Annu. Rev. Plant Physiol. Plant Mol. Biol. 1996. 47:445­76 Copyright © 1996 by Annual Reviews Inc. All rights reserved

Annu. Rev. Plant. Physiol. Plant. Mol. Biol. 1996.47:445-476. Downloaded from by University of Georgia on 04/26/06. For personal use only.


Nicholas C. Carpita

Department of Botany and Plant Pathology, Purdue University, West Lafayette, Indiana 47907

KEY WORDS: cereals, grasses, cell-wall polysaccharides, cell-wall biosynthesis, cell-wall architecture


The chemical structures of the primary cell walls of the grasses and their progenitors differ from those of all other flowering plant species. They vary in the complex glycans that interlace and cross-link the cellulose microfibrils to form a strong framework, in the nature of the gel matrix surrounding this framework, and in the types of aromatic substances and structural proteins that covalently cross-link the primary and secondary walls and lock cells into shape. This review focuses on the chemistry of the unique polysaccharides, aromatic substances, and proteins of the grasses and how these structural elements are synthesized and assembled into dynamic and functional cell walls. Despite wide differences in wall composition, the developmental physiology of grasses is similar to that of all flowering plants. Grass cells respond similarly to environmental cues and growth regulators, exhibit the same alterations in physical properties of the wall to allow cell growth, and possess similar patterns of wall biogenesis during the development of specific cell and tissue types. Possible unifying mechanisms of growth are suggested to explain how grasses perform the same wall functions as other plants but with different constituents and architecture.


INTRODUCTION ......................................... .......................................................................... THE STRUCTURAL ELEMENTS OF THE PRIMARY WALLS OF THE POACEAE ...... Cellulose .................................................. .......................................................................... Glucuronoarabinoxylans ......................... .......................................................................... The (13),(14)--D-glucans................ .......................................................................... 446 447 447 447 448





450 450 450 451 453 454

Annu. Rev. Plant. Physiol. Plant. Mol. Biol. 1996.47:445-476. Downloaded from by University of Georgia on 04/26/06. For personal use only.

454 ARCHITECTURE ......................................... .......................................................................... 458 STRUCTURAL DYNAMICS DURING CELL ELONGATION ........................................... 462 CELL-WALL BIOGENESIS IN GRASSES. .......................................................................... 465 Biosynthesis of (13),(14)-ß-D-Glucan ......................................................................... 466 Biosynthesis of Other Cell-Wall Polysaccharides .............................................................. 467 GENETIