Research ArticleResearch Article

Interplay between the NADP-Linked Thioredoxin and Glutathione Systems in Arabidopsis Auxin Signaling

Talaat Bashandy, Jocelyne Guilleminot, Teva Vernoux, David Caparros-Ruiz, Karin Ljung, Yves Meyer, Jean-Philippe Reichheld

Published February 2010. DOI: https://doi.org/10.1105/tpc.109.071225

Abstract

Intracellular redox status is a critical parameter determining plant development in response to biotic and abiotic stress. Thioredoxin (TRX) and glutathione are key regulators of redox homeostasis, and the TRX and glutathione pathways are essential for postembryonic meristematic activities. Here, we show by associating TRX reductases (ntra ntrb) and glutathione biosynthesis (cad2) mutations that these two thiol reduction pathways interfere with developmental processes through modulation of auxin signaling. The triple ntra ntrb cad2 mutant develops normally at the rosette stage, undergoes the floral transition, but produces almost naked stems, reminiscent of the phenotype of several mutants affected in auxin transport or biosynthesis. In addition, the ntra ntrb cad2 mutant shows a loss of apical dominance, vasculature defects, and reduced secondary root production, several phenotypes tightly regulated by auxin. We further show that auxin transport capacities and auxin levels are perturbed in the mutant, suggesting that the NTR-glutathione pathways alter both auxin transport and metabolism. Analysis of ntr and glutathione biosynthesis mutants suggests that glutathione homeostasis plays a major role in auxin transport as both NTR and glutathione pathways are involved in auxin homeostasis.

INTRODUCTION

Modulation of plant development in response to stress plays a major role in adaptation of plants to their environment. Redox signaling pathways are involved in many aspects of these responses, detoxifying reactive oxygen species produced during stress responses or regulating developmental processes (Vernoux et al., 2000; Xing et al., 2005; Reichheld et al., 2007, Marty et al., 2009, Benitez-Alfonso et al., 2009; for relevant reviews, see Foyer and Noctor, 2005; Potters et al., 2007; Van Breusegem et al., 2008). Key players are the NADPH-dependent glutathione/glutaredoxin system (NGS) and the NADPH-dependent thioredoxin system (NTS), which together determine protein thiol/disulfide status, a primary factor in redox signaling. Thioredoxins (TRXs) and glutaredoxins (GRXs) are ubiquitous, evolutionarily conserved proteins catalyzing disulfide reduction in vivo through a redox-active dithiol CxxC. TRX and GRX therefore play key roles in the maintenance of cellular redox homeostasis through the sensing and transfer of reducing equivalents to numerous target proteins, such as peroxidases, reductases, metabolic enzymes of glycolysis, and photosynthesis or through structural modifications of target proteins (Arnér and Holmgren, 2000; Buchanan and Balmer, 2005; Rouhier et al., 2008; Montrichard et al., 2009). Subsequently, oxidized TRX and GRX are reduced by thioredoxin reductase or glutathione, respectively (Holmgren, 1989).

The tripeptide glutathione is an important thiol/disulfide buffer whose effects are related to the balance between GSH (reduced form) and GSSG (oxidized form), determined by GSH oxidation by reactive oxygen species and other oxidants and by GSSG reduction by glutathione reductase. Genetic analyses in plants have underlined a threshold glutathione concentration necessary for root development (Vernoux et al., 2000). At higher concentrations, glutathione is a determinant of cadmium resistance (Cobbett et al., 1998), high light signaling (Ball et al., 2004), nodulation in legumes (Frendo et al., 2005), and activation of phytoalexin synthesis in the pathogen response (Parisy et al., 2007). Genetic approaches aiming to identify functions of TRX and GRX in knockout plants have largely been limited by the absence of phenotypes of single mutants presumably due to functional redundancies among members of the multigene families of TRX and GRX (Meyer et al., 2008). In a few reports, TRXs have been shown to be involved in pathogenic responses (Rivas et al., 2004; Sweat and Wolpert, 2007; Tada et al., 2008), hypersensitiviy to oxidants (Broin et al., 2000, 2002; Broin and Rey, 2003), cytochrome b(6)f biosynthesis (Lennartz et al., 2001), and plasmodesmal transport in meristems (Benitez-Alfonso et al., 2009). Recently, genetic evidence has revealed functions of GRX-like proteins in flower development (Xing et al., 2005; Xing and Zachgo, 2008; Li et al., 2009).

Indeed, in several organisms, several publications have highlighted an interplay between the NTS and NGS pathways (Carmel-Harel and Storz, 2000; Kanzok et al., 2000, 2001; Gelhaye et al., 2003; Trotter and Grant, 2003, 2005; Koh et al., 2008). Therefore, association of mutants involved in both the NTS and NGS pathways have recently revealed functions of these pathways in several aspects of plant development. In Arabidopsis thaliana, the NTS constitutes a functional backup for cytosolic glutathione reduction activity and association of the aphenotypic cytosolic glutathione reductase (gr1) and cytosolic/mitochondrial NADPH-dependent thioredoxin reductase (ntra ntrb) mutants leads to a pollen lethal phenotype (Marty et al., 2009). Otherwise, crossing of the ntra ntrb mutant with rml1 (rootmeristemless1), a mutation partially inactivating the first enzyme of glutathione biosynthesis, leads to a lack of meristematic activity in shoots of the ntra ntrb rml1 triple mutant, demonstrating overlapping functions of the NGS and NTS systems in shoot meristem activity (Reichheld et al., 2007).

The phytohormone auxin is known to be involved in various aspects of plant growth and development, including meristematic activities (reviewed in Benjamins and Scheres, 2008). Auxin has been suggested to be synthetized in young tissues (meristems, young primordia, vascular tissues, and flower buds) by at least two separate pathways (Barlier et al., 2000; Zhao et al., 2001; Cheng et al., 2006) and is also actively transported to specific tissues where it triggers a signaling cascade causing many developmental and differentiation responses (Teale et al., 2006). This transport is mediated by both influx and efflux systems to channel auxin from cell to cell in a directional manner. Influx is carried on by the AUXIN RESISTANT1 (AUX1) protein (Bennett et al., 1996). Efflux is dependent on the group of PIN-FORMED (PIN) proteins and by several MULTIDRUG RESISTANCE p-glycoproteins (Gil et al., 2001; Noh et al., 2001)

In this work, we crossed the ntra ntrb mutant with cad2 (cadmium sensitive2), a weak mutant allele of the GSH1gene. The cad2 mutant was originally identified in a genetic screen designed to isolate mutants to hypersensitive cadmium, but the phenotype in the absence of stress is nearly indistinguishable from wild-type plants (Howden et al., 1995; Cobbett et al., 1998). Interestingly, the triple homozygote ntra ntrb cad2 mutant shows stems devoid of any flower development, resembling the pin1 mutant (Gälweiler et al., 1998), pinoid (pid) (Bennett et al., 1995; Christensen et al., 2000), monopteros (Hardtke et al., 2004), and naked pins in yuc mutants1 (npy1) (Cheng et al., 2007) that all develop pin-like inflorescences and are defective either in auxin transport, signaling, or synthesis. Moreover, the ntra ntrb cad2 mutant also shows decreased apical dominance, vasculature defects, and perturbed root development, several phenotypes tightly regulated by auxin. Therefore, the phenotype of the ntra ntrb cad2 mutant highlights a potential link between the thiol/disulfide regulation and auxin signaling.

RESULTS

Isolation and Characterization of the ntra ntrb cad2 Triple Mutant

To generate an ntra ntrb cad2 triple mutant, ntra ntrb and cad2 homozygote plants were crossed. Both mutants are in the Columbia-0 (Col-0) genetic background, so we do not expect a heterosis effect in the triple mutant. Plants of the F1 generation were all found to be heterozygous for the ntra, ntrb, and cad2 mutations. In the F2, ntra ntrb homozygous plants were first selected on Murashige and Skoog (MS) medium supplemented with l-buthionine-(S,R)-sulfoximine (BSO), a specific inhibitor of glutathione synthesis. This treatment stops growth of the double ntra ntrb homozygote plants by limiting meristematic activities (Reichheld et al., 2007). After transfer to a BSO-free medium, growth of ntra ntrb plants resumed and ntra ntrb homozygote plants were further screened for the cad2 mutation by PCR. In a first set of screenings, wild-type ntra ntrb GSH1/GSH1 and heterozygote ntra ntrb GSH1/cad2 plants were identified, but no ntra ntrb cad2 homozygotes were found. Indeed, a large majority of the plants were GSH1/GSH1 due to the genetic linkage between the GSH1 and NTRB genes (20 centimorgans). Therefore, progeny of ntra/ntra ntrb/ntrb GSH1/cad2 plants were further analyzed. Phenotypically, two distinct categories of plants were found. The first showed growth and development very similar to ntra ntrb plants (Reichheld et al., 2007), while the second showed severe flower development inhibition leading to sterility, as well as shoot and root growth perturbations. These developmental phenotypes will be described in more detail below (Figures 1 to 3⇓⇓).

Figure 1.
Figure 1.

Phenotype of the ntra ntrb cad2 Mutant at Different Developmental Stages.

(A) Fifteen-day-old Col-0 (left) and ntra ntrb cad2 (right) grown in vitro.

(B) Three-week-old Col-0 (left) and ntra ntrb cad2 (right) plants grown on soil.

(C) and (D) Five-week-old Col-0 (C) and ntra ntrb cad2 (D) plants. Arrows indicate emergence of secondary rosette stems in ntra ntrb cad2, which were not observed in the wild type at this stage.

(E) and (F) Seven-week-old Col-0 (left) and ntra ntrb cad2 (right) plants (E) and 10-week-old ntra ntrb cad2 plant (F). All plants were grown under a long-day regime (16 h light/8 h dark). At all developmental stages, cad2 and ntra ntrb plants (not represented) have a similar phenotype as Col-0 plants.

(G) and (H) Five-week-old Col-0 (G) and ntra ntrb cad2 (H) rosette leaves. Collected leaves were cleared in Hoyer's solution and observed under a binocular microscope. Note the abnormal leaf shape and the less developed and irregular vasculature of ntra ntrb cad2 leaf. Bar = 1 mm.

Figure 2.
Figure 2.

Structure of the Inflorescence Meristem of the ntra ntrb cad2 Mutant.

(A) and (B) Stem apex of 5-week-old Col-0 (A) and ntra ntrb cad2 (B) plants. Bars = 1 mm.

(C) and (D) Histological structure of the inflorescence meristem stained by toluidine blue. Col-0 inflorescence (C) and ntra ntrb cad2 meristem (D). Bars = 100 μ m.

(E) and (F) Higher magnification of the inflorescence meristem of Col-0 (E) and ntra ntrb cad2 (F). Bars = 20 μ m.

Figure 3.
Figure 3.

Root Growth and Emergence of Secondary Roots in the cad2, ntra ntrb, and ntra ntrb cad2 Mutants.

(A) Daily root growth was measured on Col-0, cad2, ntra ntrb, and ntra ntrb cad2 plants grown on MS/2 vertical agar plates at 22°C, under a 16-h-light/8-h-dark regime. Primary root growth rate was calculated as the mean of daily root growth between the 8th and the 11th day after seeds were put under growth conditions. Data are means ± se; n ≥ 35.

