TY - JOUR
AU - Rolfe, Stephen A.
AB - Abstract Chlorophyll fluorescence imaging provides a non-invasive and non-destructive means with which to measure photosynthesis. This technique has been used, in combination with 14CO2 feeding, to study the spatial and temporal changes in source–sink relationships which occur in mechanically wounded leaves of Arabidopsis thaliana. Twenty-four hours after wounding, cells proximal to the wound margin showed a rapid induction of ΦII upon illumination (a measure of the efficiency of photosystem II photochemistry) whilst cells more distal to the wound margin exhibited a much slower induction of ΦII and a large, transient increase in NPQ (a measure of the rate constant for non-photochemical energy dissipation within the light-harvesting antenna). These results are indicative of an increase in sink strength in the vicinity of the wound and this was confirmed by the retention of 14C photosynthate in this region. It has been hypothesized that wound-induced cell wall (apoplastic) invertase (cwINV) activity plays a central role in generating localized increases in sink strength in stressed plant tissue and that hexose sugars generated by the sucrolytic activity of cwINV may act as a signal regulating gene expression. Enzyme activity measurements, quantitative RT-PCR, and T-DNA insertional mutagenesis have been used to determine that expression of AtcwINV1 is responsible for all induced cwINV activity in mechanically wounded leaves. Whilst inactivation of this gene abolished wound-induced cwINV activity, it did not affect localized alterations in source–sink relationships of wounded leaves or wound-regulated gene expression. The signals that may regulate source–sink relationships and signalling in wounded leaves are discussed. Arabidopsis thaliana, chlorophyll fluorescence imaging, invertase, photosynthesis, source–sink relationships, sugar signalling, wounding Introduction Wounding is a common event in the life time of a plant and elicits a spatially and temporally complex series of responses. Mechanical wounding and insect herbivory, causing crush injuries or tears, result in localized cell death, loss of water and solutes from exposed surfaces, provide a point of entry for opportunistic bacterial and fungal pathogens, and can disrupt the vascular system. Many signalling pathways are activated either directly by the wounding process or subsequent to it. These include those sensing touch, cell wall fragments, dehydration and, in the case of insect herbivores, insect-derived elicitors, and they involve numerous signalling molecules (e.g. jasmonates, NO, ethylene, abscisic acid, systemin, reactive oxygen species, and possibly electrical or hydraulic signals); (Howe, 2004; León et al., 2001; Stratmann, 2003; Wendehenne et al., 2004; Orozco-Cárdenas et al., 2001). Mechanical wounding leads to the activation of genes involved in responses to water stress, cellular repair and metabolism, and defence against pathogens (Reymond and Farmer, 1998; Reymond et al., 2000; Jacobs et al., 2003; Delessert et al., 2004). Delessert et al. (2004) found that, in leaves of Arabidopsis thaliana, many genes encoding signal transduction and regulatory components are induced rapidly by mechanical wounding, both locally within the wounded leaf and systemically throughout the plant. In contrast, genes encoding repair-associated and metabolic enzymes tend to be induced more slowly and induction is restricted to the wound itself and immediately adjacent regions, illustrating the temporal and spatial complexity of the wound response. The activation of defence and repair mechanisms places a high metabolic demand upon the wounded region: carbon skeletons are required for the synthesis of new molecules and an energy source is required to fuel the biosynthetic reactions. Respiration is increased in wounded tissues, and measurements of 13C partitioning in developing leaves of hybrid poplar (Populus deltoids×P. nigra) have demonstrated that sink strength is increased following insect herbivory or treatment with jasmonic acid (Arnold and Schultz, 2002). It is well established that mechanical wounding leads to an induction of cell wall associated (cwINV or apoplastic) invertase gene expression and activity (Sturm and Chrispeels, 1990; Zhang et al., 1996; Ohyama et al., 1998) and it has been proposed that the hydrolysis of sucrose by cwINV generates hexoses which fulfil the energy and carbon requirements of the wounded tissue (Sturm and Chrispeels, 1990). High cwINV activity is a characteristic of sink tissues, hydrolysing sucrose to drive phloem unloading (reviewed in Roitsch and Gonzaléz, 2004; Roitsch, 1999). In source leaves, cwINV activity is lower, although ectopic expression of cwINV restores many characteristics of sink tissue (von Schaewen et al., 1990; Sonnewald et al., 1991; Balibrea Lara et al., 2004). An alteration in the source–sink status of cells in the vicinity of wounds is also supported by the observation that the expression of AtSUC3 (a sucrose transporter) and AtSTP4 (a monosaccharide transporter) is induced locally by wounding (Truernit et al., 1996; Meyer et al., 2004), whereas their expression is normally restricted to sink cells and tissues. The hexose sugars generated by cwINV activity have also been proposed to play an important signalling role in leaves subject to biotic and abiotic stresses (Tang et al., 1996a; Ehness et al., 1997; Chou et al., 2000; Roitsch, 1999; Roitsch et al., 2003; Roitsch and Gonzaléz, 2004). Plants contain multiple sugar signalling pathways which can activate defence-related gene expression and repress photosynthetic gene expression and there is extensive crosstalk between the sugar and wound/stress signalling pathways (reviewed in Gibson, 2004; Halford and Paul, 2003; Smeekens, 2000). Whilst high levels of cwINV expression in transgenic plants can generate sufficient hexose sugars to trigger these responses (Herbers et al., 1996; Herbers and Sonnewald, 1998), these genes are also directly regulated by other signalling pathways, hence the contribution of sugar signalling to the regulation of gene expression in wounded leaves is unclear. High resolution chlorophyll fluorescence imaging and radioactive feeding with 14CO2 was used to examine the spatial and temporal alterations in source–sink relationships which occur within mechanically wounded leaves of Arabidopsis thaliana. The expression of the multigene family encoding cwINV in A. thaliana has been characterized in wounded leaves, and a T-DNA insertion mutant has been used to determine that AtcwINV1 was responsible for wound-induced cwINV activity. The impact of inactivating this gene on source–sink relationships and wound-regulated gene expression is explored. Materials and methods Growth of plant material Seeds of Arabidopsis thaliana (ecotype Col-0) were sown in a peat-based compost (M3 compost, Levington Horticulture Ltd., Ipswich, UK) and placed at 4 °C for 3 d to ensure even germination. The seeds were transferred to a growth room with a daytime temperature of 22 °C and a night-time temperature of 18 °C. The growth irradiance was 160 μmol m−2 s−1, provided by fluorescent lights (Fluotone Reflex, Philips Lighting UK, Guildford, UK) with a 10 h photoperiod. After 10 d seedlings were transplanted to 6×6 cm pots. All experiments were performed on the fully-expanded leaves of 6-week-old plants. Leaves were mechanically wounded by placing the plungers of 1 ml disposable syringes (Plastipak, Becton Dickinson, Madrid, Spain) on either side of the leaf and pressing gently together, resulting in 10–12 crush injuries where the serrations on the handles crossed. The wounds produced by this procedure were distributed across the leaf surface over an area of 0.8 cm2. Where a more precisely localized method of wounding was required, crush wounds were produced by pressing the end of a cocktail stick onto the leaf which was supported on a piece of plastic. Identification of T-DNA insertion mutants in AtcwINV1 The nomenclature of Sherson et al. (2003) has been adopted for naming cwINV genes of A. thaliana. A T-DNA insertion in the AtcwINV1 gene (At3g13790) was identified in the Syngenta Arabidopsis Insertion Library (SAIL) database (Sessions et al., 2002). The insertion event occurred at position 80 in the coding region of the gene of line 258_A01. Plants homozygous for the T-DNA insertion were identified by PCR using the primer pair AtBFRUCT1F (5′-CTCCACGCCTCGTGTATGTCCGGATGGT-3′) and AtBFRUCT1R (5′-GTACAGTTGTATGCTGACATATATTGGT-3′) which amplify the intact AtcwINV1 coding region, and AtBFRUCT1R and the T-DNA left border primer LB1 (5′-GCCTTTTCAGAAATGGATAAATAGCCTTGCTTCC-3′) which only produce an amplification product when the T-DNA insert is present. Plants containing the T-DNA were back-crossed to wild-type Col-0 twice, selfed, and homozygous plants re-isolated (line ΔAtcwINV1). Quantitative imaging of chlorophyll fluorescence High resolution images of chlorophyll fluorescence were captured using