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First published online May 7, 2004; 10.1104/pp.104.040220 Plant Physiology 135:287-299 (2004) © 2004 American Society of Plant Biologists The Control of Storage Xyloglucan Mobilization in Cotyledons of Hymenaea courbaril1Seção de Fisiologia e Bioquímica de Plantas, Instituto de Botânica, CEP 01061–970, Sao Paulo, SP, Brazil (H.P.S., M.S.B.); and Departamento de Alimentos e Nutrição Experimental, Faculdade de Ciências Farmacêuticas (E.P.), and Departamento de Botânica, Instituto de Biociências (H.M.), Universidade de Sao Paulo, CEP 05422–970, Sao Paulo, SP, Brazil
Hymenaea courbaril is a leguminous tree species from the neotropical rain forests. Its cotyledons are largely enriched with a storage cell wall polysaccharide (xyloglucan). Studies of cell wall storage polymers have been focused mostly on the mechanisms of their disassembly, whereas the control of their mobilization and the relationship between their metabolism and seedling development is not well understood. Here, we show that xyloglucan mobilization is strictly controlled by the development of first leaves of the seedling, with the start of its degradation occurring after the beginning of eophyll (first leaves) expansion. During the period of storage mobilization, an increase in the levels of xyloglucan hydrolases, starch, and free sugars were observed in the cotyledons. Xyloglucan mobilization was inhibited by shoot excision, darkness, and by treatment with the auxin-transport inhibitor N-1-naphthylphthalamic acid. Analyses of endogenous indole-3-acetic acid in the cotyledons revealed that its increase in concentration is followed by the rise in xyloglucan hydrolase activities, indicating that auxin is directly related to xyloglucan mobilization. Cotyledons detached during xyloglucan mobilization and treated with 2,4-dichlorophenoxyacetic acid showed a similar mobilization rate as in attached cotyledons. This hormonal control is probably essential for the ecophysiological performance of this species in their natural environment since it is the main factor responsible for promoting synchronism between shoot growth and reserve degradation. This is likely to increase the efficiency of carbon reserves utilization by the growing seedling in the understorey light conditions of the rain forest.
The presence and mobilization of xyloglucans following seed germination were first reported in the 19th century for seeds of Impatiens balsamina, nasturtium (Tropaeolum majus), and Cyclamen europaeum (Heinricher, 1888  ; Reiss, 1889). Later in the 1960s, the botanical distribution of xyloglucans in seeds was reviewed by Kooiman (1960), who used the ability of xyloglucan to stain with iodine, giving a distinctive blue color, as a form of detection.
Seed xyloglucans have a main
The reserve function of xyloglucan in cotyledons has been demonstrated for seeds of nasturtium (Edwards et al., 1985
Reid and co-workers isolated the four main enzymes responsible for xyloglucan degradation in nasturtium. They are (1) xyloglucan-specific endo-
On the basis of these results, together with other studies on the mode of action of the XTH (Edwards et al., 1986
Reis et al. (1987)
In C. langsdorffii, the mobilization of the cotyledon cell wall storage has been observed cytochemically, physiologically, and biochemically by Buckeridge et al. (1992)
In cotyledons of H. courbaril, Tiné et al. (2000)
Although some work has been performed on the mechanism of seed storage xyloglucan mobilization in seeds, very little is known about the control mechanisms involved in the process. The only report was provided by Hensel et al. (1991) In this study, we investigated some aspects of the effect of auxin on the mobilization of xyloglucan in cotyledons of growing seedlings of H. courbaril. Our results indicate that auxin is involved in xyloglucan metabolism, being produced in the shoot of the growing seedling so that the pace of growth possibly controls storage mobilization in the cotyledons.
