Journal of Experimental Botany, Vol. 53, No. 368, pp. 513-523,
March 1, 2002
© 2002 Oxford University Press
Original Papers |
Organization of cell walls in Sandersonia aurantiaca floral tissue
Crop and Food Research Ltd., Private Bag 11600, Palmerston North, New Zealand
Received 6 August 2001; Accepted 2 November 2001
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Abstract |
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Visual symptoms of the onset of senescence in Sandersonia aurantiaca flowers begin with fading of flower colour and wilting of the tissue. When fully senescent, the flowers form a papery shell that remains attached to the plant. The cell walls of these flowers have been examined to determine whether there are wall modifications associated with the late stages of expansion and subsequent senescence-related wilting. Changes in the average molecular size of pectin were limited through flower opening and senescence, although there was a loss of neutral sugar-containing side-branches from pectins in opening flowers, and the total amounts of pectin and cellulose continued to rise in cell walls of fully senescent sandersonia flowers. Xyloglucan endotransglycosylase activity increased in opening and mature flowers, but declined sharply as flowers wilted. Concomitantly, the proportion of hemicellulose polymers of increasing molecular weight increased from flower expansion up to the point at which wilting occurred. Approximately 50% of the non-cellulosic neutral sugar in mature flower cell walls was galactose, primarily located in an insoluble cell wall fraction. Total galactose in this fraction increased per flower with maturity, then declined at the onset of wilting. ß-Galactosidase activity was low in expanding tepals, but increased as flowers matured and wilted.
Key words: Cell walls, flower senescence, galactose, molecular weight, XET.
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Introduction |
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Flower opening and petal senescence are highly controlled biological events critical in the life cycle of flowering plants. They are also events of commercial value, contributing to the visual appeal and post-harvest vase-life of cut flowers. There are questions whether flower opening and the onset of senescence-indicative deteriorative changes are likely to be a developmental continuum (with opening as the first visual symptom of a senescence programme that has already started), rather than separate but coexisting processes (senescence starts only when flower maturity is reached).
There is enormous variation among flowering plants in the manner and timing of both the flower opening phase and the onset of petal senescence. As an example, Asiatic lily opening is achieved by petal cell expansion, with uneven expansion of the different epidermal layers of the petal contributing to the development of the open flower shape (Bieleski et al., 2001
). By contrast, the fabric of the morning glory flower has formed before opening, and opening is achieved by unfolding the massed petal tissue, although this also appears to require selective cell expansion (Phillips and Kende, 1980). Studies of the factors controlling cell expansion have focused primarily on cells of growing shoots. In these tissues, expansion is controlled by cellular turgor pressure and stress on the cell wall. Expansins, non-enzymic proteins which may disrupt the tight arrangement of hemicelluloses with cellulose microfibrils, are thought to be a fundamental factor in the ability of the wall to extend (Cosgrove, 1998, 1999), but modification of covalent linkages within existing cell wall polymers as well as deposition of new cell wall material ensure the wall retains strength during expansion (Cosgrove, 1997). Gene expression relating to the presence of expansins has been identified in young pea petals (Michael, 1996) and in tomato flowers (Brummell et al., 1999), and polygalacturonase gene expression has been detected in kiwifruit flowers at anthesis (Wang et al., 2000). This suggests that the cell wall may be similarly modified in opening petals although the extent is likely to be strongly reliant on the nature of the polysaccharides present.
Factors associated with petal senescence have been reviewed most recently by Rubinstein who focused on the increasing molecular evidence supporting programmed cell death in petal senescence, determined by specific gene expression changes and signalling events (Rubinstein, 2000). Petal wilt is associated with the onset of senescence in many flowers and is related to loss of membrane integrity, changes in turgor and ion leakage. Recent studies indicate that factors controlling these changes are initiated prior to visual wilt and involve new protein synthesis (Celikel and van Doorn, 1995; van Doorn et al., 1995; Panavas and Rubinstein, 1998).
