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First published online April 7, 2006; 10.1104/pp.106.078469 Plant Physiology 141:423-435 (2006) © 2006 American Society of Plant Biologists
Ascorbate Oxidase-Dependent Changes in the Redox State of the Apoplast Modulate Gene Transcript Accumulation Leading to Modified Hormone Signaling and Orchestration of Defense Processes in Tobacco1,[W]Crop Performance and Improvement Division (C.P., G.K., C.H.F.) and Plant Pathogen Interactions Division (S.J.F.), Rothamsted Research, Harpenden, Herts AL5 2JQ, United Kingdom; and Institute for Research on the Environment and Sustainability, School of Biology and Psychology, Division of Biology, Newcastle University, Newcastle upon Tyne NE1 7RU, United Kingdom (I.H., A.A., T.T., J.B.)
The role of the redox state of the apoplast in hormone responses, signaling cascades, and gene expression was studied in transgenic tobacco (Nicotiana tabacum) plants with modified cell wall-localized ascorbate oxidase (AO). High AO activity specifically decreased the ascorbic acid (AA) content of the apoplast and altered plant growth responses triggered by hormones. Auxin stimulated shoot growth only when the apoplastic AA pool was reduced in wild-type or AO antisense lines. Oxidation of apoplastic AA in AO sense lines was associated with loss of the auxin response, higher mitogen-activated protein kinase activities, and susceptibility to a virulent strain of the pathogen Pseudomonas syringae. The total leaf glutathione pool, the ratio of reduced glutathione to glutathione disulfide, and glutathione reductase activities were similar in the leaves of all lines. However, AO sense leaves exhibited significantly lower dehydroascorbate reductase and ascorbate peroxidase activities than wild-type and antisense leaves. The abundance of mRNAs encoding antioxidant enzymes was similar in all lines. However, the day/night rhythms in the abundance of transcripts encoding the three catalase isoforms were changed in response to the AA content of the apoplast. Other transcripts influenced by AO included photorespiratory genes and a plasma membrane Ca2+ channel-associated gene. We conclude that the redox state of the apoplast modulates plant growth and defense responses by regulating signal transduction cascades and gene expression patterns. Hence, AO activity, which modulates the redox state of the apoplastic AA pool, strongly influences the responses of plant cells to external and internal stimuli.
The concept that plants respond to the imposition of environmental stress by inducing defense pathways and slowing vegetative growth is widely accepted. Although the nature of the mechanisms that control these processes is poorly understood, it is considered to involve the coordinated regulation of plant hormones that modulate DELLA proteins and antioxidant defenses (Fath et al., 2002  ; Achard et al., 2006). Key changes in gene expression are engaged that decrease cell division and elongation growth and enhance pathogen resistance (Knight and Knight, 2001). In this regard, the apoplast and cell wall act as a reservoir of information on the biotic and abiotic environment surrounding the cell as well as a major conduit of information between cells. Similarly, the plasmalemma has major functions in stress perception and the subsequent appropriate control of growth and defense.
Many environmental and metabolic triggers, including pathogen attack, ozone, and physical and chemical assaults, alter the redox state of the apoplast by triggering an oxidative burst at the plasmalemma. Similarly, reactive oxygen species (ROS) produced by NADPH oxidases and other enzymes often act as second messengers in plant growth responses. For example, ROS generated by the RHD2 NADPH oxidase (atrbohC) are required for root hair cell growth in Arabidopsis (Arabidopsis thaliana; Foreman et al., 2003
Whereas much attention has focused on the roles of ROS as signals influencing plant responses to environmental stimuli and developmental cues, relatively little attention has been given to the redox buffering capacity of the apoplast and how it is modulated. Despite the presence of flavonoids and polyamines in the cell wall that can act as antioxidants, the redox buffering capacity of the apoplast is weak (Horemans et al., 2000
Whereas most of the cellular AA pool is localized within the cytoplasm, up to 5% is transported across the plasma membrane and into the apoplast, where it constitutes the major sink for gaseous oxidants such as ozone (Barnes et al., 2002
The AA pool in the apoplast is a key determinant of oxidant signal duration and strength as well as overall redox status of the apoplast (Foyer and Noctor, 2000
Because the redox buffering capacity of the apoplast is always low compared to the cytoplasm, a steep redox gradient will arise across the plasma membrane when the apoplast AA pool is oxidized. We proposed that this gradient could influence gene expression through altered calcium release and modified channel activity (Pignocchi and Foyer, 2003
We have previously shown that the whole leaf AA pool is 90% reduced in all lines used in this study. In contrast, the apoplast AA pool is only about 3% reduced in AO sense lines, 70% reduced in AO antisense lines, and 40% reduced in wild-type plants (Pignocchi et al., 2003 ).
