The Arabidopsis Rho of plants GTPase AtROP6 functions in developmental and pathogen response pathways.

How plants coordinate developmental processes and environmental stress responses is a pressing question. Here, we show that Arabidopsis (Arabidopsis thaliana) Rho of Plants6 (AtROP6) integrates developmental and pathogen response signaling. AtROP6 expression is induced by auxin and detected in the root meristem, lateral root initials, and leaf hydathodes. Plants expressing a dominant negative AtROP6 (rop6(DN)) under the regulation of its endogenous promoter are small and have multiple inflorescence stems, twisted leaves, deformed leaf epidermis pavement cells, and differentially organized cytoskeleton. Microarray analyses of rop6(DN) plants revealed that major changes in gene expression are associated with constitutive salicylic acid (SA)-mediated defense responses. In agreement, their free and total SA levels resembled those of wild-type plants inoculated with a virulent powdery mildew pathogen. The constitutive SA-associated response in rop6(DN) was suppressed in mutant backgrounds defective in SA signaling (nonexpresser of PR genes1 [npr1]) or biosynthesis (salicylic acid induction deficient2 [sid2]). However, the rop6(DN) npr1 and rop6(DN) sid2 double mutants retained the aberrant developmental phenotypes, indicating that the constitutive SA response can be uncoupled from ROP function(s) in development. rop6(DN) plants exhibited enhanced preinvasive defense responses to a host-adapted virulent powdery mildew fungus but were impaired in preinvasive defenses upon inoculation with a nonadapted powdery mildew. The host-adapted powdery mildew had a reduced reproductive fitness on rop6(DN) plants, which was retained in mutant backgrounds defective in SA biosynthesis or signaling. Our findings indicate that both the morphological aberrations and altered sensitivity to powdery mildews of rop6(DN) plants result from perturbations that are independent from the SA-associated response. These perturbations uncouple SA-dependent defense signaling from disease resistance execution.

Rho of Plants (ROPs), also known as RACs (for clarity, the ROP nomenclature will be used throughout this article), comprise a plant-specific group of Rho family small G proteins. Like other members of the Ras superfamily of small G proteins, ROPs function as molecular switches, existing in a GTP-bound "on" state and a GDP-bound "off" state. In the GTP-bound state, ROPs interact with specific effectors that transduce downstream signaling or function as scaffolds for interaction with additional effector molecules (Berken and Wittinghofer, 2008). Conserved point mutations in the G1 (P loop) Gly-15 or the G3 (switch II) Gln-64, which abolish GTP hydrolysis, or the G1 Thr-20 or G4 Asp-121 that compromise GDP/GTP exchange, can form either constitutively active or dominant negative mutants, respectively (Feig, 1999;Berken et al., 2005;Berken and Wittinghofer, 2008;Sorek et al., 2010). Primarily based on studies with neomorphic mutants, ROPs have been implicated in the regulation of cytoskeleton organization and dynamics, vesicle trafficking, auxin transport and response, abscisic acid (ABA) response, and response to pathogens (Nibau et al., 2006;Yalovsky et al., 2008;Yang, 2008;Lorek et al., 2010;Wu et al., 2011;and refs. therein).
In Arabidopsis (Arabidopsis thaliana), there are 11 ROP proteins (Winge et al., 1997). Assigning specific functions to individual members of this family is difficult, however, because ROPs are functionally redundant. A ROP10 loss-of-function mutant was reported to be ABA hypersensitive (Zheng et al., 2002), displaying enhanced expression of tens of genes in response to ABA treatments (Xin et al., 2005). However, in the absence of exogenous ABA, gene expression in the rop10 mutant was similar to that in wild-type plants (Xin et al., 2005). Loss of leaf epidermis pavement cell polarity was reported for rop4 rop2-RNAi (for RNA interference) double mutant plants (Fu et al., 2005). Mild changes in pavement and hypocotyl cell structure and microtubule (MT) organization were reported for a rop6 loss-of-function mutant (Fu et al., 2009).
The involvement of ROPs in auxin-regulated development has been addressed in several studies (Wu et al., 2011). Ectopic expression of a dominant negative ROP2 (rop2 DN ) mutant under regulation of the 35S promoter resulted in a loss of apical dominance and a reduction in the number of lateral roots. In contrast, ectopic expression of constitutively active ROP2 (rop2 CA ) caused an increase in the number of lateral roots and an enhanced decrease in primary root length in response to auxin. Consistent with these findings, the expression of a constitutively active NtRAC1 in tobacco (Nicotiana tabacum) protoplasts induced the expression of auxin-regulated genes in the absence of auxin and promoted the formation of protein nuclear bodies containing components of the proteasome and COP9 signalosome (Tao et al., 2002(Tao et al., , 2005Wu et al., 2011). The ROP effector ICR1 (for interactor of constitutively active ROP1) regulates polarized secretion and is required for polar auxin transport (Lavy et al., 2007;Bloch et al., 2008;. In the root, local auxin gradients induce the accumulation of ROPs in trichoblasts at the site of future root hair formation (Fischer et al., 2006). Recently, it was shown that interdigitation of leaf epidermis pavement cells depends on Auxin-Binding Protein1 (ABP1)-mediated ROP activation (Xu et al., 2010). Taken together, these data indicate that ROPs are involved in both mediating the auxin response and facilitating directional auxin transport. It is still unclear, however, which ROPs function in these processes.
ROP function was linked to plant defense responses in several studies. In rice (Oryza sativa), OsRAC1 is a positive regulator of the hypersensitive response, possibly through interactions with the NADPH oxidase RbohB, Required for Mla12 Resistance, and Heat Shock Protein90 (Ono et al., 2001;Thao et al., 2007;Wong et al., 2007). Interestingly, other members of the rice ROP family, namely RAC4 and RAC5, are negative regulators of resistance to the rice blast pathogen Magnaporthe grisea . Similar to rice, when expressed in tobacco, dominant negative OsRAC1 suppressed the hypersensitive response (Moeder et al., 2005). In barley (Hordeum vulgare), several constitutively active ROP/RAC mutants and a MT-associated ROPGAP1 loss-of-function mutant enhanced susceptibility to the powdery mildew Blumeria graminis f. sp. hordei (Bgh). The activated ROP-enhanced susceptibility to Bgh was attributed to disorganization of the actin cytoskeleton and was shown to depend on Mildew Resistance Locus O (MLO; Schultheiss et al., 2002Schultheiss et al., , 2003Opalski et al., 2005;Hoefle et al., 2011). In barley, three ROP proteins, HvRACB, HvRAC1, and HvRAC3, were linked to both development and pathogen response Pathuri et al., 2008;Hoefle et al., 2011).
We have analyzed the function of the Arabidopsis AtROP6 (ROP6) by characterizing its expression pattern and its regulation by auxin and the phenotype of plants that express rop6 DN under the regulation of its endogenous promoter. The utilization of the dominant negative mutant overcame functional redundancy, while expression under the regulation of the endogenous promoter enabled the analysis of ROP6 function in a developmental context. Phenotypic and gene expression analyses indicate that ROP6 functions in developmental, salicylic acid (SA)-dependent, and SAindependent defense response pathways.

