JXB Advance Access originally published online on March 19, 2008
Journal of Experimental Botany 2008 59(6):1175-1186; doi:10.1093/jxb/ern019
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© 2008 The Author(s).
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/2.0/uk/) which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.
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RESEARCH PAPER |
Regulatory involvement of abscisic acid in potato tuber wound-healing
USDA-ARS, Northern Crop Science Laboratory, 1307 18th Street North, Fargo, ND 58105, USA
* To whom correspondence should be addressed. E-mail: ed.lulai{at}ars.usda.gov
Received 30 September 2007; Revised 11 January 2008 Accepted 18 January 2008
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Abstract |
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 Top Abstract IntroductionMaterials and methodsResultsDiscussionConcluding remarksSupplementary dataReferences |
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Rapid wound-healing is crucial in protecting potato tubers from infection and dehydration. Wound-induced suberization and the accumulation of hydrophobic barriers to reduce water vapour conductance/loss are principal protective wound-healing processes. However, little is known about the cognate mechanisms that effect or regulate these processes. The objective of this research was to determine the involvement of abscisic acid (ABA) in the regulation of wound-induced suberization and tuber water vapour loss (dehydration). Analysis by liquid chromatography–mass spectrometry showed that ABA concentrations varied little throughout the tuber, but were slightly higher near the periderm and lowest in the pith. ABA concentrations increase then decrease during tuber storage. Tuber wounding induced changes in ABA content. ABA content in wound-healing tuber discs decreased after wounding, reached a minimum by 24 h, and then increased from the 3rd to the 7th day after wounding. Wound-induced ABA accumulations were reduced by fluridone (FLD); an inhibitor of de novo ABA biosynthesis. Wound-induced phenylalanine ammonia lyase activity was slightly reduced and the accumulation of suberin poly(phenolics) and poly(aliphatics) noticeably reduced in FLD-treated tissues. Addition of ABA to the FLD treatment restored phenylalanine ammonia lyase activity and suberization, unequivocally indicating that endogenous ABA is involved in the regulation of these wound-healing processes. Similar experiments showed that endogenous ABA is involved in the regulation of water vapour loss, a process linked to wax accumulation in wound-healing tubers. Rapid reduction of water vapour loss across the wound surface is essential in preventing desiccation and death of cells at the wound site; live cells are required for suberization. These results unequivocally show that endogenous ABA is involved in the regulation of wound-induced suberization and the processes that protect surface cells from water vapour loss and death by dehydration.
Key words: Abscisic acid, poly(aliphatic), poly(phenolic), potato, Solanum tuberosum L., suberin
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Introduction |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionConcluding remarksSupplementary dataReferences |
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Potato tubers incur wounds during harvest, handling, and seed cutting. Rapid healing of these wounds is essential to avoid loss of turgor and infection. However, the relevant wound-healing processes are not fully understood and little is known about the biological factors regulating them or how putative regulators may be used to hasten these processes.
Wound-induced suberization and development of a hydrophobic matrix to reduce water vapour loss at the wound site are among the major wound-healing processes that protect the tuber from infection and deter shrinkage/loss of turgor and defect development (Vogt et al., 1983; Lulai and Orr, 1995; Lulai and Corsini, 1998; Schreiber et al., 2005). In potato tubers, as the wound site begins to heal, a closing layer develops whereby walls of existing cells at the wound surface suberize (Lulai, 2007). Following formation of the closing layer, a wound periderm develops whereby files of new cells are formed and suberized directly below the suberized cells of the closing layer. A hydrophobic matrix of long chain waxes that control water vapour loss is integrated within the walls of these suberized cells (Soliday et al., 1979).
Wound-induced suberization is a complex and poorly understood process that includes induction of phenylalanine ammonia lyase (PAL), in association with phenylpropanoid biosynthesis, and biosynthesis of specific fatty acids (Bernards, 2002; Lulai, 2007). The relevant phenylpropanoid products accumulate as suberin poly(phenolic(s)) (SPP) and are assembled through a yet to be determined process on/in the cell wall and form the suberin poly(phenolic) domain (SPPD). Specific fatty acids accumulate as suberin poly(aliphatic(s)) (SPA) and are assembled forming the suberin poly(aliphatic) domain (SPAD) which is putatively cross-linked or bridged with glycerol units and laminated in a spatially separate orientation over the SPPD (Lulai and Morgan, 1992; Graca and Pereira, 2000; Bernards, 2002; Graca and Santos, 2007; Lulai, 2007). The SPPD is formed first and has special importance because it provides a barrier to bacterial infection (Lulai and Corsini, 1998). The SPAD is formed days later and also bears distinct importance because it provides a surface barrier to fungal infections. The SPAD has also been shown to block fungal advancement deep within infected tuber tissues as well as provide resistance to fungal infection in root systems (Lulai, 2005; Lulai et al., 2006; Thomas et al., 2007). The architecture of the suberin biopolymeric matrix is hypothetical (Bernards, 2002). Soluble waxes are integrated into the suberizing cell wall. The hydrophobic nature of the wax matrix provides a barrier to water vapour conductance which protects cells associated with native periderm and at wound sites from dehydration and death (Soliday et al., 1979; Vogt et al., 1983). Rapid reduction in water vapour conductance at the wound surface is essential in maintaining viable cells for subsequent development of the suberin barrier. Wax deposition had generally been associated with suberization; however, reduction of water vapour conductance must initiate and progress prior to suberization or cells will dehydrate and die before suberization can take place (Lulai and Orr, 1995; Schreiber et al., 2005; Lulai, 2007). Induction of significant decreases in water vapour conductance have been shown within 24 h of wounding; a time prior to accumulation of a single contiguous tangential layer of SPP on the closing layer and days before development of a SPAD (Lulai and Orr, 1995). The hormonal regulation of these suberization processes and resistance to water vapour conductance has not been determined.
