Journal of Experimental Botany, Vol. 51, No. 344, pp. 507-520,
March 2000
© 2000 Oxford University Press
Changes in chalazal cell walls and in the peroxidase enzymes of the crease region during grain development in barley
Biotechnology Department, Scottish Agricultural College, West Mains Road, Edinburgh, EH9 3JG, Scotland, UK
Received 13 October 1999; Accepted 25 October 1999
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Abstract |
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In an investigation of the role of peroxidase enzymes in the differentiation of the tissues of the crease region of barley, plants of winter barley cv. Halcyon were grown from anthesis onwards in controlled conditions at a constant temperature of 16 °C. Four ears were harvested at 2-d intervals from 6 d after anthesis (daa) until 50 daa. Grains from mid-ear were used for (i) fresh and dry weight determinations, (ii) extraction of crease tissue for the determination of peroxidase activity and for the separation of isozymes of peroxidase by isoelectric focusing (IEF) and (iii) detection of lignin and suberin in the tissues of the crease using autofluorescence and cytochemistry. Peroxidase activity was located histochemically in the crease tissue of cv. Chariot. Scanning electron microscopy studies were carried out on developing grains of cv. Blenheim. Maximum grain water content was achieved at 14 daa. Lignin and suberin were detected in the walls of the chalazal cells from 18 daa onwards. No changes in the staining of chalazal cell walls were detected at the end of grain filling (32 daa), but loss of autofluorescence and staining were observed at 42 daa, just prior to the final, rapid phase of grain dehydration. Peroxidase activity per fresh weight of crease tissue was high at 6 daa and low at 22 daa. It was also low between 32 and 40 daa, but it rose again from 42 daa onwards. IEF demonstrated that both anionic and cationic isozymes of peroxidase were present in crease tissue, the pattern of bands showing some marked changes during the course of grain development.
Key words: Barley, peroxidase, chalazal cell walls, grain development, microscopy.
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Introduction |
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Peroxidases are among the most studied of all plant enzymes yet the physiological functions of peroxidases are only partially understood. Using polyacrylamide gel electrophoresis to explore the possibility that isozyme polymorphism of peroxidases could be used in the identification of barley cultivars, fourteen cationic isozymes of peroxidase in mature barley kernels were detected (LaBerge et al., 1973
). In further work, it was shown that the peroxidase isozyme patterns of extracts of very immature kernels, up to 19 d post-anthesis, were quite different from the isozyme patterns of more mature kernels (LaBerge, 1975). When peroxidase activity was measured in extracts prepared by grinding whole kernels in a 12.5% (w/v) sucrose solution, it was found to rise sharply early in development, then fall and rise again during the later stages of grain maturation (LaBerge and Kruger, 1976). Most of the activity was located in the extracts of the starchy endosperm, the aleurone and the embryo, with very little activity being detected in the husk or in either the transparent or the green layer of the pericarp. More recently, three cationic peroxidases have been isolated in pure form from barley grains (Hejgaard et al., 1991). One of these, BP1, has been characterized and found to be highly tissue-specific, occurring maximally in the endosperm 15 d after flowering (Rasmussen et al., 1991). No physiological function has yet been suggested for this enzyme. It is likely, however, that some of the barley peroxidases are involved in defence mechanisms. The peroxidase activity in the germ aleurone of barley appears to function in the formation of an anti-microbial barrier when the tissue is ruptured during germination (Cochrane, 1994), and increases in two extracellular peroxidases have been observed following inoculation of barley leaves with powdery mildew (Kerby and Somerville, 1992).
Evidence for the role of peroxidase enzymes in cell differentiation has been reviewed (McDougall, 1992a). While there is good evidence that peroxidases are involved in the catabolism of IAA, the part these enzymes play in the complicated process of IAA homeostasis and hence in the control of plant growth has yet to be defined. Soluble and ionically-bound peroxidase activity in whole grains of wheat were related to stages in grain development (Chanda and Singh, 1997) and it was concluded that the close correlation between cessation of grain elongation and increase in wall peroxidase and IAA oxidase activities indicated that these enzymes have an important role in the termination of the elongation phase of wheat grain development. The involvement of peroxidases in the formation of di-ferulate linkages in matrix polysaccharides (van Huystee and Zheng, 1993; Zimmerlin et al., 1994) and in the lignification of cell walls is well established (Imberty et al., 1985; Fry, 1986), and evidence for the involvement of an anionic peroxidase in the production of suberin following wounding in potatoes has been reported (Espelie et al., 1986).
