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Cell Wall Changes in Spanish Pear During Ripening
J. Plant Physiol.
Vol. 144. pp. 541 - 548 (1994)
Cell Wall Changes in Spanish Pear During Ripening MARIA MARTIN-CABREJASl, KEITH W. WALDRON2>~, 1
2
*
and ROBERT R.
SELVENDRAN2
Universidad Autonoma de Madrid, Departimento Chimica Institute of Food Research, Norwich Research Park, Colney, Norwich NR4 7UA Author for correspondence
Received December 28, 1993 . Accepted March 4, 1994
Summary
Unripe Spanish pears (Pyrus communis, c.v. Blanquille) were allowed to ripen at 18°C for 1 week. Cell walls were prepared as alcohol-insoluble residues (AIRs) which were free of starch. On a fresh weight basis, ripening was accompanied by a decrease in cell wall arabinose and uronic acid, a proportion of which were metabolised, and an increase in xylose and glucose. To investigate the origin of the changes in cell wall polymers, the AIRs of unripe and ripe fruit were extracted sequentially with water, cyclohexanetrans-l,2-diamine-N,N,N',N'-tetraacetate (CDTA), Na2 C03 at 1°C and 20°C, 0.5,1 and 4M KOH to leave a cellulose-rich residue. The extracts and residues were analysed for their carbohydrate composition and, where appropriate, degree of pectin-methyl esterification. The water, CDTA and Na2C03 extracted polymers were rich in pectic polysaccharides. These exhibited heterogeneity in the ratios of neutral and acidic components and contained small quantities of xylose. The KOH-extracted polymers were rich in hemicelluloses, particularly xylans, but also contained sugars common to pectic polysaccharides. The cellulose-rich residues were rich in cellulosic glucose and xylose which probably originated from the highlylignified sclereids. Ripening resulted in an increase in water- and CDTA-soluble pectic polysaccharides which exhibited a lower degree of pectin-methyl esterification. These results and the changes in carbohydrate composition are discussed in the light of corresponding decreases in the yields of pectic polysaccharides of the Na2 CO) extracts and cellulose-rich residues during ripening.
Key words: Cell walls, Pyrus communis, ripening, pectic polysaccharides, hemicelluloses, cell types. Abbreviations: AIR = alcohol-insoluble residue; CDTA = cyclohexane-trans-l,2-diamine-N,N,N',N'tetraacetate.
Introduction
It is generally accepted that changes in the cell wall polymers of pears and other fruits playa major role in determining the ripening-related alterations in fruit texture. However, our understanding of these changes is hampered by limited knowledge of the structure of the cell walls in mature unripe fruit and the enzymes which modify the cell wall polysaccharides (Brady, 1987). In pear, the most commonly reported change in cell wall integrity involves the increase in the soluble pectic polysaccharides. This is usually accompanied by an overall loss of cell wall arabinose Germyn and Is© 1994 by Gustav Fischer Verlag, Stuttgart
herwood, 1956; Dick and Labavitch, 1989; Ahmed and Labavitch, 1980) and, uronic acid Germyn and Isherwood, 1956; Bartley et aI., 1982). The mechanism by which cell wall pectic polysaccharides are solubilised, and the subsequent fate of the neutral and acidic carbohydrate residues in ripening pears is unclear. Most studies have investigated either the changes in the carbohydrate composition of the water-insoluble cell wall material (Gross and Sams, 1984) or the degree and extent to which pectic polysaccharides are solubilised by quantifying the release of water-soluble polyuronide (Knee, 1973; Yamaki et al., 1979; Ahmed and Labavitch, 1980). Few workers
542
MARIA MARTIN-CABREJAS, KEITH W. WALDRON, and ROBERT R. SaVENDRAN
have attempted to investigate the nature of the parent wallbound pectic polysaccharides in which ripening-related changes occur. In previous investigations on tomato (Seymour et al., 1990), apple (Whitcombe et a!., unpublished results) and Kiwi fruit (Redgwell et al., 1990; Redgwell et al., 1991) cell wall preparations have been extracted sequentially using techniques which minimise degradation of the constituent polymers. These studies have shown that pectic polysaccharides exhibit a range of solubilities which probably have a bearing on the type of cross-links which hold them into the cell wall matrix. Furthermore, such studies have shown that ripening-related changes are often associated with certain groups of pectic polysaccharides. In ripening pears, those studies which have attempted to fractionate the cell wall have confined themselves to extracting pectic polysaccharides by water and chelating agents (Ben-Arie et a!., 1979), or have used methods of cell wall extractions including hot water and strong alkali, which will have solubilised many of the constituent pectic polysaccharides through {3eliminative degradation Germyn and Isherwood, 1956), masking many of the ripening-related changes in the solubility of cell wall polymers. Furthermore, there is a paucity of information concerning the fate of acidic and neutral moieties which are lost from the soluble and insoluble fractions of the cell walls during ripening of pears. In Bartlett pears, the uronic acid solubilised during ripening can be recovered fully in polymeric form from the aqueous phase showing that complete depolymerisation of the polygalacturonic acid backbone does not occur (Ahmed and Labavitch, 1980). In the same variety, the ripening-related loss of cell wail arabinose has been attributed to the depolymerisation of rhamnogalacturonan-like pectic polysaccharide moieties after solubilisation (Dick and Labavitch, 1989). In contrast, fruit softening during ripening of Spadona pears is accompanied by an increase in free galacturonic acid, indicating extensive depolymerisation of the pectic polysaccharides (Ben-Arie et aI., 1979). The aim of the present study has been to determine whether or not the solubilised (degraded) arabinose and uronic acid-containing carbohydrate moieties are metabolised during the ripening of Spanish pears, and to investigate the fractions of the cell wall from which they are released by making use of modern methods of plant cell wall sequential extraction (Waldron and Selvendran, 1992; Coimbra et al., 1994).
Materials and Methods Plant material Mature green unripe Spanish pears (Pyrus communis L. cv. Blanquille) were obtained from the central fruit market of Madrid (Merca Madrid). Fruits were selected for uniformity in size and were free from blemishes. Fruits (samples of 25) were used immediately or after being allowed to ripen at 18°C for 1 week in darkness. Whole pears were peeled (epidermis and approximately 2 mm of underlying tissue) and the ovary and central vascular tissue excised. The fruit flesh was sliced and diced, frozen immediately in liquid nitrogen, and stored at - 40°C until required.
Preparation ofalcohol·insoluble residue (AIR) AIR. was obtained by immersing the frozen pear tissues in boiling ethanol (final concentration 85 % (v/v) aq.), homogenising with a Waring Blender and then Ystral homogeniser (maximum speed, 5 min) and boiling for 5 min. The material was filtered through a glass sintered filter (No.3, Scientific Furnishings Ltd., Sussex, UK) and re-extracted twice by boiling for 10 min in 85 % (v/v) aq. ethanol (5 x initial fresh weight) and filtering. The residue was resuspended in absolute alcohol (twice), acetone (twice), filtering between times, and then air-dried after which it was stored in a sealed container at room temperature.
Dialysis ofalcohol extracts The 85 % (v/v) aq. ethanol extracts recovered during preparation of AIRs was reduced in volume by rotary evaporation. The resultant liquor was dialysed overnight against water at 0 DC, from which the diffusate was recovered and stored at - 30°C. The volumes of both the dialysate and diffusate were recorded. The dialysate was then dialysed exhaustively against water at O°C after which it was reduced in volume and lyophilised. In calculating the recovery of the diffusate, corrections were made for the volume of the initial dialysate.
Fractionation ofAIR This method is based on that developed for cell wall material of onion bulbs (Redgwell and Seivendran, 1986). AIR. (2 g) was sequentially extracted with (i) water (100 mL) at 20°C for 2 h; (ii) 50 mM cyclohexane-trans-1,2-diamine-N, N, N', N'-tetraacetate (CDTA, Na salt, 100mL) pH 6.5 at 20°C for 5h (CDTA-1); (iii) CDTA, (100mL) pH 6.5 at 20°C at 2h (CDTA-2); (iv) 50mM Na2COJ + 20mM NaBH4 (l00mL) at O°C for 20h (Na2COJ-l); (v) 50mM Na2CO J + 20mM NaBH4 (100mL) at 20°C for 2h (Na2CO J-2); (vi) 1M KOH + 20mM NaBH4 (loomL) at 20°C for 2h and (vii) 4 M KOH + 20 mM NaBH4 (100 mL) at 20°C for 2 h . The alkali extractions were carried out with Orfree solutions under argon. After each extraction, the solubilised polymers were separated from the insoluble residue by centrifugation. The alkali extracts were acidified to pH 5 with HOAc and all extracts were filtered through glass fibre filter paper GFC (Whatman). The cellulose-rich residue remaining after the final extraction was also acidified to pH 5. The filtered extracts and residue were dialysed exhaustively, concentrated and lyophilised.
Analysis of carbohydrate composition Sugars were released from AIR. and sub-fractions by dispersing in 72 % H 2S04 followed by dilution to 1 M and hydrolysing for 2.5 h at 100 °C (Saeman hydrolysis). Sugars were released from dialysate and diffusate fractions by hydrolysing for 2.5 h at 100°C in 1 M H 2S0 4. All samples were subjected to duplicate analysis. Neutral sugars were reduced with NaBH4 and acetylated by the method of Blakeney et al. (1983) using 2-deoxyglucose as an internal standard. Alditol acetates were quantified by gas chromatography on a Carlo Erba Vega gas chromatograph after automatic injection by a Carlo Erba A 200 autoinjector. Alditol acetates were separated with baseline resolution on a Restek RTx 225 WCOT column (15mx 0.32 mm i.d.; 0.25 u film) using an oven temperature programme of 90°C for 1 min, 45°C per min to 150°C, 150 °C for 1 min, 2°C per min to 210°C, and 210 °C for 1.5 min. The carrier gas was helium at a column head pressure of 60 Kpa. Detection was by flame ionisation. Data was collected and integrated on a Spectra Physics
543
Cell walls of ripening pears Table 1: Yields and carbohydrate composition of AIRs from unripe and ripe fruit. Sample Unripe Ripe
Carbohydrate Composition (Mol %)
Yield (% Fwt)
Rha
Fuc
Ara
Xyl
Man
Gal
Glc
UA
(~g/mg)
2.7 2.7
1.0 1.3
1.1 1.1
17.1 9.1
27.1 32.5
1.2 1.7
3.4 3.1
21.3 (4.8)* 25.7 (7.8)*
27.9 25.5
646 543
Total
* 1 M hydrolysis.