We chose to study the responses of these plants to auxin (naphthylacetic acid [NAA]) and GA3 because of the documented involvement of ROS in auxin responses (Joo et al., 2001
Effects of Altered AO on Leaf MAPK Activities and Responses to Pathogen Infection
The insensitivity to auxin treatment observed in the AO sense lines could be due to constitutive activation of auxin signal transduction pathways via a conserved MAPK signaling cascade, as previously shown (Kovtun et al., 1998
Because MAPK signaling has been shown to be involved in plant resistance to avirulent and virulent strains of Pseudomonas syringae (Desikan et al., 2001 ; Menke et al., 2004), we explored responses to pathogen challenge in the AO transgenic lines versus wild-type plants. Leaves of 7-week-old AO sense, AO antisense, and wild-type tobacco plants were inoculated with a suspension containing an equal volume of either a virulent or an avirulent strain of P. syringae (Fig. 4
). A mock inoculation with phosphate buffer alone was used as a negative control. No lesions were detected up to 48 h after infection (data not shown). At 48 h, lesions appeared on the infected leaves. The lesions were confined to the inoculation area in all lines in response to infection with the avirulent strain. When the virulent strain of P. syringae was used, the lesions in AO sense plants showed a far more diffuse spread in infection compared with wild-type or AO antisense leaves (Fig. 4). The same results were observed in three independent repetitions.
Effects on Antioxidant Enzyme Activities and Leaf Glutathione Content The activities of antioxidant enzymes were measured in leaves of wild-type, two AO sense (Sense 1 and Sense 2), and two AO antisense (Antis 1 and Antis 2) lines (Fig. 5 ). Whereas activities were similar in wild-type and antisense lines (P > 0.05), marked changes were observed in AO sense lines. In AO sense lines, significant (P < 0.01) decreases in DHA reductase (DHAR) and ascorbate peroxidase (APX) activities were recorded. No significant changes (P > 0.05) in glutathione reductase, CAT, or nonspecific peroxidase (guiacol peroxidase [GPOD]) activities were observed (Fig. 5). When glutathione was measured in the same leaves, the total amounts of the glutathione pools and relative amounts of reduced glutathione and glutathione disulfide were similar in all lines (data not shown).
Effects on Transcript Abundance To investigate whether the changes in antioxidant enzyme activities observed in AO sense lines were the result of transcriptional or posttranscriptional regulation, we performed semiquantitative reverse transcription (RT)-PCR on AO sense and antisense plants. Little difference in the abundance of transcripts encoding antioxidant enzymes such as MDHAR, DHAR, APX, and glutathione peroxidase (GPX) were observed (Fig. 6 ). Phe ammonia-lyase (PAL) transcripts were also similar in all lines. However, there was a marked decrease in transcripts encoding the plasmalemma-localized two-pore Ca2+ channel-associated gene, NtTPC1B, in AO sense lines compared to antisense and wild-type lines. The housekeeping genes tubulin A3 and 18S were used as controls.
