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NITRIC OXIDE-ASSOCIATED PROTEIN1 (AtNOA1) is essential for salicylic acid-induced root waving in Arabidopsis thaliana Xiang Zhao*, Jin Wang*, Jing Yuan, Xi-li Wang, Qing-ping Zhao, Pei-tao Kong and Xiao Zhang Institute of Plant Stress Biology, State Key Laboratory of Cotton Biology, School of Life Sciences, Henan University, Kaifeng 475004, China

Summary Author for correspondence: Xiao Zhang Tel: +86 371 23880008 Email: [email protected] Received: 18 November 2014 Accepted: 10 January 2015

New Phytologist (2015) doi: 10.1111/nph.13327

Key words: Arabidopsis thaliana, Arabidopsis thaliana NITRIC OXIDEASSOCIATED PROTEIN1 (AtNOA1), Ca2+ signal, polar auxin transport, root waving, salicylic acid (SA).


Root waving responses have been attributed to both environmental and genetics factors, but the potential inducers and transducers of root waving remain elusive. Thus, the identification of novel signal elements related to root waving is an intriguing field of research.
Genetic, physiological, cytological, live cell imaging, and pharmacological approaches provide strong evidence for the involvement of Arabidopsis thaliana NITRIC OXIDE-ASSOCIATED PROTEIN1 (AtNOA1) in salicylic acid (SA)-induced root waving.
SA specially induced root waving, with an overall decrease in root elongation in A. thaliana, and this SA-induced response was disrupted in the Atnoa1 mutant, as well as in nonexpresser of pathogenesis-related genes 1 (npr1), which is defective in SA-mediated plant defense signal transduction, but not in npr3/4 single and double mutants. The expression assays revealed that the abundance of AtNOA1 was significantly increased by application of SA. Genetic and pharmacological analyses showed that SA-induced root waving involved an AtNOA1-dependent Ca2+ signal transduction pathway, and PIN-FORMED2 (PIN2) -based polar auxin transport possibly plays a crucial role in this process.
Our work suggests that SA signaling through NPR1 and AtNOA1 is involved in the control of root waving, which provides new insights into the mechanisms that control root growth behavior on a hard agar surface.

Introduction As sophisticated sensors, plant roots change their growth patterns in response to many stimuli, including gravity, obstacles, temperature, humidity, light, water and nutrients. The flexible growth responses of roots are important for plants to adapt to their environment and gain maximum advantage for growth (Oliva & Dunand, 2007). One of the most extensively studied morphological responses is root waving. Previous research showed that root waving was produced on the surface of an agar plate inclined from the vertical. The root waving pattern is the result of positive gravitropism plus a thigmotropic effect. When the root tip, during the gravitropic reaction, encounters an impediment in penetrating the agar surface on a tilted hard agar plate, it experiences a rotation about its vertical axis, leading to a sinusoidal pattern of growth (Okada & Shimura, 1990; Thompson & Holbrook, 2004; Migliaccio et al., 2009). Moreover, the growth pattern produced by the root–agar interaction also depends on the type and concentration of salts in the medium, the concentration of ethylene in the air filling the caps, and the degree of tilt of the agar plate (Buer et al., 2000, 2003). In addition to the various environmental factors mentioned above, some genetic factors have been showed to play crucial roles *These authors contributed equally to this work. Ó 2015 The Authors New Phytologist Ó 2015 New Phytologist Trust

in the regulation of root waving growth. The initial discovery of mutants defective in both waving and auxin transport suggested a prominent role for this hormone in the establishment of root waving growth (Okada & Shimura, 1990). Subsequently, other studies demonstrated that the auxin efflux carrier PINFORMED2 (PIN2) and the influx carrier AUXIN RESISTANT1 (AUX1) were involved in the changes in root growth direction, thus establishing the model for the relationship between polar auxin transport and root waving growth (Bennett et al., 1996; Chen et al., 1998; Rashotte et al., 2000). Furthermore, several genes involved in root growth responses have been identified and characterized using some significant mutants with altered root waving. For instance, the WAVE GENE1 (WAG1) and WAG2 genes, which encode protein-serine/threonine kinases closely related to PINOID, negatively regulate root waving in Arabidopsis thaliana (Santner & Watson, 2006). WAVY GROWTH2 (WAV2) encodes a protein belonging to the BUD EMERGENCE 46 family and possesses an a/b-hydrolase domain, and the mutant wav2-1 shows a compressed wavy behavior on a tilted agar surface, possibly as a result of the regulation of G proteins and the organization of cortical microtubules in roots (Chant et al., 1991; Mochizuki et al., 2005). A mutation of ANTHRANILATE SYNTHASE a1 (ASA1) gene, which affects the tryptophan biosynthesis pathway, confers a compressed root waving phenotype (Rutherford et al., 1998). In addition, the New Phytologist (2015) 1 www.newphytologist.com

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ROOT HAIR DEFECTIVE3 (RHD3) gene, which encodes a protein containing GTP-binding motifs and regulates root hair development (Yuen et al., 2005), and WAVE-DAMPENED2 (WVD2), which encodes a novel microtubule-associated protein (Perrin et al., 2007), have been shown to regulate root waving growth. However, the function of these genes in the induction of root waving remains elusive. Salicylic acid (SA) plays diverse roles in plant growth, development, and responses to abiotic stress (Rivas-San Vicente & Plasencia, 2011). The most well-established role of SA is as a signaling molecule in the plant immune response (Vlot et al., 2009). Upon pathogen challenge, plants initiate systemic acquired resistance (SAR) through the production of the immune signal SA. Recently, NONEXPRESSER OF PATHOGENESISRELATED GENES3 (NPR3) and NPR4 have been identified as receptors for SA, and SA controls the accessibility of the receptors to their substrate NPR1 in a concentration-dependent manner, thereby regulating downstream defense signal transduction (Fu et al., 2012; Moreau et al., 2012). However, it is unknown whether NPR3/NPR4-mediated SA perception is involved in the diverse roles that this hormone plays in growth and development, or in the abiotic stress response (Moreau et al., 2012). Arabidopsis thaliana NITRIC OXIDE SYNTHASE1 (AtNOS1) was previously identified as a potential nitric oxide synthase (NOS) in A. thaliana, and was extensively implicated in plant development and phytohormone actions (Guo et al., 2003; Besson-Bard et al., 2008). Recent studies have confirmed that AtNOS1 is not a NOS, and the renamed NITRIC OXIDEASSOCIATED PROTEIN1 (AtNOA1) is a member of the circularly permuted GTPase family (cGTPase) (Moreau et al., 2008; Sudhamsu et al., 2008). In bacteria and some eukaryotes, this family of GTP-binding proteins is essential for cell growth, and is associated with an RNA/ribosome binding function (Arigoni et al., 1998; Matsuo et al., 2006). However, little is known concerning the function of this new class of A. thaliana GTPases in plants, although the biological role of AtNOA1 is currently believed to be primarily associated with chloroplast ribosome functions (Flores-P
erez et al., 2008; Gas et al., 2009; Liu et al., 2010; Yang et al., 2011). Interestingly, canonical and extra-large G proteins each participate in the control of root morphogenesis, including the root waving response (Ding et al., 2007; Pandey et al., 2008), which could shed light on the possible role of AtNOA1 in plant root growth and development. Furthermore, recent studies have revealed that AtNOA1 levels are regulated by polyunsaturated fatty acid (FA) levels (Mandal et al., 2012), and FAs and their derivatives function in root waving and development as signaling molecules (Vellosillo et al., 2007). Root waving responses have been attributed to both environmental and genetic factors, and the identification of novel signal elements related to root waving is an intriguing research field. In plant defense, SA is usually associated with ethylene (An & Mou, 2011), and ethylene modulates root waving responses (Buer et al., 2003). Moreover, SA effectively induces nitric oxide production to regulate root growth by activating NOS (AtNOA1) (Zottini et al., 2007). In the light of these findings, we designed the present study with the following questions in mind: does SA New Phytologist (2015) www.newphytologist.com

specifically induce root waving, and how does AtNOA1 modulate SA-induced root waving responses? Our results suggest that AtNOA1 is involved in SA-induced root waving by affecting cytosolic Ca2+ signaling and polar auxin transport.