a microscope-based system as described in Rolfe and Scholes (2002). The nomenclature of images of chlorophyll fluorescence captured at different points during the experiment, and the subsequent calculations performed to calculate images of various photosynthetic parameters are described in Maxwell and Johnson (2000) and Rolfe and Scholes (1995). All images were taken using an UPlanFl ×4 objective (Olympus Optical Company, London) and the output from the camera was binned 2×2 providing a field of view 3.72 by 2.96 mm with 500 by 400 pixel resolution. Plants were placed in darkness at the end of the normal photoperiod and attached leaves imaged 14–16 h later. An image of Fo was captured at the beginning of the experiment at a photon irradiance of 1.2 μmol m−2 s−1 and an image of Fm captured 40 s later using a pulsed saturating photon irradiance of 1800 μmol m−2 s−1. (Although this irradiance is not completely saturating, independent experiments indicated that values of ΦII measured were within 5% of the true values under the conditions examined.) The saturating pulse of light was 1.2 s long and the Fm image was captured 800 ms after the onset of illumination. The leaf was then illuminated with a continuous actinic photon irradiance of 80 μmol m−2 s−1 for 10 min until photosynthetic induction was complete and steady-state photosynthesis achieved. Images of chlorophyll fluorescence under actinic light (Ft) and saturating light (F′m) were taken 2 s after the actinic light was first turned on and at 20 s intervals thereafter throughout induction. Images of the photosynthetic parameters Fv/Fm, ΦII, and NPQ were calculated using Image Pro Plus 4.1 (Media Cybernetics, USA). Integrated images of photosystem II (PSII) photochemistry were calculated by summing images of ΦII captured between 22 s and 222 s after the onset of actinic illumination by which time ΦII had reached 98% of the steady-state value. Values were extracted from the images using the area selection tools of Image Pro Plus. To extract values from regions at defined distances from the wound margin, a binary threshold was applied to the unprocessed Fm image to define regions which lacked chlorophyll. The outline of this region was smoothed by applying sequential binary erosions and dilations. A series of contours were then defined at ∼80 μm intervals from the wound margin by applying a 10 pixel binary dilation and subtracting the previous undilated image. These binary images were used to create objects which defined the regions from which values were extracted from processed images of chlorophyll fluorescence parameters. Imaging of each treatment was replicated 4–5 times and representative images are shown. 14CO2 labelling Whole A. thaliana plants (wild type or ΔAtcwINV1) were placed in an open 40 l Perspex container in a fume cupboard at the start of the normal photoperiod and illuminated for 2 h at 250 μmol m−2 s−1. Four leaves were wounded on each plant 1 h or 24 h prior to radiolabelling. The container was then sealed and 14CO2 generated by adding 2 M lactic acid to 1 ml of 750 mM NaHCO3 and 18 μl 2 GBq mmol−1 NaH14CO3 (Amersham Biosciences, Chalfont St Giles, UK) producing an atmosphere containing 1000 μmol mol−1 CO2 with a specific activity of 103 MBq 14C μmol−1 CO2. Air was circulated by a small fan and after 3 min feeding the chamber was opened. Leaves were detached immediately, 3 h or 24 h after feeding, and were frozen in liquid nitrogen vapour and then freeze-dried for 72 h. Radiolabel was then localized by autoradiography (Kodak Biomax film, Sigma-Aldrich Company Ltd., Gillingham, UK) and quantified in transects 8 mm long and 0.8 mm wide using a μImager radioactive imaging system (Biospace, Paris, France). Staining for active oxygen The presence of hydrogen peroxide was used as an indicator of reactive O2 species and stained using 3, 3′-diaminobenzidine (Sigma) (DAB)-HCl, pH 3.8 according to Thordal-Christensen et al. (1997). DAB solution was transpired into mature leaves of wounded and control plants, 24 h and 48 h after wounding, by excising the petiole under water and standing the leaf upright in a tube containing 1 mg ml−1 DAB. Dark brown staining of the basal vascular system became apparent after 1 h; after 3 h leaves were cleared in 95% boiling ethanol for 10 min and photographed. Extraction and assay of soluble and cwINV activity Invertase extractions and assays were performed using a modification of the procedure described in Tang et al. (1996b). For large-scale assays, leaf tissue was excised 1, 3, 6, 10, 24, and 48 h after wounding and weighed. Samples were placed in 2 ml Micro-tubes (Sarstedt, Nümbrecht, Germany), immediately frozen in liquid nitrogen and stored at −80 °C. Soluble invertase activity was extracted by adding 1 ml ice-cold extraction buffer (20 mM sodium phosphate pH 6.5, 1 mM EDTA, 1 mM dithiothreitol, 1 mM benzamidine, and 0.1 mM phenyl methyl sulphonyl fluoride) and homogenized with a stainless steel ball bearing for 5 min in a 8000 M Mixer Mill (Glen Creston Ltd, Stanmore, Middlesex, UK). The ball bearing was removed and the sample centrifuged at 26 000 g for 10 min at 4 °C. The supernatant contained soluble invertase activity whilst the pellet, containing cwINV activity, was washed with 1 ml extraction buffer then 1 ml extraction buffer containing 50 mM NaCl. For small-scale assays 24 h after wounding, two sampling methods were used; either a single disc (0.16 cm2) was excised using a hole punch, or ten discs (each 0.0113 cm2) were excised using a 1.2 mm diameter micropunch (1.2 mm diameter Harris Micropunch, Fisher Scientific Ltd, Loughborough, UK). In either case, discs were frozen immediately in liquid nitrogen in 0.5 ml Eppendorf tubes and extractions performed as above except the buffer volumes were reduced to 0.2 ml. Soluble invertase activity was assayed by incubating aliquots of the extract in 200 mM sodium acetate buffer pH 5.5 containing 100 mM sucrose at 37 °C for 30 min. Reducing sugars produced were assayed using 3,5-dinitrosalicylic acid (DNSA) according to the method of Arnold (1965). cwINV activity was assayed by resuspending the pellet in 250 μl (large-scale assay) or 50 μl (small-scale assay) 200 mM sodium acetate buffer pH 4.5, 100 mM sucrose and incubating at 37 °C for 45 min with constant agitation. The reaction was stopped by boiling the sample for 5 min then the cell wall material was removed by centrifugation at 26 000 g for 10 min at 4 °C. Reducing sugars in the supernatant were quantified using DNSA (large scale) or an enzyme-linked assay for glucose (small scale) (Scholes et al., 1994). The enzyme-linked assay for glucose was performed using 10 μl of the supernatant in a total volume of 100 μl in a black 96-well microtitre plate. The reduction of NAD+ to NADH by glucose-6-phosphate dehydrogenase was monitored by following the increase in fluorescence at 440 nm using an excitation wavelength of 340 nm in a Fluostar Optima plate reader fluorimeter (BMG LABTECH GmbH, Offenburg, Germany). Extraction of RNA and cDNA synthesis One hundred mg of leaf tissue was taken from control and wounded plants 1, 3, 6, 10, 12, 24, and 48 h after wounding and immediately frozen in liquid nitrogen. RNA was extracted by homogenizing in RNAWIZ [Ambion (Europe) Ltd., Huntingdon, Cambridgeshire, UK] with a stainless steel ball bearing in an 8000 M Mixer Mill (Glen Creston Ltd.) for 8 min. The ball bearing was removed and 200 μl of chloroform was added. Samples were mixed, incubated at room temperature for 10 min and centrifuged at 10 000 g for 15 min at 4 °C. The upper aqueous phase was transferred to a new tube and 500 μl RNase-free water (Ambion) was added, followed by 1 ml propan-2-ol. Samples were mixed, incubated at room temperature for 10 min, and centrifuged at 10 000 g for 15 min at 4 °C. The pellet was washed with 1 ml ice-cold 70% (v/v) ethanol and centrifuged at 10 000 g for 5 min at 4 °C; the supernatant was discarded and the pellet was air dried and resuspended in 40 μl RNase-free water. RNA was quantified by UV spectroscopy. Residual DNA was removed by treating the samples with DNase I according to the manufacturer's instructions (DNA-free, Ambion). To ensure that all DNA contamination had been removed, a PCR reaction with primers amplifying a small nuclear ribonucleoprotein was performed. For first-strand cDNA synthesis, 4 μg RNA and 1 μl oligo(dT)15 primer (Promega, Southampton, UK) were heated at 70 °C for 5 min to denature the RNA and samples were then immediately placed on ice. The following reagents were then added to give a final volume of 20 μl: 4 μl 5× first-strand synthesis buffer (Invitrogen, Paisley, UK), 2 μl 10 mM dNTPs (Promega), 2 μl 100 mM dithiothreitol (Invitrogen), 0.5 μl RNasin (Promega). Samples were heated to 42 °C for 2 min, 200 U of SuperScript II Reverse Transcriptase (Invitrogen) were added, and samples incubated at 42 °C for 1 h. Synthesized cDNA was purified using a Qiaquick PCR purification kit (Qiagen, Crawley, West Sussex, UK). cDNA