ROP6 Expression Pattern and Subcellular Localization
Previously, we analyzed lipid modifications, membrane interaction dynamics, and the function of ROP6 in the regulation of cell polarity using the constitutively active rop6 CA mutant expressed under the regulation of the cauliflower mosaic virus 35S promoter. Because of the functional redundancy of ROPs, the rop6 mutant phenotype is inseparable from that of wild-type plants. To overcome this difficulty and elucidate the specific function of ROP6, its promoter was isolated (Supplemental Fig. S1) and subcloned into the LhG4/pOp transcription transactivation system (Moore et al., 1998) to drive the expression of GUS, DsRed, GFP-ROP6, GFP-rop6 CA , and GFP-rop6 DN . To verify the expression pattern and phenotypes, three independent transgenic lines were analyzed for each expression construct.
In rosette leaves, ROP6 expression was detected at the tip and in the hydathodes at the leaf margin (Fig. 1, A-C). In cauline leaves, however, expression was not confined to the leaf margins, and leaf epidermis pavement cells expressing a constitutively active GFP-rop6 CA (rop6 CA ) were rectangular with few and shallow lobes (Fig. 1F). No expression could be detected in the inflorescence meristem and in young flowers (Fig.  1D). In mature flowers, strong expression was detected in stamen filaments, pollen grains, and stigma and weak expression was detected in ovules (Fig. 1E). In the primary root, expression was detected in the root tip, was absent from the cell division zone, and increased in the stele of the transition zone. In the root differentiation zone, expression increased in lateral root initials (LRI) and was already detected in lateral root founder cells in the pericycle (Fig. 1, G-I). The expression pattern of ROP6 in the root was further confirmed by whole-mount RNA in situ hybridization. Similar to expression detected with reporter genes fused to the ROP6 promoter, following hybridization with an antisense probe, ROP6 RNA was detected in the stele, LRI, and the tip of the emerging lateral root (Fig. 2).
The expression pattern of ROP6 in rosette leaves and primary and lateral roots resembled the expression pattern of the auxin-sensitive synthetic promoter DR5 (Sabatini et al., 1999;Aloni et al., 2003;Benková et al., 2003), suggesting that ROP6 expression might be induced by auxin. Furthermore, the ROP6 promoter contains a TGTCTC auxin response element. To examine whether ROP6 expression is induced by auxin, 5-d-old seedlings were transferred to control or auxincontaining media. The GFP-rop6 CA signal was detected in some pericycle cells as early as 3 h following transfer to an auxin-containing medium (10 mM naphthaleneacetic acid [NAA]; Fig. 1J). Six hours after transfer to auxin-containing medium, GFP-rop6 CA was detected in many pericycle cells, whereas on control plates, it was detected only in a few cells (Fig. 1K). GFP-rop6 CA was visible in many dividing pericycle cells 24 h after transfer to auxin (Fig. 1L). The results of the auxin induction experiments confirm that ROP6 expression is quickly induced by auxin during lateral root development. Furthermore, the identical expression pattern, which was detected when using GUS, DsRed, GFP-ROP6, GFP-rop6 CA , or GFP-rop6 DN (Figs. 1 and 3), and the fast (3-h) response to auxin treatments make it unlikely that the activation status of the transgenic ROP protein affected the AtROP6 expression pattern and the preliminary response to auxin.
To further explore possible function(s) of ROP6 in lateral roots, we examined its subcellular localization  in the shoot and the root. A, B, D, E, G, and H, pROP6..GUS. C, F, J, K, and L, pROP6..GFP-rop6 CA . I, pROP6.. DsRed. A to C and F, In rosette leaves, ROP6 expression is primarily restricted to hydathodes, whereas in cauline leaves, it is detected throughout the epidermis. D, ROP6 expression could not be detected in the inflorescence meristem and young flowers. E, In mature flowers, expression is detected in stamen filaments and pollen grains, stigma, style, and weakly in ovules. G, In the root, expression is detected in the root cap, in and around the quiescent center; it is absent from the cell division and transition zone, and it reappears in the stele in the differentiation zone. H and I, Expression then drops again and reappears in LRI and founder cells. The arrow denotes a protoxylem file. J, Expression of ROP6 is induced by auxin and can be detected in pericycle cells as early as 3 h after induction. K, After 6 h, expression is spread to more cells in the pericycle. L, Twenty-four hours after induction, expression is spread throughout the pericycle and extensive cell divisions are detected. Bars = 100 mm (A), 50 mm (B, D, E, and G-L), and 20 mm (C and F). For more information, see Supplemental Figure S2.
in the developing LRI. In lateral root founder cells in the pericycle, either GFP-ROP6 or GFP-rop6 CA was localized in the cell at the side that is closer to the xylem pole (Fig. 3, A, B, I, and J). At later stages of lateral root development, GFP-ROP6 and GFP-rop6 CA were localized such that they aligned the cell division planes (Fig. 3, C-P) and were absent from the side of the cell pointing toward the lateral root tip (Fig. 3,M,and N). This polar localization suggests that ROP6 relays a special signal in LRI cells. Similarly, it has been found that during stomata development in maize (Zea mays), the type I ROPs ROP2 and ROP9 enhance the polarization of asymmetric cell divisions (Humphries et al., 2011). Together, these data suggest that ROP-dependent polarization could be critical for asymmetric cell divisions. In contrast to GFP-ROP6 and GFP-rop6 CA , the dominant negative GFP-rop6 DN accumulated in the cytoplasm and nuclei in root and leaves (Fig. 3, Q-S), resembling the subcellular distribution of the nonprenylated GFP-rop6 C195S mutant (Fig. 3T). Hence, nucleotide exchange is required for the recruitment of ROPs to the plasma membrane.
The Phenotype of rop6 DN Plants Next, we examined the phenotype of the pROP6.. GFP-rop6 DN (rop6 DN ) plants (Fig. 4). The rop6 DN plants were small and developed multiple inflorescence stems (Figs. 4,A and B,and 7,D and E). The rosette leaves of the rop6 DN plants were curled, and their cauline leaves were twisted (Fig. 4, A-F). The stronger effect of rop6 DN on the structure of cauline leaves compared with rosette leaves was closely correlated with its expression levels in the two leaf types (Fig. 1, A and F). Analysis of the cell structures in rosette (Figs. 4,G and H,and 7,H and I) and cauline leaves (Fig. 4, I and J) revealed that pavement cells in the rop6 DN leaves had fewer lobes and indentations. Quantitative analysis of cell structure, using a value designated the average polarity score, which is based on the ImageJ tools "circularity" and "Skeleton end points" (Sorek et al., 2010(Sorek et al., , 2011, shows that the rop6 DN cells are significantly less polar compared with wild-type cells (P # 0.001, ANOVA, Tukey-Kramer test; Fig. 7, L-N).