The loss of cell wall integrity is closely associated with tissues undergoing senescence, most notably in the ripening-related softening of many fruits. Catabolic processes such as solubilization and breakdown of cell wall polymers (Fischer and Bennett, 1991) dominate in this situation, although there are reports of new cell wall synthesis also occurring (Mitcham et al., 1989). There are also a small number of studies describing significant cell wall modification in flower petals that are undergoing changes to texture during senescence, such as quantitative loss of hemicellulose in senescing morning glory (Wiemken-Gehrig et al., 1974) and increased pectin solubilization and loss of large-sized hemicellulose polymers in senescing petals of cut carnation flowers (de Vetten and Huber, 1990; de Vetten et al., 1991). Panavas et al. found increasing activity of cell wall hydrolases in senescing daylily, indicating the potential for wall breakdown (Panavas et al., 1998). Up-regulated expression of a polygalacturonase gene has also been found in senescing kiwifruit flowers (Wang et al., 2000). With this in mind the cell walls and some cell wall-related hydrolases of Sandersonia aurantiaca flowers have been analysed during opening and subsequent senescence, with the primary aim of determining whether there are specific elements of wall modification that occur during these events, especially with reference to changes in petal texture.
Sandersonia aurantiaca (Hook.) is a liliaceous monocotyledon with a bell-shaped corolla formed from fused petals and sepals (tepals). Sandersonia flowers have been used as models for the study of senescence in ethylene-insensitive floral systems (Eason and Webster, 1995; Eason and de Vré, 1995; Eason et al., 1997, 2000a, b). The progression of bud appearance, flower opening and senescence in the sandersonia flower proceeds in a manner that provides reliable visual indicators for flower selection. These developmental stages have been linked to quantitative measurements of intensifying and fading colour, loss of chlorophyll and protein, and changes to fresh weight, dry weight and starch content (Eason and Webster, 1995; Eason et al., 1997). Pollination does not affect the onset or progression of senescence in sandersonia (JR Eason, personal communication). New protein synthesis, central to the co-ordination of senescence events in sandersonia, is thought to be linked to changes in gene expression that begin prior to the visual symptoms of deterioration (Eason et al., 1997, 2000a, b).
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Materials and methods |
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Plant material
Sandersonia tubers were grown in a plastic house as described previously (Eason and Webster, 1995
). Flowers were selected according to the criteria described earlier (Eason and Webster, 1995) and are shown in Fig. 1A, I–V. In brief, flowers in bud (included for some experiments) have aperture tips that are pointed, green and closed. Opening flowers are defined by an open, green aperture with the tips turned back, while mature flowers are completely opened and are fully orange with no trace of green coloration. The subsequent wilting phase affects the turgidity of the whole flower and there is a loss of colour intensity. Senesced flowers are pale brown, do not resist compression but are not brittle. Subsequent to this the flowers become dry and shrivelled but still remain attached to the pedicel. Eason and Webster indicated that for sandersonia the average developmental time from bud was 2.5 d (opening), 5.5 d (mature), 9.5 d (wilted), and 11.5 d (senesced) (Eason and Webster, 1995).
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Tissue preparation
Fresh tissues from the bell region of sandersonia flowers were fixed in ethanol:acetic acid:formalin:water (10:1:2:7; by vol.), equilibrated in ethanol, followed by Histoclear, then embedded in paraffin wax. Thin sections were stained with toluidine blue (1%) and examined using light microscopy.
For other procedures, the stem, reproductive tissues and nectaries were discarded and the remaining tepal tissue (encompassing the bell and aperture regions) of each flower was weighed, frozen in liquid N2 then stored at -80 °C. Frozen tissue was subsequently used for cell wall preparations or freeze-dried.