was eluted in 100 μl elution buffer and stored at −80 °C. Quantification of gene expression by RT-PCR Gene expression was determined by quantitative RT-PCR (qRT-PCR). Forward and reverse primers were designed which specifically amplified 50–150 bp regions of each target gene at a similar efficiency to each other. The specificity of each primer pair was determined by comparison against the Arabidopsis genome using BLAST (Altschul et al., 1990) and confirmed by PCR using genomic DNA as a template. Target genes and primer pairs are shown in Supplementary Tables 1 and 2 at JXB online. The small nuclear ribonucleoprotein (SnRNP D1: At3g07590) was selected as a control gene as its expression did not alter in response to wounding and the amplicon amplified at the same efficiency as the other target genes. Each RT-PCR reaction consisted of 3.4 μl cDNA, 8.5 μl SYBR Green PCR Master Mix (Applied Biosystems, Warrington, UK) and 5.1 μl primer mix (5 μl 100 μM forward and 5 μl 100 μM reverse primer, 290 μl RNase-free water). Reactions were prepared in a 96-well optical reaction plate (Applied Biosystems) and cDNA amplification performed using an ABI Prism 7700 Sequence Detector (Applied Biosystems) with the following cycling conditions: 50 °C (2 min), 95 °C (10 min) followed by 40 cycles of 95 °C (15 s) and 60 °C (1 min). Fluorescence and Ct values were measured using Sequence Detector version 1.7 (Applied Biosystems). The Ct value for each reaction was compared with a standard curve created by amplifying A. thaliana genomic DNA under the same conditions. Results were corrected for variations in cDNA input by comparison with SnRNP D1 and then converted into a number of genome-equivalent molecules per μg input RNA (assuming a genome size of 180 MBp). Statistical analyses Statistical analyses were performed using Minitab ver 13.1 (Minitab Ltd., Coventry, UK). Results Wound-induced invertase activity and gene expression Leaves of A. thaliana contain both soluble, vacuolar (vacINV), and cell wall associated, apoplastic (cwINV) acid invertase activity. When fully-expanded leaves were mechanically wounded uniformly across the leaf surface, there was no change in vacINV activity (Fig. 1A) whereas cwINV activity increased significantly (Fig. 1B). A 2-fold stimulation in cwINV activity compared with unwounded control leaves was first detected 3 h after wounding which increased to 3-fold from 24 h after wounding. To determine whether this increase in cwINV activity occurred in the immediate vicinity of the wound or whether induction was more widespread, a localized series of 10–12 crush injuries were made on one side of the leaf lamina (Fig. 2A). After 24 h, leaf discs were taken at different points from the wounded leaves and corresponding regions of control leaves and assayed for cwINV activity. cwINV activity was stimulated (2.3-fold) in regions containing the wound sites (P <0.05, One-way ANOVA) compared to equivalent regions of unwounded leaves (Fig. 2B). In addition there was a small (1.3-fold), but significant, increase in cwINV activity when all unwounded regions of the wounded leaf were compared with all equivalent regions of control leaves (P <0.005, One-way ANOVA). Fig. 1. View largeDownload slide Vacuolar (A) and cell-wall associated (B) invertase activity in fully-expanded leaves of A. thaliana. Extracts were prepared from unwounded (closed symbols) or uniformly mechanically wounded (open symbols) of wild-type (circles) and ΔAtcwINV1 (squares) plants. Results are the mean ±SE of four replicates. An asterisk indicates a significant difference (t-test, * P <0.05, ** P <0.01, *** P <0.001) between wounded and unwounded measurements. Fig. 1. View largeDownload slide Vacuolar (A) and cell-wall associated (B) invertase activity in fully-expanded leaves of A. thaliana. Extracts were prepared from unwounded (closed symbols) or uniformly mechanically wounded (open symbols) of wild-type (circles) and ΔAtcwINV1 (squares) plants. Results are the mean ±SE of four replicates. An asterisk indicates a significant difference (t-test, * P <0.05, ** P <0.01, *** P <0.001) between wounded and unwounded measurements. Fig. 2. View largeDownload slide Localized changes in cell-wall associated invertase activity 24 h after mechanically wounding leaves of wild-type A. thaliana. A series of 10–12 mechanical wounds were made in a small region of a fully-expanded leaf (A). Sampled regions are indicated by dotted circles. Scale bar=0.5 cm. cwINV activity (B) was determined above the wound site (region 1), at the wound site (region 2), and below the wound site (region 3) on the wounded side of the leaf (black bars), in corresponding regions on the opposite side of the wounded leaf (grey bars) and unwounded leaves (white bars). Results are the mean ±SE of four replicates. An asterisk indicates significant differences (one-way ANOVA, P <0.05). Fig. 2. View largeDownload slide Localized changes in cell-wall associated invertase activity 24 h after mechanically wounding leaves of wild-type A. thaliana. A series of 10–12 mechanical wounds were made in a small region of a fully-expanded leaf (A). Sampled regions are indicated by dotted circles. Scale bar=0.5 cm. cwINV activity (B) was determined above the wound site (region 1), at the wound site (region 2), and below the wound site (region 3) on the wounded side of the leaf (black bars), in corresponding regions on the opposite side of the wounded leaf (grey bars) and unwounded leaves (white bars). Results are the mean ±SE of four replicates. An asterisk indicates significant differences (one-way ANOVA, P <0.05). The Arabidopsis genome contains six genes which have been proposed to encode cwINV (AtcwINV1-6, Sherson et al., 2003) although recently two of these genes, AtcwINV3 and AtcwINV6 have been shown to encode fructan exohydrolases and not to possess significant invertase activity (De Coninck et al., 2005). As the literature contains conflicting accounts of the expression of these genes in A. thaliana leaves (Fridman and Zamir, 2003; Sherson et al., 2003), the expression of each family member was quantified by qRT-PCR and the results expressed relative to a control gene SnRP. In RNA isolated from the fully-expanded leaves of 6-week-old A. thaliana plants, significant expression of AtcwINV1 and AtcwINV3 was detected (Fig. 3, open bars). Although AtcwINV5 expression was detectable, the expression was approximately 100-fold less than that of AtcwINV1 and therefore unlikely to contribute significantly to leaf cwINV activity. No transcripts of AtcwINV2, 4 or 6 could be detected (at a detection limit of 10 genome equivalents per μg input cDNA). Fig. 3. View largeDownload slide Expression of AtcwINV1–6 genes in fully-expanded leaves of wild-type (open bars) and ΔAtcwINV1 plants (filled bars). Gene expression is shown as genome equivalents μg−1 input RNA and is the mean ±SE of three independent replicates. Results were normalized for differences in cDNA input relative to the control gene SnRNP D1. (–) Indicates that no expression was detectable. Fig. 3. View largeDownload slide Expression of AtcwINV1–6 genes in fully-expanded leaves of wild-type (open bars) and ΔAtcwINV1 plants (filled bars). Gene expression is shown as genome equivalents μg−1 input RNA and is the mean ±SE of three independent replicates. Results were normalized for differences in cDNA input relative to the control gene SnRNP D1. (–) Indicates that no expression was detectable. AtcwINV1 is responsible for all wound-induced cwINV activity in leaves The expression of AtcwINV1 increased rapidly upon mechanical wounding (Fig. 4) with maximum expression detected at the first time point sampled (1 h). Expression remained significantly greater than that of control leaves for up to 12 h after wounding. No wound-induced expression of other AtcwINV genes was detected (data not shown). As these data were consistent with the hypothesis that AtcwINV1 was the sole wound-induced cwINV, a T-DNA insertion mutant of AtcwINV1 (ΔAtcwINV1) was obtained in which complete inactivation of AtcwINV1 expression was confirmed by qRT-PCR (Fig. 3) There were no gross morphological or growth differences between the wild type and mutant line (data not shown). Fig. 4. View largeDownload slide AtcwINV1 is induced rapidly upon mechanical wounding. RNA samples were prepared from fully-expanded leaves of A. thaliana which had been mechanically wounded (open circles) and corresponding unwounded control plants (closed circles) and AtcwINV1 gene expression quantified by RT-PCR. The first time-point was 1 h after wounding. Gene expression is shown as genome equivalents μg−1 input RNA and is the mean ±SE of three independent replicates. Results were normalized for differences in cDNA input relative to the control gene SnRNP D1. Fig. 4. View largeDownload slide AtcwINV1 is induced rapidly upon mechanical wounding. RNA samples were prepared from fully-expanded leaves of A. thaliana which had been mechanically wounded (open circles) and corresponding