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Background and aims Many fruits soften during ripening, which is important commercially and in rendering the fruit attractive to seed-dispersing animals. Cell-wall polysaccharide hydrolases may contribute to softening, but sometimes appear to be absent. An alternative hypothesis is that hydroxyl radicals (•OH) non-enzymically cleave wall polysaccharides. We evaluated this hypothesis by using a new fluorescent labelling procedure to ‘fingerprint’ •OH-attacked polysaccharides.
Methods We tagged fruit polysaccharides with 2-(isopropylamino)-acridone (pAMAC) groups to detect (a) any mid-chain glycosulose residues formed in vivo during •OH action and (b) the conventional reducing termini. The pAMAC-labelled pectins were digested with Driselase, and the products resolved by high-voltage electrophoresis and high-pressure liquid chromatography.
Key Results Strawberry, pear, mango, banana, apple, avocado, Arbutus unedo, plum and nectarine pectins all yielded several pAMAC-labelled products. GalA–pAMAC (monomeric galacturonate, labelled with pAMAC at carbon-1) was produced in all species, usually increasing during fruit softening. The six true fruits also gave pAMAC·UA-GalA disaccharides (where pAMAC·UA is an unspecified uronate, labelled at a position other than carbon-1), with yields increasing during softening. Among false fruits, apple and strawberry gave little pAMAC·UA-GalA; pear produced it transiently.
Conclusions GalA–pAMAC arises from pectic reducing termini, formed by any of three proposed chain-cleaving agents (•OH, endopolygalacturonase and pectate lyase), any of which could cause its ripening-related increase. In contrast, pAMAC·UA-GalA conjugates are diagnostic of mid-chain oxidation of pectins by •OH. The evidence shows that •OH radicals do indeed attack fruit cell wall polysaccharides non-enzymically during softening in vivo. This applies much more prominently to drupes and berries (true fruits) than to false fruits (swollen receptacles). •OH radical attack on polysaccharides is thus predominantly a feature of ovary-wall tissue.
Fruit ripening is often accompanied by changes in flavour, odour, colour and texture which are attractive to the animals that will disperse the seeds. In particular, many berries, drupes and pomes soften during ripening owing to changes in cell wall organization. Plant cell walls are complex networks based on cellulose microfibrils, partly tethered by hemicelluloses, with pectic polysaccharides infiltrating the rest of the wall matrix (Albersheim et al., 2010; Fry, 2011a). During fruit softening, the matrix polysaccharides, especially the pectins, often become more readily extractable and/or decrease in molecular weight, indicating depolymerization, e.g. in avocado, plum, mango, banana and tomato (Huber and O’Donoghue, 1993; Prasanna et al., 2003; Ali et al., 2004; Ponce et al., 2010; Basanta et al., 2014). The importance of pectic depolymerization has led to the widely held view that ripening can be regarded as principally a ‘hydrolytic’ process.
Earlier, however, Blackman and Parija (1928) had suggested that ripening involves a loss of ‘organisational resistance’, i.e. fruit cells lose the ability to maintain separate compartments owing to cellular (membrane) degeneration. Although this concept lost popularity, some workers continued to interpret ripening as a form of senescence attributable to oxidation reactions (Brennan and Frenkel, 1977) and more recently to emphasize the (possibly related) decrease in water content that occurs near the onset of ripening (Frenkel and Hartman, 2012). There are indeed similarities between physiological changes (e.g. chlorophyll loss and membrane permeabilization) occurring in a ripening fruit and in a leaf or petal approaching abscission. Lipoxygenases, which often increase during ripening (Ealing, 1994; de Gregorio et al., 2000), generate hydroperoxide groups (>CH–OOH) in unsaturated fatty acid residues, accompanied by the formation of reactive oxygen species (ROS). Such lipid oxidation may permeabilize membranes, resulting in the release of certain metabolites, e.g. ascorbate (Dumville and Fry, 2003), into the apoplast (the aqueous solution that bathes the cell wall), and ROS by-products may drive other oxidative reactions.
Viewing fruit ripening as an ‘oxidative’ process is supported by evidence from several quarters. For example, in avocado (Lauraceae; Meir et al., 1991) and serviceberry (Rosaceae; Rogiers et al., 1998), lipid peroxidation was the earliest symptom of ripening, and tomato (Solanaceae) fruit ripening was accompanied by elevated H2O2 and the oxidation of lipids and proteins (Jimenez et al., 2002). In the present study, we propose a link between oxidative agents (especially the hydroxyl radical, •OH) and pectic polysaccharide degradation in softening fruit.