(B) The number of secondary root initials was measured after 10 d after germination. Col-0, cad2, ntra ntrb, and ntra ntrb cad2 plants were grown in the same conditions as in (A). Data are means ± se; n ≥ 35.

(C) Phenotype of Col-0, cad2, ntra ntrb, and ntra ntrb cad2 plants cultivated as in (A) and observed 12 d after germination. Bars = 5 mm.

(D) Phenotype of the primary root meristem of Col-0, cad2, ntra ntrb, and ntra ntrb cad2 plants cultivated as in (A) and observed 10 d after germination. The arrows show the approximate extent of the meristem based on elongation of the pericycle cell files. Bars = 40 μ m.

[See online article for color version of this figure.]

To confirm that the second class of plants contained triple ntra ntrb cad2 homozygotes, 40 plants were genotyped for the cad2 mutation. All plants having a developmental phenotype (9/40, χ2 = 0.1, P > 0.7) were homozygous for ntra, ntrb, and cad2 mutations, while wild-type-like plants had at least one wild-type copy of the GSH1 gene (31/40, χ2 = 0.03, P > 0.8), indicating that mutation of all three genes is necessary to trigger the developmental phenotype. Supporting this observation, no plants harboring the flower development phenotype were isolated in the progeny of cad2/cad2 ntra/ntra NTRB/NTRB or cad2/cad2 NTRA/NTRA ntrb/ntrb plants.

To demonstrate that the phenotypic characteristics of the ntra ntrb cad2 mutant were linked to inactivation of NTR genes, the mutant was complemented by a 3.27-kb genomic fragment containing the NTRB gene. Due to sterility of the ntra ntrb cad2 mutant, we performed complementation of ntra/ntra ntrb/ntrb GSH1/cad2 plants. In 20 transformed plants containing the transgene, the NTRB protein was recovered, showing that the transgene is functional (see Supplemental Figure 1 online). These plants were further screened for the cad2 mutation, and nine plants had a wild-type GSH1 gene (ntra/ntra ntrb/ntrb GSH1/GSH1), five were heterozygous for the cad2 mutation (ntra/ntra ntrb/ntrb GSH1/cad2), and six were triple homozygotes (ntra/ntra ntrb/ntrb cad2/cad2). All complemented plants recovered a growth rate comparable to that of wild-type plants, showing that at least the NTRB protein is sufficient to confer a normal growth rate to the plant. To study the recovery in a broader collection of complemented plants, 152 ntra/ntra ntrb/ntrb GSH1/cad2 plants derived from a self-fertilized ntra/ntra ntrb/ntrb GSH1/cad2 plant that had been transformed with the NTRB complement were grown. Thirty-four ntra ntrb cad2 homozygous plants were found in the progeny (34/152, χ2 = 0.42, P > 0.5). Most of these expressed the NTRB transgene and showed a wild-type phenotype, while some of them (6/34) were segregants that did not receive the transgene. These segregant plants still harbored the developmental phenotype.

Biochemical Characterization of the ntra ntrb cad2 Mutant

Crosstalk between NTS and NGS has been recently shown in Arabidopsis (Reichheld et al., 2007; Marty et al., 2009). To evaluate whether this crosstalk is active in the ntra ntrb cad2 mutant, we studied the level of total glutathione in the wild type, cad2, ntra ntrb, and ntra ntrb cad2 mutant plants. According to previously published data (Howden et al., 1995; Reichheld et al., 2007), the glutathione level in cad2 plants is 20% of that found in wild-type Col-0 plants, while the level in ntra ntrb was not significantly different (88%) from wild-type plants (see Supplemental Table 1 online). The total glutathione level (including GSSG and GSH) found in ntra ntrb cad2 was 32% that of wild-type plants, suggesting that glutathione level is slightly but significantly (χ2 = 41.4, P < 0.01) enhanced (1.6-fold) in the triple mutant compared with cad2 plant. In all mutants, the level of GSSG was below detection (<1% of total glutathione).

The NGS system was previously found to affect the redox state of the cytosolic TRXh3 in the ntra ntrb mutant (Reichheld et al., 2007). The redox state of TRXh3 was measured in the ntra ntrb cad2 mutant and compared with wild-type Col-0 plants and cad2 and ntra ntrb mutants (see Supplemental Figure 2 online). TRXh3 was found to be mainly in a reduced state in Col-0 and cad2 mutant, showing that the 80% decrease of glutathione level does not influence the redox state of TRXh3. As previously shown (Reichheld et al., 2007), the pool of TRXh3 is only partially oxidized in the ntra ntrb mutant due to a GSH-dependent reduction pathway. This redox state is not greatly modified in the ntra ntrb cad2 mutant, suggesting that the residual GSH pool is sufficient for the alternative reduction of TRXh3 by the NGS system.

NTR and GSH1 Genes Regulate the Architecture of the Aerial Part of the Plant and Are Required for the Proper Development of Flowers

At the rosette stage, ntra ntrb cad2 homozygote plants were smaller in size, but emergence of their leaves was not profoundly perturbed, suggesting that the shoot meristem is not highly affected. However, cotyledon and leaf shape were altered and showed abnormal vasculature (Figures 1G and 1H). Floral transition was not perturbed in the ntra ntrb cad2 mutant, and stem growth was not affected (Figures 1D and 1E). Remarkably, no floral structures appeared at the top of the stems, giving the inflorescence stem a pin-like phenotype. Secondary stems also showed this phenotype (Figure 1D). At later developmental stages, abnormal numbers of secondary stems appeared (up to 30 stems after 50 d of development), suggesting a loss of apical dominance (Figures 1E and 1F). Longitudinal cross sections were prepared to compare the structure of the pin-like with wild-type flower meristems. Flower buds initiated in the mutant, but, in contrast with wild-type plants, their development rapidly stopped (Figures 2A to 2D). However, the overall organization of the inflorescence meristem was not profoundly perturbed in the ntra ntrb cad2 mutant (Figures 2E and 2F).

The flowerless phenotype of the ntra ntrb cad2 mutant suggests that the corresponding proteins are functional in the flower meristem of the wild type. Therefore, we studied the expression pattern of the respective genes in wild-type flowers by RT-PCR. As previously shown, NTRA and NTRB genes are constitutively expressed in all plant organs, including flower buds and flowers (Reichheld et al., 2007). Here, we show that GSH1 is also ubiquitously expressed, and significant expression was found in flower buds (see Supplemental Figure 3A online). To study the pattern of expression of GSH1 gene in the inflorescence, we used a ProGSH1:GUS (for β-glucuronidase) line. Prominent GUS staining was found in the flower meristem (see Supplemental Figure 3B online).

We performed in situ hybridization to detect NTR mRNAs in flower meristems. A ubiquitous specific signal was found in the whole inflorescence, the staining being higher in the inflorescence and flower meristems (see Supplemental Figure 4 online). This expression pattern correlates with previous data obtained with ProNTRA:GUS and ProNTRB:GUS staining (Reichheld et al., 2007). Overall, these data show that NTRA, NTRB, and GSH1 genes are expressed in both inflorescence and flower meristems.

Root Development Is Affected in the ntra ntrb cad2 Mutant

We showed previously that root elongation was slowed in the ntra ntrb mutant. This was presumably due to a lower cell division rate in the root meristem (Reichheld et al., 2007). We wished to know whether this effect was enhanced in the ntra ntrb cad2 mutant. Measurement of root elongation as well as the number of secondary roots was estimated in the different mutants (Figure 3). In cad2 homozygote plants, daily root growth was reduced to 79% of that of wild-type, ntra ntrb to 69%, and ntra ntrb cad2 to 33%, showing that association of cad2 and ntra ntrb mutants had an additive effect on root growth (Figures 3A and 3C). Microscopy observation of the primary root meristem of the ntra ntrb cad2 mutant showed a pronounced decrease in size. However, the structure of the meristem was not perturbed (Figure 3D). Initiation of secondary roots was also measured in the mutants: both cad2 and ntra ntrb had about a 50% reduction in secondary roots, whereas secondary root initiation was dramatically inhibited to <4% in the ntra ntrb cad2 mutant (Figure 3B), suggesting an additive effect of ntra ntrb and cad2 mutation of secondary root initiation.

Auxin Transport Is Deficient in ntra ntrb cad2, cad2, and pad2 Mutants

The ntra ntrb cad2 mutant has a pin-like phenotype, loss of apical dominance, and a reduced root system. These phenotypes are often associated with perturbed auxin transport. We performed experiments to find out whether polarized auxin transport (PAT) is altered in the mutant. Therefore, we compared PAT between the ntra ntrb cad2 mutant and wild-type plants (Figure 4). The tritium-labeled indole-3-acetic acid (3H-IAA) basipetal transport in the inflorescence stems excised from ntra ntrb cad2 plants was reduced to 5% of that of the Col-0 plants. To estimate whether inactivation of NTR and/or GSH biosynthesis impaired auxin transport, we also measured the basipetal auxin transport in the ntra ntrb and cad2 parent mutants. While the ntra ntrb mutation had a mild effect on auxin transport (less than twofold), the cad2 mutation alone reduced the auxin transport to one-tenth of wild-type levels, indicating that the glutathione level plays a major role in PAT. To confirm this point, we measured PAT in the pad2 mutant, an independent mutant allele of the GSH1 gene containing ∼ 16% wild-type amounts of GSH in rosette plants (see Supplemental Table 1 online). This glutathione level is consistent with the 22% previously found by Parisy et al. (2007). In cad2, PAT was greatly reduced (8% of the wild type) in pad2, confirming that low GSH availability affects auxin transport. As expected, PAT was dramatically reduced in pin1 mutant inflorescence stems (3% of PAT found in the heterozygote PIN1/pin1 plants). Taken together, these data indicate that auxin transport is impaired in the ntra ntrb cad2 and that this deficiency is mainly due to lower availability of GSH.

Figure 4.
Figure 4.

Auxin Transport Analysis in Mutants.

Basipetal auxin transport was evaluated by measuring the radioactivity of stem segments of the plants after feeding of 3H-IAA. Each value represents the average of eight plants (± se, n = 8) of wild-type Col-0, homozygote mutants cad2, pad2, ntra ntrb, ntra ntrb cad2, pin1/pin1, and heterozygote PIN1/pin1. As a negative control, acropetal auxin transport was evaluated in stem segments of Col-0 plants.