unwounded control plants (closed circles) and AtcwINV1 gene expression quantified by RT-PCR. The first time-point was 1 h after wounding. Gene expression is shown as genome equivalents μg−1 input RNA and is the mean ±SE of three independent replicates. Results were normalized for differences in cDNA input relative to the control gene SnRNP D1. Unwounded leaves of the ΔAtcwINV1 T-DNA insertion mutant contained the same vacINV and 50–70% of the cwINV activity of wild-type plants (Fig. 1). However, the wound-induced increase in cwINV activity was completely abolished in the ΔAtcwINV1 line confirming that the expression of the AtcwINV1 gene was responsible for all wound-induced increases in cwINV activity. The expression of AtcwINV2-6 did not differ significantly between the wild type and knockout line (Fig. 3). Imaging photosynthetic metabolism via chlorophyll fluorescence Imaging chlorophyll fluorescence provides a non-invasive and quantitative method with which to examine localized changes in photosynthetic metabolism and has been applied widely to study plant responses to biotic and abiotic stress (reviewed in Oxborough, 2004). Recently, a number of systems have been developed which can image chlorophyll fluorescence with sufficient spatial resolution to provide information at the cellular and subcellular level (Oxborough and Baker, 1997; Kupper et al., 2000; Rolfe and Scholes, 2002; Endo et al., 2002). Energy absorbed by the light-harvesting apparatus has a number of possible fates: (i) it can be transferred to the photosynthetic reaction centres where it provides energy to drive photosynthetic electron transport; (ii) it can be dissipated by non-photochemical processes, a major component of which is normally dissipation as heat within the light-harvesting antennae; (iii) a small proportion (typically 1–5%) is re-emitted as fluorescence with a longer wavelength than that of the incident illumination. As these processes compete with each other, chlorophyll fluorescence can be used to probe underlying photosynthetic function. Quantification of chlorophyll fluorescence emission at different times during photosynthetic induction and under different illumination conditions allows a number of photosynthetic parameters to be calculated. These include ΦII, the proportion of absorbed light which is used for photosynthetic electron transport through PSII (essentially the operating efficiency of PSII measured in the light), Fv/Fm, the maximum efficiency of PSII, a useful measure of photodamage and NPQ (which is calculated as (Fm − F′m)/F′m), which describes the efficiency with which energy is dissipated non-photochemically as heat (reviewed in Maxwell and Johnson, 2000). Recently, Meng et al. (2001) demonstrated that the source–sink transition in developing tobacco leaves could be visualized by imaging the rate of induction of photosynthetic electron transport. They found a close correlation between the rate of induction of photosynthetic electron transport (derived from measurements of the induction of ΦII in leaves which had been placed in darkness for extended periods) and carbohydrate import (determined by 14CO2 feeding). They attribute the more rapid induction of photosynthetic electron transport in ‘sink’ regions of the leaf to the presence of larger pools of metabolic intermediates facilitating the rapid regeneration of Calvin cycle intermediates upon illumination. This approach has been used to study localized changes in source–sink relationships in the vicinity of wounds at cellular resolution. Figure 5 shows the strategy employed. In this example, the leaf was mechanically wounded at 08.00 h, placed in darkness at the end of the usual photoperiod (18.00 h) and chlorophyll fluorescence imaged during photosynthetic induction at 08.00 h (24 h after wounding) the following day. Figure 5A shows an unprocessed image of chlorophyll fluorescence taken during a pulse of saturating light. The wounded cells have died and appeared dark, whilst intact cells fluoresced and appeared bright. A series of contours 80 μm (∼2–3 mesophyll cells) wide were drawn around the wound margin (Fig. 5B). Images of ΦII (Fig. 5C) and NPQ (not shown) were then calculated as the leaf underwent photosynthetic induction. ΦII values were initially low as a result of the extended period of dark adaptation and upon the onset of actinic illumination rose over the following 6 min to a uniformly high value of 0.67. However, although the steady-state values of ΦII were uniform across the image, the rate of induction of ΦII varied depending on the proximity to the wound margin. Induction was more rapid close to the wound hence a bright ring, or halo, was visible around the wound in the images of ΦII. A video of the induction of ΦII in control and wounded leaves is shown in the Supplementary Video 1 at JXB online. Values of ΦII, NPQ, and Fv/Fm were then extracted from each contour to provide a quantitative measure of each photosynthetic parameter at intervals from the wound margin. Fig. 5. View largeDownload slide Unprocessed chlorophyll fluorescence image from a wild-type leaf 24 h after wounding (A). Contours used for extracting average values of calculated photosynthetic parameters (B). Images of ΦII acquired during photosynthetic induction (C). Numbers indicate the time (s) following the onset of actinic illumination. Scale bar=500 μm. Fig. 5. View largeDownload slide Unprocessed chlorophyll fluorescence image from a wild-type leaf 24 h after wounding (A). Contours used for extracting average values of calculated photosynthetic parameters (B). Images of ΦII acquired during photosynthetic induction (C). Numbers indicate the time (s) following the onset of actinic illumination. Scale bar=500 μm. Figure 6A–D shows representative images and extracted values of photosynthetic parameters of control and wounded leaves of wild type and ΔAtcwINV1 A. thaliana taken during photosynthetic induction 1 h and 24 h after wounding. In control leaves, ΦII was induced upon illumination reaching a steady-state value of 0.68 after approximately 200 s. This was accompanied by a transient increase in NPQ which reached a maximum after approximately 100 s. In leaves that were imaged 1 h after wounding (Fig. 6A, B), ΦII was induced more slowly and attained a lower steady-state value in regions immediately proximal to the wound margin. This was accompanied by a corresponding increase in the duration of the NPQ transient. Integrated images of ΦII during the first 300 s following illumination clearly indicated a reduction in photosynthetic electron transport in cells 40–120 μm from the wound margin compared to more distal regions of the wounded leaf or equivalent regions of unwounded leaves. No significant differences were observed between wild type and ΔAtcwINV1 plants: the precise size of the wound margin varied between 40–120 μm in replicate treatments. There was little effect on values of Fv/Fm with values between 0.78–0.83 (see Supplementary Fig. 1A, B at JXB online). Fig. 6. View largeDownload slide Induction of photosynthesis in control and wounded leaves of wild type (A, C) and ΔAtcwINV1 (B, D) A. thaliana. Leaves were placed in darkness overnight and chlorophyll fluorescence imaged during photosynthetic induction. Leaves were wounded 1 h (A, B) or 24 h (C, D) before imaging. Images of NPQ and ΦII were calculated at each time-point and values extracted from 80 μm contours surrounding the wound margin (or corresponding regions of control leaves). Average values for each contour plotted against time are shown as coloured lines with control values from an unwounded leaf shown as a dotted line. For each treatment, an image of integrated ΦII was calculated from images of ΦII taken during the first 300 s of induction (indicated by the grey dotted line). Microautoradiography of 14CO2 radiolabelled leaves indicated that fixed carbon was retained in regions proximal to the wound site in wild-type (E) and ΔAtcwINV1 (F) leaves. Radiolabel was quantified in a transect across the region shown. The solid line shows quantification of radiolabel in wounded leaves, the dotted line in unwounded, control leaves 24 h after feeding. Hydrogen peroxide staining of wounded wild-type (G) and ΔAtcwINV1 (H) leaves. Fig. 6. View largeDownload slide Induction of photosynthesis in control and wounded leaves of wild type (A, C) and ΔAtcwINV1 (B, D) A. thaliana. Leaves were placed in darkness overnight and chlorophyll fluorescence imaged during photosynthetic induction. Leaves were wounded 1 h (A, B) or 24 h (C, D) before imaging. Images of NPQ and ΦII were calculated at each time-point and values extracted from 80 μm contours surrounding the wound margin (or corresponding regions of control leaves). Average values for each contour plotted against time are shown as coloured lines with control values from an unwounded leaf shown as a dotted line. For each treatment, an image of integrated ΦII was calculated from images of ΦII taken during the first 300 s of induction (indicated by the grey dotted line). Microautoradiography of 14CO2 radiolabelled leaves indicated that fixed carbon was retained in regions proximal to the wound site in wild-type (E) and ΔAtcwINV1 (F) leaves. Radiolabel was quantified in a transect across the region shown. The solid line shows quantification of radiolabel in wounded leaves, the dotted line in unwounded, control leaves 24 h after feeding. Hydrogen peroxide staining of wounded wild-type (G) and ΔAtcwINV1 (H) leaves. When leaves were imaged 24 h after wounding (Figs 5, 6C, D), a much more widespread impact on the rate of induction of photosynthetic parameters was observed. Although the steady-state values of ΦII were similar in all regions of the wounded leaf, the rate of induction of ΦII was highly heterogeneous. ΦII was induced most rapidly in regions proximal to the wound margin and progressively more slowly with increasing distance from the wound margin. This progressive variation in the rate of induction of ΦII extended over 1000 μm from the wound margin. In regions where ΦII was induced most rapidly, the NPQ transient was small and reached a maximal value at 100 s. As the rate of ΦII induction slowed, the size of the NPQ transient increased, and the peak value was reached progressively later during photosynthetic induction. This effect was apparent in both wild-type and ΔAtcwINV1 plants and consistent with the induction of a localized sink for carbohydrate in the vicinity of the wound. Again there was little effect on Fv/Fm in the vicinity of the wound (see the Supplementary Fig. 1C, D at JXB online). 14CO2 feeding of wounded leaves To confirm that these alterations in the kinetics of ΦII and NPQ during photosynthetic induction were associated with the development of a localized sink for carbohydrate, control and wounded wild-type and ΔAtcwINV1 plants were fed with 14CO2, frozen at intervals following feeding, and the location of fixed carbon determined by autoradiography. Leaves were wounded 1 h or 24 h prior to feeding. Material frozen immediately after radioactive feeding showed uniform labelling irrespective of wounding. Three hours after feeding, there was still significant radiolabel in both wounded and unwounded leaves: no heterogeneity was apparent at this point (data not shown). However, 24 h after feeding, radiolabel was retained preferentially in a discrete ring (approximately 0.5–1 mm in diameter) around the wound margin (Fig. 6E, F). Radiolabel retention was apparent in both wild type and ΔAtcwINV1 plants and was 1.5–2-fold greater than equivalent regions of control plants. As the generation of active oxygen via the Mehler reaction may act as a potential sink for electrons, leaves were stained using DAB for hydrogen peroxide accumulation 24 h and 48 h after wounding. The veins of both unwounded and wounded plants stained brown indicating that hydrogen peroxide was present, but no accumulation was observed in mesophyll cells in the vicinity of the wound margin in either wild-type or ΔAtcwINV1 plants (Fig. 6G, H). Wound-responsive gene expression in wild type and ΔAtcwINV1 plants Although the chlorophyll fluorescence analysis and radioactive feeding indicated that a localized sink for carbohydrate was induced in the vicinity of the wound in both wild-type and ΔAtcwINV1 plants, the absence of wound-induced apoplastic invertase activity in the mutant plants would be expected to limit the generation of hexose sugars in the apoplast and hence potentially affect aspects of sugar-regulated gene expression. The expression of selected genes was therefore measured in control and wounded leaves of wild-type and mutant plants 3, 24, and 48 h after wounding (Fig. 7). The genes were selected using data derived from publicly-available microarray experiments. Delessert et al. (2004) examined local, adjacent, and systemic alterations in gene expression in mechanically-wounded leaves on a time-course basis (although the wound was much more severe than in the current study). Candidate genes for this study were selected which were induced (or repressed) locally or immediately adjacent to the wound, but not systemically, and which showed a delay in response after wounding (to select genes which were potentially responding after the induction of apoplastic invertase activity). Candidate genes should be of known function and were also known to be glucose-regulated using data from Price et al. (2004). Four genes were selected which were wound and glucose up-regulated, and two genes which were wound and glucose down-regulated (see Supplementary Table 2 at JXB online). Fig. 7. View largeDownload slide Gene expression in control (C) and wounded (W) leaves of wild type (WT) and ΔAtcwINV1 leaves, 3 h (open bars), 24 h (grey bars), and 48 h (solid bars) after wounding. (A) Cinnamyl alcohol dehydrogenase (At4g34230); (B) heat shock protein hsp81-2 (At5g56030); (C) 4-coumarate:CoA ligase 1 (At1g51680); (D) low-temperature-induced protein 78 (At5g52310); (E) PSII type I chlorophyll a/b binding protein (At2g34430); (F) Oxygen-evolving enhancer protein OEE3 (At4g05180). Results are the mean ±SE of three replicates and are reported as genome equivalents μg−1 input RNA. Fig. 7. View largeDownload slide Gene expression in control (C) and wounded (W) leaves of wild type (WT) and ΔAtcwINV1 leaves, 3 h (open bars), 24 h (grey bars), and 48 h (solid bars) after wounding. (A) Cinnamyl alcohol dehydrogenase (At4g34230); (B) heat shock protein hsp81-2 (At5g56030); (C) 4-coumarate:CoA ligase 1 (At1g51680); (D) low-temperature-induced protein 78 (At5g52310); (E) PSII type I chlorophyll a/b binding protein (At2g34430); (F) Oxygen-evolving enhancer protein OEE3 (At4g05180). Results are the mean ±SE of three replicates and are reported as genome equivalents μg−1 input RNA. The expression of genes encoding cinnamyl alcohol dehydrogenase (AtCAD5; Kim et al., 2004) and the heat shock protein hsp81-2 (a stress-induced member of the hsp90 family; Yabe et al., 1994) was induced 3 h after wounding compared with unwounded control leaves (Fig. 7A, B). The induction of AtCAD5 was greater, and more sustained than that of hsp81-2, but there was no significant difference in the response of wild-type and ΔAtcwINV1 plants. Although the genes encoding 4-coumarate:CoA ligase 1 and low-temperature-induced protein 78 have been reported to be both wound- and glucose-induced, no induction was seen in either the wild-type or mutant plants at any of the time-points examined (Fig. 7C, D). The glucose- and wound-repressed genes encoding the chlorophyll a/b binding protein LHB1B1 and the oxygen-evolving enhancer protein OEE3 were strongly repressed 3 h after wounding in both plants (Fig. 7E, F). In summary, the pattern of wound-responsive gene expression observed in wild-type plants in this study did not differ from that observed in ΔAtcwINV1 mutants, even though the wound-induced increase in apoplastic invertase was entirely lacking in the mutant. Discussion The genome of A. thaliana contains a small multi-gene family encoding cwINV (Sherson et al., 2003; Fridman and Zamir, 2003). In this report, quantitative measurements of gene expression have been obtained for each family member in mature leaves and it has been demonstrated that insertional inactivation of AtcwINV1 leads to a 30–50% reduction in cwINV activity of unwounded leaves, and completely abolishes the wound-induced increase in cwINV activity. It is unclear whether the residual cwINV activity of ΔAtcwINV1 leaves results from the expression of other AtcwINV family members or from residual vacINV activity which remains associated with the cell wall fraction even after extensive washing with extraction buffer containing 50 mM NaCl. Quantitative RT-PCR measurements showed that AtcwINV1, AtcwINV3 and, to a lesser extent, AtcwINV5 were expressed in mature leaves. De Coninck et al. (2005) have shown that AtcwINV3 is actually a fructan exohydrolase with no detectable sucrolytic activity and a putative vacuolar location, hence expression of this gene will not contribute to the cwINV activity of the plant. The expression of AtcwINV5 was detectable, but at an expression level 100-fold less than that of AtcwINV1, which itself is not an abundant transcript. However, differences in translational efficiency and protein stability may result in the expression of AtcwINV5 contributing to leaf cwINV activity. Clearly, insertional inactivation of AtcwINV5 and crossing with AtcwINV1 is required to address the contribution of this gene to cwINV activity in unwounded leaves. The pattern of expression of the AtcwINV genes in this report are in agreement with the results of Tymowska-Lalanne and Kreis (1998) who determined the expression of AtcwINV1 and AtcwINV2 using northern blot analysis. Sherson et al. (2003) describe unpublished results of semi-quantitative RT-PCR measurements of AtcwINV gene expression in expanding rosette leaves. The expression pattern reported is similar to that described in this report except expression