Cell Wall Storage Carbohydrates in Seeds—Biochemistry of the Seed “Gums” and “Hemicelluloses”

Cell Wall Storage Carbohydrates in Seeds-Biochemistry of the Seed “Gums” and L6Hemicelluloses”

J. S. GRANT REID Department of Biological Science University of Stirling Stirling, Scotland

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Structures of Cell Wall Storage Carbohydrates in Seeds ........................ A. Background Methodology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Structural Types and Their Distribution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Formation and Postgerminative Catabolism ......... ..... ...... A. Galactomannan Metabolism in Leguminous Seeds .......................... B. Mannan Mobilization in the Date Endosperm ........................ C. Xyloglucan Metabolism in Tropaeolum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. “Galactan” Metabolism in Lupinus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1V. Considerations of Biological Function . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . V. Perspectives ....................... .................................. .................. References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

126 126 128 132 133 144 144 146 148 152 153

I. INTRODUCTION By the late nineteenth century it was clearly recognized by botanists that the massively thickened cell walls present in many seeds contained reserve substances. Reiss (1889) and others described “reserve celluloses” which were utilized following germination, while Tschirch (1 889) and Nadelmann (1 890) were able to demonstrate that the “mucilages” in the endosperm cell walls of some leguminous seeds had a storage function. Schleiden (quoted by Vogel and ADVANCES IN BOTANICAL RESEARCH. VOL. I I

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Schleiden, 1839) reported that the thickened cell walls of some seeds could be stained blue with iodine, and he named the substance responsible for the starchlike reaction “amyloid.” Amyloids were later shown to occur widely in seeds (Winterstein, 1893; Kooiman, 1960a) and to be mobilized following germination (Godfrin, 1884). The carbohydrate nature of the cell wall reserves of seeds was inferred from their microchemical staining reactions, and proven by the positive identification of sugars released from them on acid hydrolysis. For example, the reserve cellulose of the ivory “nut,” Phytelephas macrocarpa, released “seminose” or mannose (Reiss, 1889); the mucilage of the locust “bean,” Ceratonia siliqua, gave mannose and galactose (Bourquelot and Herissey, 1899), and the amyloid of the nasturtium seed, Tropaeofummajus, yielded glucose, xylose, and galactose (Winterstein, 1893). The combined ultrastructural, physiological, and “biochemical” approach which many of the early botanists adopted to study the cell wall storage carbohydrates of seeds was extraordinarily effective. It is unfortunate that it was not carried forward with vigor into the twentieth century. The mid-twentieth century (1930- 1970) saw the introduction of a series of new techniques for the determination of the structures of complex carbohydrates (Whistler and Wolfrom, 1965; Whistler and Bemiller, 1972) and most of our present knowledge of the structures of cell wall storage carbohydrates was obtained during that period. Seeds were generally treated with alkali to extract polysaccharides of the “hemicellulose” type or with water to extract “gum” polysaccharides. Consequently the molecules with which this article is concerned are still widely classified as seed gums and hemicelluloses. In recent years (from about 1970) there has been a reawakening of interest in the physiology and biochemistry of the cell wall storage carbohydrates of seeds, and a recent review article has treated them for the first time as a single, botanically coherent group of substances (Meier and Reid, 1982). It is the purpose of this article to outline the structures and occurrence of cell wall storage carbohydrates, to give an account of current research on their metabolism, and to explore their overall biological significance in the seeds which contain them. 11. STRUCTURES OF CELL WALL STORAGE CARBOHYDRATES IN SEEDS This section indicates the principal types of carbohydrate molecules stored in the cell walls of seeds, and their distribution. To allow the assessment of the status of published structural data of various kinds the section is prefaced by a brief resum6 of the methods which have been used to determine these structures, and their limitations. A. BACKGROUND METHODOLOGY

The determination of the primary structure of a cell wall polysaccharide normally involves the following procedures:

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1. Isolation of polysaccharide material from the plant tissue 2. Purification to a degree of homogeneity acceptable for structural determination 3. Total hydrolysis to release the constituent monosaccharides for qualitative and quantitative analysis 4. Determination of molecular weight, or degree of polymerization 5. Linkage analysis to determine linkage types ( 1 += 2, 1 + 3, etc.), linkage modes (a or p), ring sizes (pyranose or furanose), and to obtain some information concerning the distribution or ordering of monosaccharide residues within the molecule The isolation of cell wall polysaccharides from seeds has normally been carried out by treating the tissue with a solvent, usually hot water or dilute alkali. There may be a pretreatment to remove lipids and/or low-molecular-weight carbohydrates or to inactivate enzymes. Hot water is unlikely to cause extensive degradation of carbohydrate macromolecules, but it can bring about irreversible changes in their noncovalent interactions. (Once gelatinized and partially solubilized, a starch granule cannot be reconstituted.) Alkali is potentially degradative. In the presence of oxygen it can bring about the sequential oxidative cleavage of monosaccharide residues from the reducing end of the molecule (Whistler and Bemiller, 1958). Alkaline oxidation of this type can be avoided (Aspinall et al., 1961), but other types of alkaline modification are inevitable. Substituents bound by ester linkage can be cleaved or can migrate (Bouveng et al., 1960), p-elimination reactions can occur at uronic acid residues, and nonglycosidic linkages between wall components may be broken. It must also be borne in mind that polysaccharides are generally polydisperse (they span a range of molecular weights) and polymolecular (they encompass a limited range of molecular structures). Incomplete extraction from the tissue can, therefore, cause unwanted fractionation: the polysaccharide material passing into solution need not be identical in molecular weight and/or structure with that which is left behind in the tissue. Once extracted and isolated, polysaccharide preparations may be purified, usually by fractional or selective precipitation by organic solvents or metal ions (Whistler and Woifrom, 1965). These methods are simple to carry out, but they suffer from two disadvantages. Insufficient purification may lead to structural studies being carried out on a mixture of distinct molecular types while overzealous purification may subfractionate a polydisperse and polymolecular native polysaccharide. The complete hydrolysis of polysaccharides is effected by acids. Different workers routinely use different hydrolysis conditions-for example, 72% H,SO, at 30°C followed by 4% H,SO, at 120°C; 0.5 M H,SO, at 100°C; 1 M trifluoroacetic acid at 121°C (Whistler and Wolfrom, 1965; Albersheim er al., 1967). Different glycosidic linkages differ greatly in their susceptibility to acid hydrolysis, while the monosaccharides released differ greatly with regard to their

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stability in an acidic environment. Acidic sugar residues (uronic acids) present a particularly serious problem. When they are involved in a glycosidic linkage they often stabilize it, yet once released into an acid medium they are highly unstable. There is no fully satisfactory method of determining the uronic acid residues in a polysaccharide quantitatively. Ideally, published data on the composition of polysaccharides should be corrected for losses during the hydrolysis procedure; in practice this is seldom done. Molecular weight determinations may be carried out by physical techniques or by chemical (end-group) analyses. The former are often subject to errors arising from the self-association of polysaccharide molecules in solution, while the latter may be subject to disproportionate interference from low-molecular-weight, partially degraded material. At the heart of any linkage analysis of a polysaccharide there is a methylation analysis (Whistler and Wolfrom, 1965). The entire molecule is subjected to a procedure to convert all free hydroxyl groups to methyl ethers, and the permethylated product is then hydrolyzed with acid to give a mixture of partially methylated monosaccharides which can be derivatized, separated, and identified. This reveals which hydroxyl groups were involved in glycosidic linkage within the intact macromolecule provided that the original methylation was complete and that no extensive demethylation took place during the subsequent hydrolysis procedure. Methylation analysis gives no information on linkage modes (aor p), nor does it provide much information on the order of residues in a polysaccharide. Additional information can be obtained from selective cleavage of particular types of glycosidic linkages by acids or enzymes (if available). On the basis of methylation analyses and associated techniques, “structures” may be postulated for the molecule. These can be tested by periodate oxidation techniques (Whistler and Wolfrom, 1965) since periodate ion, IO,-, cleaves 1,2 diols in a stoichiometric reaction to give predictable products. The application of the above methods will give a “structure” for a polysaccharide preparation, but it must be understood that such structures are not necessarily definitive. They are, of course, only average structures, but they will also to some extent reflect the method and completeness of extraction, the degree of purification, the methods of hydrolysis and quantitative analysis of sugars, the method used to determine the molecular weight, and the procedures used in the methylation analysis. This should be borne in mind, particularly when comparing polysaccharide preparations. B . STRUCTURAL TYPES AND THEIR DISTRIBUTION