Low Glutathione Availability Affects Expression of PIN and Auxin Response Genes

To further define the influence of glutathione on auxin transport, we investigated the expression of members of the PIN group of membrane proteins, which are known to mediate auxin transport processes (Figure 5). Wild-type plants were subjected to BSO treatment, a specific inhibitor of glutathione biosynthesis, mimicking mutation in the GSH1 gene. Steady state levels of several PIN mRNAs were followed at different time points after the treatment, and these levels were compared with the level of total glutathione (Figure 5A). PIN1, PIN2, PIN3, and PIN4 mRNAs were found to be decreased after BSO treatment, indicating that the decrease is mediated at the transcriptional or mRNA stability level. This decrease is somehow correlated to the decrease of GSH level (−36 and −81% after 12 and 24 h of BSO treatment, respectively), suggesting that the GSH levels influence the abundance PIN mRNAs. However, the intensity of this decrease varies from one isoform to the other (two- to fourfold after 24-h treatment), suggesting that all PIN isoforms are not similarly affected by the treatment. Visualization of PIN proteins as fusions to green fluorescent protein (GFP) previously allowed detailed studies of their abundance and distribution. Transgenic plants expressing functional ProPIN1:PIN1-GFP, ProPIN2:PIN2-GFP, and ProPIN3:PIN3-GFP constructs were subjected to BSO treatment. The PIN1, PIN2, and PIN3 constructs all showed a pronounced decrease of abundance after 24 h of treatment (Figure 5B). Interestingly, the very weak GFP signal found after BSO treatment suggests that additional factors other than PIN mRNA abundance (i.e., posttranscriptional mechanisms, modified protein localization, etc.) may be influenced by the BSO treatment. Likewise, mRNA levels of the AUX1 gene coding for the auxin influx transporter were also altered by BSO treatment. Therefore, decreased abundance of auxin transport proteins certainly contributed to the suggested decrease in auxin transport capacity in the cad2 mutant. Moreover, the abundance of mRNA of the auxin response marker gene IAA1 is decreased in BSO-treated plants, indicating that BSO treatment also affects auxin signal transduction. By contrast, expression of the cytokinin response marker gene IBC7 was not affected, suggesting that BSO specifically affects auxin responses (Figure 5A).

Figure 5.
Figure 5.

PIN Expression Is Influenced by Glutathione and NTR.

(A) Five-day-old plantlets were transferred to a medium containing 2.5 mM BSO. Steady state levels of PIN1 (light green), PIN2 (dark green), PIN3 (light blue), PIN4 (dark blue), AUX1 (red), IAA1 (orange), and IBC7 (pink) mRNAs were analyzed by quantitative RT-PCR at different time points after treatment (0, 12, and 24 h). For each gene, the level of mRNA was normalized to that of EF1 α, which is constitutive under these conditions. Data are means of two technical replicates. The level of total glutathione was measured at the same time points and is reported in each panel (black lines). Data are means of six biological repetitions ± se, n = 6.

(B) Five-day-old plantlets expressing PIN1-GFP, PIN2-GFP, and PIN3-GFP proteins under the control of their natural promoters were subjected to the same treatment as in (A). Plants either treated (BSO) or untreated (C) with 2.5 mM BSO for 24 h were analyzed by confocal microscopy. Compared with untreated plants, the abundance of PIN1-GFP, PIN2-GFP, or PIN3-GFP was reduced in plants treated by BSO. Higher magnification of the PIN1-GFP and PIN2-GFP signals (inset) suggests that the overall cellular localization of the fusion protein is not perturbed by the BSO treatment. A line expressing the plasma membrane–localized LTI6b (low temperature induced) protein fused to GFP (GFP-LTI6b) under the 35S promoter was used as a control (Cutler et al., 2000). The abundance of the GFP-LT16b protein was not affected by BSO treatment. Bar = 50 μ m.

(C) and (D) PIN1 immunolocalization in Col-0 and ntra ntrb cad2. Abundance of PIN1 protein was analyzed in 5-d-old plantlets by immunolocalization and subsequently by confocal microscopy. Col-0 (C) and ntra ntrb cad2 (D). Arrowheads indicate some stellar cells in which a very low membrane signal persisted. Bar = 50 μ m.

To study PIN1 protein expression in the ntra ntrb cad2 mutant, we immunolocalized the PIN1 protein in primary root tips (Figures 5C and 5D). Wild-type plants showed a characteristic membrane localization at the lower side of stellar cells, but the PIN1 signal was dramatically decreased in the ntra ntrb cad2 mutant. Nevertheless, a very low membrane signal persisted in some stellar cells, suggesting that accumulation of PIN1 protein is affected rather than its cellular localization.

We also studied the expression of some auxin transport and response genes in mutant plants (see Supplemental Figure 5 online). The mRNA levels of these auxin genes were not affected in cad2 and ntra ntrb mutants, as shown previously in microarray data from Ball et al. (2004) and Bashandy et al. (2009). We also did not find any significant decrease in mRNA steady state levels of auxin genes in the ntra ntrb cad2 mutant. The fact that auxin-responsive genes are hardly affected in the cad2 mutant is somehow surprising in regard to the effect of BSO on auxin gene expression. The PIN, AUX1, and IAA1 gene expression may be more affected in wild-type plants suddenly subjected to BSO treatment than in the cad2 mutant that may be adapted to a constant lower GSH homeostasis. However, this hypothesis cannot explain the apparent discrepancy between the absence of changes in the transcripts level in ntra ntrb cad2 and the marked decrease in PIN1 accumulation in the ntra ntrb cad2 root tip. As suggested above for PIN:GFP data, posttranscriptional, translational, and/or posttranslational mechanisms may account for this disagreement.

As in the BSO treatment experiments, this mild effect on mRNA levels in the ntra ntrb cad2 mutant contrasts with the very low PIN1 immunolocalization found in the ntra ntrb cad2 mutant root tip, suggesting that in addition to mRNA abundance further mechanisms (e.g., protein stability or modification of cellular localization) may be involved in the PIN1 accumulation and the auxin transport capacities. Another hypothesis is that a decrease in auxin gene mRNA level is only restricted to specific tissues (i.e., root tip or flowers) and may be masked in whole plant extracts.

Auxin Level Is Altered in the ntra ntrb cad2 Mutant

The observation showing that decreased auxin transport in the cad2 mutant did not trigger a pin-like phenotype strongly suggests that the pin-like phenotype of the ntra ntrb cad2 mutant is due not only to perturbed auxin transport. We therefore tested whether auxin level is also perturbed in the ntra ntrb cad2 mutant. Auxin content was directly measured in the ntra ntrb cad2 mutant and compared with wild-type plants. Auxin (IAA) concentration was almost threefold lower in 2-week-old ntra ntrb cad2 mutant seedlings (Figure 6A). However, the auxin level was not affected in the ntra ntrb and cad2 mutants (see Supplemental Figure 6 online). To support the lower auxin level found in the ntra ntrb cad2 mutant, we crossed the mutant with the ProDR5:GUS auxin level marker line (Ulmasov et al., 1997). GUS staining was found to be lower in both the shoot and root tips of ntra ntrb cad2 ProDR5:GUS plants compared with wild-type ProDR5:GUS plants, supporting the finding that auxin level is lower in the ntra ntrb cad2 mutant (Figures 6B to 6E). We also measured the auxin level in dissected pin-like apex (3-mm-long apex) of adult ntra ntrb cad2 mutant plants and in similar structures from the pin1 homozygote mutant and in wild-type flower buds (Figure 6A). As expected, the auxin level was found to be lower in the ntra ntrb cad2 apex versus wild-type flower buds. More interesting, this lower level was not found in the similar pin-like apex of the pin1 mutant, showing that although the ntra ntrb cad2 and pin1 mutants harbor similar pin-like structures, their respective auxin levels are different. This suggests that the cause of the pin-like phenotype in the two mutants is distinct: in ntra ntrb cad2, it is associated with decreased auxin level, while this level is not affected in the in the pin1 mutant.

Figure 6.
Figure 6.

Auxin Levels Are Altered in the ntra ntrb cad2 Mutant.

Localization of the ProDR5:GUS activity in wild-type Col-0 ([B] and [C]) and in ntra ntrb cad2 ([D] and [E]) shoots ([B] and [D]) and roots ([C] and [E]).

(A) IAA levels were determined in wild-type Col-0 and homozygous ntra ntrb cad2 mutant seedlings 15 d after germination (left panel) grown as described in Methods and Figure 1A. Data are means of three biological repetitions ± se, n = 3. The IAA level is significantly lower in the ntra ntrb cad2 homozygotes versus wild-type seedlings. IAA levels were also measured in isolated wild-type Col-0 flower buds and in stem apex of adult ntra ntrb cad2 and pin1 homozygote plants grown in soil during 5 weeks as described in Methods and Figures 1C and 1D. Auxin content was also much lower in the ntra ntrb cad2 mutant but not in the pin1 apex.

(B) to (E) Auxin reporter ProDR5:GUS gene expression in the ntra ntrb cad2 mutant.

Rescue of ntra ntrb cad2 Mutant Development Phenotypes by GSH and Exogenous IAA

To provide evidence that the pin-like phenotype of the ntra ntrb cad2 mutant is due to low availability of GSH, we attempted to reverse the mutant phenotype by application of exogenous glutathione. Reduced glutathione (GSH) was added to irrigation water (0.4 mM). We observed that flowering occurred: flowers emerged but siliques could not develop, suggesting that these flowers are sterile (Figure 7). These data show that the lack of flower development in the ntra ntrb cad2 mutant can be partially reversed by exogenous GSH and demonstrate that the flower development phenotype of the ntra ntrb cad2 mutant is due to a limited availability of GSH. It also shows that GSH is efficiently transported from roots to shoots to trigger flower development in the ntra ntrb cad2 mutant. Similarly, Li et al. (2006) had previously shown that GSH can be transported from shoot to root to trigger root development in the cad2 mutant.

Figure 7.
Figure 7.

Rescue of ntra ntrb cad2 Floral Phenotype by Exogenous Glutathione.

Induction of flower development in ntra ntrb cad2 by supplementation with exogenous glutathione.

(A) After bolting, flowerless plants were regularly irrigated with water alone (−GSH) or with exogenous glutathione 0.4 mM (+GSH).

(B) Higher magnification of ntra ntrb cad2 flowers.

[See online article for color version of this figure.]

To study in more detail the rescue of flower development in glutathione-treated ntra ntrb cad2, we studied the mRNA steady state level of auxin transport and response genes in inflorescence meristems of the ntra ntrb cad2 mutant (see Supplemental Figure 7 online). Compared with wild-type Col-0 inflorescences, mRNA levels of PIN1, PIN2, PIN3, PIN4, AUX1, and IAA1 were found in lower levels in the ntra ntrb cad2 inflorescence meristems, suggesting that auxin metabolism and transport is perturbed in the mutant inflorescence. These data contrast with those showing no significant decrease of auxin genes in plantlets (see Supplemental Figure 5 online) and may argue for a specific perturbation in auxin-responsive gene expression at the meristem level. This perturbation may be masked by the dilution of meristematic tissues with other tissues in mRNA extracted from whole plant. By contrast, the cytokinin response marker gene IBC7 was not affected in the mutant. In GSH-rescued ntra ntrb cad2 inflorescences, the level of mRNAs of all genes was almost equivalent to wild-type inflorescence, showing that exogenous GSH rescued auxin transport and metabolism as well as flower development.

To study further the link between the pin-like phenotype and perturbed auxin homeostasis in ntra ntrb cad2, we also attempted to rescue the pin-like phenotype by exogenous auxin. To this purpose, we applied locally exogenous IAA (0.1 to 10 mM) at the top of the pin-like apex, as previously described by Reinhardt et al. (2000). No initiation of flower development occurred on 30 treated plants, suggesting that exogenous auxin is not sufficient to rescue the pin-like phenotype of the ntra ntrb cad2 mutant. By contrast, when the same treatment was performed on the pin1 mutant, initiation of flower meristem-like structures was observed after 4 to 7 d at 1 mM IAA on 15/15 treated plants.