Materials and Methods Plant material and growth conditions Seeds of Arabidopsis thaliana (L.) Heynh were surface-sterilized and planted in square Petri dishes containing 1.2% (w/v) agar Murashige and Skoog (MS) medium. Sterilized seeds were kept in a cold room at 4°C for 3 d. The Petri dishes were then transferred to a cultivation chamber to let the seedlings grow for 4 d under a 16: 8 h, light: dark cycle. Arabidopsis thaliana ecotype Columbia-0 (Col-0) was used as the wild-type in this study. Atnoa1 and two complementation line seeds (B3-1 and C2-6) were kindly provided by Nigel M. Crawford (Guo et al., 2003). The resistant to inhibition by fosmidomycin (rif1-1) mutant seeds were obtained from Manuel Rodriguez-Concepcion (Gas et al., 2009). The npr1-1 (CS3726) seeds were kindly provided by Xinnian Dong (Cao et al., 1997). The npr3 (SALK_031835) and npr4 (Salk_098460) mutant seeds were ordered from the Arabidopsis Biological Resource Center (ABRC) (http://www.arabidopsis.org/abrc/). The double mutant npr3npr4 was obtained by crossing npr3 and npr4, selfing F1, and screening the F2 population for plants homozygous for mutations in both genes by reverse transcription PCR (RTPCR). The primers used for the expression analysis of the genes NPR3 and NPR4 by semiquantitative RT-PCR were as follows: NPR3, 50 -CTCGTATGGTGGCTCTAATGAA-30 (sense) and 50 -TCGAGGATGTCGTCGTCTAT-30 (antisense); NPR4, 50 GAATGGGAACAAGTGGGT-30 (sense) and 50 0 CTTCGAGTTAAGTCGGTGAA-3 (antisense). AtPIN2-GFP, DR5-GFP auxin reporter and DR5-GUS seeds (Col-0 background) were obtained from Jian Xu (Sassi et al., 2012). AtPIN3GFP and AtPIN7-GFP seeds (Col-0 background) were obtained from Zhizhong Gong (Wang et al., 2011). Pollen from the AtPIN2-GFP, AtPIN3-GFP, AtPIN7-GFP, DR5-GFP or DR5GUS transformed plant was transferred to the mature stigma of an Atnoa1 plant. F1 plants were self-pollinated and grown to the F2 generation. F2 seeds were cultured on the agar surface with SA, and seedlings without root waving were chosen and grown on soil to obtain the F3 generation. Root waving analysis of F3 lines was performed to identify the homozygous mutant lines with AtPIN2-GFP, AtPIN3-GFP, ATPIN7-GFP, DR5-GFP or DR5-GUS expression, which were used for the experiments as indicated below. Root waving analysis The A. thaliana root waving analysis was performed as described by Buer et al. (2003), with slight modifications. Four-day-old seedlings were transferred to 1.5% (w/v) agar MS medium containing SA or other agents and maintained at a 30° angle between the root growth axis and the gravity vector. Seedlings were Ó 2015 The Authors New Phytologist Ó 2015 New Phytologist Trust

New Phytologist photographed 7 d later (using a Coolpix 900 digital camera; Nikon, Tokyo, Japan). To evaluate the wave characteristics of a seedling, the first wave including a left-handed and a righthanded rotation around its general axis of growth was measured. Quantification of root length (the distance from the base of the hypocotyl to the root tip), wavelength (the distance along the root axis between the crests of two successive waves), wave amplitude (the distance along the root vertically down to the wave crest), and root axis angle were measured by using IMAGEJ software (National Institutes of Health (NIH); http:// rsb.info.nih.gov/ij/). Every experiment was repeated at least three times and c. 10 seedlings were measured each time. The significance of differences between wild-type and mutant seedlings was confirmed using Student’s t-test. Total RNA extraction, RT-PCR and quantitative real-time PCR analysis Gene expression was analyzed using RT-PCR and quantitative real-time PCR, according to previous methods (Zhao et al., 2013). The primers used for gene expression analysis by semiquantitative RT-PCR were as follows: AtNOA1, 50 GCACGGAAAGTTGTTGATAC-30 (sense) and 50 -GAGTTG GCAAAGTTGAAATA-30 (antisense); NPR1, 50 -ATGGACAC CACCATTGATGGAT-30 (sense) and 50 -TCACCGACGACG ATGAGAGAGT-30 (antisense); ACTIN2, 50 -TTCCTCATGCC ATCCTCCGTCTT-30 (sense) and 50 -CAGCGATACCTGA GAACATAGTGG-30 (antisense). For qRT-PCR analysis, ACTIN2 was used as an internal standard to normalize the data and was amplified with the primer pair 50 -AACCACTATGTTCTCAGGCATCG-30 (sense) and 50 -CCTGGACCTGCCTCATCATACT-30 (antisense). The primers used for AtNOA1 expression analysis by qRT-PCR were 50 -GGAGTTAGCGGAGTTGC-30 (sense) and 50 -GCTTGC CTGTGGTGTAG-30 (antisense). Subcellular localization and GUS staining For transient expression of AtNOA1-GFP, the full-length cDNA and N-terminal 600-bp fragment of AtNOA1 were fused upstream of GFP under the control of the super-promoter in the pHBT-GFP-NOS vector. The primers used for amplification of the full-length cDNA of AtNOA1 were 50 -ATCGGTACCAT GGCGCTACGAACACTCTCAACG-30 (sense) and 50 -GAA CCCGGGCCAAAGTACCATTTGGGTCTTACT-30 (antisense). The primers used for amplification of the N-terminal 600-bp fragment of AtNOA1 were 50 -CGCGGATCCATCCAC TATATCAACC-30 (sense) 50 - TAAGTCGACATGGCGC TACGAACAC-30 (antisense). After the plasmids had been purified by CsCl gradient centrifugation, they were introduced into leaf protoplasts following a previously described protocol (Hua et al., 2012). For translationally fused AtNOA1-GFP, the full-length cDNA of AtNOA1 was fused upstream of GFP under the control of the super-promoter in the pCAMBIA1300 vector. The primers used for amplification of the full-length cDNA of AtNOA1 were Ó 2015 The Authors New Phytologist Ó 2015 New Phytologist Trust