SP4400 integrator; re-integration was handled by a Spetraphysics Winner data handling station. Total uronic acid content was determined colorimetric ally by the method of Blumenkrantz and AboeHansen (1973) after dispersal in 72 % H 2S0 4, dilution to 1 M, and hydrolysis for 1 h at 100 °C (Selvendran et aI., 1989).
Klason lignin The sample (100 mg) was dispersed in 72 % H2S0 4 for 3 h at room temperature followed by dilution to 1 M and hydrolysing for 2.5 h at 100°C. The residue was recovered by centrifugation (12,000 x g for 10 min), washed 3 times in distilled water and lyophilised. The Klason lignin was then quantified gravimetrically.
Results and Discussion
Carbohydrate composition ofAIRs and alcohol-soluble components Unripe and ripe pears were light green in colour and weighed approximately 145 g. Unripe pears were hard and inedible whilst ripe pears had become softer and more palatable. Cortical tissue from unripe and ripe years were prepared as AIRs. In each case, the yield of the AIR was 2.7 % (Fwt). As the pears were free of starch, as shown by negative staining with I2/KI and light microscopy, the AIR was used as the source of the cell wall material. Hydrolysis of the AIRs was performed using the Saeman method or 1 M H 2S0 4• The values given in Tables 1- 3 are the means of duplicate determinations and the variations between the duplicates was less than 2 %. The results reported in Tables 1 and 2 show the same trends in changes of cell wall com-
Table 2: Total carbohydrate composition of unripe and ripe pears. Sample
Carbohydrate recovery (~glg Fwt)
Unripe AIR EtOH dialysate Diffusate Total
Rha
Fuc
Ara
Xyl
0.16
0.18
2.62
4.16
0.19
Man
Gal
Glc
UA
Total
.
3.90 0.09
17.4
0.46
0.62 0.04 0.08
5.5
0.04 0.45
0.25
0.73
3.99
6.39
19.6
om
0.5
4.3 0.04
4.6 0.14 0.97
15.8 0.21 2.82
0.62
4.34
5.7
18.8
om
0.35
0.18
3.1
4.6
0.2
0.16
1.3
4.5 0.02
0.23 0.02
Ripe AIR E,OH dialysate Diffusate
0.16
0.05
0.44
1.1
*..
Total
0.36
0.21
1.7
5.6
0.28
* **
4.37 26.1 *.* 5.5 >tltl/>" .28.1
0.28
0.12
0.72
0. 1
0.31
1.90
Values referred to with asterisks: The majority of the glucose and man nose released during hydrolysis of the diffusate will have originated from intracellular sucrose, glucose and fruc· tose. For darity, their values have been omitted from the main body of the table.
ponents as those of preliminary experiments performed during the previous growth season. The carbohydrate composition of the AIR of unripe pears was rich in xylose and glucose which, together, comprised approximately half of the cell wall carbohydrate. The majority (75 %) of the glucose can be inferred to be cellulosic in origin since it was released only by Saeman hydrolysis. Since the level of cell wall xylose was over 5 times that of the non-cellulosic glucose, the presence of large quantities of xylan hemicelluloses can be inferred (Waldron and Selvendran, 1992). Approximately 25 % of the glucose was released by 1 M H 2S0 4, and it is likely that a significant quantity of non-cellulosic and non-starch glucans were present in the AIR. This will be discussed below. The remainder of the AIR carbohydrate comprised pectic polysaccharides as indicated by the levels of rhamnose, arabinose, and uronic acid. In comparison, the AIRs of ripe pears exhibited reduced levels of arabinose and uronic acid indicating a ripening-related decrease in branched pectic polysaccharides, and higher levels of xylose and glucose. The ripening-related decrease in pectic arabinose is consistent with the changes in the cell walls of Conference pears Germyn and Isherwood, 1956), and Bartlett pears (Ahmed and Labavitch, 1980; Dick and Labavitch, 1989). The substantial quantity of xylose and glucose in the AIRs and the ripening-related increase in these moieties, is due to the presence and continued maturation of sclereids (stone cells) during the ripening of Spanish pears. This is presently the subject of a more detailed investigation.
Carbohydrate composition of the 85 % {vlv} aq.-soluble components In order to determine the possible fate of the carbohydrate moieties that were lost from the pear cell walls during ripening, the 85 % (v/v) aq. ethanol extracts were retained, reduced under vacuum, and then dialysed exhaustively (10 changes) against water at 0 The first dialysis step was carried out against approximately 10 times the volume of the dialysis bag. After stirring overnight, the volumes of the equilibrated diffusate and dialysate solutions were quantified. The diffusate was reduced under vacuum and, because it proved difficult to freeze-dry, was dissolved in a larger quantity of water from which measured aliquots were taken. The dialysate was re-dialysed exhaustively, reduced in volume and lyophilised. The carbohydrate composition of the dialysate and diffusate samples were determined after hydrolysis in 1 M H 2S0 4 • To facilitate comparison, the data presented in Table 2 has been calculated on a fresh-weight basis and the recoveries for the diffusate have been corrected for the volume of the dialysate. The majority of carbohydrate released
0c.
544
MARIA
MARTIN-CABREJAS, KEITH W. WALDRON, and ROBERT R. SELVENDRAN
Table 3: Yields and carbohydrate composition of polymers extracted from cell walls of unripe and ripe pears. Sample
Carbohydrate composition (Ilg/ mg) Rha
Fuc
Ara
Xyl
Man
Gal
Glc
UA
Total
Water Unripe Ripe
0.8 0.9
1.2 1.5
21.7 22.0
7.1 6.7
0.9
1.1
7.6 5.1
4.8 3.6
55.8 59.2
781 692
19.2 25.1
CDTA-l Unripe Ripe
0.5 2.1
0.7 1.2
15.2 16.1
5.7 5.7
1.3 0.9
3.5 2.9
5.6 3.8
6704
67.6
244 320
5.9 7.2
CDTA-2 Unripe Ripe
1.7 1.5
2.0 2.3
12.7 14.1
4.5 8.6
3.6 2.8
2.2 2.9
55.1
lOA
5704
110 113
5.2
Residue 1 Unripe Ripe
0.7 0.8
0.02 0.1
14.9 5.6
35.1 39.5
0.04 0.3
3.3 3.8
35.1 44.7
10.9 6.2
644 655
66.6 61.8
Na2C03-1 Unripe Ripe
1.2 1.5
0.03
21.2
3404
2.79 6.8
0.2 0.8
3.2 3.3
0.8 3.0
63.1
5704
729 296
10.5 5.8
Na2C03-2 Unripe Ripe
1.0 3.1
59.7 29.7
4.0 6.5
0.2
3.2 4.7
0.8 2.22
31.1 52.7
817 387
2.0 1.9
0.5 M KOH Unripe Ripe
1.2 1.64
2.1
104
23.5 15.5
35.7 3704
1.8
IMKOH Unripe Ripe
0.9 2.7
0.9 1.6
18.2 20.8
19.8 29.3
4MKOH Unripe Ripe
0.5 0.6
2.0 1.8
3.8 3.3
Residue Unripe Ripe
0.7 0.9
0.3
9.7 3.0
1.1
18.2
Yield (% air)
204
104
7.1 7.2
17.0 16.7
12.9 17.6
1000 783
1.7 1.2
2.9
1.3
4.7 7.7
20.7
1404
39.9 14.2
261 638
1.2
51.3 45.2
3.2 3.7
6.5 6.1
19.6 16.7
13.2 22.7
742 591
17.1 26.3
22.0 28.0
1.5 1.6
2.7 1.5
40.1 43.3
23.1 21.7
336 360
31.9 24.2
during hydrolysis of the diffusate comprise Glcp and Manp. These will have originated from intracellular sucrose, glucose and fructose which impart a sweet flavour to pear flesh during ripening (Burton, 1982; Hulme, 1970) and have been presented beneath Table 2. The carbohydrate composition of the dialysate and diffusate of the Ethanol extracts of both unripe and ripe pear AIRs contained sugars common to pectic polysaccharides, as shown by the levels of uronic acid, galactose and, in the case of the diffusate, rhamnose and arabinose. Xylose was also present, particularly in the diffusate, and increased during ripening. The carbohydrate of the dialysate is likely to have been oligomeric in nature, but not polymeric, since it failed to precipitate in 85 % (v/v) aq. ethanol. Both oligomers and monomers are likely to have been present in the diffusate. There was little difference in the recovery of sugars of pectic origin from un-ripe and ripe pears in the diffusate, although the uronic acid exhibited a small increase. This is consistent with studies on Spadona pears (Ben-Arie et al., 1979) in which ripening was accompanied by an increase in free galacturonic acid. However, in the present study, the total recoveries of uronic acid, galactose and arabinose (from the AIRs, diffusates and dialysates), decreased during ripening,
104
showing unequivocally that a proportion had each been metabolised after the release from the cell wall. This contrasts with studies on Bartlett pear (Ahmed and Labavitch, 1980) in which total levels of polymeric uronic acid reportedly remained constant. In that study, the authors also reported that the total level of polymeric arabinose remained constant during ripening. However, re-calculation of their results confirm that total cell wall derived polymeric arabinose underwent a significant ripening-related decrease. Free arabinose had not been quantified.
