JXB Advance Access originally published online on February 14, 2007
Journal of Experimental Botany 2007 58(6):1261-1270; doi:10.1093/jxb/erl279
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© 2007 The Author(s).
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/2.0/uk/) which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.This paper is available online free of all access charges (see http://jxb.oxfordjournals.org/open_access.html for further details)
RESEARCH PAPER |
NADPH oxidase-dependent reactive oxygen species formation required for root hair growth depends on ROP GTPase
1School of Biosciences, University of Exeter, Geoffrey Pope Building, Stocker Road, Exeter EX4 4QD, UK
2Center for Plant Cell Biology, Department of Botany and Plant Sciences, University of California, Riverside, CA 92521-0124, USA
* To whom correspondence should be addressed. E-mail: N.Smirnoff{at}exeter.ac.uk
Received 15 September 2006; Accepted 15 November 2006
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Abstract |
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Reactive oxygen species (ROS) production by an NADPH oxidase (NOX) encoded by AtrbohC/RHD2 is required for root hair growth in Arabidopsis thaliana. ROP (RHO of plants) GTPases are also required for normal root hair growth and have been proposed to regulate ROS production in plants. Therefore, the role of ROP GTPase in NOX-dependent ROS formation by root hairs was investigated. Plants overexpressing wild-type ROP2 (ROP2 OX), constitutively active (CA-rop2), or dominant negative (DN-rop2) rop2 mutant proteins were used. Superoxide formation by root hairs was detected by superoxide dismutase-sensitive nitroblue tetrazolium reduction, and ROS production in the root hair differentiation zone was detected by dihydrofluorescein diacetate oxidation. Both probes showed that ROS production was increased in ROP2 OX and CA-rop2 plants, and decreased in DN-rop2 plants, relative to wild-type plants. When CA-rop2 was expressed in the NOX loss-of-function rhd2-1 mutant, ROS formation and root hair growth were impaired, suggesting that RHD2 is required for this ROP2-dependent ROS formation.
Key words: AtrbohC/RHD2, NADPH oxidase, root hairs, ROP GTPase, superoxide
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Introduction |
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Plants can generate extracellular superoxide via plasma membrane (PM)-localized NADPH oxidases (NOXs), which are homologous to the catalytic subunit (gp91phox) of mammalian phagocyte NOX. Arabidopsis thaliana has a family of 10 NOX genes, termed Atrboh (Arabidopsis thaliana respiratory burst oxidase homologues; Keller et al., 1998; Torres et al., 1998; Foreman et al., 2003). They catalyse the extracellular formation of the superoxide anion (O2·–) from molecular oxygen using NADPH as an electron donor (Sagi and Fluhr, 2001). Extracellular superoxide readily gives rise to other reactive oxygen species (ROS) including hydrogen peroxide, by dismutation, and the hydroxyl radical via the Fenton reaction (Halliwell and Gutteridge, 1989). Hydrogen peroxide readily crosses the plant PM through water channels (Henzler and Steudle, 2000) and can subsequently act as an intracellular (Kovtun et al., 2000) and intercellular (Allan and Fluhr, 2001) signalling molecule. Investigation of mutants in members of the Atrboh family have shown that they contribute to superoxide and hydrogen peroxide formation in the oxidative burst caused by pathogen infection (Torres et al., 2002). ROS generated by NOX are also involved in mediating stomatal closure induced by the hormone abscisic acid (ABA; Kwak et al., 2003). Evidence is accumulating that NOX activity is required for plant development because mutants in various Atrboh genes are affected in growth and development. Plant size and root length are reduced in knockout mutations of AtrbohF and in the AtrbohD/F double mutant (Torres et al., 2002; Kwak et al., 2003), while root length is reduced and root hair elongation is prevented in AtrbohC knockout mutants (Foreman et al., 2003). Tomato plants expressing plant NOX RNA interference (RNAi) transgenes have highly pleiotropic developmental abnormalities (Sagi et al., 2004). ROS have been more indirectly implicated in controlling the deposition of secondary cell walls in developing cotton fibres (Potikha et al., 1999) and in maize leaf growth (Rodriguez et al., 2002) using inhibitors of NOX such as diphenylene iodonium (DPI). However, since DPI can also inhibit other enzymes, including peroxidases (Frahry and Schopfer, 1998), its use does not provide conclusive proof of NOX involvement.