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50 -CGATCTAGAATGGCGCTACGAACAC-30 (sense) and 50 -GCGGAATTCAAAGTACCATTTGGGT-30 (antisense). The vector was transformed into Agrobacterium tumefaciens strain GV3101 and then used to transform the plants. The transformed seedlings were screened on MS plates containing 30 mg l1 hygromycin. GFP fluorescence of seedling roots and hypocotyls was detected using a confocal microscope (FV1000; Olympus, Tokyo, Japan) with an excitation wavelength of 488 nm and an emission wavelength of 525 nm. Before microscopy, the roots were incubated in a 0.8 M mannitol solution for 10 min to induce plasmolysis. Quantitative analysis of plasma membrane (PM) AtPIN2-GFP fluorescence was performed as described by Du et al. (2013). DR5-GUS staining was performed as described by Willige et al. (2011), with slight modifications. Arabidopsis thaliana seedlings were incubated in GUS staining solution (100 mM Na-phosphate buffer, pH 7.0, 0.5 mM K4Fe(CN)6, 0.5 mM K3Fe(CN)6, 0.1% Triton X-100, and 1 mg/ml X-Gluc). GUS-stained seedlings were photographed using a Leica MZ16 stereomicroscope with a PLAN-APOX1 objective (Leica, Lambrecht, Germany). Polyclonal antibodies and immunodetection The 19 N-terminal amino acids of AtNOA1 were used as antigens to immunize rabbits and obtain polyclonal antibodies raised against AtNOA1, as described previously (Guo et al., 2003). Polyclonal antibodies raised against the plasma membrane H+ATPase were provided by K. Shimazaki and have been described previously (Kinoshita & Shimazaki, 1999). Immunodetection was performed according to Zhao et al. (2013). Imaging of cytosolic Ca2+ levels YC3.6 expression vectors were gifts from T. Nagai (Nagai et al., 2004). These constructs were introduced into A. tumefaciens strain LBA4404 and transformed by floral infiltration into A. thaliana Col, the Atnoa1 mutant and the rescued Atnoa1 line (C2-6), respectively. Imaging of cytosolic Ca2+ concentrations was performed as described by Monshausen et al. (2008), with slight modifications. Arabidopsis thaliana seedlings expressing the fluorescence resonance energy transfer (FRET)-based Ca2+ sensor YC3.6 (Nagai et al., 2004) were transferred to very thin halfstrength MS medium with or without SA for 7 d, and then placed on the inverted Olympus confocal laser scanning microscope plate for measurement of FRET-sensitized emission. Excitation wavelengths were 458 and 514 nm for CFP and YFP, respectively. FRET efficiency was calculated using the Olympus software employing the formula described by Monshausen et al. (2008). Aequorin reconstitution and cytosolic Ca2+ measurements Aequorin expression vectors cloned into A. tumefaciens strain GV3101 were gifts from M. R. Knight. These constructs were New Phytologist (2015) www.newphytologist.com

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introduced into A. tumefaciens strain LBA4404 and transformed by floral infiltration into A. thaliana Col, the Atnoa1 mutant and the rescued Atnoa1 line (C2-6), respectively. In vivo reconstitution of aequorin from expressed apoaequorin and coelenterazine was performed by the method of Zhao et al. (2013). In vivo Ca2+ concentrations were estimated according to the method of Baum et al. (1999). Student’s t-test was used to determine the significance of the results.

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Results Salicylic acid specifically induces root waving in Arabidopsis thaliana Exogenous SA reduced root elongation and enhanced root waving growth on an inclined agar surface in a dose-dependent manner, similar to the previously described effects of ethylene (Fig. 1a; Buer et al., 2003). To further investigate the influence of SA on root morphogenesis and waving phenotypes, we measured some parameters describing waving, such as the wave amplitude, wave length, wave tangent angle, and length of the root in the wild-type. With an increase in SA concentration, the amplitude and tangent angle of the wave increased (Fig. 1c,e), and the wave length and root elongation decreased (Fig. 1d,f), which indicates that SA increases the tightness of root waving. To confirm the specificity of SA-induced root waving, we compared the effects of SA analogs and other signal elements on root growth and development. Two SA analogs (5-sulfosalicylic acid and 3,5-dinitrosalicylic acid) had no effects on root growth (Supporting Information Fig. S1a). Similarly, hydrogen peroxide (H2O2), nitric oxide (NO), abscisic acid (ABA) and indole-3New Phytologist (2015) www.newphytologist.com

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Arabidopsis thaliana root cell protoplasts were isolated according to previous methods (Bai et al., 2009). Protoplasts with dense cytoplasm (diameter c. 15 mm) were patch clamped. The wholecell voltage-clamp currents in A. thaliana root cells were recorded with an EPC-9 amplifier (HEKA Electronik, Lambrecht, Germany) as described previously (Zhao et al., 2013). Standard solutions contained (in mM) 20 CaCl2, 0.1 DTT and 2 MES-Tris (pH 5.6) in the bath and 40 potassium glutamate, 10 tetraethylammonium chloride, 5 BAPTA (free Ca2+ adjusted to 100 mM with Ca(OH)2) and 10 HEPES-Tris (pH 7.1) in the pipette. DSorbitol was used to adjust the osmolarities of the bath and pipette solutions to 300 and 330 mmol kg1, respectively. Pipettes were pulled with a vertical puller (Narishige, Tokyo, Japan) modified for two-stage pulls. SA was added to the bath solutions at a final concentration of 30 lM. Hyperpolarizationactivated conductance appeared within a few minutes of exposure to SA. The Ca2+ current in individual cells was recorded before and 20 min after SA application. Data were analyzed using PULSEFIT (version 8.7; HEKA Electronik), IGOR (version 3.0; HEKA Electronik) and ORIGIN (version 7.0; Microcal, CA, USA) software.

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Fig. 1 Analysis of root waving patterns in response to salicylic acid (SA). (a) Images of 4-d-old seedlings of Arabidopsis thaliana wild-type (WT) grown on Murashige and Skoog (MS) medium that was supplemented with SA for 7 d. (b) Parameters describing waving: a, wave amplitude; b, wave length; c, wave tangent angle; d, primary root length. (c) Quantitative assessment of the wave amplitude. (d) Quantitative assessment of the wave length. (e) Quantitative assessment of the wave tangent angle. (f) Quantitative assessment of primary root length. The values are the average of three independent experiments (20–35 measurements each) with SD (*, P < 0.05; **, P < 0.01).

acetic acid (IAA) did not induce root waving (Fig. S1b), although these agents have been clearly identified as signal molecules in the regulation of root growth (Vieten et al., 2005; Bai et al., 2009). These results suggest that exogenous SA specifically induces the root waving responses. SA-induced root-waving responses are disrupted in Atnoa1, npr1 and not in npr3/4 single and double mutants As AtNOA1 is a functional A. thaliana circularly permuted GTPase (Moreau et al., 2008) and the root waving response is regulated by extra large and conventional G proteins (Pandey et al., 2008), we investigated phenotypic alterations in Atnoa1 mutants and in Atnoa1 mutants containing a 35S-AtNOA1 transgene (rescued Atnoa1; Guo et al., 2003) in response to SA. As shown in Fig. 2, SA-induced root waving responses were Ó 2015 The Authors New Phytologist Ó 2015 New Phytologist Trust

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introduced into A. tumefaciens strain LBA4404 and transformed by floral infiltration into A. thaliana Col, the Atnoa1 mutant and the rescued Atnoa1 line (C2-6), respectively. In vivo reconstitution of aequorin from expressed apoaequorin and coelenterazine was performed by the method of Zhao et al. (2013). In vivo Ca2+ concentrations were estimated according to the method of Baum et al. (1999). Student’s t-test was used to determine the significance of the results.