Lulai and Suttle (2004) determined that, although ethylene evolution is part of the tuber wound response, ethylene is not directly required for wound-induced suberization. Abscisic acid (ABA) regulation of responses to drought and salt stress are well known (Himmelbach et al., 2003; Efetova et al., 2007). ABA has been correlated with aquaporin gene expression, putatively protecting against desiccation, in Agrobacterium-induced tumours and with pharmacologically enhanced suberin accumulation in root tissue of Arabidopsis (Efetova et al., 2007). Earlier, pharmacological studies correlating the effect of exogenous ABA suggested that the hormone may be involved in the regulation of tuber wound-healing (Soliday et al., 1978; Cottle and Kolattukudy, 1982; Espelie and Kolattukudy, 1985) and water vapour conductance (Lulai and Orr, 1995). However, until now, this involvement has not been unequivocally determined by quantification of tuber ABA content at and after wounding and specific blockage of de novo ABA biosynthesis to determine if wound-healing processes are suppressed or down-regulated. Most importantly, it had not been determined if addition of ABA would restore wound-healing processes in ABA-depleted tissues.
The blocking of ABA synthesis through genetic mutants or through the use of directed inhibitors could provide a reliable non-correlative means of determining the role of ABA in wound-healing. The xenobiotic compound, fluridone (FLD), effectively inhibits the formation of carotenoids that are precursors in ABA biosynthesis (Gamble and Mullet, 1986). As a directed inhibitor of de novo ABA biosynthesis, FLD has been used to determine the role of ABA in various plant processes including dormancy. Although FLD affects non-ABA related bleaching in photosynthesizing tissues, it is especially useful in tissues that are not photoautotrophic and do not require protection from photo-oxidation (Gamble and Mullet, 1986; Suttle and Hultstrand, 1994; Grappin et al., 2000).
High performance liquid chromatography–mass spectrometry (HPLC-MS) techniques for quantification of ABA were combined with FLD treatment, for directed depletion of ABA in situ, to determine the involvement of ABA in the major wound-healing processes of suberization and development of resistance to water vapour conductance.
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Materials and methods |
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Tuber material and storage conditions
Certified seed potatoes (Solanum tuberosum L. cv. Russet Burbank) were used throughout this research. Either mini-tubers (10–12 g) or field-grown tubers were used. Tubers were allowed to cure and undergo further periderm maturation for 14 d at 20 °C after harvest. Thereafter, the tubers were stored in the dark at 4 °C (
95% RH) to retard deterioration and sprouting. Three days prior to use, tubers were gently hand washed and equilibrated in the dark at 20 °C (95% RH).
Statistical analysis
All experiments were repeated and data from each experimental time point were derived from three or more separate samples of tubers. Data from each experiment were analysed by standard analysis of variance (ANOVA) techniques, treatment means were compared using Fisher's protected least squares difference (LSD) at a P-value of 0.05, using Statistical Analysis System (SAS, v 8.0; Cary, NC, USA).
Tuber tissue wound-healing systems
Tubers were wounded and allowed to heal using one of two model systems: (i) the tuber disc model system in which cylinders of tissue were laterally excised from each tuber with a cork borer (11 mm diameter) and discs (3 mm thick) of parenchyma tissue cut from the cylinder and immediately treated as described for each experiment; (ii) the half-tuber model system in which mini-tubers or small (40–85 g) field-grown seed tubers were cut in half from stem to bud end and the cut surface of the half tuber immediately treated as indicated for each experiment.
Tissue sampling for determination of ABA content
Intact tubers were sampled to determine basal ABA content at harvest and during storage. Equatorial, stem, and bud end surface portions of the tuber were sliced to provide sample sections of periderm with small amounts of neighbouring cortical cells. Sampling uniformity was enhanced by using a specially fabricated device designed to slice 0.75-mm-thick sections of tissue from the tuber surface. Parenchyma and pith samples were taken from pith rays and parenchyma tissues exposed after the tuber was cut in half from stem to bud end. Parenchyma consisted of tissues located between the vascular ring and pith rays. Pith rays were identified by their typical translucent characteristic. Tissue samples were cored from the sections with a 0.6-mm-diameter cork borer as described below for ABA analysis.
ABA extraction and analysis
Each sample consisted of six plugs of tissue (0.3 g FW) cut with a 0.6-mm-diameter cork borer from sliced sections (0.75 mm thick) of intact tubers or from six wound-healing parenchyma discs (11 mmx3 mm) excised from six tubers. Samples were immediately frozen in liquid nitrogen. Two or more samples were analysed in duplicate for each time point. The tissues were thawed at 4 °C in 80% (v/v) aqueous acetone, homogenized, extracted, purified, and the ABA content quantified by HPLC-MS using an internal standard of 50 ng [2H]6-(+)-ABA (OlChemIm Ltd, Czech Republic) as described by Destefano-Beltran et al. (2006).
ABA and FLD treatments
ABA [(±) cis, trans-ABA; Sigma Chemical Co., St Louis, MO, USA] and FLD (OlChemIm Ltd) were separately dissolved (0.1 M) in dimethylsulphoxide and appropriate aliquots diluted, generally 1:1000 (0.1 mM) unless the final concentration indicated otherwise, in reagent grade water. The treatment solutions were sonicated (3x 1 min, 20 000 cycles s–1) before use to ensure uniform dispersion. Freshly cut discs were placed in the appropriate treatment solution (<30 discs 100 ml–1) and incubated on a rotary shaker ( 50 cycles min–1) for 1 h with fresh changes of treatment solutions every 20 min. After treatment, the tuber tissues were allowed to wound-heal in the dark at 20 °C (95% RH). Samples were taken for analyses at the indicated time points.