Organization of the Cytoskeleton in Wild-Type and rop6 DN Plants
ROPs have been implicated in cytoskeleton organization and dynamics, suggesting that the phenotype of rop6 DN plants could be associated with impaired actin and MT functions. Therefore, we examined the organization of MT and actin in wild-type and rop6 DN plants by immunostaining the cytoskeleton in rosette leaves (Fig. 5). In wild-type pavement cells, F-actin was organized in thin filaments and a few thicker bundles that were oriented at different angles. In addition, diffuse fluorescence likely corresponding to fine F-actin or G-actin could be seen (  The obvious developmental perturbations of rop6 DN plants prompted us to examine whether the phenotype of the rop6 DN plants is associated with changes in gene expression. Gene expression in rop6 DN , wildtype, pROP6..GFP-ROP6 (ROP6), and pROP6.. GFP-rop6 CA (rop6 CA ) plants (Supplemental Fig. S2) was analyzed using microarrays. Entire seedlings were used in the microarrays. To identify differentially expressed genes, we used the "rank product" method. The rank product analysis gives an estimate of false discovery rate while providing flexible means  . Subcellular localization of GFP-ROP6, GFP-rop6 CA , and GFP-rop6 DN during lateral root development. Subcellular localizations of GFP-ROP6 (A-H) and GFP-rop6 CA (I-P) were observed in developing lateral roots in pROP6..GFP-ROP6 and pROP6..GFP-rop6 CA seedlings. A, B, I, and J, Lateral root founder cells. GFP-ROP6 and GFP-rop6 CA were localized along the side of the cell adjacent to the xylem pole (J, arrow). C, D, K, and L, Stage I/II LRI. The majorities of the GFP-ROP6 and GFP-rop6 CA proteins moved and were localized along anticlinal cell borders (arrows). E, F, M, and N, Stage II/III LRI. The majority of GFP-ROP6 and GFP-rop6 CA were localized along anticlinal and periclinal cell borders (arrows) separating the newly formed outer and inner layers. G, H, O, and P, Emerged and mature lateral roots with organized cell files. The majority of the GFP-ROP6 and GFP-rop6 CA proteins were concentrated on proximal anticlinal cell borders (arrows). In the meristematic zone, the protein was distributed more or less evenly around the cells. Q to S, GFP-rop6 DN was observed in pROP6..GFP-rop6 DN seedlings in lateral root founder cells (Q), stage II/II LRI cells (R), and leaf epidermis pavement cells (S). The recruitment of GFP-rop6 DN to the plasma membrane is compromised, and it accumulates in the cytoplasm and nuclei in roots (arrows). T, Nonprenylated GFP-rop6 CA mutant accumulates in cytoplasm and nuclei (arrows) but, unlike GFP-rop6 DN , does not affect cell polarity. Bars = 20 mm.
to assign a significance level to each gene (Breitling et al., 2004). Differentially expressed genes are listed in Supplemental Tables S1 to S5. Pairwise plots showing relative gene expression levels revealed that the expression of many genes was changed in rop6 DN seedlings relative to wild-type, ROP6, and rop6 CA plants (Fig. 6A). Gene expression data were analyzed using hierarchical (Fig. 6B) and K-means ( Fig. 6, C-H) clustering algorithms to identify genes with similar expression patterns. The clustering analyses revealed a group of 14 genes that were strongly induced in rop6 DN plants (Fig. 6, C and F), a group of 24 genes that were mildly up-regulated (Fig. 6, D and G), and a group of 30 genes that were suppressed in rop6 DN plants relative to wild-type, ROP6, or rop6 CA transgenic plants (Fig. 6, E and H). Functional categorization of the differentially expressed genes by Gene Ontology (Martin et al., 2004) revealed that many of the up-regulated genes in the rop6 DN plants were reminiscent of SA-dependent defense responses (Supplemental Tables S1-S5; Supplemental Fig. S4). SA-mediated disease resistance is a mechanism of induced defense that confers protection against a broad spectrum of microorganisms. SA-mediated pathogen defense requires the accumulation of SA, which serves as a signaling molecule leading to the induction of pathogenesis-related proteins that contribute to pathogen resistance (Durrant and Dong, 2004;Conrath, 2006). The frequency of SA-associated genes among the up-regulated genes is statistically significant (P # 0.05) compared with their random up-regulation (Supplemental Table S4). Furthermore, expression analysis of the SA-associated genes in Genevestigator (http://www.genevestigator.ethz.ch; Hruz et al., 2008) showed that their expression is primarily associated with biotic defense responses and not with other stress responses. Examination of the gene expression patterns of all the genes that were differentially expressed in rop6 DN plants revealed that most of them are subject to regulation by biotic stress. Hence, the microarray study suggested that expression of the dominant negative rop6 DN induces a constitutive SA-mediated defense response and that most if not all of the observed changes in gene expression can be explained by activated SA signaling.
The microarray data required validation of the gene expression pattern by an independent method and raised questions that required additional experimental work. The questions that we asked were as follows. (1) Are the levels of SA higher in the rop6 DN plants compared with the controls, as would be expected from the gene expression analysis? (2) Is the developmental phenotype of the rop6 DN plants associated with a constitutive systemic acquired resistance response? (3) Are the rop6 DN plants more resistant to pathogens as a result of the constitutive systemic acquired resistance response? rop6 DN and the SA Response Validation of the microarray data was carried out by real-time quantitative reverse transcription PCR (qPCR) analysis of three genes that displayed strong differential expression in the rop6 DN plants and are often used as markers for SA-dependent defense responses: (1) NIMIN1 (for nonexpresser of PR genes1/noninducible immunity1 interacting pro-tein1; AT1G02450), (2) Pathogenesis-Related Protein5 (PR-5; AT1G75040), and (3) PR-1 (AT2G14610). The qPCR results show that NIMIN1, PR-5, and PR-1 expression levels are 4-, 12-, and 2,000-fold higher, respectively, in rop6 DN relative to wild-type plants (Fig. 7, A-C), confirming the microarray results. Importantly, the expression levels were the same when RNA was extracted from 14-d-old seedlings (Supplemental Fig.  S2), like the plants analyzed by microarrays or from mature flowering plants (Figs. 4 and 7).
To examine whether the changes in gene expression in the rop6 DN plants are indeed associated with SA, we measured the levels of total and free SA in wild-type, ROP6, rop6 CA , and two groups of rop6 DN plants. One group is characterized by strong expression of the rop6 DN protein, similar to the plants shown in Figures 3, 4, and 7, while the second group is characterized by very low expression levels of rop6 DN and has a mild developmental phenotype (these plants are resistant to both kanamycin and basta, confirming that both transfer DNA (T-DNA) inserts of the transactivation expression system are present). In each of the lines, SA levels were compared with those in plants that were inoculated with spores of Golovinomyces orontii, a host-adapted virulent powdery mildew fungus of Arabidopsis (Spanu et al., 2010). As expected from the microarray and qPCR analyses, the levels of both total and free SA in noninoculated plants is markedly elevated in rop6 DN plants (Fig. 6, I and J). In addition, noninoculated and G. orontii-infected rop6 DN plants with high transgene expression accumulate similar SA levels, but these levels are considerably higher compared with pathogen-inoculated wild-type plants.
In contrast, in wild-type, ROP6, and rop6 CA plants, SA levels only increase following inoculation with G. orontii. SA levels are also increased in the rop6 DN plants with low transgene expression levels but at lower levels compared with rop6 DN plants with high expression levels of the transgene (Fig. 6, I and J). The higher SA levels confirmed that the changes in gene expression detected in rop6 DN plants are indeed associated with the SA-induced defense response.
To examine whether the developmental phenotype of the rop6 DN plants is linked to the constitutive SA-mediated response, the rop6 DN plants were crossed to mutants in NPR1, which is required for SA-mediated signaling (Cao et al., 1997), and Salicylic Acid Induction Deficient2 (SID2)/Enhanced Disease Susceptibility16 (EDS16), which is required for pathogen-inducible SA biosynthesis (Nawrath and Métraux, 1999;Supplemental Figs. S4 and S5). It has been shown that both npr1 and sid2 plants fail to induce the expression of PR genes and display enhanced disease susceptibility to pathogens (Cao et al., 1997;Nawrath and Métraux, 1999;Kinkema et al., 2000;Stein et al., 2006). The expression of NIMIN1, PR-5, and PR-1 is suppressed in the rop6 DN npr1 and rop6 DN sid2 double mutant plants, resembling their expression in naive wild-type plants (Fig. 7, A-C). The mutant analysis confirmed that the expression of PR genes in the rop6 DN mutant plants requires SA biosynthesis and NPR1 function, as is known to occur in authentic pathogen-induced SAmediated defense responses (Durrant and Dong, 2004).
The macroscopic phenotype of both rop6 DN npr1 and rop6 DN sid2 double mutants resembles the rop6 DN single mutants. The plants were small, the rosette and cauline leaves were uneven and twisted, and the plants lost their apical dominance and developed numerous adventitious inflorescence stems (Fig. 7, D-G). Qualitative (Fig. 7, H-K) and quantitative (Fig. 7, L-N) analyses of leaf epidermis pavement cell structure showed that cells of rop6 DN npr1 and rop6 DN sid2 double mutants resembled the rop6 DN single mutant and are significantly (P # 0.001, ANOVA, Tukey-Kramer test) different from wild-type plants. The phenotypic analysis of the rop6 DN npr1 and rop6 DN sid2 double mutant plants indicates that the developmental phenotype of the rop6 DN plants can be uncoupled from the constitutive SA-dependent defense response. Hence, the overall small plant size, loss of apical dominance, twisted leaves, and deformed cells observed in the rop6 DN plants are not associated with the constitutive SA-dependent defense response.
Pathogen Response of rop6 DN Plants Mutants in SA biosynthesis or signaling enhance disease susceptibility to pathogens, while the induction of SA-dependent defense responses reduces pathogen growth, especially against biotrophic pathogens such as powdery mildews (Durrant and Dong,  2004). Hence, it was expected that the rop6 DN plants would display increased resistance to G. orontii due to the high levels of SA and PR gene expression that greatly exceed the SA levels seen in wild-type plants after G. orontii infection (Fig. 6, I and J). To test this, wild-type, ROP6, rop6 CA , rop6 DN , rop6 DN WT (plants that phenotypically resembled wild-type plants and in which no visible GFP-rop6 DN expression could be detected, although they were resistant to both the kanamycin and basta selection markers), eds1-2, sid2-1, sid2-2, rop6 DN sid2-1, and rop6 DN npr1-2 plants were inoculated with G. orontii. Five days after conidiospore inoculation, patches of sporulating powdery mildew colonies were macroscopically detectable on the leaf surface of all lines ( Fig. 8; Supplemental Fig. S6). Consistent with this, similar growth of epiphytic fungal hyphae was observed microscopically on leaves of wild-type, rop6 CA , high-expressing, and Figure 8. Susceptibility to the powdery mildew fungus. A, Wild-type (WT), GFP-rop6 DN (ROP6 DN ), GFP-rop6 DN with low GFP-rop6 DN expression and weak phenotype (ROP6 DN wild-type ), SA biosynthesis mutants eds1-2, sid2-1, and sid2-2, and double mutants of GFP-rop6 DN with sid2-1 (ROP6 DN sid2-1) and with the SA signaling mutant npr1-2 (ROP6 DN npr1-2) plants are susceptible to the powdery mildew fungal pathogen G. orontii. B, Fungal entry rates of G. orontii on GFP-rop6 DN (6DN) and into GFP-rop6 DN plants with low expression levels and weak phenotype (6DN WT) are lower compared with the entry rate on wild-type, GFP-ROP6 (R6), and GFP-rop6 CA (6CA) plants. C, Fungal entry rates of Bgh on GFP-rop6 DN (6DN) and on GFP-rop6 DN plants with low expression levels and weak phenotype (6DN WT) are higher compared with the entry rate on wild-type, GFP-ROP6 (R6), and GFP-rop6 CA (6CA) plants. The entry rates of Bgh on the GFP-rop6 CA plants are higher than on wild-type and GFP-ROP6 plants. D, G. orontii conidiospore formation was significantly lower on GFP-rop6 DN (6DN), GFP-rop6 DN sid2-1 (6DN sid2-1), and GFP-rop6 DN npr1-2 (6DN npr1-2) compared with wild-type and GFP-rop6 DN WT (6DN WT) plants. Conidiospore formation was strongly enhanced compared with the wild type on eds1-2 and slightly enhanced on sid2-1 and sid2-2 SA biosynthesis mutants. E, Treatments with the SA analog BTH (black bars) reduced G. orontii conidiospore formation and further enhanced resistance of GFP-rop6 DN plants. Gray bars show mock-treated (water) plants.
low-expressing rop6 DN plants at early stages of pathogenesis (48 h post inoculation; Supplemental Fig. S7). As expected, enhanced disease susceptibility was detected in the eds1-2 mutant that lacks a key regulatory node of plant immune responses known to limit the growth of host-adapted virulent pathogens ( Fig. 8A; Wiermer et al., 2005).
To obtain a more accurate account of infection phenotypes on the different lines, fungal entry into plant cells from each of the different lines was determined (as percentage entry at single plant-powdery mildew interaction sites). The results of this quantitative analysis show that 48 h after inoculation, the fungal entry rates into the rop6 DN plants were around 65% compared with 90% in wild-type, ROP6, and rop6 CA plants. The fungal entry rates on rop6 DN plants with low expression of the transgene were around 80%, slightly lower compared with wild-type, ROP6, and rop6 CA plants and higher compared with rop6 DN plants with high transgene expression levels (Fig. 8B). These data indicate that the constitutive SA-dependent defense response in the rop6 DN plants results in a moderate increase in preinvasive defense responses to the powdery mildew fungus, despite even higher constitutive SA levels than in wild-type plants found after pathogen challenge (Fig. 6, I and J).
We also determined preinvasion infection phenotypes after inoculation with Bgh, a nonadapted powdery mildew that fails to colonize Arabidopsis due to effective extracellular (preinvasive) resistance responses at early stages during fungal pathogenesis (Collins et al., 2003). At 48 h post inoculation, the entry rates of Bgh sporelings into wild-type Columbia (Col-0) leaf epidermal cells were around 10% (Fig. 8C). In comparison, the entry rates of virulent G. orontii at 48 h post inoculation were approximately 90% (Fig. 8B). The entry rates of Bgh into epidermal cells expressing a constitutively active rop6 CA were higher, averaging between 16% and 20% (Fig. 8C). Interestingly, in a compatible interaction in barley expressing constitutively active ROP mutants, entry rates of Bgh were also higher compared with the wild type (Schultheiss et al., 2002(Schultheiss et al., , 2003Opalski et al., 2005;Hoefle et al., 2011). Significantly, Bgh entry rates of rop6 DN epidermal cells were even higher, reaching approximately 30% (Fig.  8C). Taken together, these data suggest that the disruption in cytoskeleton organization impaired extracellular disease resistance responses of the rop6 DN plants toward Bgh, albeit the SA levels in these plants were constitutively elevated.
Next, we quantified the reproductive fitness of the host-adapted G. orontii powdery mildew by counting fungal conidiospores at a late stage after pathogen challenge (7 d after inoculation) in wild-type, rop6 DN , rop6 DN WT (low levels of rop6 DN expression), sid2-1, eds1-2, sid2-2, rop6 DN sid2-1, and rop6 DN npr1-2 plants. As expected, G. orontii reproductive fitness on eds1-2 plants was greatly increased compared with the wild type and was moderately increased on sid2-1 and sid2-2 plants ( Fig. 8D; Supplemental Fig. S8). In contrast, G. orontii reproductive fitness on rop6 DN plants was significantly lower compared with the wild type ( Fig. 8D; Supplemental Fig. S8). Importantly, reproductive fitness of G. orontii remained significantly lower than wild-type levels in rop6 DN sid2-1 and rop6 DN npr1-2 double mutants and was comparable to or only slightly higher than in rop6 DN single mutants ( Fig. 8D;  Supplemental Fig. S8). To further substantiate these findings, we tested G. orontii reproductive fitness following pretreatments with the SA analog benzothiadiazole (BTH). The BTH treatments caused decreased G. orontii reproductive fitness in wild-type, ROP6, rop6 CA , rop6 DN , and rop6 DN WT plants, demonstrating that SA can confer enhanced disease resistance to G. orontii in Arabidopsis. In rop6 DN plants, BTH treatments had an additive effect, further reducing the already lower reproductive fitness of the fungus ( Fig. 8E; Supplemental  Fig. S9). Taken together, the extended data set provides evidence that rop6 DN mediates an enhanced disease resistance phenotype to host-adapted G. orontii independently of the SA-mediated defense pathway.