It is well known that plant hormones, especially auxin and cytokinin, play important roles in adventitious shoot formation on calli. The perturbed auxin level in the ntra ntrb cad2 mutant suggested that the mutation also changed the callus regeneration capacity and/or shoot regeneration from calli. The dedifferentiation capacity of the ntra ntrb cad2 mutant to form calli was first evaluated on callus-inducing medium plates (Figure 8). Callus regeneration from explants of different tissues of the ntra ntrb cad2 mutant was compared with that of the wild type. Callus regeneration was not modified in the mutant, indicating that cell division was not perturbed. When calli were transferred onto shoot-inducing medium plates containing fixed concentrations of the synthetic cytokinin 2-BAP and different concentrations of IAA, wild-type calli dedifferentiated to form adventitious shoots (Figures 8I to 8K). Leaf primordia could be observed after 7 d and adventitious shoots after 15 d. The number of calli with adventitious shoots increased gradually, and the number of adventitious shoot were counted after 21 d (Figure 8L). However, ntra ntrb cad2 mutant calli were unable to dedifferentiate in the absence of exogenous auxin. When increased concentrations of IAA were added, shoot regeneration resumed, suggesting that auxin is limiting for shoot regeneration in the ntra ntrb cad2 mutant (Figures 8I to 8K).

Figure 8.
Figure 8.

The ntra ntrb cad2 Mutant Is Less Susceptible to the de Novo Shoot Induction System.

(A) to (H) Shoot apical meristem ([A] and [E]), stem ([B] and [F]), leaf ([C] and [G]), and root explants ([D] and [H]) were harvested from 7-week-old wild-type Col-0 ([A] to [D]) or ntra ntrb cad2 ([E] to [H]) mutant plants and transferred to an auxin-rich callus-inducing medium, which induces cell proliferation and callus formation after 3 weeks.

(I) to (K) In order to regenerate shoots, the calli induced from leaves for 3 weeks were transferred to a cytokinin-rich (10 μ M BAP) shoot-inducing medium containing increasing concentrations of auxin: 0 μ M IAA (I), 0.1 μ M IAA (J), and 1 μ M IAA (K). After 2 additional weeks, regenerated shoot meristems were observed. Note that shoot regeneration occurred in all media in Col-0 calli but only in media containing high concentrations of IAA in ntra ntrb cad2 calli. The black arrow indicated the regenerated shoots.

(L) Counting of the number of regenerated shoot per callus in Col-0 and ntra ntrb cad2 (±se, n = 30). No IAA (white bars), 0.1 μ M IAA (yellow), 1 μ M IAA (orange), and 3 μ M IAA (red).

We wanted to know whether exogenous IAA also rescued the perturbations of root development in the ntra ntrb cad2 mutant. Primary root elongation as well as the number of secondary roots was estimated in the different mutants (Figure 9). As expected, increasing concentrations of exogenous IAA inhibited primary root growth elongation in both wild-type and mutant plants. Intriguingly, low concentrations IAA (10−8 M) inhibited root elongation in the wild type, but this concentration was not effective with cad2, ntra ntrb, and ntrb ntrb cad2 mutants, suggesting that they may be less sensitive to auxin. However, this effect was less obvious in the pad2 mutant (Figure 9A). Increasing concentrations of auxin triggered secondary root emergence in all of these mutants, showing that exogenous hormone partially rescued the lack of secondary root emergence in the ntra ntrb cad2 mutant (Figures 9B and 9C). However, higher concentrations of IAA (10−6 M) were required to rescue the secondary root emergence in the ntra ntrb cad2 mutant, suggesting that either auxin is limiting for secondary root development in the mutant or the mutant is less sensitive to auxin.

Figure 9.
Figure 9.

ntra ntrb cad2 Root Development Phenotype Is Rescued by Exogenous Auxin.

Col-0, cad2, pad2, ntra ntrb, and ntra ntrb cad2 plants grown on MS/2 vertical agar plates at 22°C, under a 16-h-light/8-h-dark regime. Five days after seeds were put under growth culture, plantlets were transferred on plates containing different concentrations of IAA.

(A) Primary root growth rate was calculated as the mean of daily root growth measured between the 2nd and the 5th day after the transfer. Data are means ± se; n ≥ 30.

(B) and (C) Number of secondary roots initials was measured 2 d (B) and 6 d (C) after the transfer. Data are means ± se; n ≥ 30.

DISCUSSION

Association of ntra ntrb and cad2 Mutants Reveals Additional Developmental Phenotypes

Depending on its level, GSH deficiency produces several phenotypic consequences. In Arabidopsis, γ-glutamylcysteine synthetase (GSH1), the first enzyme of GSH synthesis, is encoded by a unique gene. Complete inactivation of this gene causes white seed formation and embryo lethality (Cairns et al., 2006). Fortunately, several alleles of the gene are available in Arabidopsis: rml1 produces a very inefficient GSH1 that produces ∼5% glutathione in comparison to the wild type. Homozygous seeds are formed with a normal embryo, but the root meristem does not grow during germination. By contrast, the shoot meristem develops, at least in the initial stage. pad2-1 harbors ∼16% glutathione and cad2, ∼20%. Both lines display normal fertility and development under standard conditions (Cobbett et al., 1998; Parisy et al., 2007). pad2-1 was shown to be sensitive to several pathogens, possibly due to inefficient production of phytoalexins and glucosinolates (Parisy et al., 2007). cad2 is hypersensitive to Cd, which may be due to a limited synthesis of phytochelatins (Howden et al., 1995). We previously characterized the ntra ntrb mutant as a mutant inactivated in both mitochondrial and cytosolic thioredoxin reductases. It does not show a pronounced phenotype. By crossing the ntra ntrb mutant with rml1, we obtained seed with embryos blocked in both the root and shoot meristems, showing that glutathione plays an overlapping function with the cytosolic thioredoxin system in controlling shoot meristem activity (Reichheld et al., 2007). In contrast with rml1, cad2 shows wild-type development. However, association of ntra ntrb and cad2 mutations revealed developmental perturbations in the triple homozygote plants that are observed in none of the parent mutants, indicating additive effects of the ntra ntrb and cad2 mutations on plant development. The most obvious impact of the ntra ntrb cad2 mutant is that flowers are not formed, the floral stems resembling pin-like structures. This developmental trait was fully complemented by expression of the NTRB transgene, indicating that this phenotype is not due to independent mutations or chromosome rearrangements. Furthermore, we show that a single NTR is sufficient for normal flower development. We have previously reported on the redundancy between NTRA and NTRB in other functions of plant development (Reichheld et al., 2007; Bashandy et al., 2009). This redundancy is supported by the fact that the two genes share overlapping biochemical activities, cellular localization (i.e., cytosolic and mitochondrial), and tissue expression, suggesting that selective pressure favors the maintenance of both NTR copies in the genome. By contrast, knockout in mammalian NTR trr1 and trr2 single mutants has been shown to lead to specific embryo lethal phenotypes due to specific cellular localization and tissue expression of the TRR1 and TRR2 (Conrad et al., 2004; Jakupoglu et al., 2005; Soerensen et al., 2008).

The flowerless phenotype of the ntra ntrb cad2 mutant also indicates that proper flower development is dependent on high availability of glutathione. This fact is confirmed by our GSH irrigation experiments showing rescue of flower development in the ntra ntrb cad2 mutant. Therefore, we assume that GSH is transported or can freely diffuse from the roots to the inflorescence through the phloem. Basipetal glutathione transit through root sink tissues has already been suggested by other reports (Li et al., 2006). The rescue of the pin-like phenotype by GSH supply also suggests that this phenotype is not triggered by a perturbed development of the vascular system as was shown in the interfascular fiberless/revoluta (ifl1/rev) mutant (Zhong et al., 1997). However, in our case, GSH-rescued ntra ntrb cad2 flowers were not fertile, suggesting that a particular fertilization step cannot be overcome by exogenous GSH supply.

Thiol Reduction Pathways Regulate Auxin Homeostasis

The ntra ntrb cad2 morphological phenotypes (pin-like phenotype with defects in vasculature and secondary root growth) suggest a deficiency in auxin metabolism, signaling, or transport. Both pin1, pid, and ifl1/rev1 mutants show a pin-like phenotype affected in auxin transport, suggesting that the primary effect of the ntra ntrb cad2 mutations may be due to perturbed auxin transport. Accordingly, we have shown that auxin transport is highly affected in the ntra ntrb cad2 mutant, the basipetal stem auxin transport being as affected as in the pin1 mutant. PIN1 immunolocalization also showed a dramatic decrease of PIN1 protein accumulation in the root meristem of the ntra ntrb cad2 mutant that may explain the perturbed auxin transport.

The PAT perturbation is probably provided by the additive effect of both ntra ntrb and cad2 mutations as both lines showed decreased 3H-IAA transport. However, the fact that the cad2 and pad2 mutants showed a pronounced PAT decrease without the shoot developmental perturbations (i.e., pin structure and loss of apical dominance) observed in the ntra ntrb cad2 mutant argue against the idea that PAT is responsible for those phenotypes. However, PAT perturbation may be responsible for the root phenotype of the cad2, ntra ntrb, and ntra ntrb cad2 mutants.

Nevertheless, our results can be most easily explained by the observation that the ntra ntrb cad2 mutant has lower auxin (IAA) concentrations. Likewise, an auxin reporter gene, ProDR5:GUS, displays lower expression level in the ntra ntrb cad2 background. Our data showing a loss of ability of ntra ntrb cad2 mutant calli to regenerate shoots in the absence of exogenous auxin suggest that either this mutant lacks an auxin threshold necessary to regenerate shoots or the mutant is less susceptible to auxin.

Feedback mechanisms between auxin transport and auxin metabolism have been largely described (reviewed in Benjamins and Scheres, 2008). The npy1 mutant has recently been isolated in a genetic screen for enhancers of the yuc1 yuc4 double mutant, a background that is partially defective in auxin biosynthesis. In contrast with the npy1 and yuc1 yuc4 double mutants, the npy1 yuc1 yuc4 triple mutant exhibited a pin-like phenotype. Interestingly, NPY1 was found to act in the same pathway as PID, a protein kinase involved in auxin transport. Therefore, the pin-like phenotype of the npy1 yuc1 yuc4 triple mutant is probably due to a combination of auxin synthesis and auxin transport (Cheng et al., 2007). Whether perturbed auxin level, auxin transport, or a combination of both effects is responsible for the pin-like phenotype in the ntra ntrb cad2 mutant remains to be determined.

The fact that expression of the cytokinin-responsive gene IBC7 is not affected in the mutant and that callus regeneration can occur in the absence of exogenous cytokinin suggests that cytokinin is not a limiting factor for the mutant.

Meristematic Functions Affected in the ntra ntrb cad2 Mutant

The pin-like structure of the ntra ntrb cad2 stems suggests some dysfunction of flower meristem development. However, vegetative-to-inflorescence meristem transition, stem growth, and emergence of secondary rosette stems are not affected, suggesting that the dysfunction is restricted to flower development. Our cytological studies of the apical meristem structures support this point: the shoot apical meristem is not profoundly perturbed, initial organs are formed but cannot develop further, and flower meristem is not observed. It remains to be shown whether these initial organs correspond to aborted flowers or leaves.