Fractionation ofpear AIR's In order to identify polysaccharide fractions of the cell wall in which the ripening-related changes occurred, the cell wall (AIR) preparations were sequentially extracted with water at 20°C, 0.05 M CDTA (20 o q, 0.05 M Na2C03 at 1 °C and 20°C, and 0.5, 1 and 4 M KOH at 20°C, essentially as described before (Redgwell and Selvendran, 1986; Waldron and Selvendran, 1992). The extraction procedures were designed to minimise /1-eliminative degradation of pectins during the initial stages of extraction and to solubilise the cell wall polymers in as close to their native form as possible.
Cell walls of ripening pears
Data on the amounts of polymers solubilised and their sugar compositions are contained in Table 3. The amounts of material extracted are based on one sequential extraction of AIR from unripe and ripe pears. The pectic polysaccharides that were not cross-linked into the cell wall were solubilised by water at 20°C. Those held in the wall by Ca2+ only were subsequently extracted by CDTA. It is probable that the bulk of the water and CDTAsoluble pectic polysaccharides are of middle lamella origin (Selvendran, 1985). They consisted mainly of uronic acid, the predominant neutral sugar being arabinose. In addition, smaller but significant quantities of galactose were present. In spite of exhaustive dialysis, the carbohydrate recovery of the CDTA extracts was low. This can be attributed to the presence of contaminating CDTA which was detected by IR spectroscopy (results not shown). The incomplete removal of CDTA in this study is probably due to the presence of Ca2+ ions in the extract which complex with the CDTA and thus prevent its complete removal by dialysis. Similar difficulties have been encountered in CDTA extracts of olive pulp, and have been discussed in depth (Coimbra et al., in press). Much of the water and CDTA-insoluble pectic polysaccharides were subsequently solubilised with dilute Na2C03 at 1°C and 20°C, presumably by hydrolysis of weak ester cross links. The initial Na2C0 3 extractions were performed at 1 °C to minimise (3-eliminative degradation of the pectins in subsequent alkali extractions. Like the water and CDTA extracts, both Na2 C03 extracts were rich in pectic polysaccharides as shown by the high levels of uronic acid, arabinose and galactose, and the occurrence of rhamnose as the main deoxy sugar. As in other tissues (Ryden and Selvendran, 1990 a; Waldron and Selvendran, 1992), heterogeneity amongst the pectic polysaccharides was evident from the differing ratios of the neutral sugar (arabinose and galactose) both to each other and to the uronic acid component. For example, water-soluble pectic polysaccharides from unripe pear AIR exhibited an arabinose/galactose ratio of 2.8, and this ratio increased though the sequence of extractions to 18.7 in the Na2C03-2 extract. Interestingly, all the pectic polysaccharides solubilised by water, CDTA and Na2C0 3 contained small, but significant, quantities of xylose. The levels of xylose in the polysaccharides (3 - 9 mol %) were not only much greater than the amounts found in equivalent cell wall extracts from ripening tomatoes (Seymour et al., 1990), apples (Whitcombe et al., unpublished results) but were greater than the amounts found in equivalent cell wall extracts from maturing asparagus (Waldron and Selvendran, 1992), potato (Ryden and Selvendran, 1990a) and runner bean (Ryden and Selvendran, 1990 b). This is discussed further in the General Discussion. In addition to pectic polysaccharides, the 0.5 M KOH extractions solubilised significant quantities of hemicelluloses as indicated by the presence of xylose and glucose. The presence of glucose, xylose, galactose and fucose in the hemicellulosic extracts indicate the presence of fucogalactoxyloglucans. Since the levels of xylose were always considerably greater than those of glucose, the presence of xylan hemicelluloses could also be inferred. Interestingly, the 1 M and 4 M KOH extracts also contained significant quantities of pectic polysaccharides. This contrasts with the results of similar fractionation studies from
545
other fruits including tomatoes (Seymour et a1., 1990), apples (Whitcombe et al., unpublished results), and olives (Coimbra et a1., in press), and vegetables such as Asparagus (Waldron and Selvendran, 1992) and potato (Ryden and Selvendran, 1990 b). In all of these, the pectic component of these extracts was low. However, large quantities of pectic polysaccharides have been found in the 4 M KOH extract of cell walls of mung bean (Gooneratne et al. , unpublished results). In addition, the cell walls from the outer pericarp of Kiwi have also yielded significant quantities of pectic polysaccharides in the 4M KOH extract (Redgwell et al., 1990); however, this may arise because no previous 0.5 or 1 M alkali extractions had been performed. The remaining a-cellulose residue consisted mainly of glucose, xylose and uronic acid. The relatively low carbohydrate recovery in the residue (approximately 35 %) could be attributed to the large quantity of lignin present which originated from the residual stone cells. This was confirmed (a) by the measurement of Klason lignin values which were 60.1 % in the a-cellulose residue of un-ripe pear, and 52.2 % in the residue from ripe pear, and (b) by visual identification by light-microscopy. Interestingly, in spite of the relatively high Klason lignin values, the stone cells in the a-cellulose residues stained light pink with phloroglucinol-HCL (Fig. 1 a). However, those in the residue which remained after the initial water and CDTA extractions stained deep red (Fig. 1 b). This suggests that many of the phloroglucinol-HCLstaining functional groups in the stone cell lignin were either removed or modified during the alkali extractions. The formation of a red colour with phloroglucinol-HCI is indicative of coniferaldehyde, cinnamic aldehydes and related benzaldehydes (Monties, 1989). It is likely that the aldehyde groups in the lignin polymer were reduced by the NaBH4 during the 0.5, 1 and 4 M KOH extractions, eliminating their ability to interact with the stain. In contrast, the ability of the sclereids to stain turquoise with toluidine-blue was not altered by the sequential extractions indicating that the appropriate toluidine-blue-staining functional groups are still present (results not shown).
a Fig. 1: Light micrograph of phloroglucinol-HCl-stained sci ere ids of ripe Spanish pear (a) from CDTA-insoluble residue; (b) from cellulose-rich residue. Bar, 100 jJ.m.
546
MARIA MARTIN-CABREJAS, KErTH W. WALDRON, and ROBERT R. SELVENDRAN
5 '01
en
Table 4: Methyl-esterification (%) of uronic acid component of selected wall fractions from unripe and ripe pears.
4
x
E
AIR Water CDTA-1 CDTA-2 CDTA-2 Residue
~3
e ell
-g,2
.!:
o
... o
94 96 74 78 96
Unripe SD
8 3 4 3 8
x
Ripe SD
65
5
46
4
62 39 102
2 2 2
.D
u
W
CI
C2
NI
N2 O·5K IK
4K CR
cell wall extract Fig. 2: Yields of carbohydrate (mg anhydrosugar g-l fresh weight) from extracts and insoluble residues of unripe and ripe pear AIRs. W: water extract; Ct and C2: CDTA extracts t and 2; Nt and N2: Na2CO) extracts 1 and 2; 0.5 K, t K, 4 K: 0.5, 1 and 4 M KOH extracts; CR: cellulose-rich residue. 0 Unripe pear; • ripe pear.
Ripening related changes In order to make useful comparisons between the carbohydrate composition of the cell wall and .extracted. polym~rs from unripe and ripe pears, and to permlt comparlsons wlth the data in Table 2, the recoveries of total carbohydrate and selected sugars are presented on a Fwt basis basis in Figur~s 2 and 3 respectively. The results in Figure 2 show that dunng ripening, there is an increase in the water and CDTA-solub.le carbohydrate. This is accompanied by a greater decrease 10 the levels of carbohydrate solubilised by the Na2C03-1 and 2 extractions and to a lesser extent by decreases in the levels of carbohydr;te remaining in the a-cell~lose residue. The r!pening-associated loss of carbohydrate lS greater than the. 10crease in water-soluble and CDTA-soluble polysacchande, and is consistent with the decrease in total AIR carbohydrate (Tables 1 and 2). Since the ripening-related changes in carbo~4 '01 01
53 ell
"0
-0
>o
2
.!:
...
.D
o
hydrate are principally due to the changes in the levels of arabinose and uronic acid, the data for the recovery of these moieties throughout the cell wall fractions has been calculated and is shown in Figure 3. The results show that the increase in water-soluble uronic acid and arabinose, and the loss of these moieties from the cell wall, arise from the turnover of the pectic polysaccharides in the Na2C03-1 extract and, to a lesser extent, the Na2C03-2 extract and cellulosic residue. In addition to the carbohydrate composition, the degree of pectin methyl esterification was also quantified. In AIR, pectin methyl esterification decreased from 94 % (± 8 %) to 65% (± 5%) during ripening (Table 4). This is comparable to the trend observed in ripening Conference pears (Knee, 1982). To assess the origin of this change, the esterification of the pectic polysaccharides in each of the cold water extracts, CDTA extracts, and CDTA-insoluble residues, were also measured. The results (Table 4) demonstrate that the lower degree of methyl esterification in AIR of ripe pear results from the low levels of esterification of the water and CDTA-soluble pectic polymers; the methyl esterification of the residue in both unripe and ripe pears was above 95 %. This indicates that a significant proportion of the N a2 C03soluble polymers of unripe pears which become CDTA- and water-soluble during ripening also become de-esterified. The origin of the ripening-related increases in xylose was also investigated. Whilst the arabinose and uronic acid recoveries from the cell wall fractions were quantitative, (greater than 95 %), the total recovery of xylose was approximately 60 %. Hence, it is not possible to conclude which fractions of the cell wall contained the polymers which increase in xylose during ripening. Previously, it was noted t~at ripening resulted in an increase in «low» molecular welght xylosecontaining oligomers (Table 2). If similar oligomers comprising xylose were released from the cell wall durin~ the fractionation process, they may have been lost dunng the subsequent dialysis stages, thus accounting for the poor xylose recovery.
u
W CI
C2
NI
N2 O·5K IK
cell wall extract
4K CR
Fig.3: Yields of uronic acid and arabinose (~g anhydros~gar g-l fresh weight) from extracts and insoluble resldues of unnpe and ripe pear AIRs. W: water extract; C 1 and C2: CDTA extracts 1 and 2; Nt and N2: Na,CO) extracts t and 2; O.5K, lK, 4K: 0.5,1 and 4M KOH extracts; CR: cellulose-rich residue. Uronic acid: 0 unripe, • ripe; arabinose: Eil unripe; E8 ripe.