Given the emerging roles of NOX-generated ROS in plant developmental processes, the regulation of NOX activity is an important consideration. The mammalian phagocyte NOX consists of a complex of different regulatory subunits, in addition to the NOX catalytic subunit, gp91phox. No homologues of these regulatory subunits have been identified in plants (Sagi and Fluhr, 2001). It is also well established that a RHO GTPase (Rac2) is required for activation of the phagocyte gp91phox (Diekmann et al., 1994; Diebold and Bokoch, 2001). Rac2 regulates the initial electron transfer from NADPH to gp91phox-associated FAD through a physical interaction between the Rac insert domain and gp91phox. The subsequent electron transfer from gp91phox-associated FAD to gp91phox-associated haem, which precedes the final electron transfer to molecular oxygen, requires Rac2 to interact with another regulatory subunit of the NADPH oxidase complex, p67phox (Diebold and Bokoch, 2001). There is also strong evidence that ROPs (RHO of plants) regulate NOX activity. ROPs form a unique subfamily of RHO, within the RAS superfamily of small GTPase molecular switches, with 11 ROP genes in the A. thaliana genome (Yang, 2002). Although there is, as yet, no evidence to show that a specific ROP activates a specific plant NOX, ROPs do influence DPI-sensitive ROS formation in planta (Yang, 2002; Agrawal et al., 2003). In rice, both ROS-mediated cell death (Kawasaki et al., 1999) and ROS formation during disease resistance are dependent on the ROP OsRac1 (Ono et al., 2001). Significantly, ROP2 is involved in DPI-sensitive ROS formation in response to hypoxia in A. thaliana roots (Baxter-Burrell et al., 2002) and active GTP-bound ROP2 activates superoxide production in vitro (Park et al., 2004). Both ROP GTPases and NOX-generated ROS have, separately, been implicated in ABA-induced stomatal closure, although no direct link has been established between any members of these two gene families (Pei et al., 2000; Lemichez et al., 2001; Zhang et al., 2001; Zheng et al., 2002; Kwak et al., 2003). There may be a high degree of functional and structural conservation between plant and animal RHO GTPases and the corresponding NOXs: maize constitutively active (CA)-rop can activate ROS formation in mammalian cells (Hassanain et al., 2000) and human Rac can activate DPI-sensitive ROS formation in plant cells (Park et al., 2000). These results suggest that ROPs are strong candidates to control plant NOXs in vivo.
The rhd2 mutant of A. thaliana has root hairs that do not elongate. In a seminal paper, Foreman et al. (2003) have shown that RHD2 encodes AtrbohC, and superoxide produced by activity of the NOX encoded by AtrbohC/RHD2 is needed for root hair elongation. As DPI-treated wild-type (WT) root hairs resemble the total loss-of-function rhd2 mutant and as ROS formation is much lower in loss-of-function rhd2 root hairs (Foreman et al., 2003), it appears that AtrbohC/RHD2 is the major source of ROS in growing root hairs. Root hairs form in a root hair differentiation zone (RHDZ) further from the root tip than the elongation zone (Dolan et al., 1994). The first visible stage of root hair morphogenesis is a localized swelling, and tip growth is subsequently established at this site (Dolan et al., 1994). Root hair tip growth requires a high intracellular Ca2+ gradient in the tip and root hairs show a tip-localized influx of extracellular Ca2+ (Bibikova et al., 1997; Wymer et al., 1997). Hyperpolarization-activated Ca2+ channels that might be involved in Ca2+ influx have been identified in the root hair tip (Véry and Davies, 2000; Demidchik et al., 2003). We have previously shown that the ROP2 GTPase is required for all stages of root hair morphogenesis and is expressed in growing root hair cells (Jones et al., 2002) by using transgenic plants expressing a CA mutant of ROP2 locked in the active GTP-bound form and a dominant negative (DN) form locked in the inactive GDP-bound form (Li et al., 2001). CA-rop2 and DN-rop2 transgenic plants have long, depolarized root hairs, and short root hairs, respectively, whereas overexpression of the native ROP2 gene (ROP2 OX) results in multiple swelling formation on the same root hair cell and highly branched root hair tips (Jones et al., 2002). Proposed targets for ROP2 action include the actin cytoskeleton (Jones et al., 2002).