Root apical meristem (RAM) and shoot apical meristem are not profoundly affected in the ntra ntrb cad2 mutant, in contrast with rml1 and ntra ntrb rml1 mutants in which RAM and both RAM and shoot apical meristematic activities are almost completely inhibited. Therefore, a comparison of the phenotype of ntra ntrb cad2 and ntra ntrb rml1 mutants suggests that a threshold glutathione concentration is necessary for root and shoot development. Moreover, the ntra ntrb cad2 mutations define another threshold necessary for proper flower development. This suggests that flower development may be much more sensitive to redox perturbation than leaf development.

Auxin is key for both shoot and root meristematic activities but is also metabolized in actively dividing tissues. Therefore, it would be interesting to know whether the perturbed auxin metabolism in the ntra ntrb cad2 mutant affects meristematic activities or if the auxin perturbation is a secondary effect of perturbed meristematic activity. Solving this question will be a major aim for future research. The fact that the pin-like phenotype was apparently not rescued by exogenous auxin favors the second hypothesis. However, the partial rescue by exogenous IAA of the root phenotypes (i.e., primary root growth and secondary root initiation) favors a direct role of auxin in root meristem activities.

Is the ntra ntrb cad2 Phenotype Trx or Grx Dependent?

Which mechanism explains the redundancy between NTRs and GSH in meristem development? NTRs are the reductants of thioredoxins and so far no other target is known in plants. Thus, NTRs allow normal flower formation of the cad2 mutant through the reduction of one or several Trx in the flower meristem, which, in turn, reduce a particular target. A high level of GSH can substitute for the absence of NTR to allow flower formation in the ntra ntrb mutant. GSH has several functions, but we have previously shown that, in the absence of NTR, a GSH-dependent pathway is able to reduce Trx. Thus, the first possibility relies on this GSH function. Nevertheless Trx h3, the Trx reduced by GSH, is not oxidized to a greater extent in ntra ntrb cad2 than in ntra ntrb. It is possible that other Trx are less efficiently reduced by the GSH pathway when the GSH content drops to the level of the cad2 mutant. We have constructed a quadruple mutant in which four cytosolic TRXh (TRX h1, h2, h3, and h5) are inactivated. This mutant does not show any phenotype. Similarly, a knockout of TRX h4 is asymptomatic. Another important function of GSH is the reduction of Grxs, the other family of disulfide reductases thath in most organisms are alternative reducers of several Trx targets, including deoxyribonucleotide reductase, a key enzyme in cell proliferation. GSH is redundant with Grx, and Trx and Grx have some common target proteins in most organisms, including Arabidopsis. Thus, the phenotype of the ntra ntrb cad2 mutant may result from inefficient reduction of one or several protein(s) targeted by both Trx(s) and Grx(s).

Two closely related CC-type glutaredoxin proteins (ROXY1 and ROXY2) were recently shown to act in flower development (Xing et al., 2005; Xing and Zachgo, 2008; Li et al., 2009). Flowers develop in these plants, but the roxy1 mutant exhibits abnormalities during petal development, while the roxy1 roxy2 double mutant shows additional anther development perturbation. Other members of the Grx family were also shown to be able to interact with TGA transcription factors. Thus, identification of the active Trxs and Grxs and their target proteins will help to understand if the hormonal alteration is the primary cause or is a consequence of the developmental perturbation.

METHODS

Plant Materials, Growth Conditions, and Treatments

Seedlings and plants from Arabidopsis thaliana ecotype Col-0 (the wild type and ntra ntrb, cad2, and pin1-7 mutant lines described by Reichheld et al. [2007], Howden et al. [1995], and Noh et al. [2001], respectively) were used for these experiments. For in vitro seedling growth, seeds were surface sterilized and plated on half-strength MS medium, including Gamborg B5 vitamins (M0231; Duchefa), 1% (w/v) sucrose, and 0.8% (w/v) plant agar. For growth in soil, seeds were sown in pots containing a mixture of soil and vermiculite (3:1, v/v) and irrigated with water. Both plants and seedlings were grown at 22°C and 70% humidity under a 16-h-light (4000 lux)/8-h-dark regime.

Supplementation of the growth medium with BSO (Sigma-Aldrich) was performed as follows. Appropriate amounts of the filter-sterilized stock solution containing these compounds were added to sterile Petri dishes, and molten MS/2 medium that had been cooled to 50°C was added while swirling the plates.

Arabidopsis plants harboring cad2 and ntra ntrb mutations, ProNTRA:GUS, and ProNTRB:GUS were described previously (Howden et al., 1995; Reichheld et al., 2007). ProGSH1:GUS plants were kindly provided by Andreas Wachter and Andreas Meyer (Heidelberg Institute for Plant Science, Heidelberg). Arabidopsis plants harboring ProDR5:GUS, ProPIN1:PIN1-GFP, ProPIN2:PIN2-GFP, ProPIN3:PIN3-GFP, and Pro35S:GFP-LTI6b were described previously (Ulmasov et al., 1997; Cutler et al., 2000; Benková et al., 2003; Yin et al., 2007).

For ntra ntrb cad2 mutant analysis, the cad2 mutation was introgressed into the ntra ntrb mutant background. For auxin reporter gene expression, the construct ProDR5:GUS was introgressed in the ntra ntrb cad2 mutant. For each cross, 10 flower buds were emasculated and pollinated with the partner mutant in both cross combination (male versus female and female versus male). For isolation to the homozygote mutants, seeds were selected on MS/2 medium supplemented by 0.5 mM BSO to select homozygous ntra ntrb mutants. Homozygous lines were further selected for the cad2 mutation. Genotyping of the cad2 allele was performed by PCR amplification using CAD2-forward and CAD2-reverse primers (see Supplemental Table 2 online). In the wild-type allele, a 132-bp fragment is amplified, and in the mutant allele, the fragment is 126 bp. The distinct fragments are separated on a 4% Metaphor Agarose (BMA).

For root growth measurements, seeds were germinated on vertical plates, and the position of the root tip was marked on the underside of the plate. Every day after the transfer, root growth was measured on 20 to 30 roots per plates using a binocular microscope (Leica MZ12) with the aid of an eyepiece reticle.

For callus formation and plant regeneration, different explants (flower buds, meristem, stem, and roots) were used for callus induction on MS medium supplemented with 2.2 μ M 2,4-D. Then, calli were transferred to three different MS-modified regeneration media supplemented with BAP and/or IAA and incubated under the same light conditions as previously described.

Microscopy Observations, Tissue Cross Sections, and Confocal Laser Scanning Microscopy

For phenotypic characterization, tissues were cleared overnight with Hoyer's solution (2.5 g arabic gum, 100 g chloral hydrate, 5 mL glycerol, and 30 mL distillate water), mounted on slides, and observed under a binocular microscope (Zeiss Axioskop2). Pictures were taken by a numeric camera (Leica DC300FX) coupled to imaging software (Leica FW4000). Cross sections of the wild type and ntra ntrb cad2 mutant meristems were prepared using a microtome (Leica RM2255) from tissues embedded in hydroxyethyl methacrylate (Technovit 7100; Heraus-Kulter) and were stained with Toluidine blue 1% in 1% Natrium Tetraborate.

Localization of fluorescent protein-tagged proteins in fresh tissue samples was performed using a Zeiss LSM 510 Meta (Carl Zeiss). GFP was excited with the argon laser at 488 nm, and the emitted fluorescence was detected between 505 and 520 nm. Images were processed using Adobe Photoshop CS3 to increase the contrast and brightness of the transmission channel.

Treatments of Plant Apices

For local treatments of apices, IAA (Sigma-Aldrich) and 1 M stock solutions in DMSO were dissolved in a prewarmed (50°C) paste consisting of lanolin with 2.5% paraffin (Merck). The paste was manually administered to Arabidopsis apices early after bolting, using a needle. IAA-treated plants were further cultivated in growth room until observation.

Gene Expression Analysis by RT-PCR, Quantitative RT-PCR, and the GUS Reporter

Total RNA was extracted from frozen plant organs using TRIzol reagent (Gibco BRL) according to the manufacturer's protocol. For RT-PCR, 5 μ g of DNase I–treated total RNA was used for first-strand synthesis of cDNA using oligo(dT) primer reverse transcription with the Moloney murine leukemia virus reverse transcriptase as described by the manufacturer's protocol (AffinityScript Multiple Temperature cDNA synthesis kit; Stratagene). PCR (25 to 34 cycles as described in corresponding figure legends) was performed as described by Laloi et al. (2004). PCR fragments were detected by GelRed (Biotium) staining and visualized with U-Genius (Syngene). For each RT-PCR experiment, two biological repetitions were performed with similar results.

For quantitative RT-PCR, the following Light Cycler run protocol was used: denaturation program (95°C, 10 min), amplification and quantification programs repeated 40 times (95°C for 15 s, 60°C for 5 s, and 72°C for 16 s), melting curve program (60 to 95°C with a heating rate of 0.1°C per second and continuous fluorescence measurement), and a cooling step to 40°C. Amplification of single highly specific products was verified (melting curve analysis). For each reaction, the crossing point was determined using the Fit Point Method of the Light Cycler Software 3.3 (Roche Diagnostics). PCR reactions were performed in duplicate, and the mean values of the crossing point were calculated. For each candidate gene, the level of transcription of the target gene was normalized using the transcription rate of the reference gene. The reference gene used in this study is the EF1 α RNA from Arabidopsis. Specific primers for quantitative RT-PCR and RT-PCR experiments were edited using the Light Cycler Probe Design Software version 1.0 (Roche Diagnostics) (see Supplemental Table 2 for primer sequences).

For ProNTRA, ProNTRB, and ProGSH1:GUS expression analyses, the constructs and experiment were performed as described by Cairns et al. (2006) and Reichheld et al. (2007), respectively.

Cloning and Transformation of Complemented Plants

For complementation of the ntra ntrb cad2 mutant with the NTRB gene, a genomic fragment of 3270 bp (1497 bp upstream from ATG to 381 bp downstream from the stop codon) was amplified by PCR using primers introducing unique XhoI and XbaI sites (see Supplemental Table 2 online) and cloned in a pCAMBIA-1300-HYG binary vector as described by Reichheld et al. (2007). The resulting plasmid was introduced into Agrobacterium tumefaciens GV3101, and the construct was transformed into ntra/ntra ntrb/ntrb GSH1/cad2 heterozyzous plants by the floral dip method (Clough and Bent, 1998). T1 seedlings were selected in vitro on MS/2 medium supplemented with 30 μ g/mL hygromycin. T2 transformed plants were screened by PCR for the cad2 mutation and for expression of the NTRB transgene. Protein gel blots were hybridized with anti-NTRB antibodies as described by Reichheld et al. (2005).