Suberization ratings and microscopy
Blocks of tissue (3 mmx8 mmx11 mm) were taken from wound-healing discs at the indicated time points and placed in Farmer's fixative composed of absolute ethanol/acetic acid (3:1, v/v). The accumulation of suberin biopolymers was determined microscopically in triplicate from 20-µm-thick sections cut with a Vibratome 1000 Plus sectioning system. Separate suberization ratings were obtained for the accumulation of SPP (autofluorescence) and SPA (toluidine blue O/neutral red; Sigma Chemical Co.) on suberizing cell walls, using methods described by Lulai and Morgan (1992) and Lulai and Corsini (1998) as modified by Lulai et al. (2006). Microscopy was performed with a Zeiss Axioskop 50 microscope configured for epifluorescent illumination as previously described (Lulai and Corsini, 1998). Digital images were obtained with a Zeiss colour AxioCam camera (Carl Zeiss Inc., Thornwood, NY, USA). Briefly, suberization ratings indicated the following accumulations: 0 = none; 5 = complete around the 1st cell layer; and 7 = complete around 1st and 2nd cell layers (Lulai and Corsini, 1998). The rating system for poly(phenolic) and poly(aliphatic) accumulation on closing layer cell walls is described in Tables S1 and S2 in Supplementary data, available at JXB online, using the current lexicon. This established rating system is further illustrated for additional detail in micrographs provided in Figs S1 and S2 in the Supplementary data at JXB online.
Water vapour loss
Two methods were used to determine water vapour loss from cut tubers. The first method employed porometric measurement of water vapour conductance from the wound-healing surfaces as outlined by Lulai and Orr (1994, 1995). Field-grown seed tubers (40–85 g) were cut in half, immediately placed in each of the respective treatment solutions (water, ABA, FLD, and ABA and FLD combined) and vacuum infiltrated at –103 kPa (15 psi) for 30 min. Treatment solutions were then drained and fresh solutions added. The tubers were vacuum infiltrated for another 30 min. The treated tubers were then placed in a controlled environment chamber (20 °C, 95% RH) and allowed to wound-heal until measurement. Triplicate water vapour conductance measurements were taken for each tuber half at each time point and the values averaged to provide a single measurement for each tuber. Vapour conductance measurements were determined with a Li-Cor model LI-1600M steady-state porometer (Li-Cor, Inc., Lincoln, NE, USA) as previously described (Lulai and Orr, 1994, 1995). Porometric measurements were corrected for boundary layer resistance as indicated by the manufacturer. This porometric approach provided the most-sensitive means of measuring real time water vapour loss during the first 4 d after wounding. In the 2nd method, mini-tubers were cut in half from stem to bud end. Corresponding halves were treated with water or FLD via vacuum infiltration, similar to that above, and weighed at the indicated time points to determine water loss. This approach allowed indirect measurement of water vapour loss beyond 4 d after wounding.
PAL activity
Tuber discs (4 g; five or six per sample) were frozen in liquid N2 and ground to a powder with a mortar and pestle. The powder was extracted with cold borate buffer (4 ml g–1, 0.025 M, pH 8.8 containing 1 mM dithiothreitol), clarified by centrifugation (10 000 g for 15 min), and the supernatant removed for assay. PAL activity was assayed spectrophotometrically following the formation of (E)-cinnamic acid from L-phenylalanine at 290 nm (
=10 000). The procedure is similar to that of Zucker (1965), but employs continuous spectrophotometric measurement.
Quantitative determination of transesterifiable products (TEPs) during wound-healing
Triplicate samples, three tuber discs per sample, were collected at the indicated time points, weighed and total surface area (mm2) of the discs was calculated. The samples were lyophilized and ground to a fine powder. Lightly soluble polar lipid was extracted with methanol (4 vols) and the powder allowed to dry. Aliquots (25 mg) of the residue were transesterified in 3 ml of 3 M methanolic HCl (Supelco, Bellefonte, PA, USA) with constant stirring (50 min at 50 °C). An internal standard of heptadecanoic acid (18 µg) was included. The residue was allowed to settle and the methanolic mixture of TEP was aspirated and collected. The residue was rinsed twice with 2 ml hexane and the extract combined with the methanolic TEP. The combined mixture of TEP was again extracted three times with 4 ml volumes of hexane. The hexane extracts were concentrated to 0.2 ml under a stream of N2 and the TEP analysed by capillary gas chromatography.
TEP were analysed using a model 5890 Hewlett Packard gas chromatograph equipped with a flame ionization detector and a HP-1/DB-1 capillary column (30 mx0.25 mmx0.25 µm film; J&W Scientific, Folsom, CA, USA). Gas chromatography was carried out under the following conditions: split injection, a helium flow rate of 10 ml min–1, injector temperature of 250 °C and detector temperature of 280 °C. The initial oven temperature of 150 °C was held for 5 min after injection, then linearly increased 5 °C min–1 up to 270 °C and then held for 5 min at 270 °C. Total TEP was quantified, using the heptadecanoic acid internal standard, by determining the total area under all peaks eluted after the solvent. The accumulation of total TEP was then normalized, using an approach similar to that used by Yang and Bernards (2006), by expressing the amount of TEP found per square millimetre of wound surface. Fully processed sample blanks were used to confirm the end point for solvent elution and the starting point for TEP quantitization.