of AtcwINV5 was not detectable (expression may have been below the limits of detection) and expression of AtcwINV6 was higher than AtcwINV1 (which may be attributable to the different developmental stage examined). Fridman and Zamir (2003) also used RT-PCR to examine expression of AtcwINV1, 2, 4, and 5 although quantification was not attempted and the developmental stage of the leaves is unclear. They found the strongest signal from AtcwINV5 (AtβFRUCT6). The quantitative RT-PCR method employed in this report allows precise quantification of relative transcript levels, although this does not necessarily equate to protein abundance. cwINV gene expression is subject to multiple post-transcriptional regulatory processes including sugar-regulated differential processing of the 3′ untranslated region (Cheng et al., 1999), cold-stress-induced exon-skipping (Bournay et al., 1996), and inhibition by proteinaceous inhibitors (Rausch and Greiner, 2004). Inactivation of the AtcwINV1 gene produced no difference in growth phenotype (either during the vegetative or reproductive phases) indicating that it was not required for normal plant development. By contrast, Tang et al. (1999) found that antisense repression of cwINV expression in carrot impaired embryo development and, in mature plants, enhanced shoot production and inhibited tap root development. This difference may reflect differences in carbohydrate metabolism between species or result from the repression of multiple cwINV genes by the antisense construct. Other cwINVs have been shown to have a critical role in reproductive development, generating a gradient of carbohydrates in developing tissues. In maize, endosperm development is impaired in the miniature-1 mutant which lacks both cwINV and vacINV (Miller and Chourey, 1992) whilst tissue-specific antisense of the cwINV Nin88 in pollen leads to male sterility in tobacco (Goetz et al., 2001). In A. thaliana AtcwINV2 is expressed solely in reproductive tissue (Tymowska-Lalanne and Kreiss, 1998) and may play a central role reproductive development. When wild-type plants were wounded, cwINV activity rose 3–4-fold over a 24 h period compared with unwounded control leaves. AtcwINV1 expression has been shown to be induced by other stresses and signals including pathogens (Chou et al., 2000; Fotopoulos et al., 2003), and carbohydrates (Tymowska-Lalanne and Kreis, 1998) and contains wound and drought/ABA sequence motifs in its promoter (Tymowska-Lalanne and Kreis, 1998; PLACE signal motif database: Higo et al., 1999). In addition, cwINV gene expression and activity has been shown to be induced by multiple plant growth regulators (reviewed in Roitsch and Gonzaléz, 2004). Although the wound-induced increase of cwINV activity in A. thaliana leaves was relatively modest, it was restricted to regions in the immediate vicinity of the wound. As the apoplast represents only a small proportion of the total cell wall volume, the fold-increase at the precise site of induction was likely to be considerably higher. The localization and timing of induction of AtcwINV1 gene expression is similar to the local, late-induced pattern of gene expression reported by Delessert et al. (2004) and is typical of many genes encoding metabolic enzymes. The precise cellular location of AtcwINV1 expression remains to be determined although studies of cwINV genes in the leaves of other species consistently indicates an association with the phloem (Ramloch-Lorenz et al., 1993; Kingston-Smith and Pollock, 1996; Hedley et al., 2000) including wound-induced activity (Zhang et al., 1996). Although wound induction of cwINV activity has been reported previously in other species, it has been shown that AtcwINV1 is solely responsible for the increase observed in wounded leaves of A. thaliana and that inactivation of this gene abolishes this increase. Thus the ΔAtcwINV1 plants provide an ideal system with which to examine the role of induced cwINV activity in localized sink induction and signalling. Chlorophyll fluorescence imaging was used to examine photosynthetic induction in wounded leaves under experimental conditions that allow this technique to probe source–sink relationships (Meng et al., 2001). The advantage of this technique is that it provides information about the primary physiological processes of the plant with cellular resolution. However, as photosynthetic processes are influenced by many environmental and developmental factors, interpretation of these results requires caution. Therefore 14CO2 labelling was used independently to verify that source–sink relationships were altered in the vicinity of the wound, and to determine whether these relationships differed between wild-type and ΔAtcwINV1 plants. When leaves were imaged immediately after wounding, there was a localized reduction in the steady-state values of ΦII in cells adjacent to the wound margin. This reduction, coupled with a relatively minor impact on the duration and magnitude of the transient increase of NPQ following the onset of illumination, suggests that these cells were damaged. Although Fv/Fm was not markedly reduced in these regions, the leaves had been maintained in darkness prior to imaging hence photodamage would not yet have occurred. There was no stimulation in the rate of induction of ΦII in leaves 1–2 h after wounding indicating that source–sink relationships had not been affected at this time. When leaves were imaged 24 h after wounding, all regions of the wounded leaf reached a similar steady-state rate of photosynthesis following photosynthetic induction and again there was no marked reduction in Fv/Fm. The cells in the vicinity of the wound were therefore capable of photosynthesizing at the same rate as neighbouring cells and were not showing evidence of photodamage. Cells which were showing reduced steady-state values of ΦII immediately after wounding had therefore either died or fully recovered. However, there was a significant localized impact on the rate of ΦII induction in wounded leaves at this time. ΦII induction was most rapid closest to the wound and became slower with increasing distance from the wound. Areas with a high rate of ΦII induction exhibited a small, rapid NPQ transient whilst areas with lower rates of ΦII induction exhibited a much larger and longer lived NPQ transient. These results are entirely consistent with a shift towards sink metabolism in the vicinity of the wound at these later time points. Cells with a greater supply of carbohydrate would contain higher concentrations of Calvin cycle intermediates allowing a rapid induction of ΦII. Those cells which had characteristics more typical of source tissue would exhibit a slower rate of ΦII induction hence energy dissipation in the light-harvesting antennae (Maxwell and Johnson, 2000) would lead to a greater and more pronounced NPQ transient. Although other explanations are possible for the alteration in the rate of photosynthetic induction, such as the Mehler reaction acting as a sink for electron transport (Asada, 1999; Heber, 2002), radioactive labelling clearly indicated that photosynthate was retained in the vicinity of the wound in a region of comparable size to that observed by fluorescence imaging (∼1 mm diameter). Recently Chang et al. (2004) proposed that hydrogen peroxide generated by photosynthetic electron transport and the Mehler reaction in mechanically wounded leaves may act as an important regulator of gene expression, particularly in leaves exposed to excess light. Leaves were wounded by crushing between forceps across the entire width of the lamina in three positions and resulted in a marked decline in ΦII immediately after wounding, which only partially recovered at later times. Under these conditions, H2O2 accumulated in veinal regions and led to a marked induction of the antioxidant enzyme, ascorbate peroxidase 2. In the current study, wounds were much less severe and photosynthetic induction experiments were performed after an extended period of darkness (14 h) and at low photon irradiance (80 μmol m−2 s−1). DAB staining indicated the presence of H2O2 in veinal regions, but there was no difference in staining pattern or intensity of control and wounded plants. In addition, there was no staining in cells in the vicinity of the wound. In fact, if energy dissipation by the Mehler reaction in the vicinity of the wounds was significant, an increase in energy dissipation via NPQ in the antenna of photosystem II would also be expected, in fact the opposite was observed. The production of reactive oxygen species would also be expected to lead to a reduction in Fv/Fm and steady-state ΦII which was not observed. Clearly, the production of localized H2O2 in wounded leaves may act as an important signalling molecule, but the extent to which this occurs will depend both upon the severity of the wound and other environmental factors. In wild parsnip leaves wounded by cabbage looper caterpillars (Trichoplusia ni) reductions in photosynthetic electron