To study differentially expressed genes in AO transgenic lines on a larger scale, we performed subtractive hybridization screening of one AO sense and one AO antisense line against the wild type. A library comprising more than 300 clones was obtained from the leaves of 4-week-old wild-type, AO sense, and AO antisense plants grown in soil. Leaf samples were harvested 9 h after the beginning of the light period. Over 70 expressed sequence tags (ESTs), which showed high scores when aligned with database sequences, were altered in AO sense and antisense lines with respect to the wild type (Fig. 7A ). Of these, 23% (18 ESTs) were involved in general metabolism, 16% (12 ESTs) constitute regulatory genes, 10% (eight ESTs) were stress related, 5% (four ESTs) were involved in photorespiration, and 4% (three ESTs) were involved in photosynthesis and electron transport (Fig. 7A). Interestingly, Gly decarboxylase transcripts were decreased in AO sense lines, whereas transcripts encoding for a Fd-NADPH reductase were increased (Fig. 7B). Moreover, Gln synthetase (GS-2), glycolate oxidase, and nitrate reductase were more highly expressed in AO antisense lines, whereas others, such as glutathione synthetase, were decreased (Fig. 7C). All transcript shifts indicated in Figure 7, B and C, were confirmed by real-time PCR (data not shown). For a full description of all gene expression changes measured in these experiments, see Supplemental Table I.
Effects on Diurnal Patterns of Transcript Abundance The above experiments indicated that key transcripts associated with photorespiration were modified by altered AO expression in transgenic plants. Because many of these transcripts show a day/night rhythm in transcript abundance, we analyzed by real-time PCR the effect of AO expression on the day/night abundance of transcripts associated with photorespiration, particularly those involved in photorespiratory hydrogen peroxide (H2O2) metabolism (Fig. 8 ). The abundance of actin transcripts was similar in all transgenic lines and was unaffected by the dark/light treatments employed here (data not shown). Actin was therefore used as an internal control. For consistency, primers for glycolate oxidase (U62485), GS-2 (X66940), and Cys synthase (AM087457) were designed on the basis of sequences identified in the subtractive screening. Real-time PCR data for these transcripts confirmed the findings shown in Figure 7, B and C. At the sampling time used for the subtractive screening (9 h in the light period), transcript abundance of glycolate oxidase and GS-2 were increased in AO antisense plants compared to wild type. In contrast, Cys synthase transcripts were found to be decreased in AO sense plants compared to wild type (Fig. 8). CAT 1, CAT 2, and CAT 3 all showed day/night rhythms in transcript abundance that were modified by altered AO expression.
The redox state of the AA pool in the apoplast has the potential to exert a profound influence on cellular redox signaling (Pignocchi and Foyer, 2003 ; Foyer and Noctor, 2005a, 2005b). The experiments discussed here were designed specifically to explore the influence of the redox state of the apoplast as defined by the abundance of the major antioxidant buffer AA on signal transduction and gene expression. We have focused on processes associated with plant growth and defense because apoplastic redox homeostasis is central to the perception and signaling related to environmental and hormonal cues. We have previously shown that the redox state of the AA pool in the apoplast can be modulated independently of the cytoplasmic pool, creating a large redox gradient across the plasma membrane (Pignocchi and Foyer, 2003; Sanmartin et al., 2003). Here, we provide evidence that oxidation of the AA pool in the apoplast specifically modulates signal transduction components, leading to changes in gene expression that alter plant responses to hormone signals and pathogen attack.