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Results |
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Tuber ABA content and the effect of storage and wounding
Tuber ABA content varied from under 50 ng g–1 FW to
200 ng g–1 FW depending on the time in storage and, to a lesser extent, the type of tuber tissue or anatomical location that was sampled (Fig. 1). Throughout the tuber, ABA content was lowest at harvest then increased 100% (pith) to 300% (periderm and cortical tissue) by the 3rd and 6th month of storage depending on the type of tissue. By the 9th month of storage, tuber ABA content decreased to approximately the levels found at harvest (Fig. 1) and in some samples lower than that at harvest (data not shown). This pattern of changes in ABA content in harvested and stored tubers occurred throughout the tuber, i.e. in all of the different types of tuber tissues analysed. This pattern was found in replicated studies from three different field locations in each of two years. The ABA content throughout the tuber varied little within each sampling time point. However, the trends found throughout storage for all field samples indicated that ABA content was slightly lower in the pith and higher in the area of the periderm and associated cortical tissues.
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The effect of wounding and FLD treatment on ABA content
Wounding had a pronounced effect on tuber ABA content (Fig. 2). The tuber disc model employed wounding on all tissue surfaces of the disc. This extensive wound trauma induced dramatic decreases in ABA content, from 122 ng g–1 FW at the time of wounding to 8–9 ng g–1 FW during the first 1–2 d after wounding. By the 3rd day after wounding, the ABA content had increased and by days 4 and 7 ABA content was over double that of the basal levels found at zero time. Throughout the period of wound-induced increases in ABA content, i.e. days 3–7, FLD, a known inhibitor of de novo ABA biosynthesis, dramatically suppressed ABA accumulations, holding ABA levels close to the minimum observed at 1–2 d after wounding.
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The effect of ABA on wound-induced PAL activity
PAL activity was below detection in non-wounded tissue, but increased greatly by 1 d after wounding (Fig. 3). At day 1, FLD treatment mildly suppressed the wound induction of PAL. FLD treatment suppressed the wound induction of PAL activity by as much as two-thirds in some tuber samples (data not shown) or as little as one-third in others as illustrated in Fig. 3. ABA treatment increased PAL activity above that of FLD-treated tissue and slightly enhanced the induction of PAL above the control at 1 d after wounding. The combining of ABA with the FLD in the treatment solution restored induction of PAL activity to that of the water-treated control (Fig. 3). These effects were ephemeral and little or no differences were observed 3 d after wounding.
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The effect of ABA on wound-induced suberization
Tuber parenchymal tissue had no accumulation of either SPP or SPA at the time of wounding (zero time) (Figs 4, 6). Control tissues, i.e. water-treated discs, began noticeably to accumulate SPP in the developing closing layer by 1 d after wounding. SPP accumulation extended around the entire first layer of existing parenchyma cells, a rating of 5, near or after the 4th day of wound-healing. By the 7th day after wounding, accumulation of SPP in tissues of the water-treated controls extended into the radial walls of the 2nd layer of parenchyma cells (a rating of 6). Although the effects were small, treatment with ABA appeared to slightly enhance SPP accumulation in comparison to the control, especially on days 2 and 3. FLD treatment resulted in two distinct changes involving the induction of closing layer cell walls for SPP accumulation (Fig. 4) and the intensity of cell wall autofluorescence (Fig. 5). FLD treatment appeared to only slightly reduce the induction of cell walls for SPP accumulation throughout most of the time-course, whereas the combining of ABA with the FLD in the treatment solution quantitatively restored induction for SPP accumulation to that of the water-treated control (Fig. 4).
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Although FLD treatment did not result in a large decrease in cell wall induction for SPP accumulation during closing layer development (Fig. 4), the treatment did result in large, reproducible reductions in autofluorescence that were irregularly located on the cell walls and in a slight discoloration of surface cell walls detectable via UV microscopy (Fig. 5, A versus C). Quantitatively, tangential and radial cell walls were clearly induced to accumulate SPP, but the intensity of related cell wall autofluorescence was irregular and diminished, indicative of reduced SPP accumulation (Fig. 5C). This irregular autofluorescence continued beyond the first layer (Fig. 5C) and into the 2nd layer of cells during closing layer development (data not shown), thus suggesting that this irregularity was not totally attributed to desiccation and cell death. The ABA-treated tissues (Fig. 5B) had slightly sharper or brighter autofluorescence than the controls (Fig. 5A). Combining ABA with the FLD treatment qualitatively restored the intensity of cell wall autofluorescence (Fig. 5D) so that it was similar to that of the control (Fig. 5A).
Accumulation of SPA occurs well after that of SPP (Figs 4, 6). There is no detectable SPA accumulation in control tissues until day 3, when traces of barely discernible fluorescence from the histochemical probe were found on some of the cell walls of the developing closing layer (Fig. 6). Further, but still relatively weak, SPA accumulations were detectable on the 1st cell layer at day 4. However, by day 7, the 1st cell layer fluoresced brightly with the full accumulation of SPA (i.e. rating 
5). This easily discernible and bright fluorescence was noted around the entire perimeter of the 1st layer of cells, indicating the completion of accumulation on the 1st layer and that accumulation on the 2nd layer was ready to ensue. Suppression of ABA biosynthesis, via FLD treatment, resulted in a large reduction in SPA accumulation that was especially evident on day 7 where accumulations were less than half that of the water-treated control. Conversely, treatment with ABA significantly increased SPA accumulation at day 3, the time point for the first sign of accumulation for this biopolymer and wound-induced ABA. Although ABA treatment continued to increase SPA accumulation through day 7, the relative increase compared with the control was not as great as at day 3. Inclusion of ABA in the FLD treatment solution increased SPA accumulation well beyond that of the water-treated control at day 3, the time point for the first weakly detectable accumulation. However, by the 4th and 7th day after wounding, FLD treatment combined with ABA treatment resulted in SPA accumulations that were similar to that of the water-treated control. FLD-mediated blockage of ABA biosynthesis resulted in large decreases in SPA accumulation in comparison to those of ABA-treated tissues.