Primary cell walls control the texture of fruit tissues. Although strong enough to withstand turgor pressure, walls are dynamic structures in which the polymers can be remodelled or degraded, resulting in wall loosening. Many enzymes and expansins have been described that act on cell-wall polymers, and the expression of these proteins has been correlated with fruit softening as well as cell expansion and abscission (reviewed by Franková and Fry, 2013). For example, glycanases and transglycanases cleave cell wall polysaccharides in mid-chain (Taylor et al., 1993; Bewley, 1997; Lazan et al., 2004; Fry et al., 2008; Schröder et al., 2009; Franková and Fry, 2011; Derba-Maceluch et al., 2014), glycosidases release mono- or disaccharides from non-reducing termini (Fanutti et al., 1991; de Veau et al. 1993; Hrmová et al., 1998; de Alcântara et al., 2006; Franková and Fry, 2012), and expansins interfere in polysaccharide–polysaccharide hydrogen bonding (Cosgrove, 2000; Harada et al., 2011; Sasayama et al., 2011).
Several polymer-hydrolysing enzymes have been studied in relation to fruit softening, with tomato as the most extensively studied system (Matas et al., 2009; reviewed by Fry, 2017). Several wall polysaccharide-modifying enzyme activities increase, especially endo-polygalacturonase (endo-PG), cellulase, xyloglucan endotransglucosylase, β-galactosidase, pectin-methylesterase and pectate lyase. Although the link between wall-hydrolysing enzymes and fruit softening seems intuitive, tests of this as a functional relationship have often yielded contradictory evidence. A major focus has been endo-PG in tomato. This enzyme is abundant in ripe tomato fruit (Tucker and Grierson, 1982), and suppression of its expression resulted in reduced depolymerization of pectin (Smith et al., 1990). Also, expression of endo-PG in the rin (ripening inhibitor) mutant caused an increased degradation of fruit pectins (Giovannoni et al., 1989). However, both these studies failed to show a related change in tomato fruit softening; no inhibition of softening was observed in endo-PG antisense fruit, and no effect on softening was induced by the expression of endo-PG in the rin mutant. Moreover, other fruits, e.g. strawberry (Pose et al., 2013), persimmon (Cutillas-Iturralde et al., 1993) and kiwifruit (Redgwell et al., 1991), show extensive pectin solubilization and/or a decrease in molecular weight even though they possess very low levels of endo-PG (e.g. Nogata et al., 1993). These observations reinforce the idea that endo-PG is not necessary for fruit softening. The proposed relationship between pectin depolymerization and fruit softening was thus not strongly supported by data, although the excessive softening associated with over-ripening can be prevented by knocking out endo-PG (in the ‘Flavr Savr’ tomato; reviewed by Krieger et al., 2008). Ripening is a robust phenomenon: knocking out any individual player (e.g. endo-PG) often fails to prevent normal softening.
In addition to proteins that remodel the wall, the highly reactive hydroxyl radical (•OH) can cause polysaccharide chain scission non-enzymically. This phenomenon is readily demonstrated in solutions of purified cell wall polysaccharides upon treatment with ascorbate in the presence of O2 plus traces of Cu2+ or Fe3+ (Fry, 1998; Yamazaki et al., 2003; Schweikert et al., 2000, 2002) and in food-related systems (Fauré et al., 2012; Mäkinen et al., 2012; Iurlaro et al., 2014).
•OH can also cleave insoluble polysaccharides that are present in situ as structural components of the cell wall: for example, when •OH was generated within the cell walls of a frozen/thawed maize coleoptile that was being held under tension, the coleoptile extended in a fashion similar to that induced by certain wall-acting proteins or in response to in-vivo auxin treatment (Schopfer, 2002). Likewise, in-vitro •OH treatment of fruit cell walls of tomato (Dumville and Fry, 2003), banana (Cheng et al., 2008b) and longan (Duan et al., 2011) promoted pectin solubilization and depolymerization.
These results indicate that •OH, if formed in the cell wall in vivo, could potentially cleave polysaccharides and thereby exert physiological effects. Indeed, it has been suggested that wall loosening induced by ROS (especially •OH) contributes to fruit ripening (Brennan and Frenkel, 1977; Fry, 1998; Fry et al., 2001; Dumville and Fry, 2003; Cheng et al., 2008a; Yang et al., 2008; Duan et al., 2011), germination (Müller et al., 2009), cell expansion (Schopfer, 2001, 2002; Rodríguez et al., 2002; Liszkay et al., 2004) and abscission (Sakamoto et al., 2008; Cohen et al., 2014). It is sometimes asserted that •OH, as a highly reactive ROS, must be biologically detrimental – for example causing mutations, membrane damage and protein denaturation – and that it would be advantageous for cells to prevent •OH formation or to scavenge it. However, the half-life of •OH within a cellular environment such as a cell wall is estimated at approx. 1ns, allowing it to diffuse no more than approx. 1nm (the length of two glucose residues in a cellulose chain) before reacting with some organic molecule (Griffiths and Lunec, 1996): a very short distance in the context of a primary cell wall, which is typically >80nm thick. Therefore, if produced at an appropriate site (within the cell wall matrix or middle lamella), •OH may have little effect on the protoplast. Moreover, in the case of a softening fruit pericarp, the cells involved are shortly destined to die in an animal’s gut, fulfilling their role in promoting seed dispersal. Therefore, any cellular damage caused to the ripe pericarp by •OH is irrelevant; likewise in other short-lived tissues such as a lysing abscission zone or a rapidly expanding coleoptile.