The requirement for both AtrbohC/RHD2 and ROP2 in root hair elongation raises the possibility that ROS formation in root hairs could depend on the ROP2 GTPase. As has been discussed above, the Rac2 GTPase activates mammalian phagocyte NOX and there is abundant circumstantial evidence for this interaction in plants. Recently, it has been shown that A. thaliana root hair growth and associated ROS localization is spatially regulated by a RhoGTPase GDP dissociation inhibitor (RhoGDI) encoded by the SCN1 gene (Carol et al., 2005). RhoGDIs are negative regulators of ROP GTPases, which sequester the GDP-bound form in the cytosol (Yang, 2002). This provides strong evidence for a link between the Rop GTPase system and ROS production. This report provides evidence that the activation state of ROP GTPase in root hairs influences ROS production and, by using crosses between CA-rop2 and rhd2-1, shows that this depends on the activity of the NOX encoded by AtrbohC/RHD2.
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Materials and methods |
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Plant material and growth conditions
Arabidopsis thaliana seed were surface-sterilized for 4 min in 10% (v/v) household bleach and for 4 min in ethanol:water:bleach mixture (7:2:1 by vol.), and rinsed twice for 2 min in sterile water. Seeds were plated either on the surface of sterile semi-solid growth medium (Jones et al., 2002) containing 0.1% (w/v) Phytagel (Sigma-Aldrich), and grown in horizontal orientation or, for phenotypic characterization, beneath the surface of semi-solid medium containing 0.5% (w/v) Phytagel (Sigma-Aldrich) and grown in vertical orientation. Plated seeds were stratified for 48 h at 4 °C then transferred to growth cabinets at 20 °C with a 24 h light regime. For crossing experiments, seedlings were transferred to soil and grown at 20 °C under a 12 h light, 12 h dark regime. The WT Columbia (Col-0) ecotype was used, except where stated in the text. ROP2 transgenic plants (Li et al., 2001) were selected on medium supplemented with kanamycin (100 µg ml–1).
Genotype analysis
Genomic DNA was extracted using a REDExtract-N-Amp Plant PCR Kit (Sigma) The CA-rop2 transgene was amplified from CA-rop2 rhd2-1 plants by polymerase chain reaction (PCR) using genomic DNA, one primer specific for the ROP2 coding region (5'-TTATAAAGTGTGTGACCGTCG) and a second primer specific for the flanking pKYLX vector sequence (5'-TCAGTAGGATTCTGGTGTGTG; expected product size 684 bp; Jones et al., 2002).
Reagents
The reagents used, along with the concentration of the stock solutions and the final concentrations, were as follows: dihydrofluorescein diacetate [H2FDA; Molecular Probes; 50 mM in dimethylsulphoxide (DMSO); 50 µM; 1 min]; hydogen peroxide (Fisher; 10 mM in growth medium; 1 mM; 5 min), DPI (Sigma; 10 mM in DMSO; 50 mM; 30 min); superoxide dismutase [SOD; Sigma; 10 000 U ml–1 in 10 mM MES buffer, 20% (v/v) glycerol; 100 U ml–1; 1 h]; and Mn-5,10,15,20-tetrakis(1-methyl-4-pyridyl)21H,23H-porphin (TMPP: Sigma; 40 mM in water; 40–500 µM; 2–12 h).