General Discussion Turnover of the carbohydrate moieties in the neutral side chains of cell wall pectic polysaccharides has been recognised in several types of cell wall, including ripening fruits (Gross and Sams, 1984; Seymour et aI., 1990; Redgwell, 1990) and the stems of dicots and monocots (Labavitch and Ray, 1974; Waldron and Selvendran, 1992). In order to understand the basis for such turnover, it is necessary to extract the cell
Cell walls of ripening pears
walls sequentially using solvents which minimise polymer degradation, and to identify the fractions in which turnover occurs. Such methods are essential, as the native pectic polysaccharides are easily susceptible to fj-eliminative degradation during extraction. Using such methods, the turnover of arabinose and uronic acid has been investigated during ripening of Spanish pears. The majority of this turnover occurs in the cold Na 2C0 3-s01uble polysaccharides, although some occurs in acellulose fraction. This contrasts with the turnover of pectic galactose during ripening of tomatoes (Seymour et al., 1990), where the loss of cell wall galactose could be attributed to its turnover in all of the cell wall fractions except for the CDTA-1 extracted polymers. However, the results reported in the present study exhibit trends that are similar to those found in ripening kiwi fruit (Redgwell et al., 1990; Redgwell et al., 1991) and apple (Whitcombe et al., unpublished results). The results showed that certain pectic polysaccharides became de-esterified during ripening. Such de-esterification has been associated with an increase in the susceptibility of polygalacturonic acid to the action of endo-polygalacturonase, as demonstrated in tomatoes (Pressey and Avants, 1982; Koch and Nevins, 1989). Ripening of tomatoes also results in a decrease in the degree of total pectin methyl esterification (Koch and Nevins, 1989). In contrast, such changes do not occur in ripening apples (Knee, 1978). However, the overall level of total pectin methyl esterification is not necessarily a true indicator of the events taking place. For example, Knee (1982) demonstrated that during ripening of Conference pears, newly synthesised methyl esterified pectic polysaccharides were continually inserted into the soluble fraction of the cell wall and subsequently turned over. The occurrence of xylose residues in the pectic polysaccharides solubilised by water, CDTA and Na2C0 3, from the cell walls of a variety of fruits and vegetables has been reported (Waldron and Selvendran, 1992; Ryden and Selvendran, 1990; Seymour et aI., 1990) and it is probable that these polysaccharides are derived from the middle lamella region (Selvendran, 1985; Bartley and Knee, 1982). The large quantity of xylose in the cellulosic residue is likely to have originated from (1-4}-linked xylan hemicelluloses from lignified sclereids (Fig. 3) which are scattered throughout the cortex. Some, however, may have been found in the walls of the cortical parenchyma, as in Guava fruit (Marcelin et aI., 1993) and olive (Coimbra et aI., unpublished results). The unusually high levels of xylose associated with the pectic polysaccharide extracts may reflect the occurrence of pectic-xylan complexes. We have detected an increase in pectic-xylanphenolic complexes during the maturation of asparagus stems (Waldron and Selvendran, 1992) and have linked this to the initial stages of lignification which occurs in the middle lamella. Similar events would be expected to be prevalent in newly-differentiated stone cells surrounding the sclereid clusters.
Acknowledgements This work was supported by the Comunidad Autonoma de Madrid, Consejeria de Educacion (M. Martin-Cabrejas) and the V .K . office of Science and technology.
547
References AHMED, A . E.and J. M LABAVlTCH: Cell wall metabolism in ripening fruit 1. Cell wall changes in ripening «Bartlett» pears. Plant Physio!' 65, 1009-1013 (1980). BARTLEY,!. M., M. KNEE, and M.-A. CASIMIR: Fruit Softening 1. Changes in cell wall composition and endopolygalacturonase in ripening pears. J. Exp. Botany 33, 1248 -1255 (1982). BARTLEY, 1. M. and M. KNEE: The chemistry of textural changes in fruit during storage. Food Chemistry 9, 47 -58 (1982). BEN-ARlE, R., L. SONEGO, and C. FRENKEL: Changes in pectic substances in ripening pears. Journal of the American Society for Horticultural Science 104, 500-505 (1979). BLAKENEY, A. B., P. J. HARRIS, R. J. HENRY, and B. A. STONE: A simple and rapid preparation of alditol acetates for monosaccharide analysis. Carbohydrate Research 113, 291-299 (1983). BWMENKRANTZ, N. and G. ASBOE-HANSEN: New method for quantitative determination of uronic acids. Analytical Biochemistry 54,484-489 (1973). BRADY, C. J.: Fruit ripening. Annu. Rev. Plant Physio!' 38, 155178 (1987). BURTON, W. G.: Post harvest physiology of food crops. Longman, London. 339 pp. (1982). COIMBRA, M. A., K. W. WALDRON, and R. R. SELVENDRAN: Isolation and characterisation of cell wall polymers from olive pulp (Olea Ouropaea L.). Carbohydrate Research (1994; in press). DICK, A. J. and J . M. LABAVlTCH: Cell wall metabolism in ripening fruit. IV. Characterisation of the pectic polysaccharides solubilised during softening of «Barlett» pear fruit . Plant Physio!. 89, 1394 -1400 (1989). G'1!.oss, K. and C. E. SAMS: Changes in cell wall neutral sugar composition during fruit ripening: a species survey. Phytochemistry 23, 2457 - 2461 (1984). HULME, A . C. (ed.): The biochemistry of fruits and their products, Vo!' 1. Academic Press, London and New York (1970). JERMYN, M. A . and F. A. ISHERWOOD: Changes in the cell wall of pear during ripening. Biochemical Journal 64, 123-132 (1956). KNEE, M.: Effects of storage treatments upon the ripening of Conference pears. J . Science of Food and Agriculture 24, 1137 -1145 (1973). KNEE, M.: Metabolism of polymethylgalacturonate in apple fruit cortical tissue during ripening. Phytochemistry 17, 1261-1264 (1978). KNEE, M.: Fruit Softening II. Precursor incorporation into pectin by pear tissue slices. J. Exp. Botany 33, 1256 -1262 (1982). KOCH, J. and D . J. NEVINS: Tomato fruit cell wall: Use of purified tomato polygalacturonase and pectin methylesterase to identify developmental changes in pectins. Plant Physio!. 91, 816-822 (1989). LABAVlTCH, J. M. and P. M. RAy: Turnover of cell wall polysaccharides in elongating pea stem segments. Plant Physio!. 54,449-502 (1974). MARCELIN, 0., P. WILLIAMS, and J.-M. BRlllouET: Isolation and characterisation of the two main cell wall types from guava (Psidium guajava L.) pulp. Carbohydrate Research 240, 233 - 234 (1993). MONTlES, B.: Lignins. In: HARBORNE, J. B. (ed.): Methods in plant biochemistry Vol. 1. Academic Press, ISBN 0-12-461011-0. pp. 113 -157 (1989). PRESSEY, R. and J. K. AVANTl: Solubilisation of cell walls by tomato polygalacturonases: effects of pectinesterases. J. Food Biochemistry 6, 57 -74 (1982). REoGWELL, R . J. and R. R. SELVENDRAN: Structural features of cellwall polysaccharides of onion (Allium cepa). Carbohydrate Research 157, 183-199 (1986).
548
MARIA MARTIN-CABREJAS, KEITH W. WALDRON, and ROBERT R. SELVENDRAN
REDGWELL, R., L. D. MELTON, and D. J. BRASCH: Cell wall changes in kiwifruit following post-harvest ethylene treatment. Phytochemistry 29, 399-407 (1990). REDGWELL, R., L. D. MELTON, and D. J. BRASCH: Cell wall polysaccharides of kiwifruit (Actinidia deliciosa): effect of ripening on the structural features of cell wall materials. Phytochemistry 209, 191-202 (1991).
RYDEN, P. and R. R. SELVENDRAN: Structural features of cell-wall polysaccharides of potato (Solanum tuberosum). Carbohydrate Research 195, 257 -272 (1990 a). RYDEN, P. and R. R. SELVENDRAN: Cell wall polysaccharides and glycoproteins of parenchymatous tissues of runner bean (PhaseoIus cocineus). Biochemical Journal 269, 393 -402 (1990 b). SELVENDRAN, R. R.: Developments in the chemistry and biochemistry of pectic and hemicellulosic polymers. J. Cell Science, Supplement 2,51-88 (1985).
SELVENDRAN, R. R., A. V. F. VERNE, and R. M. FAULKS: Methods for the analysis of dietary fibre. In: LINSKENS, H. F. and J. F. JACKSON (eds.): Modern Methods of Plant Analysis, Vol. 10. pp. 235 -259. Springer, Berlin (1989). SEYMOUR, G. B., I. J. COLQUHOUN, M. S. DUPONT, K. R. PARSLEY, and R. R. SELVENDRAN: Composition and structural features of cell wall polysaccharides from tomato fruits. Phytochemistry 29, 725-731 (1990).
WALDRON, K. W. and R. R. SELVENDRAN: Cell wall changes in immature Asparagus stem tissue after excision. Phytochemistry 31, 1931-1940 (1992).
YAMAKI, S., Y. MACHIDA, and N. KAKIUCHI: Changes in cell wall polysaccharides and monosaccharides during development and ripening of Japanese pear fruit. Plant and Cell Physiology 20, 311-321 (1979).