The production of •OH in plant cell walls most probably involves a Fenton-like reaction, whereby a transition metal ion in the reduced state reacts with hydrogen peroxide (H2O2):
Two proposals have been considered: (1) the transition metal is the Fe of the haem group in peroxidase, which can be reduced by superoxide in a Haber–Weiss cycle (Chen and Schopfer, 1999; Liszkay et al., 2003); and (2) a wall-bound transition metal (Cu and/or Fe) ion is reduced by apoplastic electron donors such as ascorbate (Fry, 1998; Vreeburg and Fry, 2005; Green and Fry, 2005; Kärkönen and Fry, 2006; Lindsay and Fry, 2007; Padu et al., 2005). The H2O2 may be generated by wall-bound oxidases (Lane et al., 1993; Asthir et al., 2002; Kärkönen et al., 2009) or superoxide dismutase (Yim et al., 1990; Ogawa et al., 1996; Kukavica et al., 2009), or by non-enzymic reduction of O2 by ascorbate (Fry, 1998). Indeed, Dumville and Fry (2003) showed that the ability of cells in a tomato fruit to secrete ascorbate, and also the tissue’s Cu content, increased during ripening: effects that would be expected to favour in-vivo •OH production.
Cheng et al. (2008a) showed that, when homogenates of frozen banana pulp harvested at different stages of ripening were incubated for 12h in phosphate buffer containing deoxyribose, in-situ generated •OH (detected by its ability to oxidize the deoxyribose to dialdehyde products) increased in parallel with softening, suggesting that ripening may be associated with •OH production in banana. Yang et al. (2008), who applied a similar method but with a shorter incubation period, also suggested that •OH production increases prior to the initiation of banana fruit softening. However, in both these studies, the source of the •OH in pulp was not clear and it could have been an artefact due to the homogenization, not reflecting reactions that occur in vivo.
•OH can cleave wall polysaccharides in vitro, but the question of whether •OH is produced in the apoplast of living tissue and actually acts in vivo on wall polysaccharides in the manner proposed is still open, a major challenge being to detect such a short-lived free radical as •OH in the walls of living cells. There are two possible experimental approaches: (1) infiltration into the apoplast of a membrane-impermeant ‘reporter’ compound that reacts with •OH to give recognizable products (Kuchitsu et al., 1995; Fry et al., 2002; Schopfer et al., 2002; Miller and Fry, 2004; Müller et al., 2009; for a review, see Vreeburg and Fry, 2005); and (2) detection of the ‘collateral damage’ done to wall polysaccharides in vivo when attacked by apoplastic •OH.
The second approach is based on the fact that the •OH radical cleaves polysaccharides by rather indiscriminate oxidative reactions. •OH-driven polysaccharide scission, proposed to contribute to fruit softening, is accompanied by concurrent reactions that introduce relatively stable oxo groups into the polysaccharide (‘collateral damage’; Fig. 1A) without necessarily cleaving it (Zegota and von Sonntag, 1977; von Sonntag 1980; Vreeburg and Fry, 2005; Vreeburg et al., 2014). Such oxo groups can serve as a chemical ‘fingerprint’ revealing recent •OH attack in the cell walls of living cells. A polysaccharide usually has only a single oxo group (its reducing terminus), but •OH attack generates oxo groups in mid-chain sugar residues, converting them to glycosulose residues (Vreeburg et al., 2014). The proportion of such glycosulose residues (non-terminal oxo groups) per 1000 sugar residues would be a valuable measure of the extent of •OH attack in vivo. Two methods are currently available for their detection:
We have used reductive tritiation with NaB3H4 to detect oxo group formation in Fenton-treated soluble polysaccharides in vitro and in presumptively •OH-exposed cell walls in vivo in ripening pears, germinating cress seeds and elongating maize coleoptiles (Fry et al., 2001, 2002; Miller and Fry, 2001; Müller et al., 2009; Iurlaro et al., 2014). In this approach, each mid-chain glycosulose residue, formed by •OH action, is reduced to yield a tritium-labelled simple sugar (aldose) residue – in some cases an aldose that does not frequently occur naturally. For example, •OH attack at carbon-3 of a xylose residue in xyloglucan will introduce an oxo group which, when treated with NaB3H4 and then acid-hydrolysed, yields a mixture of [3H]xylose and [3H]ribose (the 3-epimer of xylose); the latter is not a natural constituent of xyloglucan and is therefore a highly diagnostic fingerprint (Miller and Fry, 2001).