Detection of ROS formation in the root hair differentiation zone of the root
ROS can be detected by oxidation of various derivatives of fluorescein. H2FDA was used in these experiments because it is less sensitive to hydroxyl radicals and peroxynitrite anions than the alternatives and therefore largely reports hydrogen peroxide in a peroxidase- and esterase-dependent manner (Hempel et al., 1999). Roots of intact seedlings were incubated in 250 µl of growth medium containing 50 µM H2FDA, with or without chemical treatments added. A single 2.3 s exposure was made exactly 1 min after the root was removed from the dye to fresh medium, with or without chemical treatments added. ROS formation was quantified as mean pixel intensity (black=0, white=4095) within a region of interest that includes the outline of the root and root hairs in the RHDZ (2 mm RHDZ distal to the first visible root hair), and inside regions of interest. Background fluorescence from RHDZs in the absence of H2FDA was 
166 units of pixel intensity at the emission wavelength used. Image analysis was performed using OpenLab image analysis software (v3.0.9, Improvision). The response of the camera was linear within the range of fluorescence intensities detected. For experiments performed at different times, the mean pixel intensity measured on WT RHDZs was used as an internal fluorescence standard. Probe concentration was not a limiting factor (see Supplementary Fig. S3 at JXB online). H2FDA becomes fluorescent only after hydrolysis by esterases and oxidation to fluorescein, primarily by hydrogen peroxide, in the presence of peroxidase activity. To confirm that endogenous peroxidase activity was not a limiting factor in dye oxidation, roots were preincubated in 1 mM hydrogen peroxide. All lines tested showed increased and more uniform fluorescence intensity in the RHDZ (Supplementary Fig. S3 available at JXB online). This shows that dye oxidation is reporting hydrogen peroxide concentration and not peroxidase activity. As a negative control, WT roots were preincubated with Mn-TMPP (40–500 µM). Mn-TMPP lowered the fluorescence in the RHDZ in a dose-dependent manner (see Supplementary Fig. S3 at JXB online). All lines and treatments showed uniform fluorescence intensity with fluorescein diacetate (FDA) (Breeuwer et al., 1995; see Supplementary Fig. S3 at JXB online).
Detection of superoxide production in the root hair tip
Whole seedlings were incubated for 1.0 h (Fig. 1A–D), 5 min (Fig. 1E, F) or 0.5 h (Fig. 2) in growth medium containing 0.5 mg ml–1 nitroblue tetrazolium (NBT). To correlate NBT staining with growth (evidenced by the presence of tip-localized cytoplasm; Fig. 2G), roots of intact seedlings were incubated in growth medium containing 0.5 mg ml–1 NBT on microscope slides. Staining was stopped by the addition of an excess of 70% (v/v) methanol and roots were rinsed in fresh 70% (v/v) methanol. There was batch-to-batch variability in staining intensity with NBT (Fig. 2) but the same relative differences between treatments and lines were observed in repeated experiments. For negative control experiments, seedlings were preincubated and stained in growth medium containing SOD or DPI. The intensity of formazan precipitation in root hair tips was quantified using OpenLab image analysis software (v3.0.9, Improvision) to calculate mean pixel intensity (black=0, white=4095) within each single image and inside regions of interest fitted to the outline of the root hair tip. Pixel intensity was reduced in areas of formazan precipitation.
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Microscopy
Only root hairs in the plane of focus were measured. Growing root hairs were identified by their tip-localized cytoplasm. Seedlings were gently removed from 0.1% (w/v) phytagel plates using forceps. Root hairs grown in this way were completely intact and, when mounted on slides in a droplet of liquid growth medium, continued to grow normally. Microscopy was performed using a Zeiss Axiophot microscope fitted with a Qimaging Retiga 1300 12-bit monochrome CCD camera linked to a computer running OpenLab v3.0.9 (Improvision). Zeiss filter set 10 (excitation 450–490 nm, emission 515–565 nm) was used for H2FDA. Formazan precipitates resulting from NBT oxidation were detected using a Zeiss Axioskop 2 microscope fitted with a Zeiss Axiocam 14-bit colour digital camera linked to a computer running AxioVision v3.1 (Zeiss).
Statistical analysis.
Data were subjected to one-way analysis of variance (ANOVA). Significant differences between mean values were identified by the least significant difference test. The statistical significance of the results is indicated in the text.