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Results Salicylic acid specifically induces root waving in Arabidopsis thaliana Exogenous SA reduced root elongation and enhanced root waving growth on an inclined agar surface in a dose-dependent manner, similar to the previously described effects of ethylene (Fig. 1a; Buer et al., 2003). To further investigate the influence of SA on root morphogenesis and waving phenotypes, we measured some parameters describing waving, such as the wave amplitude, wave length, wave tangent angle, and length of the root in the wild-type. With an increase in SA concentration, the amplitude and tangent angle of the wave increased (Fig. 1c,e), and the wave length and root elongation decreased (Fig. 1d,f), which indicates that SA increases the tightness of root waving. To confirm the specificity of SA-induced root waving, we compared the effects of SA analogs and other signal elements on root growth and development. Two SA analogs (5-sulfosalicylic acid and 3,5-dinitrosalicylic acid) had no effects on root growth (Supporting Information Fig. S1a). Similarly, hydrogen peroxide (H2O2), nitric oxide (NO), abscisic acid (ABA) and indole-3New Phytologist (2015) www.newphytologist.com

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Arabidopsis thaliana root cell protoplasts were isolated according to previous methods (Bai et al., 2009). Protoplasts with dense cytoplasm (diameter c. 15 mm) were patch clamped. The wholecell voltage-clamp currents in A. thaliana root cells were recorded with an EPC-9 amplifier (HEKA Electronik, Lambrecht, Germany) as described previously (Zhao et al., 2013). Standard solutions contained (in mM) 20 CaCl2, 0.1 DTT and 2 MES-Tris (pH 5.6) in the bath and 40 potassium glutamate, 10 tetraethylammonium chloride, 5 BAPTA (free Ca2+ adjusted to 100 mM with Ca(OH)2) and 10 HEPES-Tris (pH 7.1) in the pipette. DSorbitol was used to adjust the osmolarities of the bath and pipette solutions to 300 and 330 mmol kg1, respectively. Pipettes were pulled with a vertical puller (Narishige, Tokyo, Japan) modified for two-stage pulls. SA was added to the bath solutions at a final concentration of 30 lM. Hyperpolarizationactivated conductance appeared within a few minutes of exposure to SA. The Ca2+ current in individual cells was recorded before and 20 min after SA application. Data were analyzed using PULSEFIT (version 8.7; HEKA Electronik), IGOR (version 3.0; HEKA Electronik) and ORIGIN (version 7.0; Microcal, CA, USA) software.

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Fig. 1 Analysis of root waving patterns in response to salicylic acid (SA). (a) Images of 4-d-old seedlings of Arabidopsis thaliana wild-type (WT) grown on Murashige and Skoog (MS) medium that was supplemented with SA for 7 d. (b) Parameters describing waving: a, wave amplitude; b, wave length; c, wave tangent angle; d, primary root length. (c) Quantitative assessment of the wave amplitude. (d) Quantitative assessment of the wave length. (e) Quantitative assessment of the wave tangent angle. (f) Quantitative assessment of primary root length. The values are the average of three independent experiments (20–35 measurements each) with SD (*, P < 0.05; **, P < 0.01).

acetic acid (IAA) did not induce root waving (Fig. S1b), although these agents have been clearly identified as signal molecules in the regulation of root growth (Vieten et al., 2005; Bai et al., 2009). These results suggest that exogenous SA specifically induces the root waving responses. SA-induced root-waving responses are disrupted in Atnoa1, npr1 and not in npr3/4 single and double mutants As AtNOA1 is a functional A. thaliana circularly permuted GTPase (Moreau et al., 2008) and the root waving response is regulated by extra large and conventional G proteins (Pandey et al., 2008), we investigated phenotypic alterations in Atnoa1 mutants and in Atnoa1 mutants containing a 35S-AtNOA1 transgene (rescued Atnoa1; Guo et al., 2003) in response to SA. As shown in Fig. 2, SA-induced root waving responses were Ó 2015 The Authors New Phytologist Ó 2015 New Phytologist Trust

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Research 5 (b) WT Atnoa1 rif1-1 B3-1 C2-6

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Fig. 2 Salicylic acid (SA)-induced root waving responses are disrupted in Arabidopsis thaliana nitric oxide-associated protein1 (Atnoa1) mutants. (a) RT-PCR analysis of AtNOA1 gene expression levels in A. thaliana wildtype (WT), Atnoa1, resistant to inhibition by fosmidomycin (rif1-1), B3-1 and C2-6 plants. (b) Root wave morphology of WT, Atnoa1, rif1-1, B3-1 and C2-6 seedlings treated without or with 30 lM SA for 7 d. (c–f) Quantitative assessments of the wave amplitude, wave length, wave tangent angle and length of primary roots, respectively. The values are the average of three independent experiments (25–40 measurements each) with SD (**, P < 0.01).

disrupted in the Atnoa1 mutants, whereas these responses were rescued in the rescued Atnoa1 lines (B3-1 and C2-6). Similar root waving impairment is also found in another allelic mutant line, rif1-1 (Flores-P
erez et al., 2008). These results indicate that AtNOA1 is a potential intermediate involved in SA-induced root waving. If SA is responsible for root waving and AtNOA1 is a potential intermediate in this process, then what are the responses of mutants that are defective in SA perception and signaling? Here, we investigated npr1 mutants, in which SA-mediated plant defense signal transduction is defective as a result of the disruption of the interaction of the SA receptor (NPR3/NPR4) with NPR1 (Fu et al., 2012; Moreau et al., 2012). As expected, npr1 showed a defective root phenotype in response to SA, as found for Atnoa1 and rif1-1 (Fig. 3). However, similar defects were not found in npr3/4 single and double mutants (Fig. 4), which indicates that NPR3 and NPR4 do not appear to be critical for these responses. Subcellular localization of AtNOA1 To understand the mechanisms by which AtNOA1 mediates SAinduced root waving, some knowledge of where AtNOA1 Ó 2015 The Authors New Phytologist Ó 2015 New Phytologist Trust

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Fig. 3 Salicylic acid (SA)-induced root waving responses are disrupted in nonexpresser of pathogenesis-related genes1-1 (npr1-1) mutants. (a) RTPCR analysis of Arabidopsis thaliana NITRIC OXIDE-ASSOCIATED PROTEIN1 (AtNOA1) and NPR1 gene expression levels in A. thaliana wild-type (WT), npr1-1, Atnoa1 and resistant to inhibition by fosmidomycin (rif1-1) plants. (b) Root wave morphology of WT, npr1-1, Atnoa1 and rif1-1 seedlings treated without or with 30 lM SA for 7 d. (c) Partial nucleotide and deduced amino acid sequences of NPR1 cDNAs from the wild-type and the npr1-1 single mutant. Red nucleotides indicate the nucleotide replaced in npr1-1. The amino acid and nucleotide positions are indicated on the right. (d–g) Quantitative assessments of the wave amplitude, wave length, wave tangent angle and length of primary roots, respectively. The values are the average of three independent experiments (30–45 measurements each) with SD (*, P < 0.05; **, P < 0.01).