Vol. 144. pp. 541 - 548 (1994)
Cell Wall Changes in Spanish Pear During Ripening MARIA MARTIN-CABREJASl, KEITH W. WALDRON2>~, 1
2
*
and ROBERT R.
SELVENDRAN2
Universidad Autonoma de Madrid, Departimento Chimica Institute of Food Research, Norwich Research Park, Colney, Norwich NR4 7UA Author for correspondence
Received December 28, 1993 . Accepted March 4, 1994
Summary
Unripe Spanish pears (Pyrus communis, c.v. Blanquille) were allowed to ripen at 18°C for 1 week. Cell walls were prepared as alcohol-insoluble residues (AIRs) which were free of starch. On a fresh weight basis, ripening was accompanied by a decrease in cell wall arabinose and uronic acid, a proportion of which were metabolised, and an increase in xylose and glucose. To investigate the origin of the changes in cell wall polymers, the AIRs of unripe and ripe fruit were extracted sequentially with water, cyclohexanetrans-l,2-diamine-N,N,N',N'-tetraacetate (CDTA), Na2 C03 at 1°C and 20°C, 0.5,1 and 4M KOH to leave a cellulose-rich residue. The extracts and residues were analysed for their carbohydrate composition and, where appropriate, degree of pectin-methyl esterification. The water, CDTA and Na2C03 extracted polymers were rich in pectic polysaccharides. These exhibited heterogeneity in the ratios of neutral and acidic components and contained small quantities of xylose. The KOH-extracted polymers were rich in hemicelluloses, particularly xylans, but also contained sugars common to pectic polysaccharides. The cellulose-rich residues were rich in cellulosic glucose and xylose which probably originated from the highlylignified sclereids. Ripening resulted in an increase in water- and CDTA-soluble pectic polysaccharides which exhibited a lower degree of pectin-methyl esterification. These results and the changes in carbohydrate composition are discussed in the light of corresponding decreases in the yields of pectic polysaccharides of the Na2 CO) extracts and cellulose-rich residues during ripening.
Key words: Cell walls, Pyrus communis, ripening, pectic polysaccharides, hemicelluloses, cell types. Abbreviations: AIR = alcohol-insoluble residue; CDTA = cyclohexane-trans-l,2-diamine-N,N,N',N'tetraacetate.
Introduction
It is generally accepted that changes in the cell wall polymers of pears and other fruits playa major role in determining the ripening-related alterations in fruit texture. However, our understanding of these changes is hampered by limited knowledge of the structure of the cell walls in mature unripe fruit and the enzymes which modify the cell wall polysaccharides (Brady, 1987). In pear, the most commonly reported change in cell wall integrity involves the increase in the soluble pectic polysaccharides. This is usually accompanied by an overall loss of cell wall arabinose Germyn and Is© 1994 by Gustav Fischer Verlag, Stuttgart
herwood, 1956; Dick and Labavitch, 1989; Ahmed and Labavitch, 1980) and, uronic acid Germyn and Isherwood, 1956; Bartley et aI., 1982). The mechanism by which cell wall pectic polysaccharides are solubilised, and the subsequent fate of the neutral and acidic carbohydrate residues in ripening pears is unclear. Most studies have investigated either the changes in the carbohydrate composition of the water-insoluble cell wall material (Gross and Sams, 1984) or the degree and extent to which pectic polysaccharides are solubilised by quantifying the release of water-soluble polyuronide (Knee, 1973; Yamaki et al., 1979; Ahmed and Labavitch, 1980). Few workers
542
MARIA MARTIN-CABREJAS, KEITH W. WALDRON, and ROBERT R. SaVENDRAN
have attempted to investigate the nature of the parent wallbound pectic polysaccharides in which ripening-related changes occur. In previous investigations on tomato (Seymour et al., 1990), apple (Whitcombe et a!., unpublished results) and Kiwi fruit (Redgwell et al., 1990; Redgwell et al., 1991) cell wall preparations have been extracted sequentially using techniques which minimise degradation of the constituent polymers. These studies have shown that pectic polysaccharides exhibit a range of solubilities which probably have a bearing on the type of cross-links which hold them into the cell wall matrix. Furthermore, such studies have shown that ripening-related changes are often associated with certain groups of pectic polysaccharides. In ripening pears, those studies which have attempted to fractionate the cell wall have confined themselves to extracting pectic polysaccharides by water and chelating agents (Ben-Arie et a!., 1979), or have used methods of cell wall extractions including hot water and strong alkali, which will have solubilised many of the constituent pectic polysaccharides through {3eliminative degradation Germyn and Isherwood, 1956), masking many of the ripening-related changes in the solubility of cell wall polymers. Furthermore, there is a paucity of information concerning the fate of acidic and neutral moieties which are lost from the soluble and insoluble fractions of the cell walls during ripening of pears. In Bartlett pears, the uronic acid solubilised during ripening can be recovered fully in polymeric form from the aqueous phase showing that complete depolymerisation of the polygalacturonic acid backbone does not occur (Ahmed and Labavitch, 1980). In the same variety, the ripening-related loss of cell wail arabinose has been attributed to the depolymerisation of rhamnogalacturonan-like pectic polysaccharide moieties after solubilisation (Dick and Labavitch, 1989). In contrast, fruit softening during ripening of Spadona pears is accompanied by an increase in free galacturonic acid, indicating extensive depolymerisation of the pectic polysaccharides (Ben-Arie et aI., 1979). The aim of the present study has been to determine whether or not the solubilised (degraded) arabinose and uronic acid-containing carbohydrate moieties are metabolised during the ripening of Spanish pears, and to investigate the fractions of the cell wall from which they are released by making use of modern methods of plant cell wall sequential extraction (Waldron and Selvendran, 1992; Coimbra et al., 1994).
Materials and Methods Plant material Mature green unripe Spanish pears (Pyrus communis L. cv. Blanquille) were obtained from the central fruit market of Madrid (Merca Madrid). Fruits were selected for uniformity in size and were free from blemishes. Fruits (samples of 25) were used immediately or after being allowed to ripen at 18°C for 1 week in darkness. Whole pears were peeled (epidermis and approximately 2 mm of underlying tissue) and the ovary and central vascular tissue excised. The fruit flesh was sliced and diced, frozen immediately in liquid nitrogen, and stored at - 40°C until required.
Preparation ofalcohol·insoluble residue (AIR) AIR. was obtained by immersing the frozen pear tissues in boiling ethanol (final concentration 85 % (v/v) aq.), homogenising with a Waring Blender and then Ystral homogeniser (maximum speed, 5 min) and boiling for 5 min. The material was filtered through a glass sintered filter (No.3, Scientific Furnishings Ltd., Sussex, UK) and re-extracted twice by boiling for 10 min in 85 % (v/v) aq. ethanol (5 x initial fresh weight) and filtering. The residue was resuspended in absolute alcohol (twice), acetone (twice), filtering between times, and then air-dried after which it was stored in a sealed container at room temperature.
Dialysis ofalcohol extracts The 85 % (v/v) aq. ethanol extracts recovered during preparation of AIRs was reduced in volume by rotary evaporation. The resultant liquor was dialysed overnight against water at 0 DC, from which the diffusate was recovered and stored at - 30°C. The volumes of both the dialysate and diffusate were recorded. The dialysate was then dialysed exhaustively against water at O°C after which it was reduced in volume and lyophilised. In calculating the recovery of the diffusate, corrections were made for the volume of the initial dialysate.
Fractionation ofAIR This method is based on that developed for cell wall material of onion bulbs (Redgwell and Seivendran, 1986). AIR. (2 g) was sequentially extracted with (i) water (100 mL) at 20°C for 2 h; (ii) 50 mM cyclohexane-trans-1,2-diamine-N, N, N', N'-tetraacetate (CDTA, Na salt, 100mL) pH 6.5 at 20°C for 5h (CDTA-1); (iii) CDTA, (100mL) pH 6.5 at 20°C at 2h (CDTA-2); (iv) 50mM Na2COJ + 20mM NaBH4 (l00mL) at O°C for 20h (Na2COJ-l); (v) 50mM Na2CO J + 20mM NaBH4 (100mL) at 20°C for 2h (Na2CO J-2); (vi) 1M KOH + 20mM NaBH4 (loomL) at 20°C for 2h and (vii) 4 M KOH + 20 mM NaBH4 (100 mL) at 20°C for 2 h . The alkali extractions were carried out with Orfree solutions under argon. After each extraction, the solubilised polymers were separated from the insoluble residue by centrifugation. The alkali extracts were acidified to pH 5 with HOAc and all extracts were filtered through glass fibre filter paper GFC (Whatman). The cellulose-rich residue remaining after the final extraction was also acidified to pH 5. The filtered extracts and residue were dialysed exhaustively, concentrated and lyophilised.
Analysis of carbohydrate composition Sugars were released from AIR. and sub-fractions by dispersing in 72 % H 2S04 followed by dilution to 1 M and hydrolysing for 2.5 h at 100 °C (Saeman hydrolysis). Sugars were released from dialysate and diffusate fractions by hydrolysing for 2.5 h at 100°C in 1 M H 2S0 4. All samples were subjected to duplicate analysis. Neutral sugars were reduced with NaBH4 and acetylated by the method of Blakeney et al. (1983) using 2-deoxyglucose as an internal standard. Alditol acetates were quantified by gas chromatography on a Carlo Erba Vega gas chromatograph after automatic injection by a Carlo Erba A 200 autoinjector. Alditol acetates were separated with baseline resolution on a Restek RTx 225 WCOT column (15mx 0.32 mm i.d.; 0.25 u film) using an oven temperature programme of 90°C for 1 min, 45°C per min to 150°C, 150 °C for 1 min, 2°C per min to 210°C, and 210 °C for 1.5 min. The carrier gas was helium at a column head pressure of 60 Kpa. Detection was by flame ionisation. Data was collected and integrated on a Spectra Physics
543
Cell walls of ripening pears Table 1: Yields and carbohydrate composition of AIRs from unripe and ripe fruit. Sample Unripe Ripe
Carbohydrate Composition (Mol %)
Yield (% Fwt)
Rha
Fuc
Ara
Xyl
Man
Gal
Glc
UA
(~g/mg)
2.7 2.7
1.0 1.3
1.1 1.1
17.1 9.1
27.1 32.5
1.2 1.7
3.4 3.1
21.3 (4.8)* 25.7 (7.8)*
27.9 25.5
646 543
Total
* 1 M hydrolysis.