Full structural studies have been carried out on polysaccharides isolated from the seeds of relatively few species. Nevertheless, on the basis of these and of numerous more limited investigations, it is possible to discern several major types of cell wall storage carbohydrate molecules: the mannan group of polysaccharides,

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the xyloglucans, and a galactose- and arabinose-rich class which for convenience will be referred to here as the “galactuns.” 1. The Mannun Group The mannan group comprises three distinct structural types: the “pure” mannans, the glucomannans, and the galactomannans. They are structurally related in that all three are based on 1 -+4, P-linked D-mannopyranose residues. They are distributionally related also, in that they are found only in seed endosperms as opposed to storage cotyledons or axes. “Pure” mannans, that is polysaccharides yielding over 90% mannose on hydrolysis, have been obtained from the seed endosperms of two palms, Phoenix dactylifera, the date palm, and Phytelephas macrocarpa, the ivory nut tree, and their structures have been subjected to thorough investigation over a long period (Ludtke, 1927; Klages, 1934; Aspinall et al., 1953, 1958; Meier, 1958). Both seeds have yielded two mannans (A and B) differing in their solubilities in alkali and cuprammonium solutions. All four polysaccharides have similar structures: a linear 1 -+ 4, @-linkedD-mannan backbone carries a small proportion (less than 2%) of single-unit a-D-galactopyranosyl substituents linked 1 + 6 to mannose. The mannans A and B differ in molecular weight (Meier, 1958) but it is not clear whether they are two functionally distinct molecular species or subfractions of a polydisperse native mannan. Meier’s (1958) observation that the mannans A give an X-ray diffraction pattern in the native state and that mannans B do not suggests that they are distinct. Mannan preparations from the endosperms of three other palm seeds have been investigated in less detail, but they are clearly similar in structure to the mannans of ivory nuts and dates (Mukherjee and Rao, 1962; El Khadem and Sallarn, 1967; Robic and Percheron, 1973). Yet other palm seeds are known to have endosperms which release D-mannose on hydrolysis (Lienard, 1902, cited by Herissey, 1903). It seems reasonable to assume that the cell wall storage carbohydrates in the hard endosperms of all palm seeds are mannans. Mannans with structures similar to those of the date and the ivory nut have been obtained by alkali extraction of coffee beans (Co#eu arabica) (Wolfrom et al., 1961) and of the endosperm of the seed of the umbellifer Carum r a n i (Hopf and Kandler, 1977). The endosperms of other umbelliferous seeds are known to contain reserve celluloses (Hegnauer, 1973) which are probably also mannans. Glucomannans have been obtained by alkali extraction of the seeds of Asparagus oficinalis (Goldberg, 1969; Jakimow-Barras, 1973), Endymion nutans (Goldberg, 1969), Scilla nonscripta (Thomson and Jones, 1964), Iris ochroleucu, and I . sibirica (Andrews et al., 1953). All have been subjected to structural investigation, including methylation analysis, and all are similarly constituted. A linear 1 -+ 4, P-linked backbone contains almost equal numbers of Dglucopyranosyl and D-mannopyranosyl residues. Their distribution is uncertain, and could be random. To the backbone is attached a small percentage ( 3 to 6%)

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of single-unit o-galactopyranosyl branches attached 1 + 6, probably by a linkages (Goldberg, 1969). All of the above species are from the Iridaceae and Liliaceae, other species of which are known to have seeds which are rich in mannose- and glucose-containing polysaccharides (Jakimow-Barras, 1973) or which have thick-walled endosperms (Elfert, 1894). Glucomannans may be characteristic of the hard endosperm of seeds from these families. Galactomannuns are the best characterized of all the cell wall storage carbohydrates of seeds (Whistler and Smart, 1953; Smith and Montgomery, 1959; Stepanenko, 1960; Dea and Morrison, 1975), a fact which reflects the industrial importance of some seed galactomannans (see Glicksman, 1953; Carlson et al., 1962; Saxena, 1965; Chudzikowski, 1971; Kovacs, 1973; Nurnberg and Rettig, 1974, for examples of their uses). The galactomannans are typical of the leguminous seed endosperm (Anderson, 1949) and they can be completely extracted from isolated endosperms or seed tissue with hot water. Numerous leguminous seed galactomannans have been subjected to full structural analysis, and they conform to a common structural type. A 1 44, P-linked D-mannan backbone is heavily substituted by single-unit a-D-galactopyranosyl side chains linked 1 46 to mannose. The degree of galactose substitution in galactomannans varies from about 20 to nearly 100% and is apparently genetically controlled and chemotaxonomically useful (Reid and Meier, 1970; Kooiman, 1971; Campbell, 1978). The lower degrees of substitution are characteristic of the LeguminosaeCaesalpinioideae, which are generally held to be more primitive than the Leguminosae-Faboideae. Only one endospermic leguminous seed has so far been found to contain a storage polysaccharide other than galactomannan. The seed of the Judas tree (Cercis siliquastrum: Leguminosae-Caesalpinioideae)contains a galactoglucomannan; its structure is similar to those of the galactomannans but the backbone contains D-glucopyranose residues (McCleary el al., 1976). The only nonleguminous species whose mature seeds contain galactomannans are from the Convolvulaceae (Khanna and Gupta, 1977; Kooiman, 1971). Interestingly, however, galactomannans have been isolated from the immature seeds of palm species which contain “pure” mannans at maturity (Mukherjee et al., 1961; Kooiman, 1971; Balasubramaniam, 1976). A developmental relationship between galactomannan and mannan in the palm seed endosperm has been suggested (Balasubramaniam, 1976), but has not been investigated experimentally. The mannans, the glucomannans, and the galactomannans are clearly related structurally, but in their physical properties the mannans and glucomannans differ considerably from the galactomannans. The former are almost completely insoluble in water by virtue of their cellulose-like structures, and endosperms that contain them are hard. In the cell wall they may be crystalline (Kooiman, 1960b). The highly substituted galactomannans are water soluble: presumably the galactosyl substituents effectively prevent the self-association of the main chain to give crystalline aggregates. In their properties, the mannans and