Tensile and compressive strength
The strength of fresh tepal tissue was measured by measuring the force required to pull tepal strips apart at a 90° angle to the vascular tissue. Tepal strips (approximately 15 mm long, 4 mm wide) were excised from the bell region of each of 10 flowers. Adhesive tape was attached to the sides of the strip, and the strip width and thickness measured using digital callipers. The taped edges were clamped into an Instron Universal Testing Machine (Model 4301) and the tepal pieces were pulled apart at a crosshead speed of 50 mm min-1. Load and displacement data were calculated using Instron Series IX software (Version 5.34.00). The strength required to pull the tepal tissue apart was expressed on a per strip (N mm-1) and per area (N mm-2) basis. In both cases the width of the strip was taken into account, whereas for strength per area calculations the thickness of the tepal strip was also included. Flower firmness was measured as the force (N) required to compress whole flowers through a distance 20% of the widest diameter (according to the method of Eason et al., 1997).
Cell wall compositional analyses
Ethanol-insoluble residue (EIR) was prepared by homogenizing frozen tepal tissue in ethanol and then chilling at -20 °C for 16 h. The homogenates were vacuum-filtered through several layers of organza fabric (obtained locally), then treated with buffered phenol to inactivate wall-bound enzymes (Huber, 1992). The EIR was air-dried, then ground to a powder and stored at -80 °C.
The EIR was assayed for cellulose content (Dische, 1953; Updegraff, 1969), and for total pectin (Ahmed and Labavitch, 1977). Pectin esterificiation was estimated using the protocol of Wood and Siddiqui with volumes adjusted for microassays in 96-well microtitre plates (Wood and Siddiqui, 1971).
Soluble pectins were sequentially extracted from EIR by 50 mM trans-1,2-cyclohexanediamine-N,N,N',N'-tetraacetic acid (CDTA) in 50 mM NaOAc, pH 6.0, followed by 50 mM Na2CO3 with 20 mM NaBH4. The proportion of EIR to extractant was 1 mg:0.5 ml. Both extractions were at room temperature for 4 h with constant shaking. Hemicelluloses were extracted from EIR in 6 N NaOH containing 0.13 mM NaBH4, using the method described earlier (O'Donoghue et al., 1997).
For all the extractions, uronic acids were determined by the method of Blumenkrantz and Asboe-Hansen (Blumenkrantz and Asboe-Hansen, 1973), and total carbohydrates were assayed using the phenol:sulphuric acid method (Dubois et al., 1956).
Neutral sugar analysis
Neutral sugars in the EIR were hydrolysed in 2 N trifluoroacetic acid and then derivatized to alditol acetates (Blakeney et al., 1983). The alditol acetates were quantified by gas chromatography using a BPX-70 capillary column (0.53 mmx25 m, SGE) and flame ionization detection. Allose was used as an internal standard. The temperature programme was as follows: 170 °C (2 min), 10 °C min-1 to 190 °C, then 7 °C min-1 to 240 °C (0.25 min).
In order to quantify galactose in cell wall fractions, sequential extractions of EIR, using CDTA, Na2CO3 and 6 N NaOH were prepared. CDTA and Na2CO3 extracts were dialysed against 9.0 l water (2x12 h) prior to derivatization. The 6 N NaOH-soluble material was not passed through anion exchange columns prior to analysis. Insoluble extraction residues were washed with water until neutral, then freeze-dried. All extracts and insoluble extraction residue from each developmental stage were analysed for galactose content by gas chromatography, using the derivatization method described above.
Molecular size distribution
Molecular size distribution of extracted cell wall fractions was evaluated using size-exclusion chromatography. CDTA- and Na2CO3-soluble pectin extracts (approximately 500 µg uronic acid ml-1) were applied to a Superose 6HR size exclusion column (10 mmx300 mm, Pharmacia) operating in 30 mM NaOAc, pH 6.5, containing 20 mM NaCl and 10 mM EDTA. Flow rate was 0.5 ml min-1 and 0.5 ml fractions were assayed for uronic acids (Blumenkrantz and Asbose-Hansen, 1973) and total carbohydrates (Dubois et al., 1956) in 96-well microtitre plates. Molecular size profiles are representative of four extractions of CDTA- and Na2CO3-soluble pectins from each developmental stage.