Storage Mobilization in the Cotyledons and Early Seedling Growth
The decay curve of dry mass of the cotyledons of H. courbaril decreased in two phases (Experiment I; see Fig. 7 for details of the experiments I and II). The first step was from 0 to 26 d, when protein bodies and raffinose series oligosaccharides are mobilized (Tiné et al., 2000
The addition of 10–6 M 2,4-D after 41 d significantly increased the rate and extent of xyloglucan degradation, similar to the extent observed in attached cotyledons (Fig. 2A). Exogenous 2,4-D affected xyloglucan mobilization only at the concentration of 10–6 M and in the same period as the attached cotyledons. Incubation of detached cotyledons in 2,4-D before the period of highest rate of xyloglucan mobilization (after 34 d) or in all periods at 10–4 and 10–5 M failed to evoke any detectable xyloglucan breakdown (data not shown). The drastic dry mass loss and xyloglucan mobilization in the cotyledons were directly related to the increase in dry mass of the seedling, mainly with the shoot and expansion of eophylls (Figs. 1 and 2A). This relationship was clearer from the observation of seedlings of H. courbaril growing after excision of the shoot or in the darkness. The latter prevented the increase in total leaf area (Fig. 1D) and dry masses of whole seedlings (Fig. 1B) and their shoot (Fig. 1C). Furthermore, darkness promoted a reduction of xyloglucan mobilization (Figs. 1A and 2C). Another group of seedlings, subjected to excision of the shoot at 34 d (when xyloglucan degradation starts), showed a strong delay in xyloglucan mobilization (Fig. 2C). This inhibitory effect was followed by a significant reduction of seedling growth (Fig. 1B). Approximately 15 d after excision, the shoot started to grow again, producing new leaves (Fig. 1, C and D), an event that was followed by an increase in xyloglucan mobilization (Fig. 2C).
The reduction observed in dry mass and xyloglucan during mobilization was followed by an increase in the amount of starch, mainly in detached cotyledons. Moreover, addition of 10–6 M 2,4-D to detached cotyledons at a period of advanced mobilization (41 d) promoted an even greater increase in starch synthesis (Fig. 2B), suggesting that this phenomenon might be dependent on xyloglucan mobilization. Seedlings growing in the darkness or with the excision of the shoot showed an induction of starch synthesis, accumulating relatively higher amounts of starch in the cotyledons than the control plants (Fig. 2D). In the excised seedlings, the accumulated starch was mobilized when growth of the shoot restarted. Analysis of the free sugars showed that in attached cotyledons, Fru and Glc increased at the same time as xyloglucan was degraded, peaking at 48 d and decreasing rapidly afterward (Table I, control seedling). Suc was also present during the germination period, decreasing quickly up to 26 d and increasing again during xyloglucan degradation. When seedlings were grown in the darkness or when the shoot was excised, a strong reduction in the concentration of free sugars was observed (Table I). The resumption of xyloglucan mobilization after excision of shoot was also confirmed by a parallel increase in Glc, Fru, and Suc in the cotyledons (Table I).
The analysis of detached cotyledons showed that only those isolated after 34 d were capable of maintaining an increase in free sugars, reaching higher levels when detachment occurred at 41 d (Table I). Likewise, cotyledons treated with 2,4-D at 41 d showed high free-sugar contents during xyloglucan mobilization. It is important to note that isolated cotyledons tended to show a higher Suc to monosaccharide ratio than attached cotyledons.