As an alternative to radiolabelling, we recently developed a method for fluorescently labelling mid-chain (•OH generated) oxo groups present in polysaccharides by reductive amination with 2-aminoacridone (AMAC) plus NaCNBH3, followed by N-isopropylation, to introduce fluorescent 2-isopropylaminoacridone (pAMAC) groups into the polysaccharide (Fig. 1B). The method was developed by model experiments on soluble pectic polysaccharides in vitro (Vreeburg et al., 2014). Upon subsequent Driselase digestion (Fig. 1C), the mid-chain glycosulose residues, indicative of recent •OH attack, were released as various products, of particular diagnostic value being pAMAC·disaccharide conjugates. [Note on nomenclature: The designation ‘sugar–pAMAC’ implies that the pAMAC group is linked to the former reducing group (carbon-1 in the case of an aldose) of the sugar, whereas ‘pAMAC·sugar’ indicates that the pAMAC group is attached to a different carbon of the sugar, whose C-1 remains unlabelled.] In contrast, the single reducing terminal oxo group of a poly- or oligosaccharide was released as a monosaccharide–pAMAC product. We now report the application of the pAMAC/Driselase method to demonstrate the changing abundance of •OH-attacked polysaccharides in the cell walls of various contrasting fruits during ripening to give an indication of the involvement of hydroxyl radical attack in fruit softening.
2-Aminoacridone was from Fluka (Dorset, UK). Driselase, from Sigma-Aldrich (Dorset, UK), was purified by ammonium sulphate precipitation and gel-permeation chromatography (Fry, 2000). The Luna C18 high-pressure liquid chromatography (HPLC) column [250 × 4·6mm, 5μm C18(2) 100 Å] was from Phenomenex (Cheshire, UK). The HPLC eluents were from VWR (Leicestershire, UK) or Fisher Scientific (Loughborough, UK). All other reagents were from Sigma-Aldrich or Fisher Chemicals. The PCE-PTR 200 penetrometer was from PCE Instruments UK Ltd (Southampton, UK).
Pear (Pyrus communis L.), mango (Mangifera indica L.), banana (hybrid based on Musa acuminata Colla), apple (Malus pumila Mill.), avocado (Persea americana Mill.), plum (Prunus domestica L.) and nectarine [Prunus persica (L.) Batsch] were from Sainsbury’s supermarket, Edinburgh; in each case, hard fruit not yet ready for eating were selected. Strawberry [Fragaria×ananassa (Weston) Duchesne ex Rosier (pro sp.)] was from Belhaven Fruit Farm, Dunbar, UK, and strawberry tree (Arbutus unedo L.) berries were generously provided by Sheffield Botanical Garden, UK. Fragaria and Arbutus fruit at three stages of ripening, distinguished by colour, were picked on the same day.
Fluorescent markers for high-voltage paper electrophoresis (HVPE) and HPLC were prepared as before (Vreeburg et al., 2014). In brief, a reducing sugar (0·4μmol of dry glucose, GalA, GalA2, GalA3 or GalA4) was suspended in 40μL of 0·1m AMAC in dimethylsulphoxide (DMSO)/acetic acid/pyridine (17:2:1, by vol.) followed immediately by 40μL of fresh aqueous 1m NaCNBH3. After incubation of the mixture at 20°C for 16h, 2μL of acetone and 40μL of fresh 1m NaCNBH3 were added and the mixture was incubated for another 1h at 20°C. The mixture was diluted with 5 vols of H2O and centrifuged (14000rpm, 10min). The sugar–pAMAC product in the supernatant was purified on a C18 cartridge.
Except for strawberry and Arbutus, freshly purchased hard fruits were stored in the dark in a wooden cupboard at room temperature in the laboratory. On selected days after purchase (when the fruits were hard, medium and soft, respectively), firmness was measured.
For firmness measurements, three individual fruit from each stage were randomly selected. Except with strawberry, the ‘skin’ was peeled. A 6mm diameter penetrometer probe was positioned perpendicular to the peeled fruit surface, and the sensor was pressed down until it penetrated to the sensor’s indicator mark; the force shown on the display (in Newtons) was recorded.