localizes, at a minimum, is required. Therefore, we transiently expressed constructs in which green fluorescent protein (GFP) was fused to the full-length 561 amino acid protein (NOA11-561GFP) or the N-terminal 200 amino acid fragment (NOA11-200GFP) of AtNOA1 in protoplasts of mesophyll cells. As shown in Fig. 5(a), the green fluorescence signal of NOA11-561-GFP (bottom panel) or NOA11-200-GFP (middle panel) was found in chloroplasts. By contrast, the green fluorescence signal of GFP as a control was found only in the cytoplasm (top panel), which reveals the importance of the N-terminal domain of AtNOA1 in the localization of AtNOA1 in chloroplasts. To further investigate the tissue specificity of AtNOA1 localization, we transformed an AtNOA1 null mutant of A. thaliana with a construct encoding translationally fused AtNOA1-GFP under the control of the cauliflower mosaic virus 35S promoter. The green fluorescence signal of AtNOA1-GFP was still found in New Phytologist (2015) www.newphytologist.com

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Fig. 4 Analysis of root waving patterns in nonexpresser of pathogenesisrelated genes3/4 (npr3/4) single and double mutants. (a) RT-PCR analysis of NPR3 and NPR4 gene expression levels in Arabidopsis thaliana wild-type (WT), npr3, npr4 and npr3npr4 plants. (b) Root wave morphology of WT, npr3, npr4 and npr3npr4 seedlings treated without or with 30 lM salicylic acid (SA) for 7 d. (c–f) Quantitative assessments of the wave amplitude, wave length, wave tangent angle and length of primary roots, respectively. The values are the average of three independent experiments (22–30 measurements each) with SD (*, P < 0.05; **, P < 0.01).

the chloroplasts of mesophyll and hypocotyl cells (Fig. 5b,c), which is consistent with previous findings (Flores-P
erez et al., 2008; Liu et al., 2010). Interestingly, the AtNOA1-GFP transgenic lines showed weaker GFP fluorescence in roots without SA application (Fig. 5d, left), and the application of SA significantly enhanced the green fluorescence signal which localized largely on the cell surface (Fig. 5d, middle). To differentiate between the plasma membrane and the cell wall, the root cells that contained AtNOA1-GFP were plasmolysed with 0.8 M mannitol. The SAinduced AtNOA1-GFP protein was found mainly in the plasma membrane of root cells, but not in any other organelles (Fig. 5d, right). Correspondingly, the AtNOA1-GFP construct complements the Atnoa1 mutant phenotype, restoring the SA-induced root waving (Fig. 5e). SA enhances the expression of AtNOA1 The increase in SA-induced fluorescence in AtNOA1-GFP transgenic lines described in the previous section is reminiscent of the regulation of the expression of AtNOA1. Here, quantitative realtime PCR analysis showed that SA enhanced the expression of AtNOA1 in wild-type plant root cells in a dose-dependent manner (Fig. 6a). Western blotting, with polyclonal antisera against New Phytologist (2015) www.newphytologist.com

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Fig. 5 Subcellular localization of Arabidopsis thaliana NITRIC OXIDEASSOCIATED PROTEIN1 (AtNOA1) in mesophyll cells, the hypocotyl and roots of A. thaliana seedlings. (a) AtNOA1-GFP localization in a leaf protoplast cell in a transient assay: top panel, green fluorescence signal of GFP as a control; middle panel, green fluorescence signal of AtNOA11-200GFP; bottom panel, green fluorescence signal of AtNOA11-561-GFP. (b) Confocal images of transgenic mesophyll cells showed that AtNOA1-GFP was localized in chloroplasts. (c) Confocal images of transgenic hypocotyl cells showed that AtNOA1-GFP was localized in chloroplasts. Inset, details of AtNOA1-GFP in hypocotyl cells. (d) Confocal images of transgenic root cells showed that AtNOA1-GFP was localized on the cell surface (left), and that salicylic acid (SA) enhanced protein expression (middle). After the root cells had been plasmolyzed with 0.8 M mannitol, SA-induced AtNOA1GFP was associated with the plasma membrane in root cells (right). Bar, 20 lm. (e) Root wave morphology of wild-type (WT), Atnoa1 and Atnoa1 with AtNOA1-GFP seedlings treated with or without 30 lM SA for 7 d.

the 19 N-terminal amino acids of AtNOA1 (Guo et al., 2003), confirmed the restoration of AtNOA1 expression in rescued lines, and SA significantly enhanced AtNOA1 protein expression in a time-dependent manner (Fig. 6b). Next we also examined the Ó 2015 The Authors New Phytologist Ó 2015 New Phytologist Trust

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Fig. 6 Expression pattern of Arabidopsis thaliana NITRIC OXIDEASSOCIATED PROTEIN1 (AtNOA1) in the roots of A. thaliana seedlings obtained using quantitative real-time PCR and western blotting. (a) Quantitative real-time PCR analysis of AtNOA1 gene expression levels in wild-type (WT) in response to salicylic acid (SA) at different concentrations. The values are the average of three independent experiments with SD (*, P < 0.05; **, P < 0.01). (b) Western blot analysis of AtNOA1 with anti-AtNOA1 in the roots of WT, Atnoa1, C2-6 and B3-1 (left) and in the roots of WT treated with 30 lM SA for a different number of days (right). To determine AtNOA1 levels, 20 lg (left) or 40 lg (right) of protein was loaded in each lane. H+-ATPase detected using a polyclonal antibody raised against the individual protein was used as a loading control. (c) Western blot analysis of AtNOA1 with anti-GFP in the roots of Atnoa1 with AtNOA1-GFP treated with 30 lM SA, 5-sulphosalicylic acid or 3,5-dinitrosalicylic acid and 40 lM sodium nitroprusside (SNP), respectively. To determine AtNOA1 levels, 40 lg of protein was loaded in each lane. (d) Western blot analysis of AtNOA1 with anti-AtNOA1 in the roots of WT and npr1 treated with 30 lM SA. To determine AtNOA1 levels, 40 lg of protein was loaded in each lane. The numbers on the left indicate the molecular masses. Experiments conducted three times on different occasions gave similar results.

abundance of AtNOA1 proteins using the AtNOA1-GFP transgenic lines, and found that SA significantly enhanced the AtNOA1 protein level and two SA analogs and sodium nitroprusside (SNP, a NO donor) had no effect (Fig. 6c). However, the AtNOA1 protein induction in the npr1 mutant in response to SA was blocked (Fig. 6d). Together, these results Ó 2015 The Authors New Phytologist Ó 2015 New Phytologist Trust

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Fig. 7 Salicylic acid (SA)-induced cytosolic Ca elevation was decreased in Arabidopsis thaliana nitric oxide-associated protein1 (Atnoa1) plants. (a) Roots undergoing tip growth in A. thaliana plants expressing the Ca2+ sensor YC3.6 targeted to the cytosol were imaged. Representative results of c. 15 measurements are shown. Bar, 100 lm. (b) Quantitative analysis of average cytosolic Ca2+ concentrations in representative growing roots, as indicated in the inset of (a). The increase in the fluorescence resonance energy transfer (FRET)/CFP ratio reflects an increase in the cytoplasmic Ca2+ concentration. The vertical bars indicate the average of three independent experiments (12–18 measurements each) with SD (**, P < 0.01).