In studies of grain development in wheat plants grown under controlled environment conditions, Radley demonstrated that the increase in the water content of the grain closely parallelled the increase in the volume of the grain, and that the maxima of these parameters were reached while the grain dry weight was still quite low (Radley, 1976). Grain water content remained constant for some time and then began to fall sharply a few days before the dry weight increase ceased. These observations were confirmed later (Schnyder and Baum, 1992) when it was shown that net water deposition into grains ceased when the grain had accumulated one-third of its maximum dry weight, and that dry weight accumulation ceased at approximately the same time as the onset of rapid water loss occurred. Variation among samples was such that Schnyder and Baum could not state unequivocally that the onset of rapid water loss preceded the cessation of dry weight accumulation, but they concluded nevertheless that their data supported the view that a blockage of the intake of water into the endosperm may be causally related to the cessation of dry matter accumulation in the grain. Similar patterns of grain water content and grain dry matter accumulation in developing grains of barley have been recorded (Cochrane et al., 1996).
Assimilate is imported into the developing endosperm of wheat and barley grains first via a phloem pathway and then via a post-phloem pathway (Thorne, 1985; Wang and Fisher, 1994). The phloem pathway extends from the rachis, through the base of the grain, and along the vascular bundle which is located in the chlorenchyma of the pericarp in the crease, also known as the groove or furrow, on the ventral side of the grain (Fig. 1). In the post-phloem pathway, assimilate passes through vascular parenchyma and several other layers of parenchymatous cells, the chalazal cells, the nucellar projection, the endosperm cavity and finally the crease aleurone before it enters the cells of the starchy endosperm (Hoshikawa, 1964). In a study of the ontogeny of the chalazal cells of wheat grains using light microscopy and transmission electron microscopy, it was observed that early in grain development the chalazal cells were thin-walled, but between 12 d and 18 d after anthesis, the cell walls became lignified and adcrusted with sudanophilic material except in the region of pits in which plasmodesmata were located (Zee and O’Brien, 1970). The sudanophilic wall thickening was thought to be suberin. The vacuoles in these cells contained sudanophilic deposits. Later in grain development, the adcrustation was massive, i.e. >1 µm thick. It stained pale blue-green in toluidine blue O, but was almost totally electron-lucent in grains which had undergone glutaraldehyde-osmium fixation. The cytoplasm of the chalazal cells at this stage of grain development appeared to have undergone degeneration. A comparable study of the crease region of barley during grain development (Cochrane, 1983) showed that in barley as in wheat, during the initial stage of grain development, chalazal cell walls were unthickened. These cell walls subsequently became lignified and suberin was deposited between the lignified wall and the cytoplasm, forming a barrier between the apoplast and the symplast. The sudanophilic material in the vacuoles of the chalazal cells was identified as tannin and it was suggested that the degeneration of the cytoplasm in the final stages of grain development was due to the release of this tannin from the vacuoles.
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Movement of assimilate into the endosperm is thought to follow both apoplastic and symplastic pathways until the chalazal cell walls become thickened with lignin and suberin. Except at the crease, the endosperm is encased in the testa with its inner and outer cuticular layers and so, from this stage onwards, only the symplastic pathway through the chalazal cells is available. Later in grain development this symplastic pathway appears to be much reduced by the massive wall thickening and it is finally obliterated by the release of tannins from the vacuoles. The relationship between the timing of these changes in the cytochemistry of the chalazal cells and the timing of the changes in grain water content and grain dry weight which occur during grain development has not been established.
The deposition of lignin and suberin in the chalazal cell walls suggests that peroxidase enzymes may have a significant role to play in the processes of cell differentiation which take place in the crease tissues during grain development. The aim of this work was therefore to monitor changes in total peroxidase activity and in the pattern of isozymes of peroxidase extracted from crease tissues during grain development and to correlate these with changes in the dry weight and water content of the grain and in the morphology and histochemistry of the cells of the chalazal region of the crease. Cytochemistry and autofluorescence observations were used to detect the presence of lignin and suberin in unfixed tissues. The bright ice-blue autofluorescence characteristic of barley endosperm cell walls irradiated with ultraviolet light and viewed using Leitz filter system A is due to the presence of ferulic acid (Fincher, 1976). When examined using the same light source, lignin, suberin and cutin all exhibit bright yellow autofluorescence when viewed with Leitz filter system G, but when viewed using Leitz filter system A with an additional K460 barrier filter, the fluorescence from suberin (as in cork) and from lignin (as in a wood shaving) is blue/green. When lipid-containing materials such as suberin and cutin are stained with Fluorol Yellow they emit bright yellow fluorescence when exposed to UV light (Brundrett et al., 1991). It is possible to distinguish between the colour of the fluorescence emitted by Fluorol Yellow-stained suberin and that emitted by Fluorol Yellow-stained cutin in filter system G and in filter system A. This stain thus provides a sensitive means of detecting the presence of suberin in sections of fresh tissue.
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Materials and methods |
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Chemicals
Stains used in microscopy, horseradish peroxidase, and substrates used to demonstrate and assay peroxidase activity, were all supplied by Sigma, UK. All other chemicals used were of analytical grade.