A portion (10g f. wt) of the edible part of each fruit was diced with a razor blade, immediately frozen with liquid N2 in a mortar, and ground to a fine powder with a pestle. Pre-cooled extractant (50mL; ethanol/pyridine/acetic acid/water, 75:2:2:21 by vol., containing 10mm Na2S2O3 to prevent Cu- or Fe-dependent •OH production by Fenton reactions; Fry, 1998) was added and the mixture was ground again in the mortar for another 5min. Finally, the whole homogenate was dispensed as 50 aliquots (each approx. 1·1–1·2mL, equivalent to 200 mg f. wt of fruit tissue), which were stored at –80°C.
Arbutus berries at different stages of ripening (orange, red and red–black) were immediately frozen at –80°C and later homogenized as described above.
All AMAC work was done under subdued red light; ice-cold solvents were used for the washing and precipitation steps. A portion of fruit AIR (alcohol-insoluble residue) suspension (200mg fresh fruit tissue) was thawed and centrifuged. The pellet was washed twice with 75 % ethanol, blotted to remove free ethanol, and resuspended in 261μL of a mixture comprising 45μL of 0·5 % aqueous chlorobutanol, 5μL of pyridine/acetic acid/water (2:2:1 by vol.; final pH approx. 4·0), 89μL of DMSO containing 8·9μmol AMAC and 61μL of water containing 122μmol freshly dissolved NaCNBH3; and the mixture was left for 20h at 20°C. Acetone (136μmol) and an additional 122μmol of fresh NaCNBH3 (61μL of a 2m aqueous solution) were added and the incubation was repeated for 16h at 20°C. To remove low molecular weight reagents and by-products, we added 1mL of 96 % ethanol (to precipitate any water-soluble polysaccharides), pelleted the total polymers (at 12000g for 5min), and washed the pellet twice with 1mL of 75 % ethanol. The pellet was then re-suspended by shaking in 250μL of pyridine/acetic acid/water (1:1:98 by vol.) for 10min at 20°C. The treatments with 96 and 75 % ethanol were repeated, and the final ethanolic pellet of pAMAC-labelled AIR was blotted to semi-dryness.
The blotted pellet of pAMAC-labelled AIR was de-lactonized with 100μL of 0·5m NaOH (50μmol) for 5h at 20°C, then buffered to pH 4·7 with two molar equivalents (5·75 μL) of acetic acid, washed with ice-cold 80 % ethanol, pelleted at 12000g for 5min, blotted with filter paper and immediately treated with Driselase. [The de-lactonization step facilitated subsequent Driselase digestion.] The blotted, de-lactonized pellet of pAMAC-labelled AIR (equivalent to 200 mg of fresh weight fruit) was digested in 500μL of 1 % partially purified Driselase in pyridine/acetic acid/0·5 % chlorobutanol, 1:1:98, by vol.) at 37°C for 14d, after which the solution was frozen at –20°C.
A C18-silica cartridge (500mg Supelco column; Sigma-Aldrich) was pre-conditioned with 2 vols of methanol then 2 vols of H2O. The soluble components of a whole Driselase digest (approx. 500μL) were then loaded and the column was washed with 2 × 2mL of H2O, after which bound solutes were eluted with 2 × 2mL each of 10, 20, 30, 40 and 50 % (v/v) methanol. Each fraction was dried, redissolved in 50μL of pyridine/acetic acid/water (1:1:98, by vol., pH approx. 4·7, containing 0·5 % chlorobutanol), and stored at –20°C. Fractions exhibiting the characteristic yellow–green fluorescence of pAMAC groups were pooled for further analysis. Immediately before analysis by HVPE or HPLC, a portion was dried, de-lactonized in dilute NaOH (pH >11) at 20°C for 10min, and neutralized with acetic acid.
Electrophoresis was conducted on Whatman No. 1 or No. 3 paper in a pH 6.5 buffer (pyridine/acetic acid/water, 33:1:300 by vol.) at 4·0kV for 45–50min. The papers were cooled with toluene. Methods and apparatus are described by Fry (2011b). After electrophoresis, the papers were dried and viewed under a 254nm UV lamp and fluorescence was recorded photographically (Camlab DocIt system with LabWorks 4·6 software). Fluorescent spots on paper electrophoretograms were quantified with Image J (http://rsbweb.nih.gov/ij/) as described in the Supplementary Data Fig. S3.