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Localization of superoxide formation at the tip of growing root hairs and its relationship to polarized growth
It has been shown previously that ROS are required for root hair tip growth and that intracellular ROS increase in root hair cells after the transition to tip growth (Foreman et al., 2003). However, the rapid diffusion of the dye used in their experiments (5-dichloromethyl-2',7'-dichlorodihydrofluorescein) does not allow precise localization of ROS formation. To address this question, NBT staining (Rossetti and Bonatti, 2001) was used to detect the site of superoxide production in growing WT root hairs. After reduction by superoxide, NBT forms a blue formazan precipitate indicating the site of superoxide production. Tetrazolium salts can disrupt PM integrity, leading to intracellular staining (Bernas and Dobrucki, 2000), but the addition of exogenous SOD can be used to confirm that tetrazolium salts are detecting extracellular superoxide production (Jiang and Zhang, 2003). NBT staining was concentrated at the root hair tip in young root hairs in the slow phase of tip growth (Fig. 1A; Dolan et al., 1994). Staining was more intense in longer root hairs in the fast phase of tip growth (Fig. 1B; Dolan et al., 1994). However, there was very little staining of the apex of emerging root hairs at the stage of swelling formation (Fig. 1A). Staining of tip-growing root hairs was much reduced in the presence of exogenous SOD (Fig. 1B, C). There was no detectable staining in mature root hairs (Fig. 1D). These patterns corresponded to quantitative differences in staining intensity between these developmental stages (see Supplementary Fig. S1 at JXB online). The ROS-sensitive probe H2FDA gave similar results, but staining was more diffuse because the oxidized dye rapidly diffuses throughout the cell (see Supplementary Fig. S2 at JXB online; Hempel et al., 1999). This shows that superoxide production is localized at the tips of growing root hairs. Incubating root hairs in NBT for 1 h led to intense tip-localized staining, making it impossible to detect whether the site of superoxide production was extracellular. With a shorter incubation of 5 min, formazan deposition was visible only in the cell wall, peripheral to the tip-localized cytoplasm, suggesting extracellular superoxide production (Fig. 1E, F).
The NOX inhibitor DPI inhibits both tip growth and ROS formation in root hairs (Foreman et al., 2003), but the effect of exogenous ROS scavengers on root hair growth has not been reported. WT root hairs were treated with the SOD and catalase mimic Mn-TMPP (Fig. 1I; Schopfer, 2001). Mn-TMPP decreased ROS detected by H2FDA in the RHDZ in a dose-dependent manner (see Supplementary Fig. S3 at JXB online). Mn-TMPP mimicked the rhd2-1 tip-bursting phenotype that occurs in rhd2-1 root hairs during the transition to tip growth (Table 1; Fig. 1G, H). rhd2-1 is a transcript null AtrbohC/RHD2 allele with a premature stop codon at amino acid position 597 (Foreman et al., 2003). Low concentrations (10 mM) of the antioxidant L-ascorbate also caused tip bursting (data not shown). Tip bursting induced by antioxidants indicates that AtrbohC/RHD2-dependent ROS formation is required for the integrity of the developing tip. The morphology of mature WT root hairs was unaffected by treatment with either Mn-TMPP (Fig. 1J) or L-ascorbate (data not shown). Lower doses of Mn-TMPP also reduced ROS in root hairs (see Supplementary Fig. S3 at JXB online) but, instead of bursting, caused ballooning of the root hair tip (Fig. 1K). This result shows that ROS are also essential for maintaining polarized tip growth in root hairs.
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ROS formation in the root hair differentiation zone of plants expressing ROP2 OX, CA-rop2, and DN-rop2 transgenes
If ROS formation in root hairs depends on ROP2, there should be differences in ROS formation between the RHDZs of plants expressing different ROP2 transgenes and those of WT plants. H2FDA was used to detect ROS in the RHDZ. Control experiments confirmed that H2FDA was reporting true differences in ROS formation (see Supplementary Fig. S3 at JXB online). Compared with WT RHDZs, CA-rop2 RHDZs had significantly higher ROS formation (Fig. 2A, C, one-way ANOVA P <0.05) and DN-rop2 RHDZs had significantly lower ROS formation (Fig. 2A, C, one-way ANOVA P <0.05). ROS formation in the RHDZ of ROP2 OX plants was also significantly higher than in WT plants (mean pixel intensity: WT 730±105; ROP2 OX 991±180; one-way ANOVA P <0.05; n=5). These results show that ROP2, in its GTP-bound form, activates ROS formation in the RHDZ of primary roots.