Plant Development and Hormonal Response and Transport
In the absence of other families of signaling small GTPases in plants, ROPs were suggested to function as regulators of diverse signaling pathways (Yang, 2002). The results in our work are consistent with earlier findings and implicate ROP6 in both developmental and pathogen response regulation. The expression pattern of ROP6 is highly specific and closely resembles that of auxin-regulated genes both in leaves and roots. Correspondingly, the small plant size, loss of apical dominance, and twisted leaf phenotypes of plants expressing a dominant negative rop6 DN under the regulation of the ROP6 promoter resemble mutant plants that are affected in auxin signaling, transport, and biosynthesis. Because the phenotype of rop6 T-DNA knockout mutants is indistinguishable from wildtype plants, it is likely that the rop6 DN mutant affected functions of other ROPs, which are expressed in the same tissue as ROP6. Hence, by expressing rop6 DN under the regulation of it native promoter, we overcame functional redundancy between ROPs while maintaining specificity and analyzing ROP function in a developmental context.
It has recently been proposed that ROPs are activated by auxin through a mechanism that involves ABP1 (Xu et al., 2010). Activated ROPs undergo transient S-acylation and partition into lipid rafts, a process that is required for their function (Sorek et al., , 2010 and that induces localized inhibition of endocytosis (Bloch et al., 2005). In lipid rafts, ROPs interact with their effector ICR1 (Sorek et al., 2010), which regulates the recruitment of PIN auxin efflux transporters to polar domains in the plasma membrane and is required for directional auxin transport . ICR1 transcription is induced by auxin, and similar to ROP6, it is also expressed in LRI . Our results (Fig. 3, Q-S) indicate that the dominant negative rop6 DN accumulates in the cytoplasm and nuclei, where it presumably interacts with ROPGEFs, inhibiting their function in ROP activation (Feig, 1999;Berken et al., 2005;Berken and Wittinghofer, 2008). In addition, the altered cytoskeleton organization in the rop6 DN plants (Fig. 5) may interfere with auxin transport. It has been demonstrated that actin is required for PIN recycling (Geldner et al., 2001) and MTs are required for polar PIN localization (Boutté et al., 2006;Kleine-Vehn et al., 2008;Heisler et al., 2010). Hence, the expression of rop6 DN could have inhibited ROP function in regulating auxin transport.