Plant material
Winter barley, cv. Halcyon, was grown in peat-based compost, six plants per pot (20 cm diameter) in an unheated glasshouse from 5 December 1995 to 11 March 1996, and then transferred to a glasshouse in which day/night temperatures were maintained at approximately 20/15 °C for a 16 h day. Natural daylight was supplemented with mercury vapour lamps. Two to three days prior to anthesis (29 May 1996), plants were transferred to a growth room in which in which a constant temperature of 16 °C was maintained. Day length was 16 h. Lighting was supplied by mercury vapour lamps and light intensity at ear level was approximately 140 µmol m-1 s-1.
For preliminary studies to evaluate the methodology, spring barley cvs Chariot and Blenheim were grown (April to July, 1994) in natural daylight in a glasshouse in which the day/night temperature was maintained at approximately 25/15 °C.
For the survey of grain development using scanning electron microscopy, cv. Blenheim was grown in growth rooms in a 16 h day at 13 °C throughout grain development or at 13 °C from anthesis to 14 d after anthesis (daa), 20 °C from 14 daa to 28 daa, and 13 °C from 28 daa to harvest-ripeness. The ages of these grains were expressed in °C days using 0 °C as the baseline temperature. Enzyme histochemistry was carried out on grains of cv. Chariot grown in a growth room in a 16 h day at a constant temperature of 18 °C.
Sampling procedure
Ears of cv. Halcyon were tagged at anthesis. Four ears were harvested at 5 daa, at 6 daa and thereafter at 2 d intervals until 50 daa. For each sample, no more than 2 ears were harvested from any one pot. Seven grains were removed from the centre of each ear. Five of these were used to prepare crude extracts for enzyme assays and iso-electric focusing (IEF) of peroxidase enzymes and two were used for microscopy. Thus four extracts were made at each harvest and eight grains were available for microscopy. A further five grains were removed from positions on each ear adjoining those from which the central seven grains had been removed. These five grains were used for fresh and dry weight determinations. All procedures were carried out on fresh material harvested 5 daa to 32 daa. Grains harvested for enzyme analyses and microscopy from ears 34 daa to 50 daa were frozen in liquid nitrogen and stored at -20 °C, preliminary studies having indicated that extracts made from frozen tissue had the same enzyme activity and IEF profiles as extracts made from fresh tissue obtained from the same ear.
Fresh and dry weight determinations
Five grains were weighed immediately they were removed from each of the four ears harvested on each sampling date and again after they had dried to constant weight in an oven at 70 °C.
Histochemistry
To demonstrate the location of peroxidase activity in the crease tissue, hand-cut sections of the crease region of fresh and frozen grains were transferred directly from the razor blade to vials containing one of the following solutions: (i) 0.5 M (sodium) phosphate buffer pH 6.5, (ii) 0.068 M catechol in 0.5 M (sodium) phosphate buffer pH 6.5 and (iii) 0.068 M catechol in 0.5 M (sodium) phosphate buffer pH 6.5 containing 0.003% (v/v) hydrogen peroxide. Sections were soaked in these solutions at room temperature for 10 min and then rinsed in water before being mounted on microscope slides using a glycerol/gelatin mountant.
To identify changes in cell wall composition which take place in tissues of the crease during grain development, hand-cut sections of the ventral side of fresh (5–32 daa) or frozen (34–50 daa) grains were collected in distilled water and then (i) mounted on microscope slides in 75% (v/v) glycerol to examine autofluorescence, (ii) soaked in a saturated solution of phloroglucinol in 20% (w/v) HCl to locate lignin, or (iii) soaked in a solution of Fluorol Yellow to locate suberin (Brundrett et al., 1991). Sections were examined using a Leitz Ortholux II epi-illumination fluorescence microscope fitted with a mercury vapour UV light source and filter blocks G (BP 350–460 nm, RKP 510 nm, LP 520 nm) and A (BP 340–380 nm, RKP 400 nm, LP 430 nm). An additional K460 barrier filter was used with filter block A. Images viewed using bright field microscopy were photographed using an Ektachrome 160T film. Fluorescent images were photographed using a Kodak Gold 200 ASA film.
Enzyme extraction
Using fine forceps, the crease tissue was removed from each of five caryopses. To obtain a value for the fresh weight of the crease tissue, the weight of the five caryopses was determined before and after the removal of the crease tissues. The crease tissue was homogenized in a glass hand-held homogenizer on ice in 1 ml 40 mM (sodium) phosphate buffer pH 7.0 containing 1% (v/v) Triton X-100 and 0.5 M KCl. When homogenization was complete, 0.5 ml 25 mM sucrose was added and the homogenate was centrifuged at 11 000 g for 10 min at 4 °C. More than 90% of the peroxidase activity in the homogenate was present in the supernatant. During method development it was found that the optimum pH for enzyme extraction was 7.0. To quantify possible losses of peroxidase activity during the preparation of the crude extract, horseradish peroxidase was added to the extraction medium prior to homogenization of crease tissue. Approximately 70% of the horseradish peroxidase activity was recovered in the crude extract.