indicate that AtNOA1 expression induced by SA via the regulation of NPR1 possibly plays pivotal roles in root waving growth. AtNOA1 mediates SA-induced cytosolic Ca2+ increases As root growth and development involve changes in cytosolic Ca2+ (Bai et al., 2009), we evaluated cytosolic Ca2+ changes in the wild-type, the Atnoa1 mutant and the rescued Atnoa1 lines in response to SA. As shown in Fig. 7, SA at 30 lM significantly increased cytosolic Ca2+ concentrations in Yellow Cameleon (YC) 3.60-expressing roots of wild-type plants. By contrast, SA had no effects on cytosolic Ca2+ concentrations in Atnoa1 mutant roots. A 35S-AtNOA1 transgene substantially rescued SAinduced increases in cytosolic Ca2+ concentrations in Atnoa1 mutant roots. Similar results were also obtained in A. thaliana roots containing transgenic aequorin (Fig. 8). Moreover, the SAinduced cytosolic Ca2+ increases were efficiently inhibited by LaCl3 (a channel blocker) and ethylene glycol tetraacetic acid (EGTA; an extracellular Ca2+ chelating agent). Consistently, the application of these two agents efficiently disrupted SA-induced root waving (Fig. S2). These results suggest that SA-induced New Phytologist (2015) www.newphytologist.com

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Fig. 8 Changes in cytosolic Ca2+ concentration induced by salicylic acid (SA) in the roots of Arabidopsis thaliana wild-type (WT), nitric oxideassociated protein1 (Atnoa1) and C2-6. A single seedling expressing aequorin, pretreated or not with 2 mM EGTA and 1 mM LaCl3, was kept in the dark for 30 min. Cytosolic Ca2+ concentrations in the roots of the single seedling were measured by monitoring changes in aequorin-emitted luminescence before and after application of 30 lM SA for 15 s. The bars indicate the average of four independent experiments (22–36 measurements each) with SD (**, P < 0.01).

cytosolic Ca2+ changes are dependent on AtNOA1 and that this cytosolic Ca2+ increase is mainly attributable to an extracellular Ca2+ influx pathway involved in root waving. To further confirm the Ca2+ source, we applied the patchclamp technique to record whole-cell Ca2+ currents in A. thaliana root cell protoplasts in response to SA. When the protoplasts were kept in darkness before the experiments and illuminated with dim light during the process of patch-pipette attachment, no significantly increased channel activity was recorded. After the application of exogenous SA, a time-dependent increase in Ca2+ currents was recorded in the wild-type plants (Fig. 9a). However, SA failed to induce an increase in Ca2+ currents in Atnoa1 mutants (Fig. 9b), as well as in the npr1 mutant (Fig. 9d). A 35S-AtNOA1 transgene significantly rescued the impairment in SA-activated Ca2+ currents in Atnoa1 mutants (Fig. 9c). Together, these results reinforce the finding that the AtNOA1-mediated transient increase in cytosolic Ca2+ is modulated by SA mainly through the activation of plasma membrane Ca2+ channels, and SA signaling pathways possibly play important roles in this process. AtNOA1 is involved in SA-modulated auxin distribution in roots It is well known that auxin is required to initiate the formation of lateral roots (Celenza et al., 1995) and that polar auxin transport is modulated by cytosolic Ca2+ changes (Zhang et al., 2011). Considering that the increase in cytosolic Ca2+ is responsible for SA-induced root waving (Figs 7–9, S2), we next monitored the expression of auxin in AtNOA1-related genetic lines in response to SA using the auxin response reporter DR5-GUS (Ulmasov et al., 1997), and found that the strong DR5-GUS staining observed in the root tips of the wild-type and of the Atnoa1 mutant decreased in Atnoa1 following treatment with SA but remained stable in the wild-type. A 35S-AtNOA1 transgene significantly rescued the impairment in SA-inhibited auxin New Phytologist (2015) www.newphytologist.com

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Fig. 9 Changes of calcium channel activity induced by salicylic acid (SA) in root cells of Arabidopsis thaliana. (Left) Current response of individual cell-attached patches to 1-s voltage ramps from 190 to +30 mV before and after treatment with 30 lM SA. (Right) Channel activation by SA is expressed as the current increase with 30 lM SA treatment [MI190 mV = (Ibefore  Iafter)]. Each bar corresponds to the SA-dependent current difference obtained from one experiment. Current amplitudes were measured at a command voltage of 190 mV and averaged for 10 to 15 ramps. (a–d) Changes of calcium channel activity induced by SA in root cells of wild-type (WT), nitric oxide-associated protein1 (Atnoa1), C2-6 and nonexpresser of pathogenesis-related genes1-1 (npr1-1), respectively.

expression in root tips of the Atnoa1 mutant (Fig. 10a). Consistently, SA still induced AtNOA1-dependent root waving in these DR5-GUS transgenic lines (Fig. 10b), which indicates that auxin homeostasis or transport in the root possibly plays a crucial role in root waving. To further confirm the effects of SA on auxin distribution in the roots, we analyzed the expression of DR5-GFP, a highly active auxin reporter gene, in roots (Fig. 10c). Our data showed a strong GFP signal in wild-type root tips, which appeared in the quiescent center and the central root cap cells, including columella initials and mature columella cells, and the application of SA slightly reduced the expression of DR5-GFP. By contrast, Atnoa1 roots exhibited a similar expression pattern of DR5-GFP, but the GFP signal was weaker than that in the wild-type. Interestingly, the GFP signal almost disappeared in columella root cap cells of Atnoa1 mutants in response to SA, and the SA-induced waving was accordingly impaired in these lines. A 35S-AtNOA1 Ó 2015 The Authors New Phytologist Ó 2015 New Phytologist Trust

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leading to root morphological responses (Luschnig et al., 1998). As shown in Fig. 11, AtPIN2-GFP localized apically in the epidermal cell plasma membrane and basally in the cortical cell plasma membrane, as previously reported in roots (Blilou et al., 2005). After treatment with SA, AtPIN2-GFP was greatly reduced in the wild-type root cell plasma membrane, showing less polar localization, and some AtPIN2-GFP also appeared in the cytosol (Fig. 11a–c). Mutation of AtNOA1 reduced AtPIN2GFP protein expression in root epidermal and cortical cells, and relocation of AtPIN2-GFP in the cytoplasm was not found (Fig. 11d–f). In the roots of two rescued Atnoa1 lines, the expression of a 35S-AtNOA1 transgene significantly rescued the impairment in both the polar expression of AtPIN2-GFP and SA-induced AtPIN2-GFP relocation in the cytoplasm from the plasma membrane (Fig. 11g–l). Considering the functional redundancy and complexity of PIN family members, we also examined the expression and protein localization of AtPIN3 and AtPIN7, which mainly act in the root tips (Krecek et al., 2009). As shown in Fig. 12, PIN3-GFP and PIN7-GFP in root cap columella cells have similar expression patterns without pronounced polarity. SA and mutation of AtNOA1 significantly reduced AtPIN3/7 expression, which is similar to the AtPIN2 expression changes found here. These results indicate that PIN family members possibly play a cooperative role in SA-mediated root growth and development.