In Situ Hybridization

In situ hybridization was performed as described earlier (Laufs et al., 1998). Briefly, plants were fixed in 4% formaldehyde (freshly made from paraformaldehyde) in PBS under vacuum for 2 h and 20 min and left in fixative overnight. After fixation, plants were washed, dehydrated, and embedded in paraffin wax. Paraffin sections (7 μ m thick) were cut with a disposable metal knife and attached to precoated glass slides (DAKO). Antisense probes were synthesized using digoxigenin (DIG)-UTP; Boehringer Mannheim) according to the manufacturer's instructions. Immunodetection of the DIG-labeled probes was performed using an anti-DIG antibody coupled to alkaline phosphatase as described by the manufacturer (Boehringer Mannheim). A full-length cDNA of NTRA was used as probe (see Supplemental Table 2 online).

Auxin Transport Assays

Auxin transport was measured according to Besseau et al. (2007). A 2.5-cm segment of flowering stem was excised 5 cm above the base of the stem. Segments were placed in a 1.5 mL microcentrifuge tube, and the apical ends submerged in 30 μ L of MES buffer (5 mM MES and 1% sucrose, pH 5.5) containing 1 μ M IAA and 66 nM of 3H-IAA. After 4 h of incubation in the dark, the segments were removed and the last 5 mm of the nonsubmerged ends were excised and placed in 2.5 mL of liquid scintillation cocktail (Optiphase-highsafe 2; Perkin-Elmer). The samples were slowly shaken overnight before measuring radioactivity in a scintillation counter (Beckman LS6000 SC).

Determination of IAA Concentrations

Col-0, cad2, ntra ntrb, ntra ntrb cad2, and pin1 plants were grown on MS/2 agar plates under long-day conditions. Two weeks after germination, whole seedlings or isolated shoot tissue were pooled and frozen in liquid nitrogen for quantification of auxin content. Isolated wild-type Col-0 or cad2 flower buds and shoot apical meristems of ntra ntrb cad2 or pin1 adult plants grown in soil during 7 weeks were also collected. Three replicates were analyzed for each sample.

IAA was extracted and purified as described (Andersen et al., 2008). After methylation, the samples were trimethyl-silylated and analyzed by gas chromatography–selected reaction monitoring–mass spectrometry (Edlund et al., 1995).

Measurement of Total and Oxidized Glutathione

Total glutathione content of plants was determined using the recycling enzymatic assay, as described by May and Leaver (1993). Briefly, 100 mg of fresh plant material ground in liquid nitrogen was extracted in 0.5 mL 0.1 M Na phosphate buffer, pH 7.6, and 5 mM EDTA. After microcentrifugation (10 min, 9000g), total glutathione in 0.1 mL of the supernatant was measured by spectrophotometry in a 1 mL mixture containing 6 mM dithiobis(2-nitrobenzoic acid) (Sigma-Aldrich), 3 mM NADPH, and 2 units of glutathione reductase from Saccharomyces cerevisiae (Sigma-Aldrich). Glutathione-dependent reduction of dithiobis(2-nitrobenzoic acid) was followed at 412 nm. Total glutathione levels were calculated using the equation of the linear regression obtained from a standard GSH curve. Oxidized glutathione (GSSG) was determined in the same extracts after derivatization of reduced GSH. Derivatization of 100 μ L plant extract was performed in 0.5 mL 0.5 M K phosphate buffer, pH 7.6, in the presence of 4 μ L 2-vinyl pyridine (Sigma-Aldrich) during 1 h at room temperature. After extraction of the GSH-conjugated 2-vinyl pyridine with 1 volume of diethylether, the GSSG was measured by spectrophotometry as described for total glutathione.

Redox State of TRXh3 by 4-acetamido-4-maleimidylstilbene-2,2-disulfonic acid Treatment

The redox state of TRXh3 was determined as described (Ritz and Beckwith, 2002) with minor modification. Arabidopsis plantlets were ground in liquid nitrogen, and the fine powder was dissolved in 5% trichloroacetic acid. After filtration on a 60-mm nylon net filter (NY60; Millipore) to remove tissue fragments, proteins were precipitated for 1 h on ice. Protein extracts were collected after centrifugation for 15 min, 11,000g at 4°C. Pellets were washed with ice-cold acetone, dried, and dissolved in 200 mM Tris-HCl, pH 7.5, with 2% SDS. Cys residues were alkylated by 15 mM 4-acetamido-4-maleimidylstilbene-2,2-disulfonic acid for 1 h at room temperature in the dark. Proteins were separated by 15% SDS-PAGE under nonreducing conditions as described previously (Serrato and Cejudo, 2003). TRXh3 protein was visualized by immunoblot with anti-TRXh3 antibodies.

Database Searches and Sequence Analysis

DNA sequences were analyzed using a locally installed BLAST server (Altschul et al., 1997) against public and local nucleotide and protein databases. The physical location of genes in the Arabidopsis genome was determined using the maps available at The Arabidopsis Information Resource (http://www.Arabidopsis.org/). Multiple sequence analysis was performed using the ClustalW software (http://npsa-pbil.ibcp.fr/cgi-bin/npsa_automat.pl?page=/NPSA/npsa_clustalw.html).

Accession Numbers

Sequence data for NTRA, NTRB, TRXh3, GSH1, PIN1, PIN2, PIN3, PIN4, AUX1, IAA1, IBC7, ACT2, and EF1α can be found in the GenBank/EMBL data libraries under accession numbers NP 179334 (At2g17420), NP 195271 (At4g35460), NP 199112 (At5g42980), NP 194041 (At4g23100), NP 177500 (At1g73590), NP 568848 (At5g57090), NP 177250 (At1g70940), NP 849923 (At2g01420), NP 179701 (At2g21050), NP 193192 (At4g14560), NP 172517 (At1g10470), NP 850611 (At3g18780), and NP 563801 (At1g07940), respectively.

Supplemental Data

The following materials are available in the online version of this article.

  • Supplemental Figure 1. Complementation of the ntra ntrb cad2 Mutant.

  • Supplemental Figure 2. Redox State of TRXh3 in the ntra ntrb cad2 Mutant.

  • Supplemental Figure 3. Expression of the GSH1 Gene.

  • Supplemental Figure 4. In Situ Hybridization of NTR Gene in the Inflorescence.

  • Supplemental Figure 5. Expression of Auxin Transport and Response Genes in Mutants.

  • Supplemental Figure 6. Auxin Levels Are Not Altered in the cad2 and ntra ntrb Mutants.

  • Supplemental Figure 7. Expression of Auxin Transport and Response Genes in the ntra ntrb cad2 Mutant Rescued by Glutathione.

  • Supplemental Table 1. Total Glutathione Level Is Not Altered in the ntra ntrb cad2 Mutant.

  • Supplemental Table 2. List of Oligonucleotides.

Acknowledgments

We thank B.B. Buchanan and R. Cooke for fruitful discussions and for critical reading the manuscript. We thank the ABRC for T-DNA insertional mutants, Andreas Wachter and Andreas Meyer (HIP, Heidelberg) for kindly providing us with the ProGSH1:GUS transgenic line, and Roger Granbom and Guillaume Mitta for technical assistance in IAA measurements and quantitative RT-PCR analyses, respectively. We gratefully acknowledge funding from the Centre National de la Recherche Scientifique, Genoplante (Project GNP0508) and Agence Nationale de la Recherche (ANR-Blanc-06-0047). T.V. is supported by a Career Development Award from the Human Frontier Science Program Organization and by a JC-JC grant from the Agence National de la Recherche.

Footnotes

  • The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantcell.org) is: Jean-Philippe Reichheld(jpr{at}univ-perp.fr).

  • www.plantcell.org/cgi/doi/10.1105/tpc.109.071225

  • ↵[C] Some figures in this article are displayed in color online but in black and white in the print edition.

  • ↵[W] Online version contains Web-only data.

  • Received September 24, 2009.
  • Revised January 21, 2010.
  • Accepted January 29, 2010.
  • Published February 17, 2010.