Enzyme assay
Peroxidase activity was measured using a reaction mixture consisting of 2 ml of a 0.02% (w/v) solution of ferulic acid in 0.1 M citrate/(sodium) phosphate buffer pH 6.5 and 10 µl enzyme extract. The reaction was started by the addition of 10 µl 0.3% (v/v) aqueous solution of hydrogen peroxide and was allowed to run for 10 min at 20 °C. Absorbance was monitored at a wavelength of 345 nm using the kinetics programme on a Beckman DU 65 Spectrophotometer. A blank consisting of 2 ml of the buffered ferulic acid solution plus 10 µl 0.3% (v/v) aqueous solution of hydrogen peroxide was run. Blanks containing buffer and enzyme were found to have negligible absorbance at 345 nm and were therefore not used in the assay procedure. No change in absorbance took place when buffered ferulic acid was incubated for 10 min at 20 °C in the presence of 10 µl enzyme extract. The rate of the reaction was calculated over the period 5–10 min after the addition of the hydrogen peroxide solution. During method development it was established that the absorbance maximum of the product formed during this period was at a wavelength of 345 nm, that the optimum pH for the reaction was 6.5, and that the rate of reaction was proportional to the amount of extract added to the reaction mixture.
Isoelectric focusing
The crude extract was desalted by centrifugation at 11 000 g for 20 min at 4 °C using a Microcon 30 filter. The filtrate was discarded and the recovered retentate was divided into two equal aliquots, one of which (extract A) was stored at 4 °C while the other was centrifuged at 3000 g for 30 min at 4 °C through a Micronon 100 filter. The filtrate containing molecules of Mr between 30 and 100 kDa was retained (extract B). Separation of the proteins in these extracts was performed on Pharmacia isoelectric focusing gels, pH range 3.5–9.5, on a Multiphor II Horizontal Gel System. Gels were run at 18 °C and focused for approximately 1.5 h. The volume of extract applied to the gel was between 5 µl and 35 µl and was directly proportional to the fresh weight of the crease tissue extracted. Two different substrates were used to locate peroxidase isozymes: (i) 1 mM 3-amino-9-ethyl carbazole in 0.1 M acetate buffer pH 5 and (ii) 1 mM o-dianisidine in 0.1 M acetate buffer pH 5. Gels were soaked in these solutions for 20 min and then a 0.3% (v/v) aqueous solution of hydrogen peroxide was added to give a final concentration of 1 mM hydrogen peroxide. pI standards run at the end of each gel were stained using Comassie Blue.
Scanning electron microscopy
Grains were frozen in liquid nitrogen, cut transversely at mid-grain using a cryomicrotome, freeze-dried, coated with gold and examined in a Cambridge S250 scanning electron microscope.
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Results |
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Grain development (winter barley cv. Halcyon)
Grain dry weight increased more or less linearly until 32 daa (Fig. 2
). It was not possible to determine the exact timing of the end of grain filling because of the high ear to ear variation in grain dry weight. This ear to ear variation was probably due largely to sporadic male sterility which resulted in a small but apparently significant reduction in the number of grains per ear in some plants. Grain water content increased rapidly in a linear fashion between 6 daa and 14 daa, after which age no further increase in the water content of the grain took place (Fig. 3). Between 32 daa and 42 daa the water content of the grain decreased quite rapidly and from 42 daa to 46 daa, when harvest-ripeness was achieved, the water content of the grain decreased very rapidly. The anomolous values recorded for the grains harvested at 48 daa are due to the presence of an ear which, though being recorded as 48 daa, was morphologically approximately 40 daa. In this case it appears that male sterility caused self-fertilization to fail but some days later cross-fertilization took place. In the grain percentage moisture content curve (Fig. 4), in which ear to ear variation is very low except for ears harvested at 38 daa and 48 daa, discontinuities can be seen at 12 to 14 daa, 20 to 22 daa, 34 to 36 daa, and 44 to 46 daa.
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Scanning electron microscopy (spring barley cv. Blenheim)
In grains aged 221 °C d after anthesis (°C daa), the chalazal cells had thin walls and highly hydrated contents (Fig. 5A), whereas, in grains aged 338 °C daa, the walls of the chalazal cells were slightly thickened (Fig. 5B). By 606 °C daa the walls of the chalazal cells were considerably thickened and the cell contents included what appear to be starch granules (Fig. 5C). The chalazal cells in grains aged 710 °C daa were completely filled with amorphous material and had cell walls which were massively and irregularly thickened (Fig. 5D).
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Histochemistry
Location of peroxidase activity (spring barley cv. Chariot).
Sections incubated in buffer (pH 6.5) alone (Fig. 6A), or in buffer containing hydrogen peroxide, developed no colour. Sections incubated in buffer (pH 6.5) containing catechol developed a pale brown colour in the chalazal region, while those incubated in buffer containing catechol and hydrogen peroxide, developed a dark chestnut-brown colour in the chalazal region (Fig. 6B), but no colour in any other parts of the section. When sections were incubated in the presence of the same substrates in an acetate buffer at pH 4.5, the endosperm became pink/purple.