Discussion Regulation of SA on root growth and development

C2-6

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Fig. 10 Arabidopsis thaliana NITRIC OXIDE-ASSOCIATED PROTEIN1 (AtNOA1) is involved in salicylic acid (SA)-modulated auxin distribution in the roots of A. thaliana seedlings. (a) DR5-GUS auxin reporter analysis of tissue expression of auxin in the root tips of A. thaliana wild-type (WT), Atnoa1 and C2-6. White arrows indicate different expression of DR5-GUS in root tips after 3 d of 30 lM SA treatment. (b) Root wave morphology of DR5-GUS transgenic seedlings treated with or without SA for 7 d. (c) Images of the GFP channel and differential interference contrast channel were combined to show the expression pattern of DR5-GFP in root tips of WT, Atnoa1 and C2-6 in response to treatment with SA (right panels) or no treatment with SA (left panels). White arrows indicate different expression of GFP in root tips. (d) Root wave morphology of DR5-GFP transgenic seedlings treated with or without SA for 7 d.

transgene significantly complemented these phenotypes, restoring the GFP signal in roots and SA-induced root waving (Fig. 10d). These results indicate that the generation of auxin maxima and gradients (polar transport) is probably an important event in AtNOA1-mediated root waving. AtPIN2 expression and protein localization are changed in roots in response to SA It has been reported that AtPIN2, which is mainly expressed in the root, has a root-specific role in the transport of auxin, Ó 2015 The Authors New Phytologist Ó 2015 New Phytologist Trust

SA is involved in many physiological processes in plants, and in particular plays important roles in the adaptation of plants to adverse growth conditions (Raskin, 1992). However, the mechanisms of its regulation of root growth and development remain to be determined in detail, although a previous study revealed that SA effectively modulates the growth of roots in soybean (Glycine max) (Guti
errez-Coronado et al., 1998). Here, we found that SA specifically induced root waving, with an overall decrease in root elongation, in A. thaliana (Fig. 1), which resembles the action of ethylene (Buer et al., 2003). Considering that SA, as a stress signal molecule, is usually associated with other signal molecules, such as H2O2, NO, ABA and IAA, we investigated whether these signal molecules also induced similar root waving phenotypes. This was not the case (Fig. S1). In fact, root architecture is determined by genetic factors and the integration of environmental cues (Malamy, 2005; Gruber et al., 2013). Interestingly, Castresana’s group found that 9-lipoxygenase-derived oxylipins, via a hydroperoxide pathway, induced three different root phenotypic alterations: root waving with lateral root arrest, growth arrest with loss of apical dominance, and an overall decrease of elongation, and that the phytohormone jasmonic acid (JA), which is a cyclic oxylipin, did not induce root waving (Vellosillo et al., 2007). Although SA is associated with JA in plant defense responses (Robert-Seilaniantz et al., 2011), they exhibit different effects on root morphological phenotypes, which New Phytologist (2015) www.newphytologist.com

New Phytologist indicates that different signal pathways are involved in the response to SA and JA. As SA contributes to the regulation of root waving, we infer that the perception of SA and early SA signaling events are possibly involved in this process. In this study, we found that NPR1 mutation efficiently blocked SA-induced root waving (Fig. 3), indicating the importance of NPR1 (a critical SA signal transducer) in root growth and development in response to SA. Unexpectedly, npr3/4 single and double mutants showed normal responses to SA in terms of root waving (Fig. 4). In the defense response, SA regulates the conversion of NPR1 from an oligomeric to a monomeric form, which leads to its translocation from the cytosol to the nucleus to induce defense genes (Mou et al., 2003). SA also regulates cytosolic NPR1 degradation by its binding to NPR3/NPR4 in a concentration-dependent manner (Fu et al., 2012). Thus, NPR1 homeostasis is important for plants to combat pathogens. Considering that NPR3 and NPR4 may not be SA receptors in a traditional sense (Moreau et al., 2012) and the npr3npr4 double mutant has the opposite phenotype from npr1 in that it exhibits enhanced disease resistance (Zhang et al., 2006), we infer that alternative NPRl-dependent SA signaling is possibly involved in root waving, which is different from the plant defense response. In root waving, are NPR3 and NPR4 necessary? How does SA regulate NPR1 turnover in detail? What is the spatial-temporal relationship among NPR1, AtNOA1 and signal intermediates related to root waving? These questions will be addressed in a future study. Function of AtNOA1 in root waving Given that AtNOA1 mediates NO production during plant development and phytohormone actions as an NO-associated protein (Guo et al., 2003; Moreau et al., 2008), and that SA effectively activates NOS (AtNOA1) and induces NO production in roots (Zottini et al., 2007), we examined the effects of SA on the root growth of nitrate reductase-deficient mutants (nia1 and nia2), in which NO production is impaired (Desikan et al., 2002). Unexpectedly, SA still obviously induced root wavy growth in nia1 and nia2 mutants (Fig. S3), which contrasts with the results obtained for the Atnoa1 mutant (Fig. 2). These findings, together with those of the SNP experiment (Fig. S1b), suggest that AtNOA1 functions in SA-induced root waving via an NO-independent pathway. Unlike classical small GTPases such as Rho, Ras, and Ran GTPases, which have been extensively studied, little is known about the cGTPase family of AtNOA1 (Moreau et al., 2008). However, some studies have indicated that extra large and conventional G proteins positively regulate root waving and root slanting (Pandey et al., 2008), and G-protein subunits have been shown to have distinct roles in the modulation of cell division in roots (Chen et al., 2006), which sheds light on the possible role of AtNOA1. Although some groups have investigated its activity regulation as a cGTPase in vitro (Guo et al., 2003; Moreau et al., 2008), we failed to examine the activity of this cGTPase in response to SA in vivo. However, the green fluorescence signal associated with AtNOA1-GFP expression was significantly Ó 2015 The Authors New Phytologist Ó 2015 New Phytologist Trust

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enhanced by SA application to the surface of root cells (Fig. 5d), which was reminiscent of the SA-induced increase in protein level. Because the transgene, which is controlled by the 35S promoter, should not be responsive to SA at the transcriptional level, we investigated the effect of SA on AtNOA1 expression in the nontransgenic wild-type (Fig. 6a,b) and AtNOA1-GFP transgenic Atnoa1 (Fig. 6c). Undoubtedly, SA enhanced the abundance of AtNOA1 protein (Fig. 6b,c), and mutation of NPR1 impaired the SA-induced protein increase (Fig. 6d). Moreover, the AtNOA1 protein changes were closely correlated with the root wavy phenotype alternations, indicating a role of AtNOA1 proteins regulated by SA in root waving. The localization analysis showed that AtNOA1 was targeted to chloroplasts in A. thaliana mesophyll and hypocotyl cells (Fig. 5a,b,c), which is essential for proper chlorophyll biosynthesis, Rubisco formation and plastid development (Liu et al., 2010; Yang et al., 2011). Although roots do not develop chloroplasts, root cells contain other plastid types, such as etioplasts, which become amyloplasts in some cell types such as the columella cells. Here, we unexpectedly found that AtNOA1 was expressed weakly in the root plasma membrane and SA enhanced this expression (Fig. 5d). Unfortunately, at present, we do not know how SA-induced AtNOA1 proteins localize to the plasma membrane. In A. thaliana, AtNOA1 contains guanine-binding motifs (G motifs) characteristic of small GTPases such as Rho and Ras (Moreau et al., 2008), and Rhorelated GTPase from plants (ROPs) are involved in plasma membrane-associated polar growth and Ca2+ influx (Yan et al., 2009; Lin et al., 2012; Yang & Lavagi, 2012). Therefore, we infer that AtNOA1, as a potential membrane-associated intermediate, possibly plays a crucial role in root waving signal transduction across the plasma membrane. Role of Ca2+ in SA-induced root waving As AtNOA1 localizes to the plasma membrane in roots, how does it function? Given that the cell wall is a potential Ca2+ source and that Ca2+ is involved in diverse cellular functions as a universal second messenger (Chen et al., 2008), we investigated whether AtNOA1 mediates Ca2+ signaling in SA-induced root waving. In this study, genetic and pharmacological analyses showed that the AtNOA1-mediated cytosolic Ca2+ increase was attributable to SA-activated plasma membrane Ca2+ channels, and NPR1 possibly plays a crucial role in this process (Figs 7–9). To further confirm the association between Ca2+ and root waving, we examined root phenotypic alterations following application of LaCl3 and EGTA, and found that these Ca2+-related inhibitors efficiently impaired SA-induced root wavy responses (Fig. S2). An increase in cytosolic Ca2+ and the activation of Ca2+ channels in root cells are thought to be essential for root growth and root hair development (Bibikova et al., 1999; Bai et al., 2009). Moreover, Ca2+ participates in feedback regulation of the G protein, which is involved in ion channel regulation (Aharon et al., 1998; Wang et al., 2001; Yan et al., 2009). Here, SA-induced root waving with an overall decrease in root elongation was mainly attributed to AtNOA1-mediated Ca2+ homeostasis, and in turn Ca2+ possibly regulates AtNOA1 activity (Guo et al., 2003). New Phytologist (2015) www.newphytologist.com