Hemicelluloses were similarly separated using a Superose 6HR column except that the elution buffer was 50 mM NaOAc, pH 5.0, containing 10 mM NaCl. Fractions were assayed for total carbohydrates (Dubois et al., 1956) and for xyloglucan using an iodine binding assay (Kooiman, 1960) in 96-well microtitre plates. Hemicellulose molecular size profiles are representative of two extractions from each developmental stage.
Analysis of cell wall enzyme activity
ß-Galactosidase:
Freeze-dried tissue was extracted in 50 mM NaOAc, pH 4.5 (±1 M NaCl) and extracts were assayed for ß-galactosidase activity using p-nitrophenyl-ß-D-galactopyranoside as substrate. The method of Tanimoto and Igari was used (Tanimoto and Igari, 1976), with modifications (O'Donoghue et al., 1998). Activity is expressed as U flower-1 where 1 U=1 µmol nitrophenol released h-1. Assays were performed in the presence of up to 30 µM galactose that co-solubilized in the extract.
Pectinmethylesterase (PME):
Freeze-dried tissue (20 mg) was extracted in 50 mM NaOAc pH 6.0, with 1 M NaCl, and aliquots were incubated with 0.2% citrus pectin and extraction buffer without NaCl for 16 h at 40 °C. Methanol released was assayed (Wood and Siddiqui, 1971) according to the method adapted for a microtitre plate. PME activity is expressed as U flower-1 where 1 U=1 nmol methanol released h-1.
Xyloglucan endotransglycosylase (XET):
Freeze-dried tissue (20 mg) was extracted with 1.2 ml buffer (250 mM NaOAc, 400 mM NaCl, pH 5.8) with 20 µl of 625 mg ml-1 dithiothreitol and 30 mg polyvinylpolypyrrolidone. XET was assayed using basic procedures (Fry et al., 1992; Redgwell and Fry, 1993), with the exception that the substrate consisted of asparagus xyloglucan and tritiated asparagus-derived xyloglucan oligomers buffered with 1 M NaOAc, pH 5.8, prepared and used as described previously (O'Donoghue et al., 2001). Activity was measured by the incorporation of the tritiated xyloglucan oligomer into the xyloglucan polymer using scintillation counting, and is expressed as U flower-1, where 1 U=Bq (kBq supplied. h)-1.
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Results |
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Cross-sections of toluidine blue-stained sandersonia tepal tissue are shown in Fig. 1B
, I–V, alongside photographs of whole flowers at matching developmental stages in (A). Flowers at the bud stage (B, I) had a clearly defined layer of parenchyma cells between the upper and lower epidermis. This layer became progressively disordered as the flowers began to open (B, II), reached full size (B, III) and wilted (B, IV), with large spaces becoming apparent. In fully senesced flowers (B, V) the parenchyma structure was completely collapsed and tepal tissue appeared to consist of an upper and lower epidermis, with intact vascular tissue existing within a compressed strip of the remains of the parenchyma cells.
The total fresh weight of developing and senescing sandersonia flowers is shown in Fig. 2A, with the contributory water content and dry matter flower-1 embedded in the same bar for each developmental stage, and the ratio of water:dry matter per flower as a line graph. The maturity-related increase, as well as the wilting-related decrease in average fresh weight flower-1 was due to changes in both water and dry matter content of sandersonia flowers. The proportions of water and dry matter per flower did not change from opening to wilting, but a change in this ratio occurred in senescing flowers where dry matter loss flower-1 did not keep pace with water loss. The firmness of whole tepals, measured objectively by compressibility, increased as the flowers opened and was greatest in the mature flowers (Fig. 2B). Firmness of wilted flowers was only 10% of that at maturity. There was no difference between the firmness of wilted and senesced flowers.