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Reaction to phloroglucinol/HCl (winter barley cv. Halcyon).
When mid-grain sections of caryopses aged 10–50 daa were mounted in phloroglucinol/HCl, the cherry pink colour indicative of the presence of lignin was not detected in caryopses younger than 18 daa (Fig. 6C). In 18 daa caryopses, staining was detected only at the interface between the chalaza and the nucellar projection (Fig. 6D) but in 20 daa caryopses all the walls of the chalazal region and the walls of the outermost 3–4 cell layers of the nucellar projection stained pink. This pattern of staining was seen in sections of all caryopses harvested 20 daa to 40 daa (Fig. 6E). From this stage of development onwards, the staining became progressively darker but remained confined to the chalazal cells and the outer cells of the nucellar projection. In sections of caryopses harvested 46 daa, the chalazal and nucellar projection cells appeared blackish brown and were somewhat crushed in contrast to the cells walls of the epidermis which were bright reddish-pink and were not deformed (Fig. 6F).
Autofluorescence (filter block G) (winter barley cv. Halcyon).
In transverse sections of young (10 daa) green caryopses cut at mid-grain, the chlorenchyma in the crease region fluoresced bright red and faint traces of yellow fluorescence could be detected in the cuticular layers on either side of the testa. No fluorescence was detected in the walls of the chalazal cells in caryopses younger than 18 daa (Fig. 7A). In sections of caryopses aged 18 daa, greenish yellow fluorescence was observed in the cell walls of the chalaza and the outer layers of the nucellar projection (Fig. 7B). Yellow autofluorescence was observed in the walls of the chalazal cells in sections of all caryopses harvested from 20 daa to 38 daa (Fig. 7C), the fluorescence gradually extending into the radially elongated cells of the nucellar projection, changing from greenish yellow to bright yellow and becoming more intense as the age at which the grains were harvested increased. In sections of caryopses aged 40 daa, no red fluorescence was detected in the parenchymatous cells of the crease, and all the cell walls in the pericarp fluoresced yellow. In sections of caryopses aged 42–50 daa, all the cell walls in the pericarp and nucellar projection fluoresced bright yellow, but the walls of the chalazal cells fluoresced markedly less than the cell walls of the other tissues of the crease.
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Autofluorescence (filter block A+barrier filter K460) (winter barley cv. Halcyon).
Sections of caryopses aged 10–16 d exhibited ice blue autofluorescence in all the cell walls of the crease region except those of the xylem vessels which fluoresced creamy white. This blue fluorescence was pale in the chalazal cells of caryopses aged 20 daa, but was bright and strong in the outer cells of the nucellar projection in sections of caryopses of this age. In sections of caryopses aged 22 daa to 38 daa, the autofluorescence in the cell walls of the chalazal region and the outer layers of the nucellar projection was dull green/yellow whereas that in the the radially elongated cells of the nucellar projection was bright ice blue (Fig. 7D). In sections of caryopses aged 42–50 daa, little if any autofluorescence was detected in the walls of the chalazal cells or in the walls of the cells of the outer layers of the nucellar projection. The fluorescence of the cell walls of the inner nucellar projection, the sheaf and the aleurone was bright ice blue, and that of the walls of the pericarp tissues was a pale yellowish or pinkish buff colour.
Fluorescence following staining in Fluorol Yellow (viewed using filter block G) (winter barley cv. Halcyon).
Faint yellow fluorescence was detected in the chalazal region of sections of caryopses aged 14 daa and 16 daa (Fig. 7E), but the youngest caryopses in which the bright yellow fluorescence characteristic of Fluorol Yellow could be detected with certainty in the walls of the chalazal cells were those harvested 24 daa. The intensity of the bright yellow fluorescence was very strong in the cell walls of the chalaza and in the cuticular layers on either side of the testa in caryopses aged 26–38 daa (Fig. 7F), and progressively less strong in these structures in caryopses aged from 40–50 daa (Fig. 7G).
Fluorescence following staining in Fluorol Yellow (viewed using filter block A+barrier filter K460) (winter barley cv. Halcyon).
Some yellow fluorescence was detected in the walls of the chalazal cells of caryopses as young as 18 daa and the yellow fluorescence characteristic of Fluorol Yellow staining of lipid material was bright and strong by the time caryopses reached the age of 20 daa. The yellow fluorescence was confined to the chalazal cells and the outer 2–3 cells layers of the nucellar projection (Fig. 7H). The fluorescence of the cuticular layers on either side of the testa differed from each other and from that in the walls of the chalazal cells. This pattern of fluorescence was observed in caryopses aged up to 40 daa, but in caryopses aged from 42–50 daa, the fluorescence in the walls of the chalazal cells was weak, the chalazal cells were deformed and they had contents which quenched fluorescence.