Epidermal Cell Structure and Growth
The changes in rosette leaf pavement cell morphology, which were detectable in independent transgenic lines, suggest that although ROP6 is primarily expressed at the leaf tip and hydathodes (Fig. 1, A-C), it may also be expressed in pavement cells at levels that are below detection with GUS and GFP reporter genes. Alternatively, the dominant negative rop6 DN may affect pavement cell morphology indirectly. For example, it has recently been shown that perturbations in auxin biosynthesis or distribution affect pavement cell morphology (Xu et al., 2010). Because ROP6 expression in rosette leaves corresponds to sites of high auxin response, it is possible that suppression of ROP function at these sites affects auxin response and/or distribution. Furthermore, the overall small size of the rop6 DN plants and the development of multiple inflorescence stems indicate that the dominant negative rop6 DN has non-cell-autonomous effects.
The rosette and cauline leaves of the rop6 DN plants have a wavy or twisted appearance (Fig. 4), likely owing to uneven cell growth rates in different parts of the leaf. Analysis of pavement cell growth dynamics in developing Arabidopsis cotyledons showed that cell growth is coordinated between groups of neighboring cells, and it was suggested that this coordination is required for the maintenance of flattened leaf surface (Zhang et al., 2011). Close comparison of the scanning electron microscopy images of wild-type and rop6 DN rosette and cauline leaf epidermal cells shows that the pavement cells are flatter in the wild type compared with rop6 DN (Fig. 4, compare G with H and I with J). Furthermore, the cells of the rop6 DN plants range in size and shape: some cells are rectangular or cubical, while other cells develop more lobes (Fig. 7). The bulging of the rop6 DN epidermal cells probably occurred due to isotropic and uncoordinated cell growth that may have altered the mutual pressures that cells exert on their neighbors. Pavement cells of plants expressing constitutively active ROP mutants also display isotropic growth, but they are usually more rectangular and even in their size. The leaf margins of the rop CA mutants usually bend downward, but the leaves are not wavy or twisted (Li et al., 2001;Fu et al., 2002;Bloch et al., 2005;Sorek et al., 2010). This suggests that the isomorphic cell growth cannot fully explain the uncoordinated cell growth that leads to leaf waviness and twisting. It could be that the expression of rop6 DN disrupted a non-cell-autonomous mechanism that is responsible for coordinating cell growth in leaves.