The concept that AA and AO regulate plant growth has long been accepted (Chinoy, 1984
The loss of auxin responses in AO sense plants could be explained by three possible mechanisms. First, AO has been shown to catalyze the oxidative decarboxylation of auxin in maize roots (Kerk et al., 2000
Plant growth is finely regulated by interactions between hormones (Steffens et al., 2005
Taken together, the above results suggest that AO-mediated oxidation of the apoplast desensitizes plants to hormone cues. One explanation for this effect is that constitutive AO expression mimics the action of auxin via oxidation of the apoplast. ROS generated by the activity of Rboh-encoded NADPH oxidases participate in a wide range of hormone-induced developmental responses, as well as in plant-pathogen interactions and abiotic stress responses (Torres and Dangl, 2005
The auxin-mediated signal transduction cascade is mediated by a conserved signaling cascade consisting of three protein kinases: MAPK, MAPK kinase (MAPKK), and MAPKK kinase (MAPKKK). Transient increases in protein kinase activity with characteristics of mammalian extracellular signal-regulated kinase-like MAPKs are induced by auxin in Arabidopsis roots (Mockaitis and Howell, 2000
The constitutive activation of the MAPK signaling cascades was coupled to the decreased expression of the calcium channel NtTPC1B in AO sense plants. NtTPC1B encodes a voltage-operated calcium channel, which is the major Ca2+-permeable channel activated by H2O2 (Kadota et al., 2005
We have previously shown that light-mediated control of AO expression is modified by oxidation of AA in the apoplast. Positive regulation of native AO transcript abundance by light in tobacco was completely lost when apoplastic AA was highly oxidized in transgenic plants (Pignocchi et al., 2003
Glycolate oxidase produces H2O2 at very high rates in the peroxisomes of C3 plants as a result of photorespiratory carbon flow (Noctor et al., 2002
The results presented in this study allow a new perspective of cellular redox metabolism and homeostasis in which the extracellular redox environment has a marked influence on cellular signaling cascades that influence development, antioxidant defense, and biotic interactions. To date, only apoplastic NAPDH oxidase-catalyzed production of ROS has been considered in terms of a broad role in signaling responses to infections, the abiotic environment, programmed cell death, and developmental cues (Torres and Dangl, 2005
These results emphasize the crucial importance of redox homeostasis in the apoplast in controlling signal transduction processes. Regulation of the AA pool in the apoplast could be used to modulate cross talk between different defense and growth pathways in a similar manner to that already described for ROS (Torres and Dangl, 2005
Materials
Wild-type and transgenic (T2 and T3 generation) tobacco (Nicotiana tabacum; SR1 ecotype) plants expressing pumpkin (Cucurbita maxima) AO in the sense orientation (AO sense lines P221 and P372) and partial tobacco AO in antisense orientation (AO antisense lines T161 and T271) were grown in pots (containing 3 dm3 of John Innes no. 2 compost) in controlled environment chambers at 22°C with a 16-h photoperiod and a photosynthetic photon flux density of 250 µmol m–2 s–1 at plant height or on agar plates, as described in Pignocchi et al. (2003) Cultures of Pseudomonas syringae employed in this study were tomato (Lycopersicon esculentum) pv DC3000 pLAFR (virulent strain) and pv pLAFRavrRpt2 (avirulent strain).
Ten-day-old seedlings, raised in petri dishes containing Murashige and Skoog agar culture medium (Sigma Chemicals) were sprayed to run off (using an aerosol spray bottle supplied by Nalgene) with 0.5 µM–1 NAA and 10–4 M GA3. Biomass was quantified after a further 10 d.
Frozen tissue was ground to powder under N2. One hundred-milligram samples in Eppendorf tubes were homogenized for 1 min in 0.1 mL of lysis buffer containing 20 mM HEPES-KOH, pH 7.6, 100 mM KCl, 1.5 mM EGTA, 1.0 mM EDTA, 10% glycerol, 20 mM
MAPK assays were performed in 25 µL of K buffer (25 mM HEPES, pH 7.9, 5 mM MgCl2, 0.1% 2-mercaptoethanol, 0.1 mM EDTA) with added 0.2 mg mL–1 substrate protein, MgCl2 (30 mM final), ATP (100 µM final), and 2.5 µCi [
Bacterial cultures were grown overnight at 28°C with centrifuging in King's B medium (peptone 20 g L–1; glycerol 1% v/v, K2HPO4 1.5 g L–1, pH 7.2) supplemented with tetracycline (10 µg mL–1) and rifampicin (100 µg mL–1). Cells from overnight cultures were washed and then resuspended in 10 mM MgCl2 to obtain a cell density of 0.5 x 10–6 cells mL–1; 0.1 mL of cell suspension was used for each inoculation. Inoculations were performed by infiltration into the underside of the leaves, using a 1-mL syringe without a needle. Three repetitions for each infection were performed. Inoculated plants were incubated under growth conditions for 48 h. Leaf discs of the infected leaves were harvested 2, 4, 6, 8, and 48 h after inoculation.