Peroxidase activity (winter barley cv. Halcyon)
Peroxidase activity of extracts of crease tissue plotted against the age (daa) of the grains from which the creases were obtained, showed two peaks, one at 10 daa and the other at 42–44 daa when the values were expressed on a per crease basis (Fig. 8). Expressed on both a fresh weight and a dry weight basis, the values were high at 5 daa and were progressively lower until the minimum was reached at 22 daa. Values higher than this minimum were observed between 24 and 30 daa, followed by a trough from 32–40 daa and a peak from 42–50 daa, excluding the value for 48 daa which appears from grain development curves and histochemistry to have been younger than recorded (i.e. approximately 40 daa), possibly due to male sterility at anthesis.
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Isoelectric focusing (winter barley cv. Halcyon; spring barleys cvs Blenheim and Chariot)
Extracts of crease tissue were stored at -20 °C until extracts from grains of all ages from 8 daa to 48 daa had been accumulated. The amount of extract loaded in each lane was proportional to the fresh weight of the crease tissue from which the extract was made. The extracts were run on a single gel alongside a solution containing a set of proteins of known pI. The gels were developed using either 3-amino-9-ethyl carbazole (pink bands) or o-dianisidine (orange bands) (Fig. 9), and the pI of each band was calculated (Table 1). Fewer bands were detected in the gels in which extracts which contained material <100 kDa >30 kDa had been run than in gels in which extracts >30 kDa had been run, but the differences due to the exclusion of material >100 kDa were more often in the intensity of the staining of bands rather than in the number of bands. Differences between gels developed using 3-amino-9-ethyl carbazole and those of the same series of extracts developed using o-dianisidine were slight and, where detected, were mainly in the intensity of staining or in the definition of a particular band. In studies on spring barley (data not shown) using similar IEF gels, the pattern of peroxidase isozymes of extracted crease tissue was seen to change during grain development and the isozymes of peroxidase extracted from cv. Blenheim differed slightly from those extracted from cv. Chariot.
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Discussion |
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The pattern of grain development observed in winter barley cv. Halcyon grown in a controlled environment room at a constant temperature of 16 °C (Fig. 2) was very similar to that observed in wheat cvs Kleiber and Capelle (Radley, 1976
), and cv. Achill (Schnyder and Baum, 1992). Maximum water content was reached at approximately 14 daa, i.e. when the grain had accumulated between one-third and one-half of its final dry matter content. Grain water content remained constant until 32 daa when it decreased rapidly for 10 d and then very rapidly until harvest-ripeness (12% moisture content). The beginning of grain dehydration appeared to coincide with the completion of grain filling but, because of ear-to-ear variation and the fact that anthesis dates are identified at best to ±1 d, it was not possible to establish whether dry matter accumulation ceased before or after the start of grain dehydration. Schnyder and Baum reported that cessation of dry matter accumulation and the onset of rapid water loss occurred ‘within the same sampling interval’ (Schnyder and Baum, 1992), but Radley considered that the sharp decrease in water content of wheat grains preceded the cessation of dry weight increase by a few days (Radley, 1976). The percentage moisture content values showed much less variability than either the water content or the dry matter values. This evidence supports the conclusion reached by Schnyder and Baum that percentage moisture is a valuable indicator of stages of grain development. The inflections in the percentage moisture curve occur at a number of daa which is similar to, or 2 d later than, the points of discontinuity observed in the grain water content curve, with the exception of that identified at 20/22 daa.
On the basis of the observations of the autofluorescence of the chalazal cell walls and their staining in phloroglucinol/HCl and in Fluorol Yellow, it was established that both lignin and suberin were present in the chalazal cell walls at 18 daa (320 °C daa). Prior to this stage of development, chalazal cell walls appeared to be unthickened (Fig. 5A). It was not possible to determine whether the deposition of lignin preceded the deposition of suberin. The composition of the thickened cell walls appeared to remain the same until 38 daa. In the chalazal cell walls in sections of grains older than this, both autofluorescence and the fluorescence due to Fluorol Yellow staining was much less than that observed in the chalazal cell walls of caryopses harvested between 18 and 38 daa. This might indicate a change in the chemical composition of the chalazal cell walls. Scanning electron micrographs (Fig. 5C, D) and transmission electron micrographs of chalazal cells in barley caryopses of other cultivars at a stage of development equivalent to that of the >38 daa Halcyon grains in this study (Cochrane, 1983) show massive wall thickening. From the observations presented here it may be concluded that this wall thickening is not rich in phenolic material which autofluoresces in the same way as lignin or suberin, nor does it have a high lipid content. The lack of fluorescence in the chalazal cells during the latter stages of grain maturation may be due to the deposition on the walls, or penetration into the walls, of material which quenches fluorescence. This may be the result of the massive wall thickening which takes place in these cells and/or of the liberation of tannins from the vacuoles (Cochrane, 1983). Alternatively, it may be due to degradation of the lignin and suberin, a process which could be responsible for the deformation of the chalazal cells prior to the achievement of harvest-ripeness.