The HPLC was conducted with a solvent flow rate of 1mL min–1 at room temperature on a Luna C18 silica column with solvent A (500mm acetic acid, adjusted to pH 5·0 with NaOH) and acetonitrile. All solvent compositions are given as percentage acetonitrile in solvent A, by vol. The column was pre-equilibrated for 30 min with 10 % acetonitrile. The injected sample (20μL) was eluted with: 0–5min, 10 % acetonitrile, isocratic; 5–15min: 10–12·5 % acetonitrile, linear gradient; 15–30min: 12·5 % acetonitrile, isocratic; 30–35min, 12·5–15 % acetonitrile, linear gradient; 35–40min, 15 % acetonitrile, isocratic; 40–50min, 15–25% acetonitrile, linear gradient; 50–60min, 25–10 % acetonitrile, linear gradient; 60–65min, 10 % acetonitrile, isocratic. A fluorescence detector (RF 2000, Dionex) used excitation and emission wavelengths of 442 and 520nm, respectively.
Fruit firmness at three empirically defined stages of softening (hard, medium and soft) was measured – in most cases with a penetrometer (Fig. 2). The apples did not soften perceptibly within 1 month. All other species softened considerably between the three stages selected, although we did not quantify this for Arbutus.
Our approach for detecting •OH attack on fruit cell walls at different stages of softening in vivo was based on the methodology developed for characterizing authentic •OH-treated polysaccharides in vitro (Vreeburg et al., 2014). AIR (cell wall-rich material) of unripe and ripe fruits was labelled with pAMAC, then exhaustively digested with Driselase. Fluorescent conjugates of negatively charged (mainly pectic) cell wall fragments, such as pAMAC·UA2 (dimer), would be strong evidence for •OH-attacked pectins (Fig. 1). In contrast, GalA–pAMAC (monomer) would be derived from the pectins’ reducing terminus, and could be generated by any of three proposed agents (Fig. 1).
Electrophoresis of the Driselase digestion products of pAMAC-labelled AIR (after de-lactonization) revealed at least two interesting, yellow–green-fluorescing, negatively charged spots (Fig. 3): 1AF, co-migrating with the labelled monomer GalA–pAMAC; and 2AF, approximately co-migrating with the labelled dimer GalA2–pAMAC. The anionic nature of 1AF and 2AF indicates that they were based on Driselase-digestible acidic sugar residues of the fruit cell walls, likely to be mainly GalA. Smaller amounts of putative acidic trimers were sometimes also observed, e.g. in avocado and Arbutus. An additional spot (XF), which fluoresced a less yellowish green, was seen in some species, especially in unripe banana.
In addition, strongly yellow–green-fluorescing neutral spots were present in all species; these spots could be based on any neutral, Driselase-releasable cell wall sugar residues. They showed no obvious changes in intensity throughout ripening.
Despite the approximate co-electrophoresis of 2AF with GalA2–pAMAC, spot 2AF cannot have contained GalA2–pAMAC itself since this compound is completely digested by Driselase under the conditions used. Instead, it is likely to have a constitution of the type pAMAC·UA-GalA (Fig. 1), a Driselase-resistant ‘fingerprint’ spot diagnostic of •OH attack (Vreeburg et al., 2014).
In pear, mango, banana, avocado and Arbutus, spot 1AF appreciably increased in intensity (Fig. 3; Table 1), especially when stage 3 and stage 2 (soft and medium soft fruits) are compared with stage 1 (hard fruit). It also increased during softening in plum and nectarine (Table 1; Supplementary Data Fig. S1). On the other hand, the apple and strawberry AIR samples did not show any clear evidence of an increase in 1AF at any stage.
The putative ‘fingerprint’ spot, 2AF, was detected in all species, but was very weak in apple and strawberry. The yield of 2AF increased during softening in mango, banana, avocado, Arbutus, plum and nectarine, and was always much fainter in hard, unripe fruit (Fig. 3; Table 1; Supplementary Data Figs S1 and S2). In pear, it was detected only at stage 2. The transient appearance of 2AF in pear was confirmed in one repeat experiment (Fig. S2); it is possible that a brief period of high 2AF yield was missed in an additional experiment (Fig. S1).
Some samples, e.g. of pear and mango, revealed a weak spot that approximately co-migrated with GalA3-lactone–pAMAC (Fig. 3); however, HPLC showed that this specific compound was absent (Fig. 4; see below), as expected because it is Driselase digestible.
The presence of the reducing-end-labelled monomer, GalA–pAMAC, in the digests was supported by HPLC, which we performed on representative pear, mango and banana digests (stages 3, 1 and 1, respectively; without de-lactonization) both before and after spiking with a mixture of authentic sugar–pAMACs (Fig. 4). About 5–9 fluorescent peaks were detected by HPLC, with some differences between fruit species. In pear and banana, a compound was found (Fig. 4, thick purple arrow) which co-eluted with an internal marker of authentic GalA–pAMAC, supporting the idea that this was its identity. In mango, GalA–pAMAC partially overlapped with a compound of similar retention time (Fig. 4), which remains unidentified. Thus, in these fruit species, GalA–pAMAC was present, accounting for the 1AF spot seen on electrophoretograms.