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glucomannans are ‘‘reserve celluloses” whereas the galactomannans are ‘‘seed mucilages.” 2 . The Xyloglucans The xyloglucans are amyloids, that is they can be stained blue with iodine both in situ and when extracted from the cell wall. The physical basis of the iodine coloration is not known, but it has been exploited both to survey seeds for the presence of amyloid (Kooiman, I960a) and to analyze xyloglucans quantitatively (Kooiman, 1960b). Kooiman (1960a) carried out iodine staining on endosperm and/or embryo tissue from seeds of over 2600 species and found 230 of them to be amyloid positive. In the Leguminosae-Caesalpinioideae, amyloid was restricted to the nonendospermic tribes Cynometreae, Amherstieae, and Sclerolobieae. All investigated species of the Primulales, Annonaceae, Limnanthaceae, Melianthaceae, Pedaliaceae, Thunbergiaceae, and Tropaeolaceae contained amyloid as did a number of species of Balsaminaceae, Acanthaceae, Leguminosae-Faboideae (all amyloid-positive species were nonendosperrnic), Linaceae, Ranunculaceae, Sapindaceae, and Sapotaceae. Detailed structural analyses have been carried out on “amyloids” extracted with alkali from seeds of only 4 of the 230 species, namely: Tamarindus indica (Leguminosae-Caesalpinioideae) (Kooiman, 1961), Tropueolum majus (Tropaeolaceae) (Le Dizet, 1972), Impatiens balsamina (Balsaminaceae) (Courtois and Le Dizet, 1974), and Annona muricuta (Annonaceae) (Kooiman, 1967). All have similar structures: a linear 1 + 4, P-linked glucan (cellulosic) backbone carrying substituents of two types, a-D-xylopyranosyl and P-D-galactopyranosyl(1- 2) a-D-xylopyranosyl. Both types of side chain are attached to C-6 of D-glucose. Methylation analysis has indicated that there may be some branching of the backbone and that (1 + 3) linkages may be present (Courtois et al., 1976). These results could arise from contaminating D-glucans. The ratio ga1actose:xylose:glucose is 1:2:3 in the xyloglucans isolated from Tamarindus and Tropaeolum seeds, 1 :2:4-5 in that from Impatiens, and 1:1:4 in the polysaccharide from Annona. It is not clear to what extent these differences reflect differences in methods of isolation, purification, and structure determination; nor is it clear how wide a variation in seed xyloglucan structures may yet be encountered. Even the assumption that all amyloids are xyloglucans need not necessarily be valid. 3. The Galactans The cotyledon cell walls of several lupins, notably the agriculturally important species Lupinus angustifolius, L. albus. and L. luteus, are massively thickened. Schulze and Steiger ( I 889) demonstrated that the cell wall material of L . luteus released D-galactose and a pentose on hydrolysis and named it “Paragalaktan. ” The total cell wall polysaccharides of L. angustifolius cv. Unicrop release galactose (76%) arabinose (13%), xylose (4%) and a uronic acid (7%) on hydrolysis (Crawshaw and Reid, 1984) while isolated walls have a similar composition

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(Hutcheon and Reid, unpublished). Linkage analysis of the isolated walls is underway. A water-soluble 1 + 4, P-linked D-galactan has been isolated from the cotyledons of L . albus in very low yield and using isolation techniques which would certainly lead to extensive degradation of polysaccharides (Hirst et a l . , 1947). This “lupin galactan” is clearly not representative of the water-insoluble native wall, but it indicates the presence within it of a 1 + 4, P-linked galactan component. Methylation analysis of the native walls confirms that the main galactosidic linkage is 1 + 4 (Wilkie, Reid, and Hutcheon, unpublished). 4 . Other Cell Wall Storage Carbohydrates The cell wall storage carbohydrates of so few species have been subjected to any chemical investigation that the list of major structural types given here can scarcely be considered complete. Further structural studies are necessary. Valuable ultrastructural and physiological studies have even been carried out on the postgerminative mobilization of cell wall storage carbohydrates of unknown or partially known structure. The perisperm of Yucca, for example, contains deposits of cell wall carbohydrate, the mobilization of which has been carefully documented (Homer and Arnott, 1966); yet their structure has not been determined. Similarly the mobilization of the cell wall storage carbohydrates in the endosperm of lettuce (Lactuca sativa) has been observed (Jones, 1974), one of the enzymes responsible has been identified (Halmer et a l . , 1978), and the control of the mobilization process has been investigated (Halmer and Bewley, 1979). Yet it is not clear whether the storage carbohydrate in the lettuce endosperm is a “pure” mannan, a glucomannan, a galactomannan, or some intermediate type. The control of cell wall storage carbohydrate metabolism following germination has also been studied in the coffee bean (Takaki and Dietrich, 1979) although so far only a small proportion of the cell wall material has been positively identified as “pure” mannan (Wolfrom et al., 1961; Wolfrom and Patin, 1965). The structures of quantitatively minor deposits of cell wall storage carbohydrates have received little attention, with the notable exception of the cell wall carbohydrates present in the endosperms of the commercially important cereal grains, alongside massive deposits of starch. These “cereal gums” will not be considered here (but see Meier and Reid, 1982). 111. FORMATION AND POSTGERMINATIVE

CATABOLISM This section deals in detail with a limited number of systems: galactomannan metabolism in leguminous seeds, mannan mobilization in dates, xyloglucan breakdown in nasturtium seeds (Tropaeolum majus), and “galactan” catabolism in Lupinus. All of them were first studied in the late nineteenth century and are being reinvestigated now.

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A. GALACTOMANNAN METABOLISM IN LEGIJMINOUS SEEDS

The occurrence, the formation, and the mobilization of leguminous seed galactomannans were first investigated by Nadelmann (1890). In his scholarly dissertation “Ueber die Schleimendosperme der Leguminosen” he documents the morphology of “mucilage” storage in the endosperms of several seeds chosen to represent most of the endospermic tribes in the Leguminosae. He also describes mucilage deposition during seed development in the endosperm cell walls of Trigonella foenurngraecurn, Colutea brevialata, and Tetragonolobus purpureus, and delineates the process of mucilage mobilization following germination in Trigonella foenumgraecum and Tetragonolobus purpureus. Although Nadelmann was not aware of the chemical nature of his seed “mucilages,” it was he who established a storage role for the galactomannans. In recent years there has been renewed interest in the biochemistry and physiology of galactomannan mobilization. The overall morphology and physiology of the process has been studied in two seed systems, fenugreek (Trigonella foenurngraecurn) and carob (Ceratonia siliqua), which exemplify the two extremes of galactomannan structure. The fenugreek seed contains a highly galactose-substituted galactomannan, while the galactomannan of carob is representative of the comparatively low-galactose galactomannans of the LeguminosaeCaesalpinioideae (see Section II,B, I ) . The enzymology of galactomannan degra-

Fig. 1. Cryostat sections of mature, imbibed fenugreek seeds stained with periodic acid-Schiff‘s reagent to reveal periodate-reactive polysaccharides. A, Axis; Al, aleurone layer; C, cotyledons; E, endosperm; T, part of testa. The intense staining in the endosperm is due to galactomannan. (After Reid and Bewley, 1979.)

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dation has been most fully studied with respect to the guar seed (Cyamopsis tetragonoloba), which contains a galactomannan of intermediate galactose substitution. Galactomannan formation has so far been studied only in developing fenugreek seeds. 1. Galactomannan Formation and Mobilization in the

Fenugreek Seed The mature fenugreek seed contains about 30% by weight of a high-galactose galactomannan (mannose : galactose = 53 : 47) which is localized in the endosperm. Most of the endosperm cells appear to be completely filled with the polysaccharide (Fig. 1) and must be considered to be nonliving; the only living cells in the endosperm are those of the one-cell-thick aleurone layer which surrounds the storage tissue (Reid and Meier, 1972). Studies of endosperm development have confirmed that the galactomannan which “fills” the storage cells is a cell wall polysaccharide, and germination studies have established a key role for the aleurone layer during galactomannan degradation. a. Galactomannan Formation during Endosperm Development. The time course of galactomannan deposition in the developing endosperm is shown in Fig. 2, while Figs. 3 and 4 illustrate the morphological changes which occur in the endosperm during that period. Galactomannan is deposited in the form of secondary thickenings on the cell walls of the endosperm, and the deposition continues until the galactomannan occupies the whole volume of the cell. Cytoplasm and vacuole disappear and are replaced by a mass of galactomannan. Galactomannan deposition is a cell-by-cell process; the endosperm cells next to

Fig. 2. Time course of galactomannan (0) and stachyose fenugreek seeds. (After Campbell and Reid, 1982.)