The observation that in caryopses aged 46 daa (736 °C daa) the contents of the chalazal cells appeared to quench fluorescence may be correlated with the uniform and slightly spongy appearance of the contents of the chalazal cells in scanning electron micrographs of transverse sections of frozen/freeze-dried caryopses of cv. Blenheim aged 710 °C (Fig. 5D), and in transmission electron micrographs of chalazal cells in transverse sections of grains at a similar stage in development (Cochrane, 1983). This material is thought to be formed following the release of tannin from vacuoles into the cytoplasm, and to be responsible for the cessation of symplastic transport through the chalazal cells (Cochrane, 1983; Felker et al., 1984; Lingle and Chevalier, 1985).
The earliest age at which lignin and suberin were detected in the chalazal cells was approximately 4 d after the grain had reached maximum water content. The possibility exists that the blocking of the apoplastic pathway into the endosperm is responsible for the stabilization of the water content of the grain and that the discrepancy observed in this investigation between the timing of the deposition of lignin and suberin and the achievement of maximum water content was due to the insensitivity of the methods used to detect these materials. However, at this stage of grain development, the pericarp is composed of many layers of parenchymatous cells which make a considerable contribution to the water content of the grain. It is more likely therefore that the cessation of water accumulation by the grain is due to the caryopsis reaching its maximum volume (Radley, 1976), rather than to the occurrence of changes in the pathway of assimilates into the endosperm. It is possible, however, that the discontinuity in the % moisture curve detected at 20/22 daa may be a reflection of the changes observed in the chalazal cell walls at 18/20 daa.
No changes in the composition of the chalazal cell walls were detected between 20 daa (320 °C daa) and 40 daa (640 °C daa) yet during this period, at approximately 32 daa (512 °C daa) dry matter accumulation ceased and the water content of the grain began to fall. Scanning electron micrographs show that in grains harvested 606 °C daa (Fig. 5C), the chalazal cell walls were massively thickened. The exact timing of the laying down of this additional layer of wall thickening in relation to the timing of the cessation of grain filling was not determined in this investigation. Similarly, no progress was made in determining the chemical composition of the material of which the massive wall thickening was composed. The fact that no increase in autofluorescence or in Fluorol Yellow staining was observed in the chalazal cell walls during the stage of development between 30 and 40 daa is an indication that the phenolic and lipid content of the new material differed from that of lignin and suberin. It was reported that the massive wall thickening found in chalazal cell walls of wheat caryopses at a comparable stage of development stained ‘pale blue-green’ in toluidine blue O, indicating a low phenolic content, whereas the chalazal cell walls in caryopses examined at an earlier stage in grain development stained green (Zee and O’Brien, 1970). The timing of the onset of the final phase of grain dehydration as seen on the % moisture content graph at 44–46 daa (Fig. 4) coincides with the timing of the loss of autofluorescence and Fluorol Yellow staining in the chalazal cell walls and the appearance of fluorescence-quencing material in the lumen of the chalazal cells. The release of tannins from the vacuoles in the chalazal cells thus occurred 10–12 d after the cessation of dry matter accumulation. This observation confirms the conclusion reached previously (Lingle and Chevalier, 1985) that the collapse of the chalazal cells is not responsible for the cessation of grain filling. It was shown (Cochrane, 1985) that some uptake of 14C-labelled sucrose into the endosperm of barley grains in ears grown in liquid culture continued for several days after dry matter accumulation in the grain had ceased. Thus, it is possible that a partial disruption of the symplastic pathway by the deposition of wall material in the chalazal cells causes a reduction in assimilate uptake into the endosperm and initiates the dehydration phase of grain development.
From the cytochemical and biochemical evidence presented it is clear that there is a significant amount of peroxidase activity in the crease region of developing barley grains and that this activity is mainly concentrated in the cells of the chalazal region and the outermost cells of the nucellar projection. Some brown coloration was observed in chalazal tissue in sections supplied with a buffered solution of catechol in the absence of hydrogen peroxide, indicating the possible presence in these tissues of either an o-phenol oxidase or of peroxidase(s) which can oxidize catechol in the absence of exogenously supplied hydrogen peroxide. The use of ferulic acid as the substrate in the peroxidase assay precluded interference from o-diphenol oxidases. During method development it was found that little or no activity could be extracted from crease tissue in the absence of 0.5 M KCl and so it was concluded that the peroxidases were ionically bound to the chalazal cell walls. The possibility that covalently-bound peroxidases (McDougall, 1992b) were also present in the chalazal cell walls was not explored. The large variation in the values for peroxidase activity expressed on a per crease basis may have been due in part to the difficulty experienced in dissecting complete and therefore uniform lengths of crease tissue, and in part to ear-to-ear variation. The pattern observed when the results were expressed on a fresh weight or dry weight basis showed that the initially high activity fell to a minimum at 22 daa, i.e. the age after which no further increase in staining for lignin or suberin in the chalazal cells was observed. Relatively high values were recorded between 24 and 30 daa, the period immediately prior to the end of grain filling, and again from 42–50 daa, the period immediately prior to and during the final stage of grain dehydration. There thus appears to be some relationship between peroxidase activity measured using ferulic acid as substrate and (i) the deposition of lignin and suberin and (ii) the deposition of massive wall thickening thought to take place at the end of grain filling (Zee and O’Brien, 1970; Cochrane, 1983), and (iii) the changes in cell contents and cell wall composition which occur in the final phase of grain dehydration.