References

  1. ↵
    1. Altschul S.F.,
    2. Madden T.L.,
    3. Schäffer A.A.,
    4. Zhang J.,
    5. Zhang Z.,
    6. Miller W.,
    7. Lipman D.J.
    (1997). Gapped BLAST and PSI-BLAST: A new generation of protein database search programs. Nucleic Acids Res. 25: 3389–3402.
    OpenUrlAbstract/FREE Full Text
  2. ↵
    1. Andersen S.U.,
    2. Buechel S.,
    3. Zhao Z.,
    4. Ljung K.,
    5. Novák O.,
    6. Busch W.,
    7. Schuster C.,
    8. Lohmann J.U.
    (2008). Requirement of B2-type cyclin-dependent kinases for meristem integrity in Arabidopsis thaliana. Plant Cell 20: 88–100.
    OpenUrlAbstract/FREE Full Text
  3. ↵
    1. Arnér E.S.,
    2. Holmgren A.
    (2000). Physiological functions of thioredoxin and thioredoxin reductase. Eur. J. Biochem. 267: 6102–6109.
    OpenUrlCrossRefPubMed
  4. ↵
    1. Ball L.,
    2. Accotto G.,
    3. Bechtold U.,
    4. Creissen G.,
    5. Funck D.,
    6. Jimenez A.,
    7. Kular B.,
    8. Leyland N.,
    9. Mejia-Carranza J.,
    10. Reynolds H.,
    11. Karpinski S.,
    12. Mullineaux P.M.
    (2004). Evidence for a direct link between glutathione biosynthesis and stress defense gene expression in Arabidopsis. Plant Cell 16: 2448–2462.
    OpenUrlAbstract/FREE Full Text
  5. ↵
    1. Barlier I.,
    2. Kowalczyk M.,
    3. Marchant A.,
    4. Ljung K.,
    5. Bhalerao R.,
    6. Bennett M.,
    7. Sandberg G.,
    8. Bellini C.
    (2000). The SUR2 gene of Arabidopsis thaliana encodes the cytochrome P450 CYP83B1, a modulator of auxin homeostasis. Proc. Natl. Acad. Sci. USA 97: 14819–14824.
    OpenUrlAbstract/FREE Full Text
  6. ↵
    1. Bashandy T.,
    2. Taconnat L.,
    3. Renou J.,
    4. Meyer Y.,
    5. Reichheld J.P.
    (2009). Accumulation of flavonoids in an ntra ntrb mutant leads to tolerance to UV-C. Mol. Plant 2: 249–258.
    OpenUrlCrossRefPubMed
  7. ↵
    1. Benitez-Alfonso Y.,
    2. Cilia M.,
    3. San Roman A.,
    4. Thomas C.,
    5. Maule A.,
    6. Hearn S.,
    7. Jackson D.
    (2009). Control of Arabidopsis meristem development by thioredoxin-dependent regulation of intercellular transport. Proc. Natl. Acad. Sci. USA 106: 3615–3620.
    OpenUrlAbstract/FREE Full Text
  8. ↵
    1. Benjamins R.,
    2. Scheres B.
    (2008). Auxin: The looping star in plant development. Annu. Rev. Plant Biol. 59: 443–465.
    OpenUrlCrossRefPubMed
  9. ↵
    1. Benková E.,
    2. Michniewicz M.,
    3. Sauer M.,
    4. Teichmann T.,
    5. Seifertová D.,
    6. Jürgens G.,
    7. Friml J.
    (2003). Local, efflux-dependent auxin gradients as a common module for plant organ formation. Cell 115: 591–602.
    OpenUrlCrossRefPubMed
  10. ↵
    1. Bennett M.J.,
    2. Marchant A.,
    3. Green H.G.,
    4. May S.T.,
    5. Ward S.P.,
    6. Millner P.A.,
    7. Walker A.R.,
    8. Schulz B.,
    9. Feldmann K.A.
    (1996). Arabidopsis AUX1 gene: A permease-like regulator of root gravitropism. Science 273: 948–950.
    OpenUrlAbstract/FREE Full Text
  11. ↵
    1. Bennett R.S.M.,
    2. Alvarez J.,
    3. Bossinger G.,
    4. Smyth D.R.
    (1995). Morphogenesis in pinoid mutants of Arabidopsis thaliana. Plant J. 8: 505–520.
    OpenUrlCrossRef
  12. ↵
    1. Besseau S.,
    2. Hoffmann L.,
    3. Geoffroy P.,
    4. Lapierre C.,
    5. Pollet B.,
    6. Legrand M.
    (2007). Flavonoid accumulation in Arabidopsis repressed in lignin synthesis affects auxin transport and plant growth. Plant Cell 19: 148–162.
    OpenUrlAbstract/FREE Full Text
  13. ↵
    1. Broin M.,
    2. Cuiné S.,
    3. Eymery F.,
    4. Rey P.
    (2002). The plastidic 2-cysteine peroxiredoxin is a target for a thioredoxin involved in the protection of the photosynthetic apparatus against oxidative damage. Plant Cell 14: 1417–1432.
    OpenUrlAbstract/FREE Full Text
  14. ↵
    1. Broin M.,
    2. Cuiné S.,
    3. Peltier G.,
    4. Rey P.
    (2000). Involvement of CDSP 32, a drought-induced thioredoxin, in the response to oxidative stress in potato plants. FEBS Lett. 467: 245–248.
    OpenUrlCrossRefPubMed
  15. ↵
    1. Broin M.,
    2. Rey P.
    (2003). Potato plants lacking the CDSP32 plastidic thioredoxin exhibit overoxidation of the BAS1 2-cysteine peroxiredoxin and increased lipid peroxidation in thylakoids under photooxidative stress. Plant Physiol. 132: 1335–1343.
    OpenUrlAbstract/FREE Full Text
  16. ↵
    1. Buchanan B.B.,
    2. Balmer Y.
    (2005). Redox regulation: A broadening horizon. Annu. Rev. Plant Biol. 56: 187–220.
    OpenUrlCrossRefPubMed
  17. ↵
    1. Cairns N.G.,
    2. Pasternak M.,
    3. Wachter A.,
    4. Cobbett C.S.,
    5. Meyer A.J.
    (2006). Maturation of arabidopsis seeds is dependent on glutathione biosynthesis within the embryo. Plant Physiol. 141: 446–455.
    OpenUrlAbstract/FREE Full Text
  18. ↵
    1. Carmel-Harel O.,
    2. Storz G.
    (2000). Roles of the glutathione- and thioredoxin-dependent reduction systems in the Escherichia coli and Saccharomyces cerevisiae responses to oxidative stress. Annu. Rev. Microbiol. 54: 439–461.
    OpenUrlCrossRefPubMed
  19. ↵
    1. Cheng Y.,
    2. Dai X.,
    3. Zhao Y.
    (2006). Auxin biosynthesis by the YUCCA flavin monooxygenases controls the formation of floral organs and vascular tissues in Arabidopsis. Genes Dev. 20: 1790–1799.
    OpenUrlAbstract/FREE Full Text
  20. ↵
    1. Cheng Y.,
    2. Qin G.,
    3. Dai X.,
    4. Zhao Y.
    (2007). NPY1, a BTB-NPH3-like protein, plays a critical role in auxin-regulated organogenesis in Arabidopsis. Proc. Natl. Acad. Sci. USA 104: 18825–18829.
    OpenUrlAbstract/FREE Full Text
  21. ↵
    1. Christensen S.K.,
    2. Dagenais N.,
    3. Chory J.,
    4. Weigel D.
    (2000). Regulation of auxin response by the protein kinase PINOID. Cell 100: 469–478.
    OpenUrlCrossRefPubMed
  22. ↵
    1. Clough S.J.,
    2. Bent A.F.
    (1998). Floral dip: A simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J. 16: 735–743.
    OpenUrlCrossRefPubMed
  23. ↵
    1. Cobbett C.S.,
    2. May M.J.,
    3. Howden R.,
    4. Rolls B.
    (1998). The glutathione-deficient, cadmium-sensitive mutant, cad2-1, of Arabidopsis thaliana is deficient in gamma-glutamylcysteine synthetase. Plant J. 16: 73–78.
    OpenUrlCrossRefPubMed
  24. ↵
    1. Conrad M.,
    2. et al
    . (2004). Essential role for mitochondrial thioredoxin reductase in hematopoiesis, heart development, and heart function. Mol. Cell. Biol. 24: 9414–9423.
    OpenUrlAbstract/FREE Full Text
  25. ↵
    1. Cutler S.R.,
    2. Ehrhardt D.W.,
    3. Griffitts J.S.,
    4. Somerville C.R.
    (2000). Random GFP:cDNA fusions enable visualization of subcellular structures in cells of Arabidopsis at a high frequency. Proc. Natl. Acad. Sci. USA 97: 3718–3723.
    OpenUrlAbstract/FREE Full Text
  26. ↵
    1. Edlund A.,
    2. Eklof S.,
    3. Sundberg B.,
    4. Moritz T.,
    5. Sandberg G.
    (1995). A microscale technique for gas chromatography-mass spectrometry measurements of picogram amounts of indole-3-acetic acid in plant tissues. Plant Physiol. 108: 1043–1047.
    OpenUrlAbstract
  27. ↵
    1. Foyer C.H.,
    2. Noctor G.
    (2005). Redox homeostasis and antioxidant signaling: A metabolic interface between stress perception and physiological responses. Plant Cell 17: 1866–1875.
    OpenUrlFREE Full Text
  28. ↵
    1. Frendo P.,
    2. Harrison J.,
    3. Norman C.,
    4. Hernández Jiménez M.J.,
    5. Van de Sype G.,
    6. Gilabert A.,
    7. Puppo A.
    (2005). Glutathione and homoglutathione play a critical role in the nodulation process of Medicago truncatula. Mol. Plant Microbe Interact. 18: 254–259.
    OpenUrlCrossRefPubMed
  29. ↵
    1. Gälweiler L.,
    2. Guan C.,
    3. Müller A.,
    4. Wisman E.,
    5. Mendgen K.,
    6. Yephremov A.,
    7. Palme K.
    (1998). Regulation of polar auxin transport by AtPIN1 in Arabidopsis vascular tissue. Science 282: 2226–2230.
    OpenUrlAbstract/FREE Full Text
  30. ↵
    1. Gelhaye E.,
    2. Rouhier N.,
    3. Jacquot J.P.
    (2003). Evidence for a subgroup of thioredoxin h that requires GSH/Grx for its reduction. FEBS Lett. 555: 443–448.
    OpenUrlCrossRefPubMed
  31. ↵
    1. Gil P.,
    2. Dewey E.,
    3. Friml J.,
    4. Zhao Y.,
    5. Snowden K.C.,
    6. Putterill J.,
    7. Palme K.,
    8. Estelle M.,
    9. Chory J.
    (2001). BIG: A calossin-like protein required for polar auxin transport in Arabidopsis. Genes Dev. 15: 1985–1997.
    OpenUrlAbstract/FREE Full Text
  32. ↵
    1. Hardtke C.S.,
    2. Ckurshumova W.,
    3. Vidaurre D.P.,
    4. Singh S.A.,
    5. Stamatiou G.,
    6. Tiwari S.B.,
    7. Hagen G.,
    8. Guilfoyle T.J.,
    9. Berleth T.
    (2004). Overlapping and non-redundant functions of the Arabidopsis auxin response factors MONOPTEROS and NONPHOTOTROPIC HYPOCOTYL 4. Development 131: 1089–1100.
    OpenUrlAbstract/FREE Full Text
  33. ↵
    1. Holmgren A.
    (1989). Thioredoxin and glutaredoxin systems. J. Biol. Chem. 264: 13963–13966.
    OpenUrlFREE Full Text
  34. ↵
    1. Howden R.,
    2. Andersen C.R.,
    3. Goldsbrough P.B.,
    4. Cobbett C.S.
    (1995). A cadmium-sensitive, glutathione-deficient mutant of Arabidopsis thaliana. Plant Physiol. 107: 1067–1073.
    OpenUrlAbstract
  35. ↵
    1. Jakupoglu C.,
    2. Przemeck G.K.H.,
    3. Schneider M.,
    4. Moreno S.G.,
    5. Mayr N.,
    6. Hatzopoulos A.K.,
    7. de Angelis M.H.,
    8. Wurst W.,
    9. Bornkamm G.W.,
    10. Brielmeier M.,
    11. Conrad M.
    (2005). Cytoplasmic thioredoxin reductase is essential for embryogenesis but dispensable for cardiac development. Mol. Cell. Biol. 25: 1980–1988.
    OpenUrlAbstract/FREE Full Text
  36. ↵
    1. Kanzok S.M.,
    2. Fechner A.,
    3. Bauer H.,
    4. Ulschmid J.K.,
    5. Müller H.M.,
    6. Botella-Munoz J.,
    7. Schneuwly S.,
    8. Schirmer R.,
    9. Becker K.
    (2001). Substitution of the thioredoxin system for glutathione reductase in Drosophila melanogaster. Science 291: 643–646.
    OpenUrlAbstract/FREE Full Text