(a) accumulation

in developing

Figs. 3 and 4. Cryostat sections of a fenugreek seed nearing completion of galactomannan deposition in the endosperm: sections examined by Nomarski interference contrast. Al, Aleurone layer; C, cotyledon; G ,galactornannan; PW, primary wall; T, part of testa. The endosperm cells nearest the cotyledons are already “filled” with galactomannan (Fig. 3) while the outermost cells, nearest the aleurone layer, are still depositing galactomannan (Fig. 4).

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the embryo are the first to be “filled” (Fig. 3) while those adjacent to the aleurone layer are last (Fig. 4).The mechanism which protects the aleurone cells from being “filled” with galactomannan is not understood; some galactomannan-like material is laid down on these cells but this deposition soon ceases (Meier and Reid, 1977). The galactomannan seems to be formed initially in the intracisternal space or “enchylema” of the rough endoplasmic reticulum (RER). which swells greatly and stains for periodate-reactive carbohydrate. The enchylema swells to such an extent that the cytoplasmic lamellae between the ER cisternae become pinched off to give “inside out” RER vesicles, with the ribosomes on the “inside” (poculiform ER). Where the enchylema makes contact with the plasmalemma its contents appear to be discharged into the growing wall space (Meier and Reid, 1977).

Fig. 5 . Electron micrograph of a cell almost “filled” with galactomannan ( G ) . Irregularly distributed residues of protoplasm (RP) enclose small pockets of galactomannan. Section contrasted with periodic acid, thiocarbohydrazide, and silver proteinate. (After Meier and Reid, 1977.)

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Days after onthesis

Fig. 6 . Incorporation of label from GDP-D-[U- ‘‘C]mannose into galactomannan by whole homogenates prepared from developing fenugreek seed endosperms. (Campbell and Reid, 1982.)

The degradative processes leading to the disappearance of the cytoplasm and its organelles have not yet been investigated. Some cytoplasmic material may even be sloughed off and lost into the growing cell wall (Fig. 5). It is interesting to observe that the final remnants of cytoplasm in a cell nearing the completion of galactomannan deposition remain in contact with neighboring cells, via plasmodesmata (Meier and Reid, 1977). This may be the route of cell to cell transport of cytoplasmic degradation products, which may themselves serve as substrates for galactomannan formation. During the period of galactomannan deposition (Fig. 2), the endosperm contains high levels of an enzymatic activity which catalyzes the transfer of Dmannosyl residues from GDP-D-[U-14C]mannoseto a soluble product which has been shown to be galactomannan (Campbell and Reid, 1982). Enzyme activities transferring o-galactosyl units are also present but they have not yet been thoroughly investigated. The GDPmannose : galactomannan mannosyltransferase activity peaks twice during galactomannan deposition; once at the beginning of the deposition and once at the height of galactomannan accumulation (Fig. 6 ) . The early peak corresponds largely to light particulate material ( 100,000g pellet) while the latter peak consists mainly of grossly particulate material (Fig. 7). It is possible that the early peak largely represents transferase activity still associated with endoplasmic reticulum, while the later peak is almost certainly enzyme associated with, or occluded within, large particles of galactomannan. The light particulate enzyme of the early peak sediments with ER markers and has a density of 1.06 g ml-’ (Campbell, 1978). It requires divalent metal ions for activity (Table I). The enomlous stimulation of the enzyme activity by Co2+ and Ni2+ may be due to their interaction with the galactomannan product rather than the enzyme. Both

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&&

Fig. 7. Incorporation of label from GDP-~-[U-"+C]mannose into galactomannan by enzyme preparations, obtained by differential centrifugation of endosperm homogenates. 0-0, Gross particulate enzyme (1000 g pellet); @-a,particulate enzyme (100,000 g pellet); 0.0, soluble enzyme (100,000 g supernatant). (Campbell and Reid, 1982.)

ions are known to form insoluble complexes with galactomannans (Campbell and Reid, 1982). The natural cofactor of the enzyme is probably Mg2+. b. Galactomannan Mobilization following Germination. The breakdown of galactomannan in the endosperm of the fenugreek seed begins 16 hr after germination, and is complete in a further period of 24 hr (Reid, 1971; Reid and TABLE I Effect of Cations on the Particulate GDPmannose:palactomannan Mannosyltransfera& of the Fenugreek Endosperm0 Cation None

Mg2 + Mn2 +

cu2+ Ca2 + co2+

K+

Relative incorporation level

1 23 348 9 24 524 2

" All cations were 10 mM in the assay. For details see Campbell and Reid (1982). (After Campbell, 1978.)

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Bewley, 1979). During this time mannose, galactose, and traces of mannooligosaccharides can be detected in the endosperm, but they do not accumulate to any extent. The galactornannan breakdown products are rapidly absorbed by the embryo and converted to sucrose and starch (Reid, 1971). If endosperm halves are isolated from dry fenugreek seeds and incubated under “germination” conditions, galactomannan breakdown occurs, but the end products of galactomannan degradation accumulate quantitatively and have been identified as galactose and mannose. The galactomannan is therefore depolymerized by hydrolytic as opposed to phosphorolytic cleavage, and the enzymes involved are produced within the endosperm itself (Reid and Meier, 1972). Galactomannan breakdown is paralleled in the endosperm (but nor in the embryo) by a-galactosidase and Pmannanase activities (Reid and Meier, 1973b). In isolated endosperm halves galactomannan breakdown is partially or totally inhibited by metabolic inhibitors, the site of action of which can only be the living cells of the aleurone layer. These inhibitors also suppress a-galactosidase and P-mannanase activities, suggesting a key role for the aleurone layer in the production or activation of these enzymes (Reid and Meier, 1973b). When fenugreek seeds are allowed to germinate in the presence of 80% D,O the endosperm a-galactosidase becomes density labeled (Reid and Davies, unpublished). It is probably synthesized de novo in the aleurone layer. Inhibitor studies suggest that the P-mannanase is probably also synthesized de novo (Reid et al., 1977). According to Reese and Shibata (1965) the complete hydrolytic breakdown of a leguminous galactomannan would require at least three enzymes-an a-galactosidase, a P-mannanase, and a pmannoside mannohydrolase (P-mannosidase). The fenugreek seed endosperm does contain P-mannosidase activity which increased fourfold during galactomannan breakdown (Reid and Meier, 1973b). Recent studies on galactomannan breakdown in guar (Cyamopsis tetragonoloba) seeds have shown conclusively that the P-mannoside mannohydrolase of the guar endosperm is not synthesized de novo and that it is present in association with galactomannan even in the resting endosperm (McCleary, 1983). McCleary (1982) has demonstrated that complete extraction of the P-mannoside mannohydrolase from the endosperm is possible only if the galactomannan is first depolymerized. Consequently it is probable that the P-mannoside mannohydrolase of the fenugreek endosperm is also present in an active form in the resting seed. The mode and timing of its synthesis during seed development await investigation. A role for the aleurone layer in the production of enzymes for galactomannan breakdown is indicated by ultrastructural observations. The breakdown of galactomannan is first visible next to it, in a “dissolution zone” which increases in size inward toward the cotyledons (Fig. 8). Electron microscopic examination of the aleurone layer provides evidence of intensive synthesis of secretory proteins just prior to and during galactomannan breakdown (Reid and Meier, 1972). The galactomannan of fenugreek constitutes about 30% of the total reserve material in the seed, the remainder being mainly protein and oil localized within

CELL WALL STORAGE CARBOHYDRATES IN SEEDS

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hours

Fig. 9. Dry weight changes accompanying germination of and early seedling development from fenugreek seeds. (Reid and Bewley, 1979.)

the cotyledons (Reid and Bewley, 1979). The overall pattern of reserve mobilization, shown in Fig. 9, illustrates the extraordinary rapidity of the breakdown of the endosperm reserves relative to the cotyledon reserves, The influx of material from the endosperm into the embryo during galactomannan breakdown is so rapid that it causes a transitory increase in the dry weight of the cotyledons. This strongly suggests that the mobilization of the endosperm reserves is not subject to the same regulatory constraints as are the breakdown processes in the embryo. Certainly there is no positive hormonal control by the embryo over the activity of the aleurone layer (Reid and Meier, 1972), as has been established for certain cereal grains (Yomo and Varner, 1971).