In attached cotyledons, all hydrolase activities related to xyloglucan mobilization increased during the same period of the most intense changes in dry mass and carbohydrate contents. In all treatments, the increase in the activities of XEH,
In cotyledons detached at 19 d (Fig. 7, Experiment I), the activities of the three enzymes did not increase at the same rate as in the attached cotyledons. However, the cotyledons isolated at 34 and 41 d and kept in water thereafter showed a less pronounced increase in activities of both endo- and exo-enzymes in comparison with detached cotyledons incubated in 2,4-D or attached cotyledons (Fig. 3, B, D, and F). This occurred concomitantly with the reduction of xyloglucan and with the increase of free sugars and starch in the cotyledons (Table I; Fig. 2). The enzyme activities in cotyledons detached at 41 d were also stimulated by 10–6 M 2,4-D, in particular XEH activity (Fig. 3B). The dark-grown seedlings showed reduction of all hydrolase activities in relation to the light-growth seedlings (Fig. 3, A, C, and E). This was also observed after excision of the shoot of the seedling in both experiments (Figs. 3 and 4). In order to evaluate the isolated effect of auxin on the activity of xyloglucan hydrolases in the presence of sink strength, the following treatments were used: top shoot excision or application of N-1-naphthylphthalamic acid (NPA, an inhibitor of auxin transport). Although after top shoot excision no differences in hydrolase activities had been observed, when NPA was used, with the exception of XEH, all hydrolase activities were strongly reduced (Fig. 4). The exception of this reduction was observed with XEH, which has not been significantly reduced by NPA (Fig. 4A). This reduction in hydrolase activities may be explained by the fact that auxin is produced by the entire shoot, and in our experiments, although excision of the top shoot decreased the level of auxin in the cotyledons, NPA also promoted a strong reduction in indole-3-acetic acid (IAA) contents in the cotyledons in relation to the treatments of top shoot excised and control seedlings (see below).
Aiming to test the hypothesis that there is a relationship between auxin increase and xyloglucan mobilization in the cotyledons of H. courbaril, we sought to measure the endogenous levels of auxin in detached or in attached cotyledons of seedlings submitted to shoot (all parts above cotyledons) and top shoot excision (all parts above the eophylls; see Experiment II in Fig. 7) and auxin transport inhibition (NPA). Our results showed that auxin levels increased at the same period of the xyloglucan mobilization in the cotyledons of H. courbaril (Fig. 5 ). Levels of IAA increased rapidly in cotyledons of intact (control) seedlings (approximately 8-fold), but in cotyledons of seedlings that had their shoot excised at 34 d, the highest IAA level was reached several days later and coincided with the observed regrowth of the shoot (compare Figs. 5A and 1D). On the other hand, in cotyledons detached at 34 d, no increase in IAA was observed (Fig. 5A). A similar behavior was observed for attached cotyledons from seedlings in the dark, from shoot excised kept in the light, or NPA treated (Fig. 5, A and B). Although these results suggest that the cotyledons of H. courbaril appear not to be able to produce endogenous auxin, this possibility cannot be excluded since in the light-kept cotyledons (NPA or shoot excised), a small increase in IAA was observed throughout the experiment (Fig. 5B).
According to Bewley and Black (1994) , there are two possible mechanisms by which the embryonic axis can control storage mobilization in seeds. One is through the action of the axis as a sink organ, metabolizing the products of storage compounds (e.g. Suc), avoiding feedback mechanisms by their end products, and another is through the delivery of a signal (e.g. a hormone) to modulate storage mobilization. Our results can be used as a direct evidence for the hypothesis of the existence in developing seedlings of H. courbaril of a hormonal signal (auxin) as the principal control of xyloglucan mobilization in this tree species. However, other mechanisms related to metabolic events in the shoot organs (eophyll and leaves), as well as in the cotyledons, appear to participate in a chain of events that are related to the modulation of xyloglucan catabolism and use of its products during seedling growth.
Our results suggest that the embryonic axis apparently plays a role in xyloglucan mobilization by establishment of a sink for its degradation products, mainly in the expanding leaves. In this process, as previously proposed by Chory (1993)
Our results suggest that accumulation of starch might be dependent on xyloglucan mobilization. Indeed, transitory starch has been observed previously in seeds that mobilize galactomannan (Reid, 1971
According to Rolland et al. (2002)
In the cotyledons of H. courbaril, the accumulation of starch may be therefore considered as a temporary consequence of the relationship between source (xyloglucan degradation) and sink (synthesis/transport of Suc to the shoot) intensities. By storing starch transitorily, the seedling may thus avoid potentially adverse effects of accumulation of high concentration of reducing sugars (Geiger et al., 2000
At the H. courbaril cotyledon cell wall level, the balance between XET and XEH activities is dependent on the oligosaccharide concentration present in the apoplast (Tiné et al., 2000
A possible explanation for the effect of light is the observation of presence of active photosynthesis (only electron transport but not CO2 assimilation) in cotyledons of H. courbaril (M.P.M. Aidar, U. Lutgge, L.I.V. Amaral, M.S. Buckeridge, unpublished data). According to recent observations by Rolletschek et al. (2003) Altogether, these observations suggest that xyloglucan mobilization is probably controlled by multiple factors at different levels in cotyledon tissue. There appear to exist points of control acting as local effects of reaction products in the wall (e.g. oligosaccharides and free Gal), up to a more indirect environmental effect of light on the cotyledons themselves.