Concerning the labelled acidic dimers, HPLC of the pear and mango samples gave no peak exactly co-eluting with authentic internal marker GalA2–pAMAC. The pear sample gave a peak that eluted 1·2min later than this marker. A small unidentified peak that did approximately co-elute with GalA2–pAMAC was found in the banana digest; however, this cannot have been GalA2–pAMAC itself because this substance is completely digested by Driselase under the conditions used (Vreeburg et al., 2014).
Samples of the material in the 1AF zone were eluted from electrophoretograms and submitted to HPLC analysis, typically giving 2–4 peaks (Fig. 5). In at least four fruits (pear, banana, avocado and Arbutus), a major peak co-eluting with GalA–pAMAC was again detected, whereas in mango, apple and strawberry (the three species which showed the faintest 1AF spots on the electrophoretogram), the corresponding peak was extremely minor. A small peak of GalA-lactone–pAMAC accompanied the GalA–pAMAC in pear, banana, avocado and Arbutus, supporting its identity since lactonization/de-lactonization is reversible and would be expected to occur between the electrophoresis step and the HPLC. Spot 1AF from the electrophoretogram yielded in addition HPLC peaks other than GalA-lactone–pAMAC and GalA–pAMAC (Fig. 5), with some differences between fruit species. In particular, unidentified peaks Y and Z (see Fig. 5) were observed: Y in pear, Z in banana and apple, and both in Arbutus, strawberry and possibly avocado.
Spot 2AF, deduced to be pAMAC·UA-GalA, which co-electrophoresed with authentic GalA2–pAMAC but was Driselase-stable, yielded HPLC peaks that only approximately co-eluted with authentic GalA2–pAMAC and GalA2-lactone–pAMAC in all species tested (Fig. 6). Both the acidic and lactone forms of the proposed pAMAC·UA-GalA were present in these eluates because of their interconversion, which is more rapid than in the case of the GalA2–pAMAC ↔ GalA2-lactone–pAMAC interconversion (Vreeburg et al., 2014).
Electrophoretogram spot XF, a greenish-fluorescing compound migrating slightly faster than authentic GalA2-lactone–pAMAC and observed in banana (and possibly Arbutus), was resolved by HPLC into several small peaks (Fig. 7). These peaks, however, did not match the HPLC peak ‘X’ found after pAMAC labelling of in-vitro •OH-treated pectin (Vreeburg et al., 2014), even though both have similar migration and fluorescence properties on the electrophoretogram. Both X and XF remain to be identified.
A fluorescent fingerprinting method, recently developed for demonstrating hydroxyl radical attack on polysaccharides in vitro (Vreeburg et al., 2014), has now been applied to the cell wall polysaccharides of several fruit species at different stages of softening, providing useful information on •OH attack in vivo. The fluorescent labelling procedure can yield information comparable with a radiolabelling approach used earlier (Fry et al., 2001, 2002; Müller et al., 2009; Iurlaro et al., 2014), and the two approaches are largely interchangeable. However, the fluorescent pAMAC group introduced into the polysaccharides in the new method provides a means of further characterization of these cell wall components by means of a wide variety of accessible chromatography and electrophoresis techniques, including fluorophore-assisted electrophoresis (Goubet et al., 2002). Specifically, pAMAC introduces a pH-dependent charge into the •OH-attacked, plant-derived polymer residue, which facilitates further characterization of the products. Radiolabelling is generally more sensitive and is very straightforward to quantify, but not all laboratories are authorized to use it.
At least two informative fluorescent spots (1AF and 2AF) were visualized on electrophoretograms (Fig. 3). Spot 1AF, including predominantly GalA–pAMAC (Fig. 1), increased in intensity between stages 1 and 3 of softening in most fruits (Table 1). This would correspond to an increasing number of d-GalA reducing termini during fruit ripening, which could be caused by the pectic polysaccharide-cleaving actions of not only •OH (as a result of attack at C-1 or C-4 of homogalacturonan – see fig. 1 of Vreeburg et al., 2014) but also endo-PG and/or pectate lyase. Both endo-PG and pectate lyase, proposed fruit softening agents, can attack a homogalacturonan chain, creating one new reducing terminus per cleavage event and this reducing terminus would become pAMAC labelled. Several studies have reported increases in endo-PG activity (though seldom definitively distinguished from pectate lyase activity) in pear (Pressey and Avants, 1976), banana (Pathak and Sanwall, 1998; Ali et al., 2004) and avocado (Huber and O’Donoghue, 1993). Increasing pectate lyase activity has been measured during ripening in banana (Payasi and Sanwal, 2003; Payasi et al., 2006). In addition, pectate lyase mRNA accumulation was reported in several ripening fruits including banana (Dominquez-Puigjaner et al., 1997; Marín-Rodríguez et al., 2003) and mango (Chourasia et al., 2006). Therefore, spot 1AF obtained from fruit AIR was not exclusive evidence of •OH attack, but may offer a valuable fingerprint indicating the total pectic chain scission occurring in vivo.