Cytoskeleton Organization
Thick GFP-mTalin-labeled F-actin bundles have previously been observed in plants expressing a dominant negative ROP2 (rop2 DN ) under the regulation of the 35S promoter. It was further shown that the abundance of diffuse F-actin was reduced following transient expression of rop2 DN . Interestingly, the same study showed that the expression of rop2 CA had the opposite effect, causing an increase in the abundance of diffuse F-actin (Fu et al., 2002). Using a different experimental system, our data similarly show that the abundance of diffuse F-actin was low and that F-actin was primarily organized in thick bundles in the rop6 DN plants ( Fig. 5; Supplemental Fig. S3). The effect of constitutively active rop6 CA on actin in pavement cells was not studied in this work. However, previous studies in root hairs showed that the expression of constitutively active rop6 CA or rop11 CA led to the formation of thick actin bundles and eliminated the zone of fine F-actin at the root hair tip (Molendijk et al., 2001;Bloch et al., 2005Bloch et al., , 2011. This may indicate that activated ROPs possibly have different effects on actin organization in different cell types. An alternative and tempting explanation is that ROP cycling between active and inactive states is required for F-actin turnover and the formation of fine F-actin. The constitutively active mutants are locked in the GTP-bound state (Berken and Wittinghofer, 2008;Sorek et al., 2010). Hence, when signaling requires the binding and release of effectors, it should be impaired when the dissociation of the GTPase from target proteins is compromised. Thus, the bundling of actin filaments may reflect an inhibition of the ROP switch mechanism. How ROPs regulate actin nucleation and stability is yet an open question (Hussey et al., 2006;Deeks and Hussey, 2009).
The MT analysis in this work demonstrates that in many fully developed pavement cells, MTs were organized in transverse or oblique orientation relative to the long cell axis (Fig. 5; Supplemental Fig. S3). Earlier studies with plants expressing rop2 DN showed that the cortical MTs have transverse orientation during early stages of pavement cell growth but are randomly organized in mature cells (Fu et al., 2002). The discrepancy between our data and the previous study could be explained by differences in the experimental system, such as the different ROPs used in each experiment, different expression systems, the level of transgene expression, and plant growth conditions. The observed changes in MT organization could have resulted from changes in tissue biomechanics or from the direct influence by ROPs or both. In cells of the shoot apical meristem, the orientation of cortical MTs was shown to correspond to biomechanical stress from the surrounding tissue (Hamant et al., 2008). It was further shown that both cortical MT orientation and the localization of PIN1 auxin efflux carrier likely respond to the same biomechanical regulator (Heisler et al., 2010). In developing metaxylem cells, the ICR1 homolog MIDD1/RIP3/ICR5 is recruited to specific plasma membrane domains by ROP11, where it interacts with and induces the severing of cortical MTs, leading to secondary cell wall pit formation (Oda et al., 2010;Oda and Fukuda, 2012). Interestingly, adaxial pavement cells of icr1 mutant plants have a cubical shape lacking lobes and indentations, resembling ROP mutants (Lavy et al., 2007). During pavement cell growth, ROP6 is activated by auxin and stabilizes MTs via its effector RIC1 (Fu et al., 2005;Xu et al., 2010). Hence, ROPs may modulate MTs via ICR and RIC family proteins. Analysis of pavement cell growth dynamics showed little correlation between MT organization and orientation and cell structure (Zhang et al., 2011). Clearly, more work is required to understand the complex relation between ROP function, cell growth, MT organization, and cell wall formation and flexibility.

The SA Response
The microarray studies were carried out on young seedlings before the developmental abnormalities induced by the expression of rop6 DN became apparent (Supplemental Fig. S2). The logic behind this experimental design was that by looking at changes in gene expression prior to the stage where the phenotype of the plant becomes apparent, it should be possible to define whether the dominant negative ROP induces changes in gene expression and possibly identify the key genes whose alteration leads to the developmental aberrations. However, the microarray analysis revealed that the most significant changes in gene expression induced by rop6 DN are associated with SA-dependent defense responses. Down-regulation of lignin biosynthesis by silencing the expression of hydroxycinnamoyl-CoA:shikimate hydroxycinnamoyl transferase (HCT) in alfalfa (Medicago sativa) and Arabidopsis led to retarded plant growth, which has been associated with a constitutive SA response. Growth of HCT-silenced plants was regained in HCT-RNAi sid2 but not in HCT-RNAi npr1 double mutants (Gallego-Giraldo et al., 2011a, 2011b. In contrast to the HCT-silenced plants, the phenotype of the rop6 DN npr1 and rop6 DN sid2 double mutants, in which the SA-dependent defense response is blocked, resembled that of rop6 DN single mutant plants, indicating that the developmental perturbations induced by rop6 DN are unrelated to the constitutive SA-dependent response.

ROPs and Pathogen Response
As described in the introduction, ROP function was linked to plant defense responses in several studies. Results in this work  suggest that the possible suppression of SA-dependent defense response execution by activated ROPs or by loss of MT-associated ROPGAP1 function Huesmann et al., 2011) may also contribute to the enhanced susceptibility of the respective mutants. Loss-offunction mlo mutant alleles in Arabidopsis and barley are resistant to host-adapted powdery mildews, and this preinvasive resistance depends on polarized exocytosis regulated by PEN1/ROR2 syntaxins (Collins et al., 2003;Consonni et al., 2006;Kwon et al., 2008). Infection by the Bgh powdery mildew was shown to induce a redistribution of MLO, ROR2, and other proteins involved in the defense response to plasma membrane microdomains, which could be lipid rafts, just below sites of attempted fungal ingress (Bhat et al., 2005). The actin and MT cytoskeleton are disorganized in dominant negative ROP mutants (Fu et al., 2002;this work), possibly affecting directed secretion and, in turn, plant defense responses to the powdery mildew fungus. Hence, impaired extracellular resistance responses of the rop6 DN plants toward Bgh may have resulted from the disorganized cytoskeleton. Consistent with this interpretation, pharmacological inhibitors and genetic interference by ectopic expression of an actin-depolymerizing factor-encoding gene, HvADF, impaired extracellular resistance responses to Bgh entry in barley (Miklis et al., 2007), and silencing of the HvRAB-and HvRAC1-interacting protein kinase HvRBK1 disrupted MT organization and enhanced susceptibility to Bgh (Huesmann et al., 2012). Of note, however, preinvasive resistance responses to Bgh at early stages during pathogenesis are essentially independent of an intact SA defense pathway (Consonni et al., 2006;Stein et al., 2006). This could explain why extracellular resistance responses to Bgh in rop6 DN plants are impaired despite constitutively elevated SA levels.
The BTH treatments ( Fig. 8; Supplemental Fig. S9) demonstrated that SA-dependent defense responses can confer resistance to G. orontii in Arabidopsis. Hence, we cannot exclude a small contribution of the SA-dependent defense response to the reduced reproductive fitness of G. orontii seen on rop6 DN plants. However, the double mutants of rop6 DN with sid2 or npr1 show that the reduced reproductive fitness of G. orontii on rop6 DN plants was largely a rop6 DNdependent and SA-independent defense response.