Considering the second mechanism proposed by Bewley and Black (1994)
Altogether, the results confirm that the IAA present in the cotyledons during xyloglucan mobilization is produced in and transported from the developing shoot, which is considered the main site of auxin biosynthesis (Bartel, 1997
The levels of IAA in the cotyledons seem to be strongly associated with polar auxin transport, as we confirmed by NPA treatment, and not with hydrolysis of IAA conjugates as has been described for seedlings of Pinus sylvestris (Ljung et al., 2001
Recently, studies have demonstrated the importance of polar auxin transport in many aspects of plant development, such as the expansion of hypocotyl of cucumber (Shinkle et al., 1998
The single work in which a relationship between auxin and storage cell wall metabolism had been reported was in cotyledons of nasturtium (Hensel et al., 1991
Considering the results in terms of the effects of sink and auxin on carbohydrate metabolism in the cotyledons, it can be suggested that both light and aerial parts have a strong relationship with the catabolism of the storage cell wall xyloglucan and transient accumulation of starch and Suc, as summarized in Figure 6 . As H. courbaril is a shade-tolerant species (Souza and Válio, 1999
Shinkle et al. (1998) demonstrated that red light is capable of inducing polar transport of auxin in Arabidopsis and cucumber, respectively. However, Kraepiel et al. (2001) showed indirectly that in tomato, the transport was independent of light. Our results showed that darkness strongly inhibited polar auxin transport in seedlings of Hymenaea. Also, in experiments where seedlings were grown under different conditions of red to far red ratios, mobilization was faster in increasing proportions of red light (H.P. Santos and M.S. Buckeridge, unpublished data). This suggests that seedlings of H. courbaril respond to light in a similar way to that of Arabidopsis and cucumber. Considering the results of Shinkle et al. (1998) on the influence of light on auxin polar transport, the hormonal control observed could play an important ecophysiological role in seedlings of H. courbaril by promoting synchronism between growth and reserve degradation under variable light conditions (Fig. 6). This is likely to increase the efficiency of carbon reserve utilization by the growing seedling in the understorey of the rain forest.
Altogether, our results corroborate the hypothesis that the presence of cell wall polysaccharides as storage compounds in cotyledons reflects an ancient functional transfer during evolution, as has been suggested by Buckeridge et al. (2000)
Material and Experimental Conditions In all experiments we employed size-selected seeds (5.5–6.0 g) of Hymenaea courbaril, collected in São João da Boa Vista county (22°00'S; 47°18'W), São Paulo, Brazil. These seeds were stored for 4 years under dry and cold conditions (relative humidity 35%, 8°C) in the Botanical Institute of São Paulo, Brazil. Seeds were scarified with sandpaper on the lateral position in relation to the embryo, surface-sterilized for 15 min in 10-fold-diluted commercial hypochlorite bleach, rinsed, and placed on trays between two sheets of wet paper (at 30°C) until germination was visible (0.5 cm of radicle). Elapsed time in all experiments was registered in relation to the beginning of imbibition, in days (days after imbibition of seeds). Germinated seeds were placed in pots (1.5 L) with a washed sand:vermiculite mixture (2:1, v/v), and the pots were placed in a growth chamber, under shelves equipped with 10 fluorescent lamps (60 W) and four incandescent lamps (40 W) reaching around 200 µE m–2 s–1 of photosynthetic active light intensity. The photoperiod throughout the experiments was 12-h-light/12-h-dark cycle with constant temperature (25°C) and relative humidity (60%). Every 15 days, 50 mL of a complete Hoagland solution was added to each pot to avoid mineral deficiency.