2 . Galactomannan Mobilization in the Carob Seed Structurally the galactomannan of carob or locust “bean,” Ceratonia siliqua (Leguminosae-Caesalpinioideae)typifies the low-galactose galactomannans of the Caesalpinioideae. Its degree of galactose substitution is only 20%, whereas that of fenugreek galactomannan is almost 100%.The carob seed is very much larger than that of fenugreek, but its gross anatomy is similar, the galactomannan-rich endosperm completely surrounding the embryo. In the carob seed, however, the endosperm is particularly massive, accounting for about 60%of the dry weight of the seed. The endosperm of the carob seed does not exhibit the same degree of anatomical specialization as that of fenugreek: there is no clear division of it into aleurone layer and storage tissue, and all the cells have living Fig. 8. Top: Cryostat section of part of a fenugreek seed after imbibition but prior to galactomannan mobilization: stained with periodic acid-Schiff‘s reagent to reveal galactomannan. C, Part of a cotyledon; G ,galactomannan; A, Aleurone layer. Note that all the storage cells of the endosperm are “filled” with galactomannan. (After Reid, 1971 .) Midd/e: Section comparable with that above, but during galactomannan mobilization. Note that the dissolution zone (D) in the endosperm is adjacent to the aleurone layer. (After Reid, 1971.) Bottom; Section comparable with those above but after galactomannan mobilization. Only a remnant of the endosperm (E) remains. (After Reid, 1971 .)

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protoplasts. The galactomannan is present in the thickened cell walls. Galactomannan formation in the carob endosperm has not been studied, but its mobilization has been investigated by Seiler (1977). In many respects the physiology of galactomannan utilization in carob resembles that in fenugreek. Again, the enzymes involved are a-galactosidase, pmannanase, and p-mannosidase, and they are present within the endosperm itself. The a-galactosidase at least is synthesized de novo following germination. The pattern of galactornannan mobilization in carob shows an interesting variation from that in fenugreek. Initially some wall dissolution occurs around all the endosperm cells, presumably because they all secrete hydrolytic enzymes. But

Fig. 10. Electron micrograph of the outer part of the carob seed endosperm in the early stages of galactomannan breakdown. AZ, Outer cell layer of endosperm; FS, fibrillar wall layer next to cell lumen-probably not composed of galactomannan; GM, galactomannan. Note the limited digestion of galactomannan (arrows). (After Seiler, 1977.)

CELL WALL STORAGE CARBOHYDRATES IN SEEDS

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this soon ceases (Fig. LO); bulk galactomannan mobilization then takes place from the cotyledons outward toward the testa. It seems probable that enzyme production in the endosperm cells is inhibited by the accumulation of breakdown products, and the inhibition is relieved only as the breakdown products are transported into the cotyledons. 3 . Enzymatic Interactions in Galactomannan Hydrolysis Although it is clear that an a-galactosidase, a (3-mannanase, and a P-mannoside mannohydrolase contribute to galactomannan hydrolysis in fenugreek and carob seeds, none of the fenugreek or carob seed enzymes has been purified to homogeneity. The corresponding enzymes from the guar seed (Cyamopsis fefragonoloba)have, however, been purified and their molecular and catalytic properties have been investigated (McCleary, 1982, 1983). Furthermore, the cooperative interaction of the three enzymes in the hydrolysis of galactomannan has been elegantly demonstrated in vizro using the purified enzymes (McCleary , 1983). To determine the relative importance of these three enzymes in galactomannan hydrolysis and sugar uptake by guar cotyledons, McCleary (1983) incubated washed endosperm-free embryos with galactomannan alone or in the presence of the enzymes singly and in combination. Five embryos were incubated in the presence of a quantity of galactomannan equivalent to that normally present in the endosperms of five guar seeds. If enzymes were added, the amounts were equivalent to the amounts present in’five seeds at the time of the maximum rate of galactomannan breakdown. The galactomannan substrate was guar galactomannan which had been pretreated with a pure fungal 9-mannanase to reduce its viscosity without altering its mannose : galactose ratio. When embryos were incubated in the presence only of galactomannan, and of galactomannan plus the (3-mannoside mannohydrolase, there was no uptake of carbohydrate. Similarly, galactomannan plus P-mannanase gave no uptake of carbohydrate, since the limit galactomannan was resistant to P-mannanase attack. The a-galactosidase gave quantitative removal of the galactose from the galactomannan, and the galactose was rapidly taken up by the cotyledons. When a-galactosidase plus (3mannanase were used, the galactomannan was hydrolyzed to galactose and a series of manno-oligosaccharides, ranging from mannobiose to mannopentaose; the galactose was rapidly absorbed by the embryo, but the uptake of the oligosaccharides occurred more slowly and was incomplete after 24 hr. (In a separate series of experiments it was shown that the cotyledons took up mannobiose and mannotetraose intact, but that higher oligosaccharides were probably hydrolyzed at the surface of the cotyledons by a cotyledonary (3-mannosidemannohydrolase with properties similar to that of the endosperm.) Only when all three enzymes were present was the galactomannan completely hydrolyzed and carbohydrate uptake complete within 48 hr-the time taken in vivo (McClendon et a l . , 1976; McCleary, 1983).

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J. S . GRANT REID B . MANNAN MOBILIZATION IN THE DATE ENDOSPERM

The only mannan-containing seed in which storage carbohydrate mobilization has been studied in detail is that of Phoenix dactylifera, the date palm. In his classic paper of 1862 Sachs describes the anatomy of the date seed, in its resting state and at different times after germination, noting the enormously thick walls of the endosperm cells and their content of aleurone and oil. He describes the germination of the tiny cone-shaped embryo and the mode of its absorption of the endosperm’s reserves. The cotyledon of the embryo acts as a haustorium and grows inward into the endosperm, absorbing the reserves in a narrow zone in front of it. Sachs recognized that the cell walls of the date endosperm constitute a major substrate reserve, and noted that they are completely broken down, with the exception of the thin primary walls which accumulate in the dissolution zone. Sachs’ highly accurate drawings of the date seed and its germination are reproduced unmodified in Fig. 1 1 . The pioneering work of Sachs was confirmed and extended by Keusch (1968) who demonstrated unequivocally that the mannan reserves of the date endosperm are broken down in the dissolution zone surrounding the advancing haustorium, and showed that the end products of the breakdown process are mannose, manno-oligosaccharides and traces of galactose. Using 14C-labeled D-mannose, Keusch was able to demonstrate the uptake of mannose by the haustorium and its rapid conversion to sucrose. The enzymes responsible for mannan hydrolysis were not directly investigated by Keusch (1968); he did however demonstrate that the breakdown of the polysaccharide was hydrolytic rather than phosphorolytic and reasoned that a pmannanase and a (3-mannosidase mannohydrolase had to be involved. We have detected these enzymes in endosperm homogenates (De Mason, Reid, and Sexton, unpublished). The p-mannanase activity is present exclusively in the “dissolution zone” while the p-mannoside mannohydrolase activity is present throughout the endosperm, even prior to germination. The site of production of the P-mannanase is not yet clear. Keusch (1968) incubated isolated, washed haustoria for periods of 15 to 40 hr in solutions containing ivory nut mannan A, 0.01 M Na/K phosphate buffer, toluene, and ammonium molybdate, and observed extensive breakdown of the mannan. The conclusion was drawn that it is the haustorium which secretes the enzymes necessary for mannan breakdown. The endosperm cells of the date are living (DeMason, Reid, and Sexton; unpublished), and there is no reason why they should not be capable of synthesizing and/or secreting p-mannanase. The question of the origin of the hydrolytic enzymes is receiving further attention. C. XYLOGLUCAN METABOLISM IN TROPAEOLUM

Apart from the early reports of Heinricher (1888), Reiss (1889), and others that ‘‘amyloids” in a variety of seeds including Impatiens balsamina, Tropaeolum

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Fig. 11. Reproduction of Sachs’ (1862) line drawings of date seed germination. Drawing No. 4 shows the anatomy of the endosperm (E) during reserve mobilization. p, Primary cell wall; WS, weakened zone; PZ, residual compressed primary walls; Ep, epithelium of haustorium.