CONCLUSION
Our results imply that ROPs regulate highly complex defense responses to powdery mildews that can be detected at early and late stages of the infection process. ROPs either enhance or reduce early preinvasive disease resistance responses to the tested host-adapted and nonadapted powdery mildews, presumably via their effects on the cytoskeleton and intracellular trafficking. At later stages of fungal pathogenesis, interference with ROP function reduces fungal reproductive success by mechanisms that remain to be elucidated. The mechanism underlying the constitutive activation of SA accumulation/signaling in rop6 DN plants remains unclear. One possibility is that perturbation of the cytoskeleton or default secretion by rop6 DN mimics a pathogen attack. Alternatively, similar to HCT-RNAi plants (Gallego-Giraldo et al., 2011b), release of cell wall materials or changes in cell wall integrity may activate SA responses in rop6 DN plants. Changes in cell wall integrity may have also contributed to the susceptibility of the rop6 DN plants to Bgh. Irrespective of this, our results implicate ROPs as important regulators of plant development as well as in SA-dependent and SA-independent defense execution.

Molecular Cloning
All plasmids and primers are listed in Supplemental Tables S6 and S7, respectively. Plasmid construction was carried out using standard molecular cloning techniques. AtROP6 was cut from pSY700 with SacI and then was subcloned to pGFP-MRC to create pSY811. PCR-based site-directed mutagenesis (QuikChange kit; Stratagene) was used to create a constitutively active mutant (Atrop6 CA/DN ) by substituting Gly-15 with Val and Thr-30 with Asn. pSY811 was used as template with the following primer sets: SYP189 + SYP190 to create pSY812 (GFP-rop6 CA ) and SYP602 + SYP603 to create pSY807 (GFP-rop6 DN ). To subclone GFP-ROP6, GFP-rop6 CA , and GFP-rop6 DN into the pOP effector plasmids, pSY811, pSY812, and pSY807 were digested with XhoI to isolate the GFP-ROP6, GFP-rop6 CA , and GFP-rop6 DN fragments. The three fragments were, in turn, subcloned into pOP-Bj36 to create pSY836, pSY819, and pSY835, respectively. For expression in plants, pSY836, pSY819, and pSY835 were digested with NotI, and the resulting pOP-GFP-ROP6/rop6 CA / rop6 DN fragments were subcloned into pMLBART plant binary vector to create pSY832, pSY818, and pSY831, respectively. For construction of the ROP6 promoter LhG4 driver plasmid, a 1,047-bp fragment upstream of AtROP6 (pROP6) was amplified by PCR from Arabidopsis (Arabidopsis thaliana Col-0) genomic DNA using SYP615 and SYP616. The amplified AtROP6 was subcloned into pGEM-T (Promega) and, in turn, into pBj36 (Eshed et al., 2001) with SalI and KpnI (Acc65I) to create the pSY810 promoter LhG4 driver plasmid. For expression in plants, pSY810 was digested with NotI, and the LhG4-pAtROP6 cassette was subcloned into a pART27 plant binary vector to create pSY813. All fragments were fully sequenced to verify that no PCR-generated errors were introduced.

Plant Material
All transgenic Arabidopsis lines used in this study are listed in Supplemental Table S8. Arabidopsis plants were grown as described previously (Lavy et al., 2002). For plant transformation, Arabidopsis plants were transformed with Agrobacterium tumefaciens strain GV3101/pMp90 harboring the appropriate plasmids using the floral dip method (Clough and Bent, 1998). A minimum of three independent transgenic lines for each plasmid were analyzed. Transcription-transactivation lines were obtained by crossing pROP6::LhG4 driver lines with pOp effector lines to obtain pROP6..GUS, pROP6..DsRed, pROP6..GFP-ROP6, pROP6..GFP-rop6 CA , and pROP6..GFP-rop6 DN . An Arabidopsis T-DNA insertion line (SALK_91737) in the ROP6 gene was obtained from the Arabidopsis Biological Research Center at the University of Ohio. The existence of the insertion was verified with SYP179 and SYP163 ROP6-specific primers and a T-DNA left border primer (Lba1; SYP179).

Auxin Induction
For auxin induction, seeds were germinated in the presence of the auxin transport inhibitor N-1-naphthylphthalamic acid followed by transfer to growth medium containing 10 mM auxin (NAA), as described previously (Himanen et al., 2004). Alternatively, seeds were germinated on medium lacking N-1-naphthylphthalamic acid and were, in turn, treated with 10 mM NAA.

GUS Staining
GUS staining was carried out as described previously (Weigel and Glazebrook, 2002).

Whole-Mount in Situ Hybridization
Tissue preparation and whole-mount in situ hybridization were performed essentially as described previously Hejátko et al., 2006). A 244-bp fragment from the 39-untranslated region of ROP6 was amplified by PCR from Arabidopsis genomic DNA and was used as template to create genespecific digoxigenin-labeled antisense/sense RNA probes. The digoxigeninlabeled probes were prepared by in vitro transcription according to the manufacturer's instructions (Roche).