  37. ↵
    1. Kanzok S.M.,
    2. Schirmer R.H.,
    3. Turbachova I.,
    4. Iozef R.,
    5. Becker K.
    (2000). The thioredoxin system of the malaria parasite Plasmodium falciparum. Glutathione reduction revisited. J. Biol. Chem. 275: 40180–40186.
    OpenUrlAbstract/FREE Full Text
  38. ↵
    1. Koh C.S.,
    2. Navrot N.,
    3. Didierjean C.,
    4. Rouhier N.,
    5. Hirasawa M.,
    6. Knaff D.B.,
    7. Wingsle G.,
    8. Samian R.,
    9. Jacquot J.,
    10. Corbier C.,
    11. Gelhaye E.
    (2008). An atypical catalytic mechanism involving three cysteines of thioredoxin. J. Biol. Chem. 283: 23062–23072.
    OpenUrlAbstract/FREE Full Text
  39. ↵
    1. Laloi C.,
    2. Mestres-Ortega D.,
    3. Marco Y.,
    4. Meyer Y.,
    5. Reichheld J.P.
    (2004). The Arabidopsis cytosolic thioredoxin h5 gene induction by oxidative stress and its W-box-mediated response to pathogen elicitor. Plant Physiol. 134: 1006–1016.
    OpenUrlAbstract/FREE Full Text
  40. ↵
    1. Laufs P.,
    2. Grandjean O.,
    3. Jonak C.,
    4. Kiêu K.,
    5. Traas J.
    (1998). Cellular parameters of the shoot apical meristem in Arabidopsis. Plant Cell 10: 1375–1390.
    OpenUrlAbstract/FREE Full Text
  41. ↵
    1. Lennartz K.,
    2. Plücken H.,
    3. Seidler A.,
    4. Westhoff P.,
    5. Bechtold N.,
    6. Meierhoff K.
    (2001). HCF164 encodes a thioredoxin-like protein involved in the biogenesis of the cytochrome b(6)f complex in Arabidopsis. Plant Cell 13: 2539–2551.
    OpenUrlAbstract/FREE Full Text
  42. ↵
    1. Li S.,
    2. Lauri A.,
    3. Ziemann M.,
    4. Busch A.,
    5. Bhave M.,
    6. Zachgo S.
    (2009). Nuclear activity of ROXY1, a glutaredoxin interacting with TGA factors, is required for petal development in Arabidopsis thaliana. Plant Cell 21: 429–441.
    OpenUrlAbstract/FREE Full Text
  43. ↵
    1. Li Y.,
    2. Dankher O.P.,
    3. Carreira L.,
    4. Smith A.P.,
    5. Meagher R.B.
    (2006). The shoot-specific expression of gamma-glutamylcysteine synthetase directs the long-distance transport of thiol-peptides to roots conferring tolerance to mercury and arsenic. Plant Physiol. 141: 288–298.
    OpenUrlAbstract/FREE Full Text
  44. ↵
    1. Marty L.,
    2. Siala W.,
    3. Schwarzländer M.,
    4. Fricker M.D.,
    5. Wirtz M.,
    6. Sweetlove L.J.,
    7. Meyer Y.,
    8. Meyer A.J.,
    9. Reichheld J.P.,
    10. Hell R.
    (2009). The NADPH-dependent thioredoxin system constitutes a functional backup for cytosolic glutathione reductase in Arabidopsis. Proc. Natl. Acad. Sci. USA 106: 9109–9114.
    OpenUrlAbstract/FREE Full Text
  45. ↵
    1. May M.J.,
    2. Leaver C.J.
    (1993). Oxidative stimulation of glutathione synthesis in Arabidopsis thaliana suspension cultures. Plant Physiol. 103: 621–627.
    OpenUrlAbstract
  46. ↵
    1. Meyer Y.,
    2. Siala W.,
    3. Bashandy T.,
    4. Riondet C.,
    5. Vignols F.,
    6. Reichheld J.P.
    (2008). Glutaredoxins and thioredoxins in plants. Biochim. Biophys. Acta 1783: 589–600.
    OpenUrlPubMed
  47. ↵
    1. Montrichard F.,
    2. Alkhalfioui F.,
    3. Yano H.,
    4. Vensel W.H.,
    5. Hurkman W.J.,
    6. Buchanan B.B.
    (2009). Thioredoxin targets in plants: The first 30 years. J. Proteomics 72: 452–474.
    OpenUrlCrossRefPubMed
  48. ↵
    1. Noh B.,
    2. Murphy A.S.,
    3. Spalding E.P.
    (2001). Multidrug resistance-like genes of Arabidopsis required for auxin transport and auxin-mediated development. Plant Cell 13: 2441–2454.
    OpenUrlAbstract/FREE Full Text
  49. ↵
    1. Parisy V.,
    2. Poinssot B.,
    3. Owsianowski L.,
    4. Buchala A.,
    5. Glazebrook J.,
    6. Mauch F.
    (2007). Identification of PAD2 as a gamma-glutamylcysteine synthetase highlights the importance of glutathione in disease resistance of Arabidopsis. Plant J. 49: 159–172.
    OpenUrlCrossRefPubMed
  50. ↵
    1. Potters G.,
    2. Pasternak T.P.,
    3. Guisez Y.,
    4. Palme K.J.,
    5. Jansen M.A.K.
    (2007). Stress-induced morphogenic responses: Growing out of trouble?. Trends Pharmacol. Sci. 12: 98–105.
    OpenUrl
  51. ↵
    1. Reichheld J.P.,
    2. Khafif M.,
    3. Riondet C.,
    4. Droux M.,
    5. Bonnard G.,
    6. Meyer Y.
    (2007). Inactivation of thioredoxin reductases reveals a complex interplay between thioredoxin and glutathione pathways in Arabidopsis development. Plant Cell 19: 1851–1865.
    OpenUrlAbstract/FREE Full Text
  52. ↵
    1. Reichheld J.P.,
    2. Meyer E.,
    3. Khafif M.,
    4. Bonnard G.,
    5. Meyer Y.
    (2005). AtNTRB is the major mitochondrial thioredoxin reductase in Arabidopsis thaliana. FEBS Lett. 579: 337–342.
    OpenUrlCrossRefPubMed
  53. ↵
    1. Reinhardt D.,
    2. Mandel T.,
    3. Kuhlemeier C.
    (2000). Auxin regulates the initiation and radial position of plant lateral organs. Plant Cell 12: 507–518.
    OpenUrlAbstract/FREE Full Text
  54. ↵
    1. Ritz D.,
    2. Beckwith J.
    (2002). Redox state of cytoplasmic thioredoxin. Methods Enzymol. 347: 360–370.
    OpenUrlCrossRefPubMed
  55. ↵
    1. Rivas S.,
    2. Rougon-Cardoso A.,
    3. Smoker M.,
    4. Schauser L.,
    5. Yoshioka H.,
    6. Jones J.D.G.
    (2004). CITRX thioredoxin interacts with the tomato Cf-9 resistance protein and negatively regulates defence. EMBO J. 23: 2156–2165.
    OpenUrlAbstract/FREE Full Text
  56. ↵
    1. Rouhier N.,
    2. Lemaire S.D.,
    3. Jacquot J.
    (2008). The role of glutathione in photosynthetic organisms: Emerging functions for glutaredoxins and glutathionylation. Annu. Rev. Plant Biol. 59: 143–166.
    OpenUrlCrossRefPubMed
  57. ↵
    1. Serrato A.J.,
    2. Cejudo F.J.
    (2003). Type-h thioredoxins accumulate in the nucleus of developing wheat seed tissues suffering oxidative stress. Planta 217: 392–399.
    OpenUrlCrossRefPubMed
  58. ↵
    1. Soerensen J.,
    2. Jakupoglu C.,
    3. Beck H.,
    4. Förster H.,
    5. Schmidt J.,
    6. Schmahl W.,
    7. Schweizer U.,
    8. Conrad M.,
    9. Brielmeier M.
    (2008). The role of thioredoxin reductases in brain development. PLoS One 3: 1813.
    OpenUrlCrossRef
  59. ↵
    1. Sweat T.A.,
    2. Wolpert T.J.
    (2007). Thioredoxin h5 is required for victorin sensitivity mediated by a CC-NBS-LRR gene in Arabidopsis. Plant Cell 19: 673–687.
    OpenUrlAbstract/FREE Full Text
  60. ↵
    1. Tada Y.,
    2. Spoel S.H.,
    3. Pajerowska-Mukhtar K.,
    4. Mou Z.,
    5. Song J.,
    6. Wang C.,
    7. Zuo J.,
    8. Dong X.
    (2008). Plant immunity requires conformational charges of NPR1 via S-nitrosylation and thioredoxins. Science 321: 952–956.
    OpenUrlAbstract/FREE Full Text
  61. ↵
    1. Teale W.D.,
    2. Paponov I.A.,
    3. Palme K.
    (2006). Auxin in action: Signalling, transport and the control of plant growth and development. Natl. Rev. 7: 847–859.
    OpenUrlCrossRef
  62. ↵
    1. Trotter E.W.,
    2. Grant C.M.
    (2003). Non-reciprocal regulation of the redox state of the glutathione-glutaredoxin and thioredoxin systems. EMBO Rep. 4: 184–188.
    OpenUrlAbstract
  63. ↵
    1. Trotter E.W.,
    2. Grant C.M.
    (2005). Overlapping roles of the cytoplasmic and mitochondrial redox regulatory systems in the yeast Saccharomyces cerevisiae. Eukaryot. Cell 4: 392–400.
    OpenUrlAbstract/FREE Full Text
  64. ↵
    1. Ulmasov T.,
    2. Murfett J.,
    3. Hagen G.,
    4. Guilfoyle T.J.
    (1997). Aux/IAA proteins repress expression of reporter genes containing natural and highly active synthetic auxin response elements. Plant Cell 9: 1963–1971.
    OpenUrlAbstract/FREE Full Text
  65. ↵
    1. Van Breusegem F.,
    2. Bailey-Serres J.,
    3. Mittler R.
    (2008). Unraveling the tapestry of networks involving reactive oxygen species in plants. Plant Physiol. 147: 978–984.
    OpenUrlFREE Full Text
  66. ↵
    1. Vernoux T.,
    2. Wilson R.C.,
    3. Seeley K.A.,
    4. Reichheld J.P.,
    5. Muroy S.,
    6. Brown S.,
    7. Maughan S.C.,
    8. Cobbett C.S.,
    9. Van Montagu M.,
    10. Inzé D.,
    11. May M.J.,
    12. Sung Z.R.
    (2000). The ROOT MERISTEMLESS1/CADMIUM SENSITIVE2 gene defines a glutathione-dependent pathway involved in initiation and maintenance of cell division during postembryonic root development. Plant Cell 12: 97–110.
    OpenUrlAbstract/FREE Full Text
  67. ↵
    1. Xing S.,
    2. Rosso M.G.,
    3. Zachgo S.
    (2005). ROXY1, a member of the plant glutaredoxin family, is required for petal development in Arabidopsis thaliana. Development 132: 1555–1565.
    OpenUrlAbstract/FREE Full Text
  68. ↵
    1. Xing S.,
    2. Zachgo S.
    (2008). ROXY1 and ROXY2, two Arabidopsis glutaredoxin genes, are required for anther development. Plant J. 53: 790–801.
    OpenUrlCrossRefPubMed
  69. ↵
    1. Yin X.,
    2. et al
    . (2007). Ubiquitin lysine 63 chain forming ligases regulate apical dominance in Arabidopsis. Plant Cell 19: 1898–1911.
    OpenUrlAbstract/FREE Full Text
  70. ↵
    1. Zhao Y.,
    2. Christensen S.K.,
    3. Fankhauser C.,
    4. Cashman J.R.,
    5. Cohen J.D.,
    6. Weigel D.,
    7. Chory J.
    (2001). A role for flavin monooxygenase-like enzymes in auxin biosynthesis. Science 291: 306–309.
    OpenUrlAbstract/FREE Full Text
  71. ↵
    1. Zhong R.,
    2. Taylor J.J.,
    3. Ye Z.H.
    (1997). Disruption of interfascicular fiber differentiation in an Arabidopsis mutant. Plant Cell 9: 2159–2170.
    OpenUrlAbstract/FREE Full Text
Back to top

Table of Contents

Print
Download PDF
Email Article
Citation Tools
Request Permissions
Interplay between the NADP-Linked Thioredoxin and Glutathione Systems in Arabidopsis Auxin Signaling
Talaat Bashandy, Jocelyne Guilleminot, Teva Vernoux, David Caparros-Ruiz, Karin Ljung, Yves Meyer, Jean-Philippe Reichheld
The Plant Cell Feb 2010, 22 (2) 376-391; DOI: 10.1105/tpc.109.071225
del.icio.us logo Digg logo Reddit logo Twitter logo CiteULike logo Facebook logo Google logo Mendeley logo

Jump to section

In this issue

View this article with LENS