Spot 2AF was concluded to be a Driselase limit digestion product of the type pAMAC·UA-GalA (Fig. 1), i.e. a ‘fingerprint’ indicating recent in-vivo mid-chain •OH attack. The precise chemical identity of the compound(s) present in spot 2AF has not been established. 2AF clearly did not include the reducing-terminus-labelled disaccharide, GalA2–pAMAC, since this compound does not withstand 14d of Driselase treatment (Vreeburg et al., 2014), and GalA2–pAMAC was not observed in pear and mango by HPLC analysis (Fig. 4). It probably includes pAMAC·GalA-GalA and/or its 2-, 3- and 4-epimers (pAMAC·taluronate-GalA, pAMAC·guluronate-GalA and pAMAC·glucuronate-GalA, respectively). We would expect all these structures to be Driselase resistant because the range of activities present in Driselase probably does not include α-taluronidase, α-guluronidase and α-glucuronidase, and because the pAMAC group would block the action of α-galacturonidase.
The intensity of spot 2AF increased appreciably as hard fruit (stage 1) matured into softer fruit (stages 2 and 3) in mango, banana, avocado and Arbutus. The observation in banana may possibly be related to the ripening-dependent increase in the reported ability of banana fruit homogenates to generate ‘endogenous’ •OH post-mortem (Cheng et al., 2008a; Yang et al., 2008). In pear, the increase in spot 2AF was transient, peaking in stage 2; this suggests that the glycosulose residues from which 2AF is generated (Fig. 1) were unstable in vivo. A related observation in pear (increase in 3H-labelled products released when fruit cell walls were NaB3H4 labelled and then Driselase digested) was reported by Fry et al. (2001), where the unidentified 3H-labelled products were proposed to be ‘fingerprints’ of •OH attack.
The increase in yield of 2AF during softening depended on the type of fruit under consideration. In true fruits (those whose edible portion is derived from the ovary wall; including mango, banana, avocado, Arbutus, plum and nectarine), there was an increase in 2AF that correlated with softening. In contrast, it showed little if any increase in apple or strawberry and increased only transiently in pear, which are all false fruits. In false fruits, the edible tissue is derived from the receptacle, not the ovary wall. Therefore, differences in developmental origin of the edible tissue may dictate the mechanism adopted for cell wall modification during ‘fruit’ softening.
It was reported nearly 40 years ago that in pear fruit, endogenous peroxides (and thus potentially also •OH generated from them) correlate with softening (Brennan and Frenkel, 1977). Later it was found that the polysaccharides of softening pears exhibit radiochemical ‘fingerprints’ diagnostic of recent •OH attack (Fry et al., 2001). Furthermore, of two investigated cultivars of muskmelon, the one whose microsomal membranes produced less •OH in vitro had a longer shelf-life (Lacan and Baccou, 1998). Taken together, the available evidence supports the view that fruit softening, often viewed as broadly a ‘hydrolytic’ phenomenon, is at least partly ‘oxidative’ – a suggestion raised by Brennan and Frenkel (1977) but often ignored. We hope that interest in this concept will be revived by the present study and explored in greater depth. Although several of the fluorescent ‘fingerprint’ compounds were not fully identified in the present study and deserve further analysis, our new fluorescent labelling method will provide useful information and can be used in conjunction with other approaches to add to our knowledge and understanding of the occurrence and rate of •OH attack relative to endo-PG and pectate lyase action in fruit cell walls.
Supplementary data are available online at www.aob.oxfordjournals.org and consist of the following. Figure S1: electrophoretic resolution of total Driselase digests of pAMAC-labelled cell walls from three fruit species. Figure S2: electrophoretic resolution of total Driselase digests of pAMAC-labelled cell walls from banana and pear. Figure S3: method for quantification of fluorescent spots on paper electrophoretograms.
We thank Mrs Janice Miller and Mr Tim Gregson for excellent technical assistance, and the UK Biotechnology and Biological Sciences Research Council for a grant (ref. 15/D19626) in support of this work. O.B.A. thanks the Ministry of Higher Education, Malaysia, for a studentship, and Universiti Kebangsaan Malaysia for a grant to continue the work (ref. GGPM-2013-032).