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) at each stage (LSD=0.7). In each case, df=15, 
=0.05. (B) Compression strength (N) of sandersonia flowers at each developmental stage (LSD=0.063, df=15, Tepal tissue strength per tissue strip was highest in expanding and fully opened, mature flowers, but declined as flowers began to wilt (Table 1
). The most fragile tissues were those of senesced tepals. Tepal tissue from post-mature flowers was thinner than that of opening or mature flowers (data not shown). When the tepal thickness is accounted for in the strength calculation, there was no change in the tissue strength per unit area (fixed width of the strip [4 mm]xthickness) until flowers were fully senesced, at which point the strength per unit area more than doubled.
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The amount of EIR flower-1 increased as flowers became fully mature (Fig. 3A
), but decreased as flowers began to wilt and finally senesced. Starch levels in these preparations were 
8 µg mg-1 EIR in opening flowers and <3 µg mg-1 EIR in the succeeding stages (data not shown). The amount of cellulose and pectin (flower-1 basis) increased as flowers expanded to maturity, remained at these levels in wilted flowers, but increased again in fully senesced flowers (Fig. 3B, C). There were no changes in esterified pectin content (% total pectin basis) until flowers became fully senescent. Levels decreased from 35.9% (±0.6) at the onset of wilting to 30.4% (±0.6) in the senesced tepals. Hemicellulose levels ranged from 1–2 mg per flower and no distinct trends in hemicellulose content were apparent in the developmental stages sampled (data not shown).
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The quantity of CDTA-soluble pectin flower-1 was similar in all developmental stages until flowers were fully senescent, when there was an almost 25% loss of this pectin fraction (Fig. 3D
). On the other hand, the amount of Na2CO3-soluble pectin flower-1 declined gradually in successive stages with a significant drop in this fraction as flowers began to wilt (Fig. 3E). The pectin accumulation in senesced flowers must be due to pectins other than those soluble in CDTA or Na2CO3.
The predominant non-cellulosic neutral sugar in cell walls of opening and mature sandersonia flowers was galactose (46–50 mol%, Table 2). The levels of arabinose, xylose and glucose were lower (between 10–12 mol% each). As the flowers began to wilt, the most notable feature was the reduction in the proportion of galactose present. This trend continued as the flowers approached complete senescence.
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Throughout sandersonia flower development the cell wall galactose was primarily located in polysaccharides that were insoluble in CDTA, Na2CO3 or 6 N NaOH (Table 3
). The amount of galactose in this insoluble residue (flower-1 basis) almost doubled in mature flowers, but decreased significantly as the flowers wilted and senesced. Cell wall galactose was partitioned in much lower amounts into polymers soluble in CDTA, Na2CO3 and 6 N NaOH. As sandersonia flowers matured there was some accumulation of galactose into CDTA-soluble polymers, but an almost 35% loss of galactose from Na2CO3-soluble polymers. The amount of galactose decreased in all three solubilized fractions with the onset of wilting and senescence, but these losses did not approach the amount of galactose lost from insoluble residues at the same stages.