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majus, and Cyclamen europaeum are mobilized following germination, there is little information in the literature relevant to the biochemistry of the breakdown of seed xyloglucans. Gould et al. (1971) observed that a “pectic” xyloglucan is broken down following germination in mustard seed cotyledons; xyloglucan, however, is not a major storage material in mustard. We are investigating xyloglucan mobilization in the seeds of Tropaeolum majus, and have established that the cell walls of the cotyledons lose their ability to undergo “amyloid” staining following germination (Ward, Reid, and Sexton, unpublished). Furthermore, the cotyledons of the germinated seeds develop xyloglucan-degrading enzymatic activities which vary par1 passu with the breakdown of the polysaccharide. These enzymes include endoxyloglucanase, f3-galactosidase, a-xylosidase, and P-glucosidase. The endoxyloglucanase has been purified to homogeneity (Edwards et at., 1984). D. “GALACTAN” METABOLISM IN LUPINUS

The physiological role of the cotyledon cell walls of lupin seeds was a subject of scientific controversy in the late nineteenth century! Nadelmann (1890), on the basis of microscopic observations, concluded that the thickened cell walls in the cotyledons of L. angustifolius, L. albus, and L. luteus were mobilized following germination and were substrate reserves. Elfert (1894) vigorously refuted this claim, asserting that Nadelmann’s observations were erroneous; he concluded that the changes in wall morphology which followed germination in Lupinus were simply a “metamorphosis” of the wall brought about by cotyledon expansion. Schulze (1895-1896) pointed out that neither Nadelmann nor Elfert appeared to be aware of an earlier paper (Schulze and Steiger, 1889) in which it had been shown that the seed of L . luteus contained a water- and alkali-insoluble carbohydrate material (“Paragalaktan”) which was probably present in the cotyledon cell walls and which was mobilized following germination. The observations were later extended to L . angustifolius (Schulze, 1895-1896). “Paragalaktan,” which released galactose and arabinose on acid hydrolysis, was, according to Schulze, clearly a storage material. In more recent times the ideas of both Elfert and of Nadelmann and Schulze have received support. Matheson and Saini (1977) conducted an investigation of the polysaccharides in the cotyledons of L. luteus following germination, with particular attention to the “pectic” fractions which were soluble in hot water and oxalate/EDTA solutions. They noted a net depletion of galactose- and arabinosecontaining polysaccharides, and concluded that the later stages of cotyledon expansion were accompanied by the selective hydrolysis of certain wall polymers. Although Matheson and Saini (1977) noted that depletion of wall polysaccharide was accompanied by the transitory accumulation of a glucan which they later indicated was starchlike (Saini and Matheson, 1981), they do not seem to have considered that the galactose- and arabinose-containingpolysaccharides of

-i 15

P

*;=EL.

15

-

10-

5-

\\

Nonstarch polysaccharides

\\

o Galactose residues A

Arabinose residues

o Xylose residues

Y\

'--4=-f:,y\ -7

- e- _ - _m-4%: -~

Fig. 12. Mobilization of major stored reserves in cotyledons of Lupinus angusrifolius cv. Unicrop. Open symbols, day/night conditions; filled symbols, continuous darkness. (After Crawshaw and Reid, 1984.)

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TABLE I1 Monosaccharide Residues in Storage Mesophyll Cell Walls Isolated from Cotyledons of Lupinus angustifolius cv Unicrop before and after Reserve Mobilization“ Neutral monosaccharide residues Arabinose

Xylose

Galactose

Glucose

Rhamnose

Uronic acid

Walls from 16hr-imbibed seeds6

12

15

66

4

2

8

Walls from 14day-germinated seedsc

16

20

30

23

11

35

a

Hutcheon and Reid (unpublished). Before mobilization of cell wall storage polysaccharides. After mobilization of cell wall storage polysaccharides.

L. luteus might be reserves. Evidence in support of a reserve function for the cell wall polysaccharides of Lupinus has come from Parker’s (1 976) observation that the cell wall thickenings in L . albus and L. angustifolius cotyledons disappear following germination and from cognate biochemical studies being carried out in the author’s laboratory. Fig. 12 shows the changes in the major stored reserves in the cotyledons of L. angustifofius following germination, while Table I1 shows the composition of storage mesophyll cell walls isolated from the cotyledons before and after reserve mobilization. Clearly nonstarch carbohydrates localized in the cell wall constitute a major reserve in L. angustifolius. The linkages present in the cell walls and the enzymes responsible for their hydrolysis are currently under investigation. IV. CONSIDERATIONS OF BIOLOGICAL FUNCTION Although the cell wall storage carbohydrates of seeds are utilized as substrate reserves following germination it is nevertheless pertinent to raise the question of their overall biological function in the seeds which contain them. Are they exclusively storage macromolecules, or do they have other functions? This question is by no means original. It was effectively posed by Nadelmann (1890) who set out to investigate whether or not the mucilages present in leguminous seed endosperms were reserve substances in addition to being involved in water imbibition. Nadelmann went on to demonstrate that the mucilages have a reserve function and concluded that they are first and foremost (“in erster Linie”) reserves. Marloth (1883) commented on the possibility that a “hard endosperm” with thick cell walls serves to protect seeds from mechanical damage. Gould et al. (197 1) have also suggested a protective function for storage polysaccharides

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of the celI wall type. Unlike starch, which is stored intracellularly, the cell wall storage carbohydrates are interposed between living cells and the external environment. It is perhaps not unreasonable to expect that they might play a direct role in the seeds’ interaction with that environment. It is the author’s personal opinion (a physical scientist’s view, perhaps) that the most telling argument in favor of a nonstorage role for the cell wall storage carbohydrates of seeds can be based on their bulk properties as materials. Mannans and glucomannans are crystalline, insoluble materials which confer extraordinary hardness on seeds which contain them in their endosperms. On the other hand the galactomannans and xyloglucans are hard only in the unimbibed state. They are essentially hydrophilic molecules. As materials these polysaccharides, particularly the galactomannans, are commercially important because of their complex interactions with water (Dea and Momson, 1975). It was the naive(?) expectation that these same interactions might be important to the germinative strategy of seeds which prompted us to investigate the role of galactomannan in the water relations of fenugreek seed during germination (Reid and Bewley , 1979). Figure 13 shows the movement of water which accompanies the imbibition of the fenugreek seed. Clearly the seed as a whole exhibits a normal water uptake curve-initial hydration of the tissue, followed by a lag phase culminating in the completion of germination and a further uptake of water. Analysis of the individual contributions of the endosperm, cotyledons, and radicle, however, shows

hours

Fig. 13. Pattern of water uptake by fenugreek seeds. Dry seeds were placed on wet cotton at cotyledons; W-W, axis. time = 0. 0-0,Whole seed; 0-0, endosperm and testa; 0-0, Quadruplicate batches of six seeds were analyzed: error bars represent 2 X SEM; G , completion of germination (radicle breakthrough). (After Reid and Bewley, 1979.)

