive active forms interact with the PRONE domain of RopGEF1 by BiFC analyses. Supplemental Figure S9. Abnormal accumulation of cell wall components and membrane invagination at the apex of tri-gdi pollen tubes. Supplemental Figure S10. Deposition of callose was comparable between wild-type and tri-gdi pollen tubes. Supplemental Figure S11. Overexpression of GDI3 reduced pollen germination and tube growth. Supplemental Figure S12. Coexpression of GDI1 suppressed the disrupted RabA4b distribution induced by overexpressing PIP5K11 in pollen tubes. Supplemental Movie S1. Dynamic distribution of CRIBRIC1-mRFP in a wild-type pollen tube. Supplemental Movie S2. Dynamic distribution of CRIBRIC1-mRFP in a trigdi pollen tube. Supplemental Movie S3. Dynamic distribution of YFP-RabA4b in a wildtype pollen tube. Supplemental Movie S4. Dynamic distribution of YFP-RabA4b in a growing tri-gdi pollen tube. Supplemental Table S1. Oligos used in this study. Supplemental Table S2. Vectors used in this study.

ACKNOWLEDGMENTS We thank Profs. Liam Dolan for the scn1-1 seeds, Yvon Jaillais for the PI(4,5) P2 sensor line P15Y, Jian-Wei Pan for the CLC antibody, Shan-Jin Huang for the ProLAT52:Lifeact-mEGFP transgenic seeds, Sheila McCormick for critical reading and language editing of this article, and Xian Sheng Zhang for giving us access to the microscope facilities of his laboratory, and INRA and the ABRC for other plant materials or vectors. The authors declare that there is no conflict of interest. Received October 13, 2015; accepted December 9, 2015; published December 11, 2015.

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