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Interplay between SA and auxin transport in root waving As the root waving pattern is the result of positive gravitropism plus a thigmotropic effect (Okada & Shimura, 1990; Thompson & Holbrook, 2004), how do plant roots respond to this gravistimulation? In plants, the gravitropic response involves perception of the gravistimulus by the root cap columella cells, transduction of that physical information into physiological signals, transmission of the signals to the distal and central elongation zones, and a curvature response (Chen et al., 1998). Although the molecular mechanisms underlying these changes are poorly understood, there is considerable evidence showing that polar auxin transport or distribution is involved in the response to gravity. Here, histological examination of DR5-GUS activity showed a decrease in root tips, especially in Atnoa1 mutants, in response to SA (Fig. 10a), and a further DR5-GFP assay indicated that this decrease in green fluorescence signal was found primarily in columella root cap cells (Fig. 10c). Notably, AtNOA1 was critical for SA-dependent root waving. Yet, when the pattern and level of expression of DR5-GFP were analyzed, SA was found to remain functional even in the Atnoa1 mutant background. A potential explanation is that SA and AtNOA1 have an additive effect, rather than these molecules functioning successively in a single pathway. In fact, SA also regulates the length of roots, as well as root waving (Fig. 1a; Armengot et al., 2014). Although we still do not know the relationship between auxin maxima (or transport) and root waving, the changes in auxin distribution in columella root cap cells, which are recognized as gravity sensing sections (Evans, 1991), possibly result from the alteration of auxin polar transport. It is well established that the PIN-FORMED (PIN) auxin efflux facilitator network controls growth and patterning in A. thaliana roots by modulating the auxin maximum and polar auxin transport (Blilou et al., 2005; Krecek et al., 2009). AtPIN2, a member of the PIN protein family in A. thaliana, plays an important role in the control of root morphological responses, including waving, by regulating the redistribution of auxin (Okada & Shimura, 1990; M€ uller et al., 1998). The function of AtPIN2 in waving formation is not known; however, several independently identified mutant alleles of AtPIN2, ethylene insensitive root1 (eir1), agravitropic1 (agr1), and wavy6 (wav6), indicate a role of AtPIN2 in gravity-dependent auxin transport and root waving (Chen et al., 1998; Luschnig et al., 1998; Utsuno et al., 1998). In this study, AtPIN2-GFP in the wild-type root epidermal and cortical cells decreased in the plasma membrane and appeared in cytosolic sections after treatment with SA, and polar distribution and recycling (or degradation) of AtPIN2-GFP were impaired in Atnoa1 mutants (Fig. 11), which indicates that a GTPase-dependent auxin signaling pathway regulates the subcellular distribution of AtPIN2 in A. thaliana roots (Lin et al., 2012). Notably, considering that Atnoa1 displayed a much weaker PIN2-GFP signal at the plasma membrane in epidermal and cortical cells than the wild-type (Fig. 11a,d), one would expect a decrease in the rate of shootward auxin transport, resulting in auxin accumulation in the root cap. Considering that SA promotes PIN2 internalization, at least in the wild-type New Phytologist (2015) www.newphytologist.com

(Fig. 11a,b), one would expect an even slower rate of shootward transport upon SA treatment, with more auxin in the root tip. Yet, such an accumulation was not observed. Instead, a lower level of expression of the auxin reporters in the cap of mutant roots relative to wild-type was found. Furthermore, SA treatment led to a further decrease in signal intensity, especially in the mutant (Fig. 10c). These perplexing observations suggest that something more complex is occurring here, either in the root cap columella cells themselves, or in the rootward transport of auxin toward the root cap. Here, we examined the expression pattern of AtPIN3 and AtPIN7 in root tips, and found that similar expression changes occurred in mutant roots relative to wild-type (Fig. 12). AtPIN3 and AtPIN7 localized mainly in root cap columella cells without pronounced polarity, which is different from AtPIN2 polar localization. The accumulation of auxin in root tips results from the cooperative effects of PIN-mediated shootward and rootward transport of auxin. Altered rootward transport of auxin from shoot to root tip may also be involved in SA- and AtNOA1-modulated auxin distribution in the roots, leading to less auxin arriving in the tip of Atnoa1 mutant roots relative to wild-type. In future studies, quantification of both shootward and rootward streams of auxin transport in the root tips of control (untreated) and SA-treated wild-type and Atnoa1 mutant seedlings, along with an analysis of auxin concentrations, would improve our understanding of the mechanisms involved in these responses.

Acknowledgements We thank Nigel M. Crawford (University of California, CA, USA) for providing the Atnoa1, B3-1 and C2-6 seeds, Manuel Rodriguez-Concepcion (Centre for Research on Agricultural Genomics, Spain) for providing the rif1-1 seeds, Xinnian Dong (Duke University, NC, USA) for providing the npr1-1 seeds, Zhizhong Gong (China Agricultural University, China) for providing the AtPIN3-GFP and AtPIN7-GFP seeds, Jian Xu (Huazhong Agricultural University, China) for providing the AtPIN2-GFP, DR5-GFP and DR5-GUS seeds, Marc R. Knight (University of Cambridge, UK) for providing aequorin expression vectors cloned into Agrobacterium tumefaciens strain GV3101, and Takeharu Nagai (Laboratory for Cell Function and Dynamics, Japan) for providing YC3.6 expression vectors. We also thank Dr Chunpeng Song, Pengcheng Wang and Yuchen Miao for their technical support and helpful discussions. This work was supported by the National Natural Science Foundation of China (grant nos. 31170271 and 31101023) and by Genetically Modified Organisms Breeding Major Projects of China (grant no. 2013ZX08005-004).

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Supporting Information Additional supporting information may be found in the online version of this article. Fig. S1 Salicylic acid (SA) specifically induces root waving in Arabidopsis thaliana. Fig. S2 Effects of various inhibitors on root waving induced by salicylic acid (SA) in Arabidopsis thaliana wild-type (WT), Atnoa1, B3-1 and C2-6. Fig. S3 Analysis of root waving induced by salicylic acid (SA) in Arabidopsis thaliana wild-type (WT), Atnoa1, rif1-1, nia1 and nia2. Please note: Wiley Blackwell are not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing material) should be directed to the New Phytologist Central Office.

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