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Molecular size distribution
The average molecular size distribution profiles of CDTA-soluble pectins extracted from sandersonia tepals are shown in Fig. 4
. In opening flowers, most of the uronic acid content eluted as one peak, beginning very close to the void volume of the column and with extensive tailing (Fig. 4A). Analysis of the fractions for total carbohydrates using phenol:sulphuric acid, revealed an extra dimension to this pectin profile (Fig. 4B). Phenol:sulphuric acid detects both neutral and acidic sugars, although it is more sensitive to the neutral sugar component. A second peak, eluting between the 500 kDa and 70 kDa size markers was evident, but not fully resolved from the first, largest peak. This may represent the neutral sugar branch component of a small group of pectins that were water-soluble and co-extracted in CDTA. A third peak eluted very close to the inclusion volume of the column. Neutral sugar analysis of the CDTA-soluble extracts, without prior trifluoroacetic acid hydrolysis, confirmed the presence of sugar monomers that would have eluted in this peak. The mol% of neutral sugars ranged from 80% glucose in opening flowers to 50% glucose/25% rhamnose in senesced flowers. To investigate whether there could be neutral sugar-containing oligomers present in CDTA-soluble extracts (suggested by the broadness of the third peak), fractions corresponding to this peak were collected, concentrated, then separated on a Superdex Peptide column (Pharmacia; carbohydrate exclusion limit, 10 kDa, inclusion limit 0.5 kDa). The majority of carbohydrate eluted at the inclusion volume, but a small proportion from extracts of all developmental stages did fractionate (data not shown) suggesting the presence, albeit minor, of carbohydrate oligomers.
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The size distribution of the uronic acid component of CDTA-soluble pectins from sandersonia did not alter appreciably with increasing maturity, wilting or advanced senescence of the tepals. On the other hand there were significant changes in both the mid- and smallest-sized polymer groups whose distributions became more prominent as flowers matured and wilted, and broader/flatter as flowers approached full senescence.
The molecular size profiles of polymers soluble in Na2CO3 were distinctly different from CDTA-soluble pectins (Fig. 5). The uronic acid component of these fractions eluted in one very broadly distributed peak. The shape of the leading edge of this peak suggests that there were void-volume-sized polymers present in opening flowers (Fig. 5A), whereas in subsequent stages (Fig. 5B–D) there were proportionately more polymers in the lower size ranges. This was confirmed by the phenol:sulphuric acid analysis of the column fractions which showed the gradual disappearance of the earliest-eluting polymers of relatively large molecular size. A second peak, as a shoulder to the main body of the distribution, became less evident in the same way. A comparison of total carbohydrate and uronic acid distributions of this Na2CO3-soluble extract suggests that the group of relatively large polymers contained a high proportion of neutral sugars, presumably as side-branches to the main chain of uronic acids. In senesced sandersonia flowers, Na2CO3-soluble pectins had similar total carbohydrate and uronic acid distributions, suggesting that neutral sugars were either equivalently dispersed over the pectin chains, or that they were absent altogether.
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The molecular weight distribution of hemicelluloses extractable in 6 N NaOH is shown in Fig. 6
. There were two predominant size groups of hemicelluloses in sandersonia, the largest eluting between the column void volume and the 500 kDa dextran marker and the other eluting between the 40 and 9.3 kDa size markers. A third, less distinct, size cluster was centred around the 73 kDa marker. As flowers developed from opening to wilting, the peak containing the largest-sized polymers moved to a position of larger average molecular size. The elution position of the smallest-sized group remained unchanged, although the relative amounts present were altered. The intermediate cluster became less distinguishable in wilted flowers. In fully senesced flowers there was a downshift in the peak containing the largest-sized polymers so that the elution profile resembled that of hemicelluloses from mature tepals. The xyloglucan molecular size distribution closely followed that of the large-sized hemicelluloses (Fig. 6B), with average molecular size increasing until flowers wilted, then decreasing in flowers that were fully senesced.
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) and senesced (
) sandersonia tepals, fractionated on Superose 6HR. Distributions are expressed as percentage units of the total detected. Tick marks at the top of each graph indicate the elution positions of dextran size markers, in the order 5x103 kDa, 500 kDa, 73 kDa, 40 kDa, and 9.3 kDa.
Cell wall hydrolases
Both colorimetric and molecular size analyses by size-exclusion chromatography revealed an inability of fresh tissue extracts from any of the sandersonia samples to hydrolyse either partially de-esterified citrus pectin or carboxymethylcellulose, indicating very low or no endo-polygalacturonase and ß-1,4-glucanase-type activity, and this was not pursued further.