SP4400 integrator; re-integration was handled by a Spetraphysics Winner data handling station. Total uronic acid content was determined colorimetric ally by the method of Blumenkrantz and AboeHansen (1973) after dispersal in 72 % H 2S0 4, dilution to 1 M, and hydrolysis for 1 h at 100 °C (Selvendran et aI., 1989).
Klason lignin The sample (100 mg) was dispersed in 72 % H2S0 4 for 3 h at room temperature followed by dilution to 1 M and hydrolysing for 2.5 h at 100°C. The residue was recovered by centrifugation (12,000 x g for 10 min), washed 3 times in distilled water and lyophilised. The Klason lignin was then quantified gravimetrically.
Results and Discussion
Carbohydrate composition ofAIRs and alcohol-soluble components Unripe and ripe pears were light green in colour and weighed approximately 145 g. Unripe pears were hard and inedible whilst ripe pears had become softer and more palatable. Cortical tissue from unripe and ripe years were prepared as AIRs. In each case, the yield of the AIR was 2.7 % (Fwt). As the pears were free of starch, as shown by negative staining with I2/KI and light microscopy, the AIR was used as the source of the cell wall material. Hydrolysis of the AIRs was performed using the Saeman method or 1 M H 2S0 4• The values given in Tables 1- 3 are the means of duplicate determinations and the variations between the duplicates was less than 2 %. The results reported in Tables 1 and 2 show the same trends in changes of cell wall com-
Table 2: Total carbohydrate composition of unripe and ripe pears. Sample
Carbohydrate recovery (~glg Fwt)
Unripe AIR EtOH dialysate Diffusate Total
Rha
Fuc
Ara
Xyl
0.16
0.18
2.62
4.16
0.19
Man
Gal
Glc
UA
Total
.
3.90 0.09
17.4
0.46
0.62 0.04 0.08
5.5
0.04 0.45
0.25
0.73
3.99
6.39
19.6
om
0.5
4.3 0.04
4.6 0.14 0.97
15.8 0.21 2.82
0.62
4.34
5.7
18.8
om
0.35
0.18
3.1
4.6
0.2
0.16
1.3
4.5 0.02
0.23 0.02
Ripe AIR E,OH dialysate Diffusate
0.16
0.05
0.44
1.1
*..
Total
0.36
0.21
1.7
5.6
0.28
* **
4.37 26.1 *.* 5.5 >tltl/>" .28.1
0.28
0.12
0.72
0. 1
0.31
1.90
Values referred to with asterisks: The majority of the glucose and man nose released during hydrolysis of the diffusate will have originated from intracellular sucrose, glucose and fruc· tose. For darity, their values have been omitted from the main body of the table.
ponents as those of preliminary experiments performed during the previous growth season. The carbohydrate composition of the AIR of unripe pears was rich in xylose and glucose which, together, comprised approximately half of the cell wall carbohydrate. The majority (75 %) of the glucose can be inferred to be cellulosic in origin since it was released only by Saeman hydrolysis. Since the level of cell wall xylose was over 5 times that of the non-cellulosic glucose, the presence of large quantities of xylan hemicelluloses can be inferred (Waldron and Selvendran, 1992). Approximately 25 % of the glucose was released by 1 M H 2S0 4, and it is likely that a significant quantity of non-cellulosic and non-starch glucans were present in the AIR. This will be discussed below. The remainder of the AIR carbohydrate comprised pectic polysaccharides as indicated by the levels of rhamnose, arabinose, and uronic acid. In comparison, the AIRs of ripe pears exhibited reduced levels of arabinose and uronic acid indicating a ripening-related decrease in branched pectic polysaccharides, and higher levels of xylose and glucose. The ripening-related decrease in pectic arabinose is consistent with the changes in the cell walls of Conference pears Germyn and Isherwood, 1956), and Bartlett pears (Ahmed and Labavitch, 1980; Dick and Labavitch, 1989). The substantial quantity of xylose and glucose in the AIRs and the ripening-related increase in these moieties, is due to the presence and continued maturation of sclereids (stone cells) during the ripening of Spanish pears. This is presently the subject of a more detailed investigation.
Carbohydrate composition of the 85 % {vlv} aq.-soluble components In order to determine the possible fate of the carbohydrate moieties that were lost from the pear cell walls during ripening, the 85 % (v/v) aq. ethanol extracts were retained, reduced under vacuum, and then dialysed exhaustively (10 changes) against water at 0 The first dialysis step was carried out against approximately 10 times the volume of the dialysis bag. After stirring overnight, the volumes of the equilibrated diffusate and dialysate solutions were quantified. The diffusate was reduced under vacuum and, because it proved difficult to freeze-dry, was dissolved in a larger quantity of water from which measured aliquots were taken. The dialysate was re-dialysed exhaustively, reduced in volume and lyophilised. The carbohydrate composition of the dialysate and diffusate samples were determined after hydrolysis in 1 M H 2S0 4 • To facilitate comparison, the data presented in Table 2 has been calculated on a fresh-weight basis and the recoveries for the diffusate have been corrected for the volume of the dialysate. The majority of carbohydrate released
0c.
544
MARIA
MARTIN-CABREJAS, KEITH W. WALDRON, and ROBERT R. SELVENDRAN
Table 3: Yields and carbohydrate composition of polymers extracted from cell walls of unripe and ripe pears. Sample
Carbohydrate composition (Ilg/ mg) Rha
Fuc
Ara
Xyl
Man
Gal
Glc
UA
Total
Water Unripe Ripe
0.8 0.9
1.2 1.5
21.7 22.0
7.1 6.7
0.9
1.1
7.6 5.1
4.8 3.6
55.8 59.2
781 692
19.2 25.1
CDTA-l Unripe Ripe
0.5 2.1
0.7 1.2
15.2 16.1
5.7 5.7
1.3 0.9
3.5 2.9
5.6 3.8
6704
67.6
244 320
5.9 7.2
CDTA-2 Unripe Ripe
1.7 1.5
2.0 2.3
12.7 14.1
4.5 8.6
3.6 2.8
2.2 2.9
55.1
lOA
5704
110 113
5.2
Residue 1 Unripe Ripe
0.7 0.8
0.02 0.1
14.9 5.6
35.1 39.5
0.04 0.3
3.3 3.8
35.1 44.7
10.9 6.2
644 655
66.6 61.8
Na2C03-1 Unripe Ripe
1.2 1.5
0.03
21.2
3404
2.79 6.8
0.2 0.8
3.2 3.3
0.8 3.0
63.1
5704
729 296
10.5 5.8
Na2C03-2 Unripe Ripe
1.0 3.1
59.7 29.7
4.0 6.5
0.2
3.2 4.7
0.8 2.22
31.1 52.7
817 387
2.0 1.9
0.5 M KOH Unripe Ripe
1.2 1.64
2.1
104
23.5 15.5
35.7 3704
1.8
IMKOH Unripe Ripe
0.9 2.7
0.9 1.6
18.2 20.8
19.8 29.3
4MKOH Unripe Ripe
0.5 0.6
2.0 1.8
3.8 3.3
Residue Unripe Ripe
0.7 0.9
0.3
9.7 3.0
1.1
18.2
Yield (% air)
204
104
7.1 7.2
17.0 16.7
12.9 17.6
1000 783
1.7 1.2
2.9
1.3
4.7 7.7
20.7
1404
39.9 14.2
261 638
1.2
51.3 45.2
3.2 3.7
6.5 6.1
19.6 16.7
13.2 22.7
742 591
17.1 26.3
22.0 28.0
1.5 1.6
2.7 1.5
40.1 43.3
23.1 21.7
336 360
31.9 24.2
during hydrolysis of the diffusate comprise Glcp and Manp. These will have originated from intracellular sucrose, glucose and fructose which impart a sweet flavour to pear flesh during ripening (Burton, 1982; Hulme, 1970) and have been presented beneath Table 2. The carbohydrate composition of the dialysate and diffusate of the Ethanol extracts of both unripe and ripe pear AIRs contained sugars common to pectic polysaccharides, as shown by the levels of uronic acid, galactose and, in the case of the diffusate, rhamnose and arabinose. Xylose was also present, particularly in the diffusate, and increased during ripening. The carbohydrate of the dialysate is likely to have been oligomeric in nature, but not polymeric, since it failed to precipitate in 85 % (v/v) aq. ethanol. Both oligomers and monomers are likely to have been present in the diffusate. There was little difference in the recovery of sugars of pectic origin from un-ripe and ripe pears in the diffusate, although the uronic acid exhibited a small increase. This is consistent with studies on Spadona pears (Ben-Arie et al., 1979) in which ripening was accompanied by an increase in free galacturonic acid. However, in the present study, the total recoveries of uronic acid, galactose and arabinose (from the AIRs, diffusates and dialysates), decreased during ripening,
104
showing unequivocally that a proportion had each been metabolised after the release from the cell wall. This contrasts with studies on Bartlett pear (Ahmed and Labavitch, 1980) in which total levels of polymeric uronic acid reportedly remained constant. In that study, the authors also reported that the total level of polymeric arabinose remained constant during ripening. However, re-calculation of their results confirm that total cell wall derived polymeric arabinose underwent a significant ripening-related decrease. Free arabinose had not been quantified.