Glutathione was measured as described by Noctor and Foyer (1998)
CAT activity was determined in a reaction mixture containing 66 mM potassium phosphate (pH 7), 0.7 mM H2O2, and 2% (v/v) extract. The reaction was followed at 240 nm, employing an extinction coefficient for H2O2 of 0.039 mM–1 cm–1. Control assays for each replicate were performed in the presence of aminotriazole. GPOD activity was determined in a reaction mixture containing 66 mM potassium phosphate (pH 6.1), 8 mM Guiacol, 10% (v/v) extract, and 2 mM H2O2. GPOD activity was determined as the rate of formation of tetraguiacol at 470 nm (extinction coefficient of 26.6 mM–1 cm–1). Glutathione reductase activity was measured in a reaction mixture containing 66 mM potassium phosphate buffer (pH 7.8), 150 µM NADPH, 500 µM GSSG, and 5% (v/v) extract. The reaction was followed at 340 nm (extinction coefficient for NADPH of 6.22 mM–1 cm–1). Controls were performed in the absence of GSSG to assess nonenzymatic oxidation of NADPH. DHAR activity was measured in a reaction mixture containing 66 mM potassium phosphate buffer (pH 6.5), 3 mM GSH, 250 µM DHA, and 5% (v/v) extract. Reaction was followed at 265 nm (extinction coefficient for AA of 14 mM–1 cm–1). Two control reactions were performed: one without GSH to assess non-GSH-dependent production of AA, and the other one, without extract, to assess non-DHAR production of AA.
APX activity was determined by monitoring oxidation of AA at 290 nm ( All enzyme activities were measured using a Pye-Unicam SP8700 UV/visible spectrophotometer (Pye-Unicam). All reactions were developed at 25°C in 1-mL quartz cuvettes. Data were expressed as specific activity, with protein content determined using the method of Bradford.
Total RNA was extracted by using the RNeasy plant mini kit (Qiagen), according to the supplier's recommendation. Residual DNA was removed with DNase I, amp grade (Gibco-BRL). The absence of DNA contamination in the samples was confirmed by a saturating PCR of 40 cycles using actin-specific (X63603) primers (5'-CGCGAAAAGATGACTCAAATC-3' and 5'-AGATCCTTTCTGATATCCACG-3'), which give a 687-bp product with genomic DNA and a 533-bp product with cDNA. One microgram of total RNA was reverse transcribed using 0.5 µg oligo(dT)12–18 (Gibco-BRL) and 200 units SuperScript II (Gibco-BRL), following the supplier's recommendation. cDNA samples were standardized by PCR for actin and tubulin content using the gene-specific primers. On the basis of the published sequences, the following gene-specific primers were designed and used for amplification: MDHAR, 5'-GACAGAAACTTCAAATAGCCG-3' and 5'-GAACATGTTGATCATTCTCGC-3'; DHAR (AY074787), 5'-ATCTGTGTCAAGGCTGCTG-3' and 5'-ACTTCCTGCGAAACAACGG-3'; cAPX (U15933), 5'-CTGGAGGACCTGATGTTC-3' and 5'-CGTCTAATAACAGCTGCC-3'; GPX (AB041518), 5'-CCAATCTAGCAAGCCTCAA-3' and 5'-ATGCAGACAAATCCAGAGC-3'; PAL (X78269), 5'-GCGATAGACTTGAGGCATT-3' and 5'-GATCCTGTTGTTTGGAGAACC-3'; NtTPC1B (AB124647), 5'-CCAACGGAGAATGGATTCG-3' and 5'-CAGCATGGAGAAAGGAGCA-3'; tubulin A3 (AJ421413), 5'-TCCTCATATGCTCCtGTC-3' and 5'-AGCAGACAAGCATTCTAC-3'; and 18S, 5'-GACGAACAACTGCGAAAG-3' and 5'-CATCTAAGGGCATCACAG-3'. For semiquantitative RT-PCR, the cycle number was kept within the linear range (30 cycles) and the conditions were 3 min at 94°C, a cycle of 45 s at 94°C, 30 s at 52°C, 45 s at 72°C, followed by 10 min at 72°C, using 0.5 µL of the RT reaction and 0.2 µM of each oligonucleotide primer in a total volume of 25 µL. The identity of the PCR products was verified by single-strand sequencing (ABI PRISM, 310 genetic analyzer; Perkin-Elmer). RT-PCR products were loaded on 2% (w/v) agarose gel containing 0.5 µg mL–1 ethidium bromide and the band intensities quantified with the Eagle Eye II (Stratagene).