ß-Galactosidase activity (flower-1 basis, Table 4), low in buds and opening flowers, was significantly higher in mature flowers. There was a further, almost doubling of activity as tepals wilted, then a sharp decrease as the flowers entered the final stage of senescence. With 1 M NaCl included in the extraction buffer, this activity pattern was retained, but all levels were elevated, indicating the presence of a proportion of tightly wall-bound enzyme.
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PME activity was low in sandersonia flowers and assays required long incubation times in order to obtain detectable activity. Nevertheless, PME activity (flower-1 basis, Table 5
) was greatest in opening and mature flowers then declined as flowers wilted. The predominance of XET activity (flower-1 basis, Table 5) was also associated with opening of sandersonia flowers. There was low XET activity in buds, but an almost 5-fold increase occurred as sandersonia flowers swelled and the aperture expanded. XET activity was highest in fully mature flowers, followed by sharp declines in wilted and senesced flowers.
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Discussion |
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Some of the cell wall characteristics of sandersonia flowers have been described as the flowers open and then senesce. It is interesting that the opening of sandersonia flowers is accompanied by considerable disruption to the cell organization in the parenchyma layer of the tepal tissue, similar to opening daylily (Panavas et al., 1998
). This ‘looseness’ may permit lateral expansion of the flower tissue. The parenchyma cells also appear to break down considerably, so that in senesced flowers only epidermal cells and vascular tissue are intact. The main interest was in determining whether there were distinctive changes, occurring after Sandersonia flowers reached maturity, that might contribute to the textural changes associated with wilting, and whether there were elements of cell wall modification, initiated during flower opening, that continued unchecked through to senescence. The cell walls of only two other species, daylily and carnation, have been studied in great detail with respect to this period in flower development. Sandersonia, daylily and carnation flowers differ greatly in the combinations of ethylene sensitivity, visual senescence symptoms and speed of deterioration after opening. There is new evidence that there are few common features with respect to cell wall characteristics of their opening and senescence.
There was little indication of pectin hydrolysis in sandersonia flowers, and there was detectable endo-polygalacturonase activity at any developmental stage. Increasing polygalacturonase activity was reported in senescing daylily petals (Panavas et al., 1998), indicating the potential for pectin backbones to be hydrolysed in these flowers, but carnation petals have little pectin hydrolysis during senescence (de Vetten and Huber, 1990). The carnations of that study, however, were cut and then supplied with sucrose solution rather than senescing naturally, attached to the parent plant.
A sharp increase was found in XET activity associated with sandersonia opening. It is likely that, as with growing shoots, XET is part of a secondary response of wall modification that supports the initial stress relaxation in expanding cell walls, and reforms linkages to retain wall strength. The increase in the average molecular size of xyloglucan is likely to be due to progressive XET action on existing polymers, since extractable hemicellulose levels did not change appreciably. Between wilting and full senescence, the xyloglucan of sandersonia flowers began to degrade, but this was quite limited in comparison to the extensive enzyme-mediated degradation of hemicelluloses from carnation that starts even prior to the attainment of full bloom (de Vetten et al., 1991). No evidence was found of ß-1,4-glucanase activity (carboxymethylcellulose hydrolysis) at any point in sandersonia floral development, unlike daylily where cellulase activity was greatest in opening petals (Panavas et al., 1998).
Galactose formed about half the non-cellulosic neutral sugar component of pre-senescent sandersonia flowers. The reduction in the mol% of galactose in wilted flowers paralleled the increase in ß-galactosidase activity. Daylily too, has increased ß-galactosidase activity specific to the onset of senescence-related deterioration (Panavas et al., 1998). Although galactose is not the predominant non-cellulosic neutral sugar in carnation flowers compared to sandersonia, galactose loss is also associated with senescence (de Vetten and Huber, 1990). The loss of galactose from the cell walls of many ripening fruit species has been reported repeatedly and there have been wide-ranging comparative studies of the occurrence of this phenomenon (Gross and Sams, 1984; Redgwell et al., 1997), the significance of which has not yet been satisfactorily resolved. Most galactose from ripening fruit cell walls is lost from polymers soluble only in strong (4 N) alkali or from insoluble, highly branched pectic polysaccharides (Redgwell et al., 1997). This also appears to be the case in sandersonia, but with the major quantities of galactose located in, and lost from, uncharacterized polymers that are insoluble by standard means.