Quantitative determination of TEP, transesterifiable suberin products (TESP), and conjugates
Transesterification of wound-healing tissues liberated TEP putatively consisting of lipid and lipid conjugates present at the time of wounding and during the time-course (Table 1). Little change in TEP was detected until after the 4th day of the wound-healing time-course. Both water-treated and FLD-treated tissues possessed very little TEP, 3–4 µg mm–2 of wound surface, from zero time to the 4th day after wounding. By the 7th day after wounding, large amounts of TEP (15 µg mm–2) had accumulated in control tissues. However, at this same time point, the FLD-treated tissues, i.e. those with suppressed ABA biosynthesis, had accumulated little TEP, 4.8 µg mm–2, nearly the same accumulation as that found in the controls at 0–4 d after wounding. The increase in TEP at the 7th day after wounding (15 µg mm–2–3 µg mm–2=12 µg mm–2) represents the wound-induced increase in lipid/aliphatic material and associated conjugates at that time and is indicative of TESP found in the SPAD.
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Water vapour loss
The accumulative water vapour loss, based on percentage weight loss, increased from
2% at day 2 after wounding to 14% by day 7 after wounding (Table 2). The corresponding tuber halves that were treated with FLD to suppress ABA biosynthesis lost significantly more water vapour throughout the time-course than the control (4.59% and 1766 % at day 2 and day 7, respectively).
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Porometrically determined water vapour loss provided a sensitive real-time means of specifically measuring the differential decrease in the amount of water vapour conducted through the wound surface during initial healing, and facilitated analyses of a full combination of treatments (Fig. 7). Initial water vapour conductances for all treatments at 4 h and 6 h after wounding were high, above 1.8 mol M–2 s–1, respectively, and the measurements for the treatments were somewhat mixed because of the short healing time at this point. However, even at these early time points, when erratic conductances may be obtained, measurements involving ABA treatments were beginning to show less water vapour loss than the controls and FLD-treated tissues. By 1 d after wounding, the vapour conductance values had decreased greatly for all treatments. However, the FLD-treated tissues had notably greater water vapour loss throughout the 1–4 d time-course. Most noticeable, at 1 d after wounding, the vapour conductance of FLD-treated tissues was approximately twice that of water-treated control tissues and three times that of ABA-treated tissues. Although the differences narrowed as wound-healing progressed and approached the asymptote of the vapour conductance curve, the water-treated control and ABA-treated tissues had lower water vapour loss than the FLD-treated tissues throughout the 4 d wound-healing period. Combining ABA with FLD in the treatment restored the biological control mechanism for reduction of water vapour loss.
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Discussion |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionConcluding remarksSupplementary dataReferences |
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The regulatory roles of ABA include many biological processes in a range of plant tissues (Leung and Giraudat, 1998; Marion-Poll and Leung, 2006). However, the basal levels, wound-related changes, and wound-healing involvement of endogenous ABA in potato tuber are poorly understood. Current techniques combined with HPLC-MS analyses provided a means of assessing basal ABA content, wound-related changes in ABA content, and the involvement of ABA in wound-healing. Replicated experiments show that ABA content within the potato tuber is reasonably uniform and that any variation between types of tissues is small (Fig. 1). Periderm and associated cortical tissues have slightly greater ABA content than pith tissues. This differential in ABA content is consistent with gas chromatographic analysis of cortical and pith tissues from small growing tubers (Krauss, 1981). The higher ABA content in tissues near the surface, i.e. the periderm and associated cortical tissues, may be important in regulating wound-healing processes in areas of the tuber that are most vulnerable to injury and intrusion from growth cracks, insects, pathogens, etc. Interestingly, the results further show that ABA content is low at harvest, and increases significantly by the 3rd and 6th month of storage. By the 9th month of storage, ABA content then declines to approximately that found at harvest or less in some cases. These findings show unexpected changes in ABA homeostasis from harvest through storage, i.e. an increase followed by a decrease in basal ABA content. Results differ from earlier studies, which were designed for dormancy analyses and showed a post-harvest decline in basal ABA content (Biemelt et al., 2000). The ABA-dormancy studies employed different genotypes and sampling protocols that were used across a different time-course than the current study. Physiological reasons and implications for the changes in basal ABA content over the 9 month time-course may be hypothesized and varied. For example, the decreased ABA content at 9 months of storage may play a role in the decreased PAL induction documented in aged tubers (Kumar and Knowles, 2003). Also, these changes in tuber ABA content are consistent with reports of ABA involvement in dormancy (Suttle and Hultstrand, 1994; Suttle, 1995; Destefano-Beltran et al., 2006).
Extensive wound trauma, via the tuber disc model, introduced another dynamic where ABA content decreased upon wounding then began to increase by the 2nd or 3rd day (Fig. 2). These results suggest that ABA degradation exceeds biosynthesis for a period after wounding, but that ABA biosynthesis is then induced to exceed degradation and ABA content increases. These conclusions are supported by the inhibitor studies, where FLD treatment blocked biosynthesis of ABA precursors, carotenoids, and effectively held ABA content to near the minima created by the wound-induced degradation of basal ABA. This approach to inhibition of ABA biosynthesis is especially effective in tubers of S. tuberosum because these tissues have a low carotenoid content, are not photoautotrophic, and do not require protection from photo-oxidation (Grappin et al., 2000; Morris et al., 2004). The low carotenoid content combined with FLD-directed blockage of carotenoid biosynthesis facilitates the wound-induced depletion of ABA precursors and effectively inhibits ABA biosynthesis. The ability to inhibit ABA biosynthesis provides a direct means of determining the involvement of endogenous ABA in a range of wound-related reactions including those processes required for suberin accumulation and the reduction of water vapour loss which is controlled by wax accumulation.