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J. S . GRANT REID

I

I

I

L

U

I

1

f

I

-.\a' -o-n

I

drying time (hours)

and naked Fig. 14. Drying curves for whole fenugreek seeds (O-O), testa-free seeds (0-O), embryos (m-m) at 52% relative humidity. Triplicate batches of 20 seeds were processed. (After Reid and Bewley, 1979.)

that the uptake of water in the initial hydration phase is predominantly into the endosperm: it represents 30% of the dry weight of the seed, but takes up over 60% of the water. Nadelmann's (1890) premise that the leguminous seed endosperm imbibes water is correct! Figures 14 and 15 show the analogous changes in water content which occur when a fully imbibed seed is allowed to dehydrate in a drying atmosphere: the seed as a whole loses water more or less linearly with time (Fig. 14) but the same is not true of the individual tissues (Fig. 15). Water is initially lost only from the

2LO -

0

drying time (hard

cotyledons

d r y ' q time (hars)

Fig. 15. Patterns of water loss from imbibed, whole fenugreek seeds (A) testa-free seeds (B),and naked embryos (C) at 52% relative humidity. Data derived from single batches of 10 seeds, testa-free seeds, or naked embryos, dissected and processed at each time point. (After Reid and Bewley, 1979.)

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endosperm: the cotyledons and axis lose no water until several hours after the initiation of the drying process. In the absence of the endosperm the cotyledons and axis lose water immediately (Fig. 15). The physical basis of the ability of the endosperm of the fenugreek seed to protect the embryo against desiccation can be deduced from Fig. 16, in which the water potential ($,) of the endosperm is shown in relation to water loss from the endosperm and the embryo. Clearly the endosperm is capable of losing water with little change in its water potential, until its water content falls to 100% of its dry weight. Thereafter further water loss is accompanied by a rapid change in water potential. The embryo does not lose water initially because it is not directly subjected to the very low water potentials of the surrounding atmosphere. It is challenged only by the water potential of the endosperm which for a period of some hours remains above - 1.5 MPa, a value which living, turgid plant tissues can resist (Wiebe, 1966). The endosperm’s hydrodynamic properties can be directly attributed to its high content of galactomannan [cf. Fig. 8 (top) and compare Figs. 16 and 17). The biological ‘role of the galactomannan of the fenugreek seed is therefore complex. During imbibition it is responsible for the uptake of relatively large amounts of water, and its distribution around the embryo. During germination it effectively buffers the embryo against water loss. Following germination it acts as a substrate reserve for the developing seedling. To assess the relative importance of the storage and nonstorage functions of the fenugreek and galactomannan, it would be necessary to study the seed’s overall germinative strategy in its natural environment. Nevertheless, it is interesting to note that there is no purely nutritional reason for a proportion of the substrate reserves of the fenugreek seed to take the form of galactomannan (Reid and Bewley, 1979). In contrast, a clear structure-function relationship exists for the role of galactomannan in the water relations of germination.

d r y q time Iholas) Fig. 16. Water loss at 52% relative humidity from the endosperm testa (W-M) and from the embryo (0-0) of the hydrated fenugreek seed in relation to the water potential of the endospermltesta (0-0). At each time point six seeds were used for dry weight determination and four seeds for the determination of JI, of the endosperm. (After Reid and Bewley, 1979.)

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J. S. GRANT REID

drying time (hours) Fig. 17. Water content (0-0) and water potential (0-0) of a purified sample of fenugreek seed galactomannan (Campbell, 1978) hydrated and then allowed to dry at 25% relative humidity. (After Reid and Bewley, 1979.)

V. PERSPECTIVES There is now enough interest in the physiology and biochemistry of the cell wall storage carbohydrates of seeds to allow the confident prediction that the enzymatic mechanisms of their breakdown will soon be understood with respect to the major seed systems. It is to be hoped that the cell wall-degrading enzymes themselves will be purified to homogeneity and their specificities studied in detail, as is already happening in the galactomannan field. Germinated seeds could provide a convenient source of moderate quantities of highly purified enzymes capable of specifically degrading complex plant cell wall polysaccharides. Such enzymes would be of incalculable value in probing the structures of native cell walls, in determining the fine structures of cell wall polysaccharide preparations and in positively identifying products of polysaccharide biosynthesis in vitro. There is, for example, a striking structural resemblance between the xyloglucan storage carbohydrates of seeds and the xyloglucans which are now assumed to comprise the main noncellulose component of primary cell walls in dicotyledonous plants (Albersheim, 1976). Similarly there are indications (Wilkie, Reid, and Hutcheon, unpublished) that the “galactan” complex of lupin cell walls is structurally related to the “pectic” galactan-rhamnogalacturonan complex of primary cell walls (Albersheim, 1976). Xyloglucan- and galactan-containing seeds could provide enzymes to investigate primary cell wall structure, and will certainly provide insight into the types of enzymes which bring about the modification or turnover of primary cell walls which accompanies plant cell growth.

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The enzymology of cell wall storage polysaccharide biosynthesis has received relatively little attention. With the exception of the GDPmannose : galactomannan mannosyltransferase of the developing fenugreek seed endosperm (Campbell and Reid, 1982) there is no information concerning the properties of the enzymes catalyzing the formation of the major cell wall storage carbohydrates. To acquire such information should be relatively straightforward. Over a defined period of time virtually all the cells of developing endosperms of cotyledons are dedicated to the production of a single type of cell wall polysaccharide; levels of the relevant biosynthetic enzymes should therefore be relatively high. Furthermore, the structural features of synthetic products formed in virro should be easily compared with those of the carbohydrates known to be present in vivo. Once acquired such information would be relevant not only to seed systems but also to our understanding of the biosynthesis of analogous noncellulose polysaccharides in vegetative tissues. Perhaps the most interesting question surrounding the cell wall storage carbohydrates is that of their overall biological function. Do they all, like the galactomannan of fenugreek (Reid and Bewley, 1979), have a nonstorage function which is dependent upon their physicochemical properties? The answer will be provided only by the more widespread adoption of a combined biochemical and ecological approach to the study of seed “storage’ ’ carbohydrates. ACKNOWLEDGMENTS The financial support of the Agricultural Research Council and of Unilever Ltd. is gratefully acknowledged.

REFERENCES Albersheim, P. (1976). In “Plant Biochemistry” (J. Bonner and J. E. Vamer, eds.), 3rd ed., pp. 225-274. Academic Press, New York. Albersheim, P., Nevins. D. J.. English, P. D., and Karr, A. (1967). Curbohydr. Res. 5, 340-345. Anderson, E. (1949) Ind. Eng. Chem. 41, 2887-2890. Andrews, P., Hough, L., and Jones, J. K. N. (1953). J . Chem. Soc. 1186-1192. Aspinall, G. 0.. Hirst, E. L., Percival, E. G. V., and Williamson, I. R. (1953). J . Chem. Soc. 3 184-3 188. Aspinall, G. 0 . . Rashbrook, R. B., and Kessler, G . (1958). J . Chem. Soc. 215-221. Aspinall, G. 0.. Greenwood, C. T., and Sturgeon, R. J. (1961). J . Chem. SOC.3667-3674. Balasubramaniam, K. (1976). J. Food Sci. 4, 1370-1373. Bourquelot, E., and Herissey, H. (1899). C. R. Hebd. Seances Acad. Fr. 129, 228. Bouveng, H. O., Garegg, P. J . , and Lindberg, B. (1960). Acta Chem. Scand. 14, 742-748. Campbell, J. (1978). Ph.D. thesis, University of Stirling, Scotland. Campbell, J., and Reid, J. S. G . (1982). Planta 155, 105-1 1 I . Carlson. W. S., Ziegenfuss, E. M . , and Overton, J. D. (1962). Food Techno/. 16, 50-54. Chudzikowski, R. J. (1971). J. Soc. Cosrner. Chem. 22, 43-60. Courtois, J.-E., and Le Dizet, P. (1974). C. R . Hebd. Seances Acud. Fr. Ser. C 278, 81-83. Courtois, J.-E., Le Dizet, P., and Robic, D. (1976). Curbohydr. Res. 49, 439-449.