Fractionation ofpear AIR's In order to identify polysaccharide fractions of the cell wall in which the ripening-related changes occurred, the cell wall (AIR) preparations were sequentially extracted with water at 20°C, 0.05 M CDTA (20 o q, 0.05 M Na2C03 at 1 °C and 20°C, and 0.5, 1 and 4 M KOH at 20°C, essentially as described before (Redgwell and Selvendran, 1986; Waldron and Selvendran, 1992). The extraction procedures were designed to minimise /1-eliminative degradation of pectins during the initial stages of extraction and to solubilise the cell wall polymers in as close to their native form as possible.
Cell walls of ripening pears
Data on the amounts of polymers solubilised and their sugar compositions are contained in Table 3. The amounts of material extracted are based on one sequential extraction of AIR from unripe and ripe pears. The pectic polysaccharides that were not cross-linked into the cell wall were solubilised by water at 20°C. Those held in the wall by Ca2+ only were subsequently extracted by CDTA. It is probable that the bulk of the water and CDTAsoluble pectic polysaccharides are of middle lamella origin (Selvendran, 1985). They consisted mainly of uronic acid, the predominant neutral sugar being arabinose. In addition, smaller but significant quantities of galactose were present. In spite of exhaustive dialysis, the carbohydrate recovery of the CDTA extracts was low. This can be attributed to the presence of contaminating CDTA which was detected by IR spectroscopy (results not shown). The incomplete removal of CDTA in this study is probably due to the presence of Ca2+ ions in the extract which complex with the CDTA and thus prevent its complete removal by dialysis. Similar difficulties have been encountered in CDTA extracts of olive pulp, and have been discussed in depth (Coimbra et al., in press). Much of the water and CDTA-insoluble pectic polysaccharides were subsequently solubilised with dilute Na2C03 at 1°C and 20°C, presumably by hydrolysis of weak ester cross links. The initial Na2C0 3 extractions were performed at 1 °C to minimise (3-eliminative degradation of the pectins in subsequent alkali extractions. Like the water and CDTA extracts, both Na2 C03 extracts were rich in pectic polysaccharides as shown by the high levels of uronic acid, arabinose and galactose, and the occurrence of rhamnose as the main deoxy sugar. As in other tissues (Ryden and Selvendran, 1990 a; Waldron and Selvendran, 1992), heterogeneity amongst the pectic polysaccharides was evident from the differing ratios of the neutral sugar (arabinose and galactose) both to each other and to the uronic acid component. For example, water-soluble pectic polysaccharides from unripe pear AIR exhibited an arabinose/galactose ratio of 2.8, and this ratio increased though the sequence of extractions to 18.7 in the Na2C03-2 extract. Interestingly, all the pectic polysaccharides solubilised by water, CDTA and Na2C0 3 contained small, but significant, quantities of xylose. The levels of xylose in the polysaccharides (3 - 9 mol %) were not only much greater than the amounts found in equivalent cell wall extracts from ripening tomatoes (Seymour et al., 1990), apples (Whitcombe et al., unpublished results) but were greater than the amounts found in equivalent cell wall extracts from maturing asparagus (Waldron and Selvendran, 1992), potato (Ryden and Selvendran, 1990a) and runner bean (Ryden and Selvendran, 1990 b). This is discussed further in the General Discussion. In addition to pectic polysaccharides, the 0.5 M KOH extractions solubilised significant quantities of hemicelluloses as indicated by the presence of xylose and glucose. The presence of glucose, xylose, galactose and fucose in the hemicellulosic extracts indicate the presence of fucogalactoxyloglucans. Since the levels of xylose were always considerably greater than those of glucose, the presence of xylan hemicelluloses could also be inferred. Interestingly, the 1 M and 4 M KOH extracts also contained significant quantities of pectic polysaccharides. This contrasts with the results of similar fractionation studies from
545
other fruits including tomatoes (Seymour et a1., 1990), apples (Whitcombe et al., unpublished results), and olives (Coimbra et a1., in press), and vegetables such as Asparagus (Waldron and Selvendran, 1992) and potato (Ryden and Selvendran, 1990 b). In all of these, the pectic component of these extracts was low. However, large quantities of pectic polysaccharides have been found in the 4 M KOH extract of cell walls of mung bean (Gooneratne et al. , unpublished results). In addition, the cell walls from the outer pericarp of Kiwi have also yielded significant quantities of pectic polysaccharides in the 4M KOH extract (Redgwell et al., 1990); however, this may arise because no previous 0.5 or 1 M alkali extractions had been performed. The remaining a-cellulose residue consisted mainly of glucose, xylose and uronic acid. The relatively low carbohydrate recovery in the residue (approximately 35 %) could be attributed to the large quantity of lignin present which originated from the residual stone cells. This was confirmed (a) by the measurement of Klason lignin values which were 60.1 % in the a-cellulose residue of un-ripe pear, and 52.2 % in the residue from ripe pear, and (b) by visual identification by light-microscopy. Interestingly, in spite of the relatively high Klason lignin values, the stone cells in the a-cellulose residues stained light pink with phloroglucinol-HCL (Fig. 1 a). However, those in the residue which remained after the initial water and CDTA extractions stained deep red (Fig. 1 b). This suggests that many of the phloroglucinol-HCLstaining functional groups in the stone cell lignin were either removed or modified during the alkali extractions. The formation of a red colour with phloroglucinol-HCI is indicative of coniferaldehyde, cinnamic aldehydes and related benzaldehydes (Monties, 1989). It is likely that the aldehyde groups in the lignin polymer were reduced by the NaBH4 during the 0.5, 1 and 4 M KOH extractions, eliminating their ability to interact with the stain. In contrast, the ability of the sclereids to stain turquoise with toluidine-blue was not altered by the sequential extractions indicating that the appropriate toluidine-blue-staining functional groups are still present (results not shown).
a Fig. 1: Light micrograph of phloroglucinol-HCl-stained sci ere ids of ripe Spanish pear (a) from CDTA-insoluble residue; (b) from cellulose-rich residue. Bar, 100 jJ.m.
546
MARIA MARTIN-CABREJAS, KErTH W. WALDRON, and ROBERT R. SELVENDRAN
5 '01
en
Table 4: Methyl-esterification (%) of uronic acid component of selected wall fractions from unripe and ripe pears.
4
x
E
AIR Water CDTA-1 CDTA-2 CDTA-2 Residue
~3
e ell
-g,2
.!:
o
... o
94 96 74 78 96
Unripe SD
8 3 4 3 8
x
Ripe SD
65
5
46
4
62 39 102
2 2 2
.D
u
W
CI
C2
NI
N2 O·5K IK
4K CR
cell wall extract Fig. 2: Yields of carbohydrate (mg anhydrosugar g-l fresh weight) from extracts and insoluble residues of unripe and ripe pear AIRs. W: water extract; Ct and C2: CDTA extracts t and 2; Nt and N2: Na2CO) extracts 1 and 2; 0.5 K, t K, 4 K: 0.5, 1 and 4 M KOH extracts; CR: cellulose-rich residue. 0 Unripe pear; • ripe pear.
Ripening related changes In order to make useful comparisons between the carbohydrate composition of the cell wall and .extracted. polym~rs from unripe and ripe pears, and to permlt comparlsons wlth the data in Table 2, the recoveries of total carbohydrate and selected sugars are presented on a Fwt basis basis in Figur~s 2 and 3 respectively. The results in Figure 2 show that dunng ripening, there is an increase in the water and CDTA-solub.le carbohydrate. This is accompanied by a greater decrease 10 the levels of carbohydrate solubilised by the Na2C03-1 and 2 extractions and to a lesser extent by decreases in the levels of carbohydr;te remaining in the a-cell~lose residue. The r!pening-associated loss of carbohydrate lS greater than the. 10crease in water-soluble and CDTA-soluble polysacchande, and is consistent with the decrease in total AIR carbohydrate (Tables 1 and 2). Since the ripening-related changes in carbo~4 '01 01
53 ell
"0
-0
>o
2
.!:
...
.D
o
hydrate are principally due to the changes in the levels of arabinose and uronic acid, the data for the recovery of these moieties throughout the cell wall fractions has been calculated and is shown in Figure 3. The results show that the increase in water-soluble uronic acid and arabinose, and the loss of these moieties from the cell wall, arise from the turnover of the pectic polysaccharides in the Na2C03-1 extract and, to a lesser extent, the Na2C03-2 extract and cellulosic residue. In addition to the carbohydrate composition, the degree of pectin methyl esterification was also quantified. In AIR, pectin methyl esterification decreased from 94 % (± 8 %) to 65% (± 5%) during ripening (Table 4). This is comparable to the trend observed in ripening Conference pears (Knee, 1982). To assess the origin of this change, the esterification of the pectic polysaccharides in each of the cold water extracts, CDTA extracts, and CDTA-insoluble residues, were also measured. The results (Table 4) demonstrate that the lower degree of methyl esterification in AIR of ripe pear results from the low levels of esterification of the water and CDTA-soluble pectic polymers; the methyl esterification of the residue in both unripe and ripe pears was above 95 %. This indicates that a significant proportion of the N a2 C03soluble polymers of unripe pears which become CDTA- and water-soluble during ripening also become de-esterified. The origin of the ripening-related increases in xylose was also investigated. Whilst the arabinose and uronic acid recoveries from the cell wall fractions were quantitative, (greater than 95 %), the total recovery of xylose was approximately 60 %. Hence, it is not possible to conclude which fractions of the cell wall contained the polymers which increase in xylose during ripening. Previously, it was noted t~at ripening resulted in an increase in «low» molecular welght xylosecontaining oligomers (Table 2). If similar oligomers comprising xylose were released from the cell wall durin~ the fractionation process, they may have been lost dunng the subsequent dialysis stages, thus accounting for the poor xylose recovery.
u
W CI
C2
NI
N2 O·5K IK
cell wall extract
4K CR
Fig.3: Yields of uronic acid and arabinose (~g anhydros~gar g-l fresh weight) from extracts and insoluble resldues of unnpe and ripe pear AIRs. W: water extract; C 1 and C2: CDTA extracts 1 and 2; Nt and N2: Na,CO) extracts t and 2; O.5K, lK, 4K: 0.5,1 and 4M KOH extracts; CR: cellulose-rich residue. Uronic acid: 0 unripe, • ripe; arabinose: Eil unripe; E8 ripe.