Superoxide formation in the root hair tip of plants expressing ROP2 OX, CA-rop2, and DN-rop2 transgenes
Since NBT reduction at the growing tip of root hairs reports extracellular superoxide formation, this method was used to compare plants expressing ROP2 transgenes. Compared with WT root hair tips (Fig. 2B, D), both CA-rop2 (Fig. 2B, D; P <0.001) and ROP2 OX root hair tips (Fig. 2B, D; P <0.001) had significantly higher superoxide production, whereas DN-rop2 root hair tips had significantly lower superoxide production (Fig. 2B, D; P <0.02). DPI inhibited tip-localized staining in all lines tested and, like Mn-TMPP (Fig. 1I), induced depolarization of the root hair tip (Fig. 2D). The tips of multiple root hairs and branched root hairs in ROP2 OX root hair cells (Jones et al., 2002) do not always grow simultaneously, but the actively growing tips can be recognized because they have tip-localized cytoplasm and are not vacuolated (Grierson and Schiefelbein, 2002). Interestingly, NBT staining revealed superoxide production only in a single tip of some multiple ROP2 OX root hairs (Fig. 2E) and branched ROP2 OX root hairs (Fig. 2F). This was confirmed by observing ROP2 OX root hair tips undergoing staining in the presence of NBT. Formazan deposition in the cell periphery (Fig. 1E, F) only occurred in root hair tips with tip-localized cytoplasm, indicating that tip-localized superoxide production was associated with active tip growth (Fig. 2G). These results show that ROP2, in its GTP-bound form, activates superoxide formation in the tip of the growing root hair.
The effect of ROP2 transgenes on root hair morphogenesis in the AtrbohC/RHD2 loss-of-function mutant background
ROP2 is involved in root hair tip morphogenesis (Jones et al., 2002), but it is not clear when ROP2 acts during root hair morphogenesis in relation to the NOX gene AtrbohC/RHD2 (Foreman et al., 2003). It is possible that ROP2 acts earlier than AtrbohC/RHD2 as green fluorescent protein (GFP)::ROP2 localizes to the future site of swelling formation and both ROP2 OX and CA-rop2 stimulate swelling formation (Jones et al., 2002), whereas AtrbohC/RHD2 is not required for swelling formation (Schiefelbein and Somerville, 1990; Parker et al., 2000; Foreman et al., 2003). To determine at which stage during ROP2-induced root hair morphogenesis AtrbohC/RHD2 is required, long-haired CA-rop2 plants (Jones et al., 2002) were crossed with short-haired rhd2-1 plants. F3 lines were isolated where all individuals were kanamycin resistant, and had rhd2-1-like root hairs and short rhd2-1-like roots [Fig. 3A; Table 2; primary root length (mm ±SD) WT Col-0, 35.8±7.7 n=22; rhd2-1, 15.8±3.7 n=23; CA-rop2 rhd2-1, 15.4±5.6 n=10). PCR confirmed that these plants carried the CA-rop2 transgene (Fig. 3B). Like CA-rop2 plants (Jones et al., 2002), but unlike rhd2-1 plants (data not shown), CA-rop2 rhd2-1 plants had some root hair cells with multiple swelling formation (Fig. 3C). However, CA-rop2 rhd2-1 root hairs showed a similar frequency of tip bursting to rhd2-1 root hairs (Table 1). This result shows that, although CA-rop2 was able to activate multiple swelling formation in the rhd2-1 genetic background, CA-rop2 rhd2-1 root hairs were tip growth defective. These results show that ROP2 acts earlier than AtrbohC/RHD2 during root hair morphogenesis as a functional AtrbohC/RHD2 gene is required for CA-rop2-induced root hair elongation, but not for CA-rop2-induced multiple swelling formation.
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Based on these results, one might predict that root hair morphogenesis would be disrupted even earlier in a ROP2 loss-of-function mutant than in rhd2-1 (i.e. before swelling formation), enabling the exact sequence of gene action to be determined. However, ROP2 knockout or RNAi lines do not have a noticeable root hair phenotype (data not shown), suggesting that ROP2 acts in a functionally redundant manner with other ROPs during root hair morphogenesis.
The effect of ROP2 transgenes on ROS production in the AtrbohC/RHD2 loss-of-function mutant background
As the rhd2-1 mutation disrupted ROP2-dependent tip growth, it is possible that this mutation also disrupts ROP2-dependent ROS formation in the RHDZ and growing root hair tip. ROS formation was similar in both the rhd2-1 RHDZ and the CA-rop2 rhd2-1 RHDZ, and both were significantly lower than in the WT RHDZ (Fig. 2A, C, one-way ANOVA P <0.05). These results show that a functional AtrbohC/RHD2 gene is required for ROP2-dependent ROS formation in the RHDZ. It was not possible to assess superoxide production in tip-growing root hairs of rhd2-1 or CA-rop2 rhd2-1 plants as root hair cells in these plants burst during the transition to tip growth. As might be predicted, there was very little NBT staining in the apices of emerging root hairs at the stage of swelling formation in rhd2-1 or CA-rop2 rhd2-1 plants (data not shown).