Tuber wounding quickly induces processes leading to de novo biosynthesis of PAL (Zucker, 1968; Bernards et al., 2000) which catalyses the first committed step in phenylpropanoid biosynthesis required for SPP accumulation (Bernards, 2002). Limited PAL activity exists prior to induction by wounding. The amount of wound-induced PAL activity is dependent upon tuber age and decreases with the length of time that potatoes are held in storage (Kumar and Knowles, 2003). Rapid wound-responses result in the biosynthesis of measurable amounts of PAL by the 1st day and through the 3rd day after wounding (Fig. 3). The involvement of ABA in PAL regulation was suggested by the reduction in PAL activity via FLD-mediated blockage of ABA biosynthesis and the reversal of this decrease by ABA supplementation. Notably, 1 d after wounding the difference in activity between FLD- and ABA-treated tissues is significant. However, the blockage of ABA biosynthesis did not dramatically decrease PAL induction. Combined, these results suggest a possible role for basal ABA, or other regulatory mechanism(s), in mediating rapid induction for PAL biosynthesis directly upon wounding.
SPP accumulation on closing-layer cell walls of tissues treated with FLD, ABA, and ABA combined with FLD were consistent with that of PAL inductions from corresponding treatments (Fig. 4). These roughly parallel responses are consistent with the critical role of PAL induction and associated phenylpropanoid biosynthesis in construction of the SPPD. Again, these results indicate a role for basal ABA or other regulatory mechanism(s) in the rapid induction and accumulation of SPP that were not affected by FLD treatment. The blockage of ABA biosynthesis also resulted in irregular fluorescence on the cell walls in the wound-induced closing layer (Fig. 5). Some segments of cell walls in the developing closing layer fluoresced dimly, showing that they were induced to accumulate SPP, but the dim or nearly muted fluorescence indicated that the SPP accumulation was low or did not occur. Other neighbouring cell walls appeared to fluoresce more normally, indicating SPP accumulation. Restoration of SPP accumulation by inclusion of ABA confirmed involvement in this phase of suberization (Fig 5D). These results are the first known reported indications of quantitative and qualitative differences for the induction and accumulation of SPP on closing layer cell walls.
The time-course for SPA accumulation is distinct from that of SPP and, depending on the genotype and wound conditions, begins about 2 d later as indicated here and in earlier studies (Lulai and Corsini, 1998; Lulai, 2007). These distinctions suggest that SPP and SPA formation are co-ordinately regulated. The later induction of SPA accumulation during the wound-healing time-course is more dramatically affected by addition of ABA and blockage of ABA biosynthesis, indicating that wound-induced ABA biosynthesis, and not basal ABA, is primarily involved in regulation of SPA accumulation (Fig. 6). Also, SPA accumulation cannot occur, even with addition of ABA, until the cell walls are fully primed for assembly and covalent lamination of aliphatic monomers. After the cells are primed to accept assembly and lamination of the aliphatic monomers, ABA treatment significantly enhanced SPA accumulation, as noted at and after the 3rd day after wounding. These observations are consistent with the changes in ABA content found in corresponding wounded tissues and the FLD-mediated inhibition of ABA biosynthesis in these tissues during the same time period. Most important, the pronounced suppression of suberization, as a result of FLD-mediated blockage of ABA biosynthesis, is reversed by inclusion of ABA in the FLD treatment solution. This reversal confirms that endogenous ABA plays a regulatory role in suberization and indicates that wound-induced ABA biosynthesis, and not basal ABA, is required for regulation of SPA accumulation. The wound-induced biosynthesis of TESP and inhibition of TESP accumulation in tissues with blocked ABA biosynthesis are consistent with the cellular data and further confirm the involvement of ABA in regulating suberization (Table 1). This regulation of SPA accumulation is important because the SPAD forms the final surface barrier to fungal infection (Lulai and Corsini, 1998) and, recently, the formation of SPA barriers has been shown to block fungal advancements deep within the parenchyma of infected tubers and to reduce fungal infection in soybean roots (Lulai et al., 2006; Thomas et al., 2007). SPP and SPA accumulations are co-ordinated. Results clearly show that ABA plays a role in regulating these accumulations, but that other regulatory mechanisms may also be involved.