General Discussion Turnover of the carbohydrate moieties in the neutral side chains of cell wall pectic polysaccharides has been recognised in several types of cell wall, including ripening fruits (Gross and Sams, 1984; Seymour et aI., 1990; Redgwell, 1990) and the stems of dicots and monocots (Labavitch and Ray, 1974; Waldron and Selvendran, 1992). In order to understand the basis for such turnover, it is necessary to extract the cell
Cell walls of ripening pears
walls sequentially using solvents which minimise polymer degradation, and to identify the fractions in which turnover occurs. Such methods are essential, as the native pectic polysaccharides are easily susceptible to fj-eliminative degradation during extraction. Using such methods, the turnover of arabinose and uronic acid has been investigated during ripening of Spanish pears. The majority of this turnover occurs in the cold Na 2C0 3-s01uble polysaccharides, although some occurs in acellulose fraction. This contrasts with the turnover of pectic galactose during ripening of tomatoes (Seymour et al., 1990), where the loss of cell wall galactose could be attributed to its turnover in all of the cell wall fractions except for the CDTA-1 extracted polymers. However, the results reported in the present study exhibit trends that are similar to those found in ripening kiwi fruit (Redgwell et al., 1990; Redgwell et al., 1991) and apple (Whitcombe et al., unpublished results). The results showed that certain pectic polysaccharides became de-esterified during ripening. Such de-esterification has been associated with an increase in the susceptibility of polygalacturonic acid to the action of endo-polygalacturonase, as demonstrated in tomatoes (Pressey and Avants, 1982; Koch and Nevins, 1989). Ripening of tomatoes also results in a decrease in the degree of total pectin methyl esterification (Koch and Nevins, 1989). In contrast, such changes do not occur in ripening apples (Knee, 1978). However, the overall level of total pectin methyl esterification is not necessarily a true indicator of the events taking place. For example, Knee (1982) demonstrated that during ripening of Conference pears, newly synthesised methyl esterified pectic polysaccharides were continually inserted into the soluble fraction of the cell wall and subsequently turned over. The occurrence of xylose residues in the pectic polysaccharides solubilised by water, CDTA and Na2C0 3, from the cell walls of a variety of fruits and vegetables has been reported (Waldron and Selvendran, 1992; Ryden and Selvendran, 1990; Seymour et aI., 1990) and it is probable that these polysaccharides are derived from the middle lamella region (Selvendran, 1985; Bartley and Knee, 1982). The large quantity of xylose in the cellulosic residue is likely to have originated from (1-4}-linked xylan hemicelluloses from lignified sclereids (Fig. 3) which are scattered throughout the cortex. Some, however, may have been found in the walls of the cortical parenchyma, as in Guava fruit (Marcelin et aI., 1993) and olive (Coimbra et aI., unpublished results). The unusually high levels of xylose associated with the pectic polysaccharide extracts may reflect the occurrence of pectic-xylan complexes. We have detected an increase in pectic-xylanphenolic complexes during the maturation of asparagus stems (Waldron and Selvendran, 1992) and have linked this to the initial stages of lignification which occurs in the middle lamella. Similar events would be expected to be prevalent in newly-differentiated stone cells surrounding the sclereid clusters.
Acknowledgements This work was supported by the Comunidad Autonoma de Madrid, Consejeria de Educacion (M. Martin-Cabrejas) and the V .K . office of Science and technology.
547
References AHMED, A . E.and J. M LABAVlTCH: Cell wall metabolism in ripening fruit 1. Cell wall changes in ripening «Bartlett» pears. Plant Physio!' 65, 1009-1013 (1980). BARTLEY,!. M., M. KNEE, and M.-A. CASIMIR: Fruit Softening 1. Changes in cell wall composition and endopolygalacturonase in ripening pears. J. Exp. Botany 33, 1248 -1255 (1982). BARTLEY, 1. M. and M. KNEE: The chemistry of textural changes in fruit during storage. Food Chemistry 9, 47 -58 (1982). BEN-ARlE, R., L. SONEGO, and C. FRENKEL: Changes in pectic substances in ripening pears. Journal of the American Society for Horticultural Science 104, 500-505 (1979). BLAKENEY, A. B., P. J. HARRIS, R. J. HENRY, and B. A. STONE: A simple and rapid preparation of alditol acetates for monosaccharide analysis. Carbohydrate Research 113, 291-299 (1983). BWMENKRANTZ, N. and G. ASBOE-HANSEN: New method for quantitative determination of uronic acids. Analytical Biochemistry 54,484-489 (1973). BRADY, C. J.: Fruit ripening. Annu. Rev. Plant Physio!' 38, 155178 (1987). BURTON, W. G.: Post harvest physiology of food crops. Longman, London. 339 pp. (1982). COIMBRA, M. A., K. W. WALDRON, and R. R. SELVENDRAN: Isolation and characterisation of cell wall polymers from olive pulp (Olea Ouropaea L.). Carbohydrate Research (1994; in press). DICK, A. J. and J . M. LABAVlTCH: Cell wall metabolism in ripening fruit. IV. Characterisation of the pectic polysaccharides solubilised during softening of «Barlett» pear fruit . Plant Physio!. 89, 1394 -1400 (1989). G'1!.oss, K. and C. E. SAMS: Changes in cell wall neutral sugar composition during fruit ripening: a species survey. Phytochemistry 23, 2457 - 2461 (1984). HULME, A . C. (ed.): The biochemistry of fruits and their products, Vo!' 1. Academic Press, London and New York (1970). JERMYN, M. A . and F. A. ISHERWOOD: Changes in the cell wall of pear during ripening. Biochemical Journal 64, 123-132 (1956). KNEE, M.: Effects of storage treatments upon the ripening of Conference pears. J . Science of Food and Agriculture 24, 1137 -1145 (1973). KNEE, M.: Metabolism of polymethylgalacturonate in apple fruit cortical tissue during ripening. Phytochemistry 17, 1261-1264 (1978). KNEE, M.: Fruit Softening II. Precursor incorporation into pectin by pear tissue slices. J. Exp. Botany 33, 1256 -1262 (1982). KOCH, J. and D . J. NEVINS: Tomato fruit cell wall: Use of purified tomato polygalacturonase and pectin methylesterase to identify developmental changes in pectins. Plant Physio!. 91, 816-822 (1989). LABAVlTCH, J. M. and P. M. RAy: Turnover of cell wall polysaccharides in elongating pea stem segments. Plant Physio!. 54,449-502 (1974). MARCELIN, 0., P. WILLIAMS, and J.-M. BRlllouET: Isolation and characterisation of the two main cell wall types from guava (Psidium guajava L.) pulp. Carbohydrate Research 240, 233 - 234 (1993). MONTlES, B.: Lignins. In: HARBORNE, J. B. (ed.): Methods in plant biochemistry Vol. 1. Academic Press, ISBN 0-12-461011-0. pp. 113 -157 (1989). PRESSEY, R. and J. K. AVANTl: Solubilisation of cell walls by tomato polygalacturonases: effects of pectinesterases. J. Food Biochemistry 6, 57 -74 (1982). REoGWELL, R . J. and R. R. SELVENDRAN: Structural features of cellwall polysaccharides of onion (Allium cepa). Carbohydrate Research 157, 183-199 (1986).
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MARIA MARTIN-CABREJAS, KEITH W. WALDRON, and ROBERT R. SELVENDRAN
REDGWELL, R., L. D. MELTON, and D. J. BRASCH: Cell wall changes in kiwifruit following post-harvest ethylene treatment. Phytochemistry 29, 399-407 (1990). REDGWELL, R., L. D. MELTON, and D. J. BRASCH: Cell wall polysaccharides of kiwifruit (Actinidia deliciosa): effect of ripening on the structural features of cell wall materials. Phytochemistry 209, 191-202 (1991).
RYDEN, P. and R. R. SELVENDRAN: Structural features of cell-wall polysaccharides of potato (Solanum tuberosum). Carbohydrate Research 195, 257 -272 (1990 a). RYDEN, P. and R. R. SELVENDRAN: Cell wall polysaccharides and glycoproteins of parenchymatous tissues of runner bean (PhaseoIus cocineus). Biochemical Journal 269, 393 -402 (1990 b). SELVENDRAN, R. R.: Developments in the chemistry and biochemistry of pectic and hemicellulosic polymers. J. Cell Science, Supplement 2,51-88 (1985).
SELVENDRAN, R. R., A. V. F. VERNE, and R. M. FAULKS: Methods for the analysis of dietary fibre. In: LINSKENS, H. F. and J. F. JACKSON (eds.): Modern Methods of Plant Analysis, Vol. 10. pp. 235 -259. Springer, Berlin (1989). SEYMOUR, G. B., I. J. COLQUHOUN, M. S. DUPONT, K. R. PARSLEY, and R. R. SELVENDRAN: Composition and structural features of cell wall polysaccharides from tomato fruits. Phytochemistry 29, 725-731 (1990).
WALDRON, K. W. and R. R. SELVENDRAN: Cell wall changes in immature Asparagus stem tissue after excision. Phytochemistry 31, 1931-1940 (1992).
YAMAKI, S., Y. MACHIDA, and N. KAKIUCHI: Changes in cell wall polysaccharides and monosaccharides during development and ripening of Japanese pear fruit. Plant and Cell Physiology 20, 311-321 (1979).