Isozymes of peroxidase exhibit substrate specificity (Spencer et al., 1993). It is therefore not possible to make direct comparisons between the total activities observed using ferulic acid as substrate and the patterns of isozymes obtained in IEF because, in order to locate peroxidase activity in the IEF gels, substrates which formed brightly-coloured, insoluble products had to be used. The cationic isozymes exhibited isoelectric point values similar to those ehibited by peroxidases from barley leaf tissue (Saeki et al., 1986), but they are not necessarily the same isozymes because the method used to extract the peroxidase isozymes from leaf tissue would have released little if any of the peroxidase activity ionically-bound to cell walls. Five of the peroxidase components of the IEF gels of leaf extract were found to have the same amino acid composition and so were not considered to be ‘real isozymes’, their differences in pI values being ascribed to differences in glycosyl content (Saeki et al., 1986). Likewise, the peroxidase components of the IEF gels of the crease region of developing barley grains may not all be ‘real isozymes’. Three cationic peroxidases (BP1, BP2 and BP3) have been isolated in pure form from barley grains (Hejgaard et al., 1991). These appear to be more or less tissue specific, BP1 (pI ~8.5) and BP2 (pI>10) being located in the endosperm, and BP3 (pI>10) being detected in embryonic tissue culture medium (Hejgaard et al., 1991). BP2 has also been located in the scutellum of the embryo (Theilade et al., 1993). No peroxidase activity was detected in the starchy endosperm or the aleurone when hand-cut sections of fresh caryopses were incubated in catechol and hydrogen peroxide at pH 6.5 (Fig. 6B) but a marked response occurred in the endosperm when sections were incubated at pH 4.5. This reaction was probably due to the presence of BP1, the pH optimum of which is 3.8 (Theilade et al., 1993). The tissue specificity of the three BP isozymes is such that they would be very unlikely components of extracts of crease tissue. If, however, some were present through contamination of the crease tissue with endosperm tissue, their pH specificity and pI values are such that they would not have contributed to the total peroxidase activity and only BP1 would have been detected in the IEF gels.
The most obvious role for peroxidases in crease tissues of developing barley grains is in the formation of lignin and suberin in the walls of the chalazal cells. Both anionic peroxidases and cationic peroxidases were found to be present at the appropriate stage of grain development. The anionic peroxidase associated with suberization during wound-healing in potatoes has an isoelectric point of 3.15 (Espelie and Kolattukudy, 1985). Both anionic and cationic wall-associated peroxidases have been shown to be involved in lignin deposition (McDougall, 1992b). Isozymes with pI 8.25 and 8.55 increased in activity at, or just after, the end of the grain-filling period, and an isozyme with pI 8.35 appeared at 40 daa, i.e. immediately prior to the final, rapid phase of grain dehydration. These isozymes may be involved in the changes which take place in the chalazal cell walls and may be functionally related to the process of grain dehydration. Additionally, or alternatively, they may be involved in the deposition in the cell walls of the crease parenchyma of the material which, though lignin-like in its autofluorescence, fails to stain in either phloroglucinol/HCl or in Fluorol Yellow.
Following inoculation of barley leaves with the powdery mildew pathogen (Erysiphe graminis DC.:Fr. f. sp. hordei Em. Marchal), increases in two peroxidase isozymes (pI 5.2 and 8.5) were observed (Kerby and Somerville, 1992). One of these (pI 8.5) has since been shown to be located in cell walls in which phenolic material is deposited in response to fungal infection (Scottcraig et al., 1995). It is very likely that some of the peroxidase isozymes extracted from crease tissues are involved in non-specific mechanisms of defence against pathogen invasion. Other possible roles for peroxidases present in the crease tissues of developing barley grains are the oxidation of IAA (Pfanz, 1993), and the formation in cell walls of the hydrogen peroxide used in di-ferulate cross-linking and in lignification (McDougall, 1992a).
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Acknowledgments |
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We thank John Findlay of the University of Edinburgh Science Faculty Electron Microscopy Facility for preparing the grains for scanning electron microscopy, the Nuffield Foundation for the award of an undergraduate research bursary to EG, and the Scottish Office Agriculture, Food and Environment Department for financial support.
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1 Present address and to whom correspondence should be sent: 19 Blackford Hill Rise, Edinburgh, EH9 3HB, Scotland, UK.
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