A rapid biological response to reduce water vapour loss at wound sites is crucial in protecting exposed cells from desiccation and death. A desiccated layer of dead or incompetent cells will not protect the tuber from pathogens and will not suberize (Lulai et al., 2006; Lulai, 2007). Soluble waxes integrated into the suberin matrix are largely responsible for formation of the barrier that controls water vapour loss and protects cells from desiccation (Soliday et al., 1979). Although weight loss is often used as an accumulative indicator of water vapour loss, porometry is a more sensitive means of determining real-time water vapour conductance/loss during the 1st and 2nd days after wounding and can provide useful information up to the 4th day after wounding (Lulai and Orr, 1995). Exogenous ABA had been shown to accelerate development of resistance to wound-related water vapour loss (Lulai and Orr, 1995); however, the regulatory involvement of this hormone had not been unequivocally determined via reduced ABA concentrations and determination of resulting effects. The current study confirms that addition of ABA enhances the rate of reduction of water vapour loss during wound-healing, and that the largest reductions in water vapour loss occur prior to significant SPP and SPA accumulations. These results are consistent with work which showed that rapid accumulation of long chain waxes control water vapour loss (Soliday et al., 1979; Schreiber et al., 2005). FLD-mediated blockage of ABA biosynthesis retarded the processes protecting against water vapour loss by 50% compared with that of controls, most noticeably at day 1, but continuing to the 3rd day of wound-healing (Fig. 7). Importantly, this retardation of reduction in water vapour loss was reversed by including ABA with the FLD treatment, thus confirming involvement of the hormone in regulation of tuber water vapour loss during wound-healing. The results also suggest that the remaining uninhibited reduction in water vapour loss in FLD-treated tissues was regulated by basal ABA or other mechanisms. The immediate availability of basal ABA in conjunction with readily synthesized ABA and/or other constitutive regulatory mechanisms are necessary for rapid activation and biosynthesis of the water vapour barriers to protect the cells from dehydration and death. Weight loss measurements showed that the effect of blockage of ABA biosynthesis on reduction of water vapour loss continued through days 4 and 7 after wounding (Table 2); these data are consistent with the results of Schreiber et al. (2005) who showed that wax deposition continued well after water vapour loss had reached minimum levels. Collectively, these results are consistent with the relationship between loss of turgor and ABA biosynthesis in other plant tissues (Marion-Poll and Leung, 2006). The results from this research may be used to raise questions about regulation of the elevated water vapour loss in tubers with suberin-related periderm disorders, such as tuber pink eye, and suggest that the cellular response for wax deposition to protect against dehydration in these tissues is impaired or not induced (Lulai et al., 2006). The question then arises as to whether or not ABA concentrations in these tissues are below some threshold required to regulate water vapour conductance and suberin biosynthesis or if some other signal is also required to induce and/or regulate these processes.
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Concluding remarks |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionConcluding remarksSupplementary dataReferences |
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In summary, results from the addition of ABA and inhibition of ABA biosynthesis on the respective increase and retardation of suberization and the development of resistance to water vapour loss suggest critical regulatory roles for endogenous ABA in these processes. The reversal of this retardation by addition of ABA clearly proves that ABA is involved as a regulatory agent in these wound-healing processes. Importantly, wounding induces tangential and radial cell walls for closing-layer development. SPP accumulation on the induced cell walls involves regulation by wound-induced ABA and possibly basal ABA and other mechanisms, whereas ABA involvement in SPA accumulation is dependent on wound-induced ABA. This proof and description of ABA involvement in these wound-healing processes is also of great importance in addressing regulatory issues for a range of suberization-related physiologies, including those induced by pathogens and physiological disorders (Lulai, 2005; Lulai et al., 2006).
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Supplementary data |
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The following supplementary material is available at JXB online.
Table S1. Rating system for the accumulation of the suberin poly(phenolic) (SPP) domain as it develops on suberizing cell walls during wound-healing.
Table S2. Fluorescence rating system for the accumulation of the suberin poly(aliphatic) (SPA) domain as it develops on suberizing cell walls during wound-healing.
Figure S1. Examples of SPP accumulation ratings using guidelines from Table S1.
Figure S2. Examples of SPA accumulation ratings using guidelines from Table S2.
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Acknowledgements |
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The technical assistance of Ms Linda Huckle and statistical analysis by Dr Larry Campbell and Mr Mark West are gratefully acknowledged.
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Footnotes |
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Mention of company or trade name does not imply endorsement by the United States Department of Agriculture over others not named.
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Abbreviations |
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ABA, abscisic acid; ANOVA, analysis of variance; FLD, fluridone; HPLC-MS, high-performance liquid chromatography–mass spectrometry; LSD, least squares difference; PAL, phenylalanine ammonia lyase; SE, standard error; SPA, suberin poly(aliphatic(s)); SPAD, suberin poly(aliphatic) domain; SPP, suberin poly(phenolic(s)); SPPD, suberin poly(phenolic) domain; TEP, transesterifiable products; TESP, transesterifiable suberin products.
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References |
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Bernards MA. Demystifying suberin. Canadian Journal of Botany (2002) 80:227–240.
Bernards MA, Susag LM, Bedgar DL, Anterola AM, Lewis NG. Induced phenylpropanoid metabolism during suberization and lignification: a comparative analysis. Journal of Plant Physiology (2000) 157:601–607.[ISI][Medline]
Biemelt S, Hajirezaei M, Hentschel E, Sonnewald U. Comparative analysis of abscisic acid content and starch degradation during storage of tubers harvested from different potato varieties. Potato Research (2000) 43:371–382.[CrossRef][ISI]
Cottle W, Kolattukudy PE. Abscisic acid stimulation of suberization: induction of enzymes and deposition of polymeric components and associated waxes in tissue cultures of potato tuber. Plant Physiology (1982) 70:775–780.
Destefano-Beltran L, Knauber D, Huckle L, Suttle J. Effects of postharvest storage and dormancy status on ABA content, metabolism, and expression of genes involved in ABA biosynthesis and metabolism in potato tuber tissues. Plant Molecular Biology (2006) 61:687–697.[CrossRef][ISI][Medline]
Efetova M, Zeier J, Riederer M, Lee C-W, Stingl N, Mueller M, Hartung W, Hedrich R, Deeken R. A central role of abscisic acid in drought stress protection of Agrobacterium-induced tumors on Arabidopsis. Plant Physiology (2007) 145:853–862.