This study was performed with two complementary experiments, which are summarized in Figure 7 . The first one compared the xyloglucan mobilization process in detached and attached cotyledons. The attached cotyledons were followed in intact seedlings (control), in the darkness, and with excision of the shoot above the cotyledon insertion (shoot excised). With these procedures, we intended to characterize the xyloglucan mobilization process in intact plants or in plants growing without light stimulus or shoot sink, as well as to check whether and when the isolated cotyledons are able to start/maintain the mobilization process. With these experiments, it has been possible to test whether cotyledons are able to respond to endogenous factors, including auxin. The control plants were grown in pots as described above and without growth restrictions. For the dark treatment, the pots were placed into a black paper box under the same growth chamber conditions when the emergence of seedlings started, approximately 19 d. The shoot excision was performed at about 34 d, when the development of eophylls (the first green leaves developed by seedlings) started, by cutting the entire shoot above the cotyledons insertion (epicotyls and eophylls).
Detachment of cotyledons was performed at 7, 19, 26, 34, and 41 d after the beginning of imbibition, from plants grown as in the control treatment. The detachment dates were chosen on the basis of the results obtained by Tiné et al. (2000)
The second experiment (Fig. 7) was performed only with attached cotyledons in seedlings following different treatments in order to distinguish between the effects of sink strength, endogenous IAA, and light on xyloglucan mobilization. The treatments were: intact seedlings (intact), shoot excised seedlings (excised), shoot excised seedlings with light-protected cotyledons (excised LPC), shoot apex excised seedlings (top shoot excised), and intact plants treated with NPA at 200 µM (NPA 200 µM). The intact seedlings were grown under the same conditions of light, temperature, and humidity as used for the control plants in the first experiment. Shoot excision was performed as in the first experiment. However, for seedlings that had their top shoot excised, the plants were grown without the apex of the last internode (the second), the remaining sink being the expansion of eophylls. In light-protected cotyledons treatment, the cotyledons were wrapped with aluminum foil before the start of xyloglucan mobilization (approximately 26 d). At the same time, the NPA treatment was performed by applying a ring of lanolin paste containing 200 µM NPA (concentration chosen according to Reed et al., 1998 In all experiments, treatment samples of seedlings (Experiment I) and/or cotyledons (Experiments I and II) were collected once a week. After measurements of leaf area, all materials were stored at –80°C.
The seedlings of the first experiment were divided into leaves, stems (nodes and internodes), and roots. Leaf area was analyzed using a digital system (Skye Instruments, Llandrindod Wells, UK) and together with the other seedling parts dried at 70°C (72 h) and weighed to determine dry mass. The analyses were performed separately on five seedlings per sample.
Ten cotyledons per sample (Experiment I) were divided into two groups, in which five were used for dry mass and carbohydrate determinations, and the other five were used for enzyme analyses. Samples of the first group were dried at 70°C (72 h), weighed (dry mass), powdered, and divided into three subsamples (100 mg each). These were subjected to xyloglucan extraction in 30 mL of water at 80°C for 8 h. After filtration through nylon cloth, centrifugation was performed (10,000g, 30 min, 5°C), followed by precipitation with three volumes of ethanol. The precipitate was stored overnight at 5°C, collected by centrifugation, freeze-dried, and weighed. The water-soluble polysaccharides from the cotyledons produced a freeze-dried fluffy material that comprised more than 95% xyloglucan (Buckeridge et al., 1992
The same powdered cotyledons used for xyloglucan extraction were submitted to starch and soluble sugar measurements. The starch analyses were performed according to an enzymatic technique described by Arêas and Lajolo (1980) To measure the soluble sugars (Suc, Glc, and Fru) the alcohol supernatants were dried, suspended in water (1 mL), filtered (Millipore 0.25; Bedford, MA), and analyzed by high performance anion-exchange chromatography on a CarboPak PA-1 column (Dionex, Sunnyvale, CA) using a gradient elution from 0 (water) to 200 mM NaOH in water (20 min). Sugars were detected by a pulsed amperometric detector (PAD; Dionex). Detector responses were compared with the standards of Glc, Fru, and Suc at 25, 50, 75, 100, 150, and 200 µM. The standard curve for each sugar was used to calculate carbohydrate contents in the cotyledons.