Fixation and Immunostaining
Fixation and immunostaining were carried out essentially as described previously (Chaimovitsh et al., 2012). To stabilize actin and MT, specimens were fixed for 5 min with 500 mM m-maleimidobenzoyl-hydroxylsuccinimide ester in MT-stabilizing buffer (MTSB) containing 100 mM PIPES, pH 6.89, 5 mM EGTA, 5 mM MgSO 4 $7H 2 O, and 1% Triton X-100 (v/v) buffer and, in turn, for 1 h in freshly prepared 8% paraformaldehyde in MTSB. Fixation was carried out at room temperature in the dark. Samples were then rinsed three times in MTSB and treated with the following enzyme mixture: 2% cellulase Onzuka R-10 and 1% (w/v) pectinase, protease inhibitor cocktail (Sigma), and 20 mM phenylmethylsulfonyl fluoride for 10 min. After washing in MTSB for 30 min, the samples were squashed on coverslips coated with poly-L-Lys (1 mg mL 21 ; Sigma-Aldrich). Then, the cells were retreated for 10 min with enzyme mixture (2% cellulase Onzuka R-10 and 1.5% [w/v] pectinase) with protease inhibitors. This step was followed by a rinse in phosphate-buffered saline (PBS) with 1% Triton X-100 (v/v) for 30 min and then incubation in PBS containing 1% (w/v) bovine serum albumin for 20 min. To reduce aldehyde-induced autofluorescence, samples were treated for 5 min with 10 mg mL 21 sodium borohydride in PBS and then washed in PBS for 20 min. For immunofluorescence detection of MTs, specimens were incubated overnight at room temperature with primary sheep anti-ab-tubulin polyclonal antibodies (Cytoskeleton) and mouse anti-actin monoclonal (clone C4) antibodies (ICN Biomedical) at a dilution of 1:100. Samples were then washed in PBS and incubated with antisheep secondary antibodies conjugated to Alexa Fluor 488 and anti-mouse secondary antibodies conjugated to Alexa 555 (Molecular Probes) diluted 1:100 for 1 h at room temperature. Mounting was done in 50% glycerol/water (v/v).

Plant DNA and RNA Isolation and qPCR
Plant genomic DNA was isolated with the GenElute Plant Genomic DNA miniprep kit according to the manufacturer's instructions (Sigma). Total RNA was isolated with the SV Total RNA isolation kit according to the manufacturer's instructions (Promega). The RNA from the qPCR experiments was extracted from both 14-d-old seedlings and mature flowering plants. Complementary DNA first-strand synthesis was carried out using Moloney murine leukemia virus reverse transcriptase (Promega). Quantifications with the primer sets (Supplemental Table S7) were carried out by qPCR using an ABI Prism 7700 StepOnePlus Instrument (Applied Biosystems). Study samples were run in triplicate on eight-well optical PCR strips (Applied Biosystems) in a final volume of 10 mL. Primers were designed using the Roche Universal Probe Library (https://www.roche-applied-science.com/sis/rtpcr/upl/ index.jsp). The PCR cycles were run as follows: 10-min initial denaturation at 95°C, followed by 40 subsequent cycles of 15 s of denaturation at 95°C, and 1 min of annealing and elongation at 60°C. The specificity of the unique amplification product was determined by melting-curve analysis according to the manufacturer's instructions (Applied Biosystems). Relative quantities of RNA were calculated by the comparative threshold cycle method (Applied Biosystems User Bulletin 2: ABI PRISM 7700 Sequence Detection System; http://www.appliedbiosystems.com). DNA dilution series were prepared to calculate an amplification efficiency coefficient for each gene. The relative levels of RNA were calculated according to the amplification efficiency coefficient and normalized against ACTIN8 and UBQ21 gene standards (Czechowski et al., 2005), the level of which was taken as 1. The stability of the standards in each experiment was verified with the geNorm analysis tool (http://medgen.ugent.be/jvdesomp/genorm/) and was calculated as M # 0.7. The analysis was repeated with three independent biological replicates.

Array Hybridization and Evaluation
Seedlings of pAtROP6..GFP-ROP6, pAtROP6..GFP-ROP6 CA , and Col-0 were grown on 0.53 Murashige and Skoog medium for 14 d in growth chambers at 21°C under long-day photoperiods (16 h of light, 8 h of darkness). A total of 100 mg of plant tissue was harvested from seedlings, and RNA was extracted using Trizol (Invitrogen) according to the manufacturer's instructions. The entire experiment was performed three times, providing independent biological replicates. Affymetrix Arabidopsis ATH1 GeneChips were used in the experiment. Labeling of samples, hybridizations, and measurements were performed as described (Hennig et al., 2004). Signal values were derived from the Affymetrix *.cel files using the GCRMA algorithm (Psarros et al., 2005). Data were processed with the statistical package R (version 1.9.1). Differentially expressed genes were determined using the rank-product method (Breitling et al., 2004) implemented in R. Genes were considered as being differentially expressed at P # 0.05.

Light, Fluorescence, and Confocal Microscopy
Low-resolution imaging was performed with an SV-11 stereomicroscope (Zeiss). Wide-field fluorescence imaging was performed with an Axioplan-2 Imaging fluorescence microscope (Zeiss) equipped with an AxioCam cooled CCD camera, using either 203 dry or 633 water-immersion objectives with numerical aperture values of 0.5 and 1.2, respectively. Confocal imaging was performed with a Leica TCS-SL confocal laser scanning microscope equipped with 203 multi-immersion and 633 water-immersion objectives with numerical aperture values of 0.7 and 1.2, respectively. GFP was visualized by excitation with an argon laser at 488 nm, a 500-nm beam splitter, and a spectral detector set between 505 and 530 nm. DsRed was visualized by excitation with an argon laser set to 514 nm, a 456/514-nm double dichroic beam splitter, and a spectral detector set between 530 and 560 nm. Scanning electron microscopy was carried out with JEOL JSM-840A scanning electron microscope. Image analysis was performed with Zeiss AxioVision, ImageJ, Leica LCS, and Adobe Photoshop.

SA Measurements, Fungal Entry Rates, Conidiospore Counts, and BTH Treatments
Measurements of free and conjugated SA levels were carried out by analytical HPLC essentially as described previously (Bartsch et al., 2006). Golovinomyces orontii and Blumeria graminis f. sp. hordei conidiospore inoculations were carried out as described previously (Consonni et al., 2006). Conidiospore counts were carried out as described previously (Wessling and Panstruga, 2012). BTH treatments were carried out essentially as described previously (Lawton et al., 1996). Briefly, 4-week-old plants were sprayed with 42.6 mg L 21 water-solubilized BTH 24 h prior to inoculation by G. orontii conidiospores. Control plants were sprayed with water (BTH mock control). For all infection assays, plants were grown in a phytochamber with the following settings: 65% humidity; temperature of 22°C from 9 AM to 7 PM and 20°C from 7 PM to 9 AM; and 10/14-h light/dark cycles. Light intensity was 100 mE m 22 s 21 .

Supplemental Data
The following materials are available in the online version of this article.
Supplemental Figure S1. Nucleotide sequence of the AtROP6 promoter.
Supplemental Figure S3. Immunostaining of actin and microtubule in wild-type and GFP-rop6DN leaf epidermis cells.
Supplemental Figure S4. Immunostaining of actin and microtubule in wild-type and GFP-rop6DN leaf epidermis cells.
Supplemental Figure S6. Susceptibility to the powdery mildew fungus.
Supplemental Figure S9. Susceptibility to G. orontii following SA analog BTH treatments.
Supplemental Table S4. Gene Ontology of differentially expressed genes by GoToolBox.