Espelie KE, Kolattukudy PE. Purification and characterization of an abscisic acid-inducible anionic peroxidase associated with suberization in potato (Solanum tuberosum). Archives of Biochemistry and Biophysics (1985) 240:539–545.[CrossRef][ISI][Medline]
Gamble PE, Mullet JE. Inhibition of carotenoid accumulation and abscisic acid biosynthesis in fluridone-treated dark-grown barley. European Journal of Biochemistry (1986) 160:117–121.[ISI][Medline]
Graca J, Pereira H. Suberin structure in potato periderm: glycerol, long-chain monomers, and glyceryl and feruloyl dimers. Journal of Agriculture and Food Chemistry (2000) 48:5476–5483.[CrossRef]
Graca J, Santos S. Suberin: a biopolyester of plants' skin. Macromolecular Bioscience (2007) 7:128–135.[CrossRef][ISI][Medline]
Grappin P, Bouinot D, Sotta B, Miginiac E, Jullien M. Control of seed dormancy in Nicotiana plumbaginifolia: post-imbibition abscisic acid synthesis imposes dormancy maintenance. Planta (2000) 210:279–285.[CrossRef][ISI][Medline]
Himmelbach A, Yang Y, Grill E. Relay and control of abscisic acid signaling. Current Opinion in Plant Biology (2003) 6:470–479.[CrossRef][ISI][Medline]
Krauss A. Abscisic and gibberellic acid in growing potato tubers. Potato Research (1981) 24:435–439.[CrossRef][ISI]
Kumar GN, Knowles NR. Wound-induced superoxide production and PAL activity decline with potato tuber age and wound healing ability. Physiologia Plantarum (2003) 117:108–117.[CrossRef]
Leung J, Giaudat J. Abscisic acid signal transduction. In: Annual Review of Plant Physiology and Plant Molecular Biology—Jones RL, Somerville CR, Walbot V, eds. (1998) 49:198–222.
Lulai EC. Non-wound induced suberization of tuber parenchyma cells: a physiological response to the wilt disease pathogen Verticillium dahliae. American Journal of Potato Research (2005) 82:433–440.
Lulai EC. Skin-set, wound-healing and related defects. In: Potato biology and biotechnology: advances and perspectives—Vreugdenhil D, ed. (2007) Amsterdam: Elsevier. 471–496.
Lulai EC, Corsini DL. Differential deposition of suberin phenolic and aliphatic domains and their roles in resistance to infection during potato tuber (Solanum tuberosum L.) wound-healing. Physiological and Molecular Plant Pathology (1998) 53:209–222.[CrossRef][ISI]
Lulai EC, Morgan WC. Histochemical probing of potato periderm with neutral red: a sensitive cytofluorochrome for the hydrophobic domain of suberin. Biotechnic Histochemistry (1992) 67:185–195.[CrossRef][ISI][Medline]
Lulai EC, Orr PH. Techniques for detecting and measuring developmental and maturational changes in tuber native periderm. American Potato Journal (1994) 71:489–505.[CrossRef][ISI]
Lulai EC, Orr PH. Porometric measurements indicate wound severity and tuber maturity affect the early stages of wound-healing. American Potato Journal (1995) 72:225–241.[CrossRef][ISI]
Lulai EC, Suttle JC. The involvement of ethylene in wound-induced suberization of potato tuber (Solanum tuberosum L.): a critical assessment. Postharvest Biology and Technology (2004) 34:105–112.[CrossRef][ISI]
Lulai EC, Weiland JJ, Suttle JC, Sabba RP, Bussan AJ. Pink eye is an unusual periderm disorder characterized by aberrant suberization: a cytological analysis. American Journal of Potato Research (2006) 83:409–421.
Marion-Poll M, Leung J. Abscisic acid synthesis, metabolism and signal transduction. In: Plant hormone signaling. Annual Plant Reviews—Hedden P, Thomas TG, eds. (2006) 24. Oxford: Blackwell Publishing. 1–35.
Morris WL, Ducreus L, Griffiths DW, Stewart D, Davies HV, Taylor MA. Carotenogenesis during tuber development and storage in potato. Journal of Experimental Botany (2004) 55:975–982.
Schreiber L, Franke R, Harmann K. Wax and suberin development of native and wound periderm of potato (Solanum tuberosum L.) and its relation to peridermal transpirations. Planta (2005) 220:520–530.[CrossRef][ISI][Medline]
Soliday C, Dean BB, Kolattukudy PE. Suberization: inhibition by washing and stimulation by abscisic acid in potato disks and tissue culture. Plant Physiology (1978) 61:170–174.
Soliday CL, Kolattukudy PE, Davis RW. Chemical and ultrastructural evidence that waxes associated with the suberin polymer constitute the major diffusion barrier to water vapor in potato tuber (Solanum tuberosum L.). Planta (1979) 146:607–614.[CrossRef][ISI]
Suttle JC. Postharvest changes in endogenous ABA levels and ABA metabolism in relation to dormancy in potato tubers. Physiologia Plantarum (1995) 95:233–240.[CrossRef]
Suttle JC, Hultstrand JF. Role of endogenous abscisic acid in potato microtuber dormancy. Plant Physiology (1994) 105:891–896.[Abstract]
Thomas R, Fang X, Ranathunge K, Anderson TR, Peterson CA, Bernards MA. Soybean root suberin: anatomical distribution, chemical composition, and relationship to partial resistance to Phytophthora sojae. Plant Physiology (2007) 144:299–311.
Vogt E, Schönherr J, Schmidt HW. Water permeability of periderm membranes isolated enzymatically from potato tubers (Solanum tuberosum L.). Planta (1983) 158:294–301.[CrossRef][ISI]
Yang W-L, Bernards MA. Wound-induced metabolism in potato (Solanum tuberosum) tubers: biosynthesis of aliphatic domain monomers. Plant Signaling and Behavior (2006) 1:59–66.
Zucker M. Induction of phenylalanine deaminase by light and its relation to chlorogenic acid synthesis in potato tuber tissue. Plant Physiology (1965) 40:779–784.
Zucker M. Sequential induction of phenylalanine ammonia-lyase and a lyase-inactivating system in potato tuber disks. Plant Physiology (1968) 43:365–374.
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