The compressibility test described here quantifies the firmness of the whole flower, objectively confirming the increasing turgidity of mature flowers and onset of wilt. This test does not provide information on the strength of the tepal fabric and an attempt was made to quantify this by measuring the force required to pull apart tissue strips of fixed dimensions, at right angles to the vascular strands. Tissue strips were composed of differing total amounts of water and dry matter at each developmental stage, with the cell wall being an increasing proportion of dry matter in post-mature flowers. In addition, while tissue strip length and width dimensions were fixed, it was found that tissue from post-mature flowers was reduced in thickness, presumably due to tissue shrinkage. It is likely that the similar proportions of water and dry matter in opening and mature flowers were responsible for the similar tensile strength in these strips, even when tissue thickness is taken into account. As flowers opened to full maturity there was a decreasing proportion of carbohydrate associated with long-chain Na2CO3-soluble pectin. The concurrent loss of galactose from this pectin fraction suggests a loss of side-branches during opening, but this change did not impact on the strength of the tepal fabric.
The strength of tepal strips was reduced in wilting and senesced sandersonia flowers compared to those of fully mature flowers. On a per unit area basis (accounting for the differences in tepal thickness), however, the tensile strength of the tissue remained unchanged in wilting flowers and significantly increased in senesced flowers. Significant cell wall polymer changes accompanied these alterations to tepal strength. There was significant loss of galactose per flower during wilting and senescence, most notably from an insoluble cell wall fraction, and the polymers undergoing this change may play a key role in cell wall cohesiveness. There were also increased amounts of cellulose and pectin flower-1 as the sandersonia flowers progressed from wilting to fully senesced, and while there was no evidence of substantial solubilization or degradation of cell wall pectins there was a slight downshift in the average molecular size of xyloglucan in this last stage. Cell walls in senescing flowers may also be stiffened in other ways (e.g. increased phenolic-based linkages, non-vascular lignification) so that the tepal shell that remains was, with thickness accounted for, much stronger even than that of the opening flower. This accumulation of cell wall material in the tepal tissue may be reflective of the development of the sandersonia tepal tissue as a protective cover for the developing ovary, and it would be interesting to identify whether it was a shared feature of the large group of monocotyledonous flowers that senesce without petal abscission (van Doorn, 2001).
Finally, as to whether the changes occurring in the cell walls of senescing sandersonia flowers are distinct events or consequences of the considerable expansion required for full opening, it was found that only one change, that of galactose mobilization, spanned the opening–senescence spectrum, although different polymers of the cell wall were involved at different times. There was continuous loss of galactose from Na2CO3-soluble pectins, but the major loss of galactose occurred as flowers began to wilt, originating from highly insoluble, but as-yet unidentified, polysaccharides. The characterization of these polymers, as well as the wilting-associated role of ß-galactosidase activity in galactose mobilization, will be addressed in future work.
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Acknowledgments |
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Our thanks to Jocelyn Eason, Bub Vizi and Greg Kent for providing the flowers for this work, Leigh de Vré and Jason Johnston for assistance with the tensile strength tests and Wilhelmina Borst and Katie Jarmai-Graf for assistance with histology. This work was supported by the New Zealand Foundation for Research, Science and Technology.
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Notes |
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1 To whom correspondence should be addressed. Fax: +6463517050. E-mail: odonoghuee{at}crop.cri.nz
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References |
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