transport were much more widespread (Zangerl et al., 2002) presumably as a result of more widespread systemic responses triggered by elicitors in insect saliva. Although the induction of cwINV activity and AtcwINV1 gene expression in wounded leaves was consistent with the proposed role in altering source/sink status, both chlorophyll fluorescence imaging and radioactive feeding indicated that the increased sink status was still evident in ΔAtcwINV1 plants. As wound-induced cwINV activity is absent from these plants it is concluded that induced cwINV activity is not a requirement for localized alterations in source/sink status. The question naturally arises therefore of what does lead to this alteration? The sucrose transporter AtSUC3, the hexose transporter AtSTP4, and, to a lesser extent AtSTP3, have all been reported to be induced in wounded leaves (Meyer et al., 2004; Truernit et al., 1996; Büttner et al., 2000). In addition, Delessert et al. (2004) report that a number of enzymes associated with carbohydrate metabolism including sucrose synthase, β-glucosidase, and fructokinase are induced locally in wounded leaves. The export of photosynthate from cells in the vicinity of the wound will be influenced by alterations in sucrolytic activity or sucrose/hexose transporter activity in those cells or in cells of the surrounding vasculature. Addressing this question will require a systematic analysis of localized changes in metabolism in wounded tissues, determination of the cellular location of alterations in expression and enzyme activity of wound-induced metabolic enzymes, and inactivation of candidate genes to assess the impact on localized source–sink relationships. AtcwINV1 may still play a role in these processes, potentially scavenging sucrose released into the apoplast from disrupted cells or the vascular system. No differences in gene expression were observed between wounded wild-type and ΔAtcwINV1 plants. The expression of cinnamyl alcohol dehydrogenase and heat shock protein were induced, and chlorophyll a/b binding protein and OEE3 repressed by wounding as described by Delessert et al. (2004). However, 4 coumarate coA ligase and low-temperature-induced protein 78 were not wound induced. This may result from differences in the severity of the wound. In the current study, 10–12 small crush wounds were made across the leaf surface. By contrast, Delessert et al. (2004) used a pair of forceps to create a much larger wound extending across the entire width of the leaf in a manner more analogous to that described by Chang et al. (2004). As the inactivation of AtcwINV1 did not disrupt localized alterations in source–sink relationships, the role of sugar sensing in wound-regulated gene expression remains to be determined. AtcwINV1 is itself sugar regulated (Tymowska-Lalanne and Kreis, 1998) and thus may be responding to locally generated sugar signals, but it is also responsive to ABA and drought, signals which regulate the relatively slow, local expression of a number of wound-responsive genes reported by Delessert et al. (2004) and are thought to play an important role in mechanically wounded leaves (Reymond et al., 2000). Ethylene may also play an important role as those genes which are repressed locally, including those encoding components of the photosynthetic apparatus, are ethylene responsive. In this report, the long-standing hypothesis that cwINV activity plays a central role in regulating the source–sink status of wounded leaves and has a potential role in generating hexose sugars which act as signals regulating gene expression has been addressed. It has been demonstrated that inactivation of AtcwINV1 does not affect these processes and that chlorophyll fluorescence imaging, supported by radioactive labelling, provides a useful tool with which to examine source–sink relationships at the cellular level. RSQ was supported by a BBSRC studentship (02BIP08366). The authors are grateful to Dr R Vasey (University of Sheffield) for help in quantitative RT-PCR measurements and to Professor WP Quick for access to the μImager radioactive imaging system. References Altschul SF, Gish W, Miller W, Meyers EW, Lipman DJ. 1990. Basic Local Alignment Search Tool. Journal of Molecular Biology  215, 403–410. Google Scholar Arnold TM, Schultz JC. 2002. Induced sink strength as a prerequisite for induced tannin biosynthesis in developing leaves of Populus. Oecologia  130, 585–593. Google Scholar Arnold WN. 1965. β-fructofuranosidase from grape berries. Biochimica et Biophysica Acta  110, 134–147. Google Scholar Asada K. 1999. The water–water cycle in chloroplasts: scavenging of active oxygens and dissipation of excess photons. Annual Review of Plant Physiology and Plant Molecular Biology  50, 601–639. Google Scholar Balibrea Lara ME, Garcia MCG, Fatima T, Ehness R, Lee TK, Proels R, Tanner W, Roitsch T. 2004. Extracellular invertase is an essential component of cytokinin-mediated delay of senescence. The Plant Cell  16, 1276–1287. Google Scholar Bournay AS, Hedley PE, Maddison A, Waugh R, Machray GC. 1996. Exon skipping induced by cold stress in a potato invertase gene transcript. Nucleic Acids Research  24, 2347–2351. Google Scholar Büttner M, Truernit E, Baier K, Scholz-Starke J, Sontheim M, Lauterbach C, Huss VAR, Sauer N. 2000. AtSTP3, a green leaf-specific, low affinity monosaccharide-H+ symporter of Arabidopsis thaliana. Plant, Cell and Environment  23, 175–184. Google Scholar Chang CC-C, Ball L, Fryer MJ, Baker NR, Karpinski S, Mullineaux PM. 2004. Induction of ASCORBATE PEROXIDASE 2 expression in wounded Arabidopsis leaves does not involve known wound-signalling pathways but is associated with changes in photosynthesis. The Plant Journal  38, 499–511. Google Scholar Cheng WH, Taliercio EW, Chourey PS. 1999. Sugars modulate an unusual mode of control of the cell-wall invertase gene (Incw1) through its 3′ untranslated region in a cell suspension culture of maize. Proceedings of the National Academy of Sciences, USA  96, 10512–10517. Google Scholar Chou HM, Bundock NJ, Rolfe SA, Scholes JD. 2000. Infection of Arabidopsis thaliana leaves with Albugo candida (white blister rust) causes a reprogramming of host metabolism. Molecular Plant Pathology  1, 99–113. Google Scholar De Coninck B, Le Roy K, Francis I, Clerens S, Vergauwen R, Halliday AM, Smith SM, Van Laere A, Van den Ende W. 2005. Arabidopsis AtcwINV3 and 6 are not invertases but are fructan exohydrolases (FEHs) with different substrate specificities. Plant, Cell and Environment  28, 432–443. Google Scholar Delessert C, Wilson IW, Van der Straeten D, Dennis ES, Dolferus R. 2004. Spatial and temporal analysis of the local response to wounding in Arabidopsis leaves. Plant Molecular Biology  55, 165–181. Google Scholar Ehneß R, Roitsch T. 1997. Co-ordinated induction of mRNAs for extracellular invertase and a glucose transporter in Chenopodium rubrum by cytokinins. The Plant Journal  11, 539–548. Google Scholar Endo R, Osama K, Kondo J. 2002. Microscopic image instrumentation of chlorophyll a fluorescence from in situ microalgae. EcoEngineering  14, 17–22. Google Scholar Fotopoulos V, Gilbert MJ, Pittman JK, Marvier AC, Buchanan AJ, Sauer N, Hall JL, Williams LE. 2003. The monosaccharide transporter gene, AtSTP4, and the cell-wall invertase, Atβfruct1, are induced in Arabidopsis during infection with the fungal biotroph Erysiphe cichoracearum. Plant Physiology  132, 821–829. Google Scholar Fridman E, Zamir D. 2003. Functional divergence of a syntenic invertase gene family in tomato, potato, and Arabidopsis. Plant Physiology  131, 603–609. Google Scholar Gibson SI. 2004. Sugar and phytohormone response pathways: navigating a signalling network. Journal of Experimental Botany  55, 253–264. Google Scholar Goetz M, Godt DE, Guivarc'h A, Kahmann U, Chriqui D, Roitsch T. 2001. Induction of male sterility in plants by metabolic engineering of the carbohydrate supply. Proceedings of the National Academy of Sciences, USA  98, 6522–6527. Google Scholar Halford NG, Paul MJ. 2003. Carbon metabolite sensing and signalling. Plant Biotechnology Journal  1, 381–398. Google Scholar Heber U. 2002. Irrungen, Wirrungen? The Mehler reaction in relation to cyclic electron transport in C3 plants. Photosynthesis Research  73, 223–231. Google Scholar Hedley PE, Maddison AL, Davidson D, Machray GC. 2000. Differential expression of invertase genes in internal and external phloem tissues of potato (Solanum tuberosum L.). Journal of Experimental Botany  51, 817–821. Google Scholar Herbers K, Meuwly P, Frommer WB, Métraux JP, Sonnewald U. 1996. Systemic acquired resistance mediated by the ectopic expression of invertase: possible hexose sensing in the secretory pathway. The Plant Cell  8, 793–803. Google Scholar Herbers K, Sonnewald U. 1998. Altered gene expression brought about by inter- and intracellularly formed hexoses and its possible implications for plant–pathogen interactions. Journal of Plant Research  111, 323–328. Google Scholar Higo K, Ugawa Y, Iwamoto