154

J. S. GRANT REID

Crawshaw, L., and Reid, J. S . G. (1984). Plunta 160, 449-454. Dea, 1. C. M., and Morrison, A. (1975). Adv. Curbohydr. Chern. Biochem. 31, 241-312. Edwards, M., Dea, I. C. M., Bulpin, P. V., and Reid, J. S. G. (1984). Plunra (in press). Elfert. T. (1894). Bibl. Bor. 30, 1-25. El Khadem, H., and Sallam, M. A. E. (1967). Curbohydr. Res. 4, 387-391. Glicksman, M. (1953). “Gum Technology in the Food Industry.” Academic Press, New York. Godfrin, M. J . (1884). Ann. Sci. Nut. 19, 5-158. Goldberg, R. (1969). Phyrochemisrry 8, 1783-1792. Could, S. E. B, Rees, D. A,, and Wight, N. J. (1971). Biochem. J. 124, 47-53. Halmer, P., and Bewley, J. D. (1979). Plunta 144, 333-340. Halmer, P., Bewley, J. D., and Thorpe, T. (1978). Plunta 139, 1-8. Hegnauer, R. (1973). “Chemotaxonomie der Pflanzen,” Vol. 6. Birkhauser, Basel. Heinricher, E. (1888). Flora (Jenu) 71, 163-185. Herissey, H. (1903). Rev. Cen. Bor. 15, 345-392, 406-417, 444-464. Hirst, E. L., Jones, J. K. N., and Walder, W. 0. (1947). J . Chem. Soc. 1225-1229. Hopf, H., and Kandler, 0. (1977). Phytochemistry 16, 1715-1717. Homer, H. T., and Amott, H. J. (1966). Bor. Gar. 127, 48-64. Jakimow-Bmas, N. (1973). Phytochemistry 12, 1331-1339. Jones, R. L. (1974). Planfa 121, 131-146. Keusch, L. (1968). Plunru 78, 321-350. Khanna, S. N., and Gupta, P. C. (1967). Phytochemistry 6, 605-609. Klages, F. (1934). Ann. Chem. 509, 159-181; 512, 185-194. Kooiman, P. (1960a). Acra Bor. Neerl. 9, 208-219. Kooiman, P. (1960b). K. Ned. Akud. Wet. C 63, 634-645. Kooiman, P. (1960~).Rec. Trav. Chim. Pays-Bas 79, 675-678. Kooiman, P. (1961). Rec. Trav. Chim. Pays-Bus 80, 849-865. Kooiman, P. (1967). Phytochemistry 6, 1665-1673. Kooiman. P. (1971). Curbohydr. Res. 20, 329-337. Kovacs, P. (1973). Food Technol. 27, 26-30. Le Dizet. P. (1972). Curbohydr. Res. 24, 505-509. Ludtke, M. (1927). Ann. Chem. 456, 201-224. McCleary, B. V. (1982). Curbohydr. Res. 101, 75-92. McCleary, B. V. (1983). Phytochernistry 22, 649-658. McClendon, J. H., Nolan, W. G., and Wenzler, H. F. (1976). Am. J. Bor. 63, 790-797. Marloth, R. (1883). Bor. Juhrb. Sysr. Pflanzengesch. 4, 225-265. Matheson, N. K., and Saini, H. S. (1977). Phytochemistry 16, 59-66. Meier, H. (1958). Biochim. Biophys. Acru 28, 229-240. Meier, H., and Reid, J . S. G. (1977). Plunta 133, 234-248. Meier, H., and Reid, J. S. G. (1982). Encycl. Planr Physiof. New Ser. 13A, 418-471. Mukhejee, A. K., and Rao, C. V. N. (1962). J . fndiun Chem. Soc. 10, 687-692. Mukherjee, A. K., Choudhury, D., and Bagchi, P. (1961). Can. J. Chem. 39, 1408-1418. Nadelmann, H. (1890). Juhrb. Wiss. Bor. 21, 1-83. Numberg, E., and Rettig, E. (1974). Drugs Made Ger. 17, 26-31. Parker, M. L. (1976). Ph.D. thesis, University of Wales, Bangor. Reese, E. T., and Shibata, Y. (1965). Can. J . Microbiol. 11, 167-183. Reid, J. S. G. (1971). Planta 100, 131-142. Reid, 1. S. G., and Bewley, J. D. (1979). Plunra 147, 145-150. Reid, J. S . G., and Meier, H. (1970). 2. Pji’anzenphysiol. 62, 89-92. Reid, J . S. G . , and Meier, H. (1972). Pluntu 106, 44-60. Reid, J. S. G., and Meier, H. (1973a). CuryotogiuSuppl. 25, 219-222. Reid, J. S. G., and Meier, H. (1973b). Planfa 112, 301-308.

CELL WALL STORAGE CARBOHYDRATES IN SEEDS

155

Reid, J. S. G., Davies, C., and Meier, H. (1977). Plunra 133, 219-222. Reiss, R. (1889). Landwirrsch. Jahrb. 18, 71 1-765. Robic, D.. and Percheron, F. (1973). Phytochemisfry 12, 1369-1372. Sachs, J. (1862). Bor. Zrg. 20, 241-246, 250-252. Saini, H. S., and Matheson, N. K. (1981). Phytochemistry 20, 64-646. Saxena, V. K. (1965). Res. I d , 10, 101-106. Schulze, E. (1895-96). Hoppe-Seyler’s 2. Physiol. Chem. 21, 392-41 1 . Schulze, E., and Steiger, E. (1889). Landwirtsch. Versuchs-Stn. 36, 391-476. Seiler, A. (1977). Planra 134, 209-221. Smith, F., and Montgomery, R. (1959). “Chemistry of Plant Gums and Mucilages.” Van NostrandReinhold, Princeton, New Jersey. Stepanenko, B. N. (1960). Bull. SOC. Chim. Biol. 42, 1519-1536. Takaki, M., and Dietrich, S. M. C. (1979). Rev. Bras. Bot. 2, 125-127. Thompson, J. L., and Jones, J. K. N. (1964). Can. J . Chem. 42, 1088-1091. Tschirch, A. (1889). “Angewandte Pflanzenanatomie.” Urban & Schwarzenberg, Vienna. Vogel, T., and Schleiden, M. J. (1839). Poggendofs Ann. Phys. Chem. 327-330. Whistler, R. L., and Bemiller, I. N. (1958). Adv. Carbohydr. Chem. 13, 289-329. Whistler, R. L., and Bemiller, J. N. (1972). “Methods Carbohydr. Chem. Vol. 8.” Academic Press, New York, London. Whistler, R. L., and Smart, C. L. (1953). “Polysaccharide Chemistry”, Academic Press, New York. Whistler, R. L., and Wolfrom, M. L. (1965). Methods Carbohydr. Chem. 5. Wiebe, H. H. (1966). Plant Physiol. 41, 1439-1442. Winterstein, E. (1893). Hoppe-Seyler’s 2. Physiol. Chem. 17, 353-380. Wolfrom, M. L., and Patin, D. L. (1965). J. Org. Chem. 30, 4060-4063. Wolfrom, M. L., Laver, M. L., and Patin, D. L. (1961). J. Org. Chem. 26, 4533-4535. Yomo, H . , and Varner, J. E. (1971). Curr. Top. Dev. Biol. 6, 111-144.