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Localization and quantification of ROS production by the RHDZ and root hairs
Foreman et al. (2003) showed that ROS production was reduced in loss-of-function rhd2 roots and root hairs, measured in excised root apices. They used lucigenin, which largely indicates superoxide production, and fluorescence microscopy to visualize oxidation of 5-dichloromethyl-2',7'-dichlorodihydrofluorescein, which largely detects intracellular hydrogen peroxide. In this report, their observations have been supplemented by use of NBT reduction to detect the location of extracellular superoxide formation. NBT reduction was strongly localized at the tip of actively growing root hairs and absent from non-growing root hairs. Observations of the staining pattern showed that the formazan was precipitated in the cell wall and that SOD was able to decrease NBT reduction. SOD will not rapidly penetrate the cell wall or PM, providing further evidence for extracellular superoxide formation. NBT reduction was also sensitive to DPI. Carol et al. (2005) have also used NBT reduction to detect superoxide production by root hairs. Without supporting genetic evidence, DPI cannot be used to confirm NOX activity as it inhibits other flavin-containing enzymes (Frahry and Schopfer, 1998). However, as AtrbohC/RHD2 has already been shown to be the major source of DPI-sensitive ROS in root hairs (Foreman et al., 2003), the use of DPI here is informative and suggests that the SOD-sensitive and DPI-sensitive superoxide production observed here is probably derived from AtrbohC/RHD2 activity.
To detect differences in ROS production between genotypes, it was necessary carefully to standardize the ROS assays, particularly because ROS formation by root hairs varied considerably according to their developmental stage. The average fluorescence intensity in the RHDZ was determined after staining with H2FDA. The fluorescence signals came largely from root hairs, although the staining intensity of the root axis showed the same difference between strains. Control experiments were carried out with FDA (which becomes fluorescent after hydrolysis of the acetate group) to ensure that there were no differences between genotypes in dye loading, esterase activity, and pH that could affect the fluorescence intensity. Addition of hydrogen peroxide to H2FDA-stained samples resulted in the same fluorescence intensity between genotypes, showing that peroxidase activity, needed for dye oxidation, was not a limiting factor. Application of these control assays, which are rarely reported by other workers, allows confidence in the detection of differences in ROS-dependent dye oxidation between strains. Pixel intensity in the root hair tip was determined after staining in NBT. NBT reduction was sensitive to the NOX inhibitor DPI in all genotypes. Similar differences between genotypes were observed in each of the two independent ROS assays.
The role of ROP2 in ROS formation by growing root hairs
A number of investigations have linked ROPs to ROS formation during programmed cell death and in response to hypoxia (Kawasaki et al., 1999; Baxter-Burrell et al., 2002; Park et al., 2004). The present results establish the dependence of NOX activity encoded by AtrbohC/RHD2 on ROP2 GTPase. The evidence for this is that CA-rop2 and ROP2 OX plants had increased ROS formation in the RHDZ and superoxide production in the root hair tip, and that DN-rop2 plants had decreased ROS formation. Although ROP2 is probably not the only ROP required for tip growth of root hairs (see below), the results from ROP2 OX plants show that the activating effect of CA-rop2 on ROS formation is not due to non-specific activation of unrelated ROPs (Hakeda-Suzuki et al., 2002). Additionally, in the loss-of-function rhd2-1 genetic background, CA-rop2 root hairs were defective in both tip growth and superoxide production, but not in ROP2-mediated multiple swelling formation. This result shows that ROP2 acts earlier than AtrbohC/RHD2 during root hair morphogenesis. Foreman et al. (2003) have shown that the AtrbohC/RHD2 NOX is the primary source of ROS formation in growing root hairs. The results presented here show that AtrbohC/RHD2 is also likely to be the source of ROP2-induced superoxide production in root hairs. Further work is required to determine the nature of the relationship between ROPs and AtrbohC/RHD2 in growing root hairs. Such a relationship might occur indirectly through one of the 11 Arabidopsis ROP-interactive CRIB motif-containing proteins (RICs; Wu et al., 2001) that function as adaptor proteins connecting ROPs with their downstream targets. Alternatively, ROP2 might interact directly with AtrbohC/RHD2. Indeed, the rice ROP, OsRac1, shows a physical interaction with rice NOX in a yeast two-hybrid assay (Pinontoan et al., 2003). Furthermore, the related mammalian RHO GTPase, Rac2, interacts physically with the NOX catalytic subunit gp91phox in vitro (Diebold and Bokoch, 2001).