M, Korenaga T. 1999. Plant cis-acting regulatory DNA elements (PLACE) database: 1999. Nucleic Acids Research  27, 297–300. Google Scholar Howe GA. 2004. Jasmonates as signals in the wound response. Journal of Plant Growth Regulation  23, 223–237. Google Scholar Jacobs AK, Lipka V, Burton RA, Panstruga R, Strizhov N, Schulze-Lefert P, Fincher GB. 2003. An Arabidopsis callose synthase, GSL5, is required for wound and papillary callose formation. The Plant Cell  15, 2503–2513. Google Scholar Kim S-J, Kim M-R, Bedgar DL, Moinuddin SGA, Cardenas CL, Davin LB, Kang C, Lewis NG. 2004. Functional reclassification of the putative cinnamyl alcohol dehydrogenase multigene family in Arabidopsis. Proceedings of the National Academy of Sciences, USA  101, 1455–1460. Google Scholar Kingston-Smith AH, Pollock CJ. 1996. Tissue level localization of acid invertase in leaves: an hypothesis for the regulation of carbon export. New Phytologist  134, 423–432. Google Scholar Küpper H, Šetlík I, Trtílek M, Nedbal L. 2000. A microscope for two-dimensional measurements of in vivo chlorophyll fluorescence kinetics using pulsed measuring radiation, continuous actinic radiation, and saturating flashes. Photosynthetica  38, 553–570. Google Scholar León J, Rojo E, Sánchez-Serrano JJ. 2001. Wound signalling in plants. Journal of Experimental Botany  52, 1–9. Google Scholar Maxwell K, Johnson GN. 2000. Chlorophyll fluorescence: a practical guide. Journal of Experimental Botanty  51, 659–668. Google Scholar Meng QW, Siebke K, Lippert P, Baur B, Mukherjee U, Weis E. 2001. Sink–source transition in tobacco leaves visualized using chlorophyll fluorescence imaging. New Phytologist  151, 585–595. Google Scholar Meyer S, Lauterbach C, Niedermeier M, Barth I, Sjolund RD, Sauer N. 2004. Wounding enhances expression of AtSUC3, a sucrose transporter from Arabidopsis sieve elements and sink tissues. Plant Physiology  134, 684–693. Google Scholar Miller ME, Chourey PS. 1992. The maize invertase-deficient miniature-1 seed mutation is associated with aberrant pedicel and endosperm development. The Plant Cell  4, 297–305. Google Scholar Ohyama A, Nishimura S, Hirai M. 1998. Cloning of cDNA for a cell wall-bound acid invertase from tomato (Lycopersicon esculentum) and expression of soluble and cell wall-bound invertases in plants and wounded leaves of L. esculentum and L. peruvianum. Genes and Genetic Systems  73, 149–157. Google Scholar Orozco-Cárdenas ML, Narvaez-Vasquez J, Ryan CA. 2001. Hydrogen peroxide acts as a second messenger for the induction of defense genes in tomato plants in response to wounding, systemin, and methyl jasmonate. The Plant Cell  13, 179–191. Google Scholar Oxborough K. 2004. Imaging of chlorophyll a fluorescence: theoretical and practical aspects of an emerging technique for the monitoring of photosynthetic performance. Journal of Experimental Botany  55, 1195–1205. Google Scholar Oxborough K, Baker NR. 1997. An instrument capable of imaging chlorophyll alpha fluorescence from intact leaves at very low irradiance and at cellular and subcellular levels of organization. Plant, Cell and Environment  20, 1473–1483. Google Scholar Price J, Laxmi A, St Martin SK, Jang JC. 2004. Global transcription profiling reveals multiple sugar signal transduction mechanisms in Arabidopsis. The Plant Cell  16, 2128–2150. Google Scholar Ramloch-Lorenz K, Knudsen S, Sturm A. 1993. Molecular characterization of the gene for carrot cell wall β-fructosidase. The Plant Journal  4, 545–554. Google Scholar Rausch T, Greiner S. 2004. Plant protein inhibitors of invertases. Biochimica et Biophysica Acta: Proteins and Proteomics  1696, 253–261. Google Scholar Reymond P, Farmer EE. 1998. Jasmonate and salicylate as global signals for defense gene expression. Current Opinion in Plant Biology  1, 404–411. Google Scholar Reymond P, Weber H, Damond M, Farmer EE. 2000. Differential gene expression in response to mechanical wounding and insect feeding in Arabidopsis. The Plant Cell  12, 707–719. Google Scholar Roitsch T. 1999. Source–sink regulation by sugar and stress. Current Opinion in Plant Biology  2, 198–206. Google Scholar Roitsch T, Balibrea ME, Hofmann M, Proels R, Sinha AK. 2003. Extracellular invertase: key metabolic enzyme and PR protein. Journal of Experimental Botany  54, 513–524. Google Scholar Roitsch T, Gonzaléz MC. 2004. Function and regulation of plant invertases: sweet sensations. Trends in Plant Science  9, 606–613. Google Scholar Rolfe SA, Scholes JD. 1995. Quantitative imaging of chlorophyll fluorescence. New Phytologist  131, 69–79. Google Scholar Rolfe SA, Scholes JD. 2002. Extended depth-of-focus imaging of chlorophyll fluorescence from intact leaves. Photosynthesis Research  72, 107–115. Google Scholar Scholes JD, Lee PJ, Horton P, Lewis DH. 1994. Invertase: understanding changes in the photosynthetic and carbohydrate-metabolism of barley leaves infected with powdery mildew. New Phytologist  126, 213–222. Google Scholar Sessions A, Burke E, Presting G, et al. 2002. A high-throughput Arabidopsis reverse genetics system. The Plant Cell  14, 2985–2994. Google Scholar Sherson SM, Alford HL, Forbes SM, Wallace G, Smith SM. 2003. Roles of cell-wall invertases and monosaccharide transporters in the growth and development of Arabidopsis. Journal of Experimental Botanty  54, 525–531. Google Scholar Smeekens S. 2000. Sugar-induced signal transduction in plants. Annual Review of Plant Physiology and Plant Molecular Biology  51, 49–81. Google Scholar Sonnewald U, Brauer M, von Schaewen A, Stitt M, Willmitzer L. 1991. Transgenic tobacco plants expressing yeast-derived invertase in either the cytosol, vacuole or apoplast: a powerful tool for studying sucrose metabolism and sink source interactions. The Plant Journal  1, 95–106. Google Scholar Stratmann JW. 2003. Long distance run in the wound response: jasmonic acid is pulling ahead. Trends in Plant Science  8, 247–250. Google Scholar Sturm A, Chrispeels MJ. 1990. cDNA cloning of carrot extracellular β-fructosidase and its expression in response to wounding and bacterial-infection. The Plant Cell  2, 1107–1119. Google Scholar Tang GQ, Luscher M, Sturm A. 1999. Antisense repression of vacuolar and cell wall invertase in transgenic carrot alters early plant development and sucrose partitioning. The Plant Cell  11, 177–189. Google Scholar Tang X, Rolfe SA, Scholes JD. 1996 a. The effect of Albugo candida (white blister rust) on the photosynthetic and carbohydrate metabolism of leaves of Arabidopsis thaliana. Plant, Cell and Environment  19, 967–975. Google Scholar Tang XW, Ruffner HP, Scholes JD, Rolfe SA. 1996 b. Purification and characterisation of soluble invertases from leaves of Arabidopsis thaliana. Planta  198, 17–23. Google Scholar Thordal-Christensen H, Zhang Z, Wei Y, Collinge DB. 1997. Subcellular localization of H2O2 in plants. H2O2 accumulation in papillae and hypersensitive response during the barley–powdery mildew interaction. The Plant Journal  11, 1187–1194. Google Scholar Truernit E, Schmid J, Epple P, Illig J, Sauer N. 1996. The sink-specific and stress-regulated Arabidopsis STP4 gene: enhanced expression of a gene encoding a monosaccharide transporter by wounding, elicitors, and pathogen challenge. The Plant Cell  8, 2169–2182. Google Scholar Tymowska-Lalanne Z, Kreis M. 1998. Expression of the Arabidopsis thaliana invertase gene family. Planta  207, 259–265. Google Scholar von Schaewen A, Stitt M, Schmidt R, Sonnewald U, Willmitzer L. 1990. Expression of yeast-derived invertase in the cell wall of tobacco and Arabidopsis leads to accumulation of carbohydrate and inhibition of photosynthesis and strongly influences growth and phenotype of transgenic tobacco plants. EMBO Journal  9, 3033–3043. Google Scholar Wendehenne D, Durner J, Klessig DF. 2004. Nitric oxide: a new player in plant signalling and defence responses. Current Opinion in Plant Biology  7, 449–455. Google Scholar Yabe N, Takahashi T, Komeda Y. 1994. Analysis of tissue-specific expression of Arabidopsis thaliana HSP90-family gene HSP81. Plant and Cell Physiology  35, 1207–1219. Google Scholar Zangerl AR, Hamilton JG, Miller TJ, Crofts AR, Oxborough K, Berenbaum MR, de Lucia EH. 2002. Impact of folivory on photosynthesis is greater than the sum of its holes. Proceedings of the National Academy of Sciences, USA  99, 1088–1091. Google Scholar Zhang L, Cohn NS, Mitchell JP. 1996. Induction of a pea cell-wall invertase gene by wounding and its localized expression in phloem. Plant Physiology  112, 1111–1117. Google Scholar © The Author [2005]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved. For Permissions, please e-mail: journals.permissions@oxfordjournals.org
TI - Imaging photosynthesis in wounded leaves of Arabidopsis thaliana
JF - Journal of Experimental Botany
DO - 10.1093/jxb/erj039
DA - 2005-12-09
UR - https://www.deepdyve.com/lp/oxford-university-press/imaging-photosynthesis-in-wounded-leaves-of-arabidopsis-thaliana-f9S7TurphS
SP - 55
EP - 69
VL - 57
IS - 1
DP - DeepDyve
ER -