Samples of cotyledons were weighed, cut into small pieces, and pooled to compose three subsamples (0.5 g each). These subsamples were homogenized for 15 s (Ultra-Turrax T25, IKA- Labortechnik, Staufen, Germany ) with 5 mL of sodium acetate buffer (500 mM, pH 5.0). The homogenized subsamples were kept at 5°C for 20 min, and after centrifugation (10,000g, 10 min) the supernatants were separated. Protein concentration was estimated according to Bradford (1976)
The determination of
The
For determination of XEH, we adapted the method described by Sulová et al. (1995)
The levels of free endogenous IAA were determined by two distinct techniques. In the first experiment, an ELISA technique was employed in which the IAA was detected by a specific antibody (Peres et al., 1997
For IAA determination by ELISA, the same samples as for enzyme activity were used. These samples were from: (1) attached cotyledons in control (34, 41, and 48 d); (2) in darkness (34, 41, and 48 d); (3) in shoot excised seedlings (41, 54, 61, and 68 d); and (4) detached cotyledons (34 and 41 d). Each previously ground sample was divided into three aliquots comprising subsamples of 1 g that were individually powdered using liquid nitrogen and submitted to an alcohol extraction. This was performed by addition of 3 mL of methanol per subsample and stirring for 60 h in the darkness at 4°C. During this extraction step 100 µL of [3H]IAA (0.5 µCi mL–1) was added to each sample as an internal standard in order to determine the extraction and purification yield of IAA. The extracts were filtered in nitrocellulose (0.45-mm Millex-HV and 0.22-mm Millex-GS [Millipore]), followed by Sep-Pak C-18 column, previously preconditioned with 80% (v/v) methanol. The filtered sample was dried, suspended in 0.2 mL L–1 formic acid, pH 3.0, and submitted to IAA purification by HPLC on a C-18 column using a gradient elution with 0.2 mL L–1 formic acid, pH 3.0, and methanol (45 min). The radioactive fractions were pooled, dried, methylated with diazomethane, resuspended in water, and distributed onto a microplate of ELISA spectrophotometer preconditioned with the specific antibody to methylated-IAA, according to Peres et al. (1997)
The GC-SIM-MS analyses of free endogenous IAA were performed according to Chen et al. (1988)
We thank the FAPESP for granting a doctoral fellowship to H.P.S. (98/02775–8) and CNPq for a research fellowship to M.S.B. We thank Mrs. A.M. Baroni, L.I.V. Amaral, and V. Tamaki for technical assistance in the auxin procedures. We also thank the colleagues Jocelyn Rose and Yvan Krapiel for the useful suggestions. Received February 1, 2004; returned for revision March 21, 2004; accepted March 23, 2004.
1 This work was supported by Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP; BIOTA-FAPESP 98/05124–8). 
2 Present address: EMBRAPA Uva e Vinho, Rua Livramento, 515, Caixa Postal 130, CEP 95700–000, Bento Gonçalves, RS, Brazil. Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.104.040220. * Corresponding author; e-mail msbuck{at}usp.br; fax 55 11 50733678.
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ABSTRACT
RESULTS
4)-glucan backbone branched with 
(1
-nitrophenyl galactopyranoside, had a pH optimum at 3.2, whereas the other hydrolases detected were active at 4.5. The results obtained for 