Total RNA was extracted using Tri reagent (Helena Biosciences). mRNA was isolated from total RNA using the PolyAtract kit (Promega). Subtractive hybridization was carried out using the PCR-select cDNA subtraction kit (BD Biosciences, CLONTECH), following the manufacturer's instructions. Forward and reverse subtractions were carried out using wild-type mRNA as the tester and each transformant line as the driver and vice versa to recover up- and down-regulated genes. The amplified, differentially expressed cDNAs were cloned using a TOPO-TA cloning system (Invitrogen). Plasmids bearing DNA insertions were extracted using a kit (Qiagen) and sequenced using M13 forward and reverse promoters (Macrogen). Sequence identification was performed using a BLAST search of the National Center for Biotechnology Information (NCBI) gene bank data resource.
Leaves were harvested from 5-week-old plants throughout the day/night cycle at 6, 9, and 15 h (day), and 18 and 21 h (night). Leaf samples were frozen in liquid N2, homogenized into a fine powder, and RNA was extracted from each sample and cDNA synthesized as described by Pastori et al. (2003)
All datasets were subjected to ANOVA and significant (P < 0.05) differences between individual means established using a Student's t test employing GENSTAT 5 (Payne et al., 1993 Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers U60495, M14419, U62485, AM087457, AJ421411, X66940, U93244, U07627, and Z36977.
The authors are grateful to Prof. Muneharu Esaka (Hiroshima University, Higashi-Hiroshima, Japan) for kindly providing the AO cDNAs. We also thank David D'Haese and Anne Borland (Institute for Research on Environment and Sustainability, Newcastle University, UK) for constructive discussions. Received February 2, 2006; returned for revision March 24, 2006; accepted March 27, 2006.
1 This work was supported by a combination of funding from the Biotechnology and Biological Sciences Research Council, the National Environmental Research Council, and the European Union (Marie Curie training site HPMT–CT–2001–00219). 
2 Present address: The Sainsbury Laboratory, John Innes Centre, Norwich, Research Park NR4 7UH, UK.
3 Present address: Departament de Fisiologia Vegetal, Universitat de Barcelona, Avenida Diagona 645, 08028 Barcelona, Spain. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Christine H. Foyer (christine.foyer{at}bbsrc.ac.uk).
[W] The online version of this article contains Web-only data. Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.106.078469. * Corresponding author; e-mail christine.foyer{at}bbsrc.ac.uk; fax 44–1582–763010.
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ABSTRACT
RESULTS
), AO sense (
), and AO antisense (
) plants.
-glycerophosphate, 50 mM NaF, 10 mM 
-32P]ATP. Substrates used were bovine MBP (Sigma-Aldrich). Five micrograms of protein extract were used for each assay. Reactions were incubated for 30 min at 30°C, then terminated by the addition of 5x SDS-PAGE loading buffer, and resolved by SDS-PAGE. Unincorporated [
= 2.88 mM–1 cm–1), as described by Nakano and Asada (1981)