ROS production is essential for the integrity of the root hair tip, as evidenced by tip bursting in the rhd2-1 mutant and in WT root hairs exposed to the ROS scavengers Mn-TMPP and L-ascorbate. ROS are also required for maintaining polarized growth (i.e. the shape of the root hair), as both Mn-TMPP and DPI altered root hair shape. On the other hand, increased ROS can also cause root hair depolarization: CA-rop2 has increased superoxide production in the root hair tips and they are depolarized. It has also been reported that total loss-of-function rhd2 plants treated with exogenous ROS have depolarized root hairs (Foreman et al., 2003). Therefore, it is apparent that precise control of ROS formation or localization is needed for normal root hair growth.
ROPs act in functionally redundant pathways (Yang, 2002). For instance, three pollen-expressed ROPs have similar regulatory effects on pollen tube growth (Kost et al., 1999; Li et al., 1999). Expression of CA-rop4 and CA-rop6 transgenes disrupted root hair tip growth (Molendijk et al., 2001), although it is not known whether, like ROP2 (Jones et al., 2002), these ROP genes are expressed in root hairs. Based on transcriptomic profiling of ROP expression in the RHDZ, T-DNA ‘knockout’ and RNAi lines for ROP2 and several other candidate root hair ROPs have been screened but none has obvious root hair phenotypes (M Jones, N Smirnoff, unpublished data). This suggests that, as in pollen tubes, several ROPs act in a functionally redundant manner. Preliminary results with double and triple mutant combinations of loss-of-function rop mutants suggest that at least three ROP genes are involved in root hair morphogenesis (M Jones, N Smirnoff, unpublished data).
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TopAbstractIntroductionMaterials and methodsResultsDiscussionConclusionsSupplementary dataReferences |
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The ROP2 GTPase, probably acting in a functionally redundant manner with other ROPs, is a fundamental switch involved in root hair morphogenesis. The results presented here show that the GTP-bound form of ROP2 also activates superoxide production in the growing root hair tip. The effects of ROP2 on both superoxide production and tip growth require the presence of a functional AtrbohC/RHD2 gene. A further layer of complexity is added when one considers the regulation of ROP2 itself. RhoGDI, a negative regulator of ROP, also affects spatial regulation of root hair growth and superoxide production (Carol et al., 2005). RhOGDIs have been shown to interact physically with ROP (Bischoff et al., 2000). Carol et al. (2005) showed in pull-down assays that ROP interacts with RhoGDI and this is required for its activity. Transcription of another negative regulator of ROPs, ROP GTPase-activating protein 4 (GAP4), is activated by ROP2-induced DPI-sensitive ROS formation (Baxter-Burrell et al., 2002). ROP GAPs promote the intrinsic GTPase activity of ROPs, switching them to their inactive form (Wu et al., 2000). It seems reasonable to speculate that one or more of the five ROP GAPs in the Arabidopsis genome might also regulate ROP2. Further work is required to understand how the ROP GTPase system interacts with AtrbohC/RHD2 to modulate its NADPH oxidase activity.
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Supplementary data |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionConclusionsSupplementary dataReferences |
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The supplementary data available at JXB online include more detailed analysis of ROS formation in relation to root hair morphogenesis and control experiments for validation of the detection of ROS using dihydrofluorescein diacetate.
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Acknowledgements |
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This research was funded by a Biotechnology and Biological Sciences Research Council grant to NS.
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Abbreviations |
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ABA, abscisic acid; ANOVA, analysis of variance; CA, constitutively active; DMSO, dimethylsulphoxide; DN, dominant negative; DPI, diphenylene iodonium; FDA, fluorescein diacetate; H2FDA, dihydrofluorescein diacetate; Mn-TMPP, Mn-5,10,15,20-tetrakis(1-methyl-4-pyridyl)21H,23H-porphin; NOX, NADPH oxidase; PCR, polymerase chain reaction; PM, plasma membrane; RHDZ, root hair differentiation zone; RhoGDI, RhoGTPase GDP dissociation inhibitor; RNAi, RNA interference; ROS, reactive oxygen species; SOD, superoxide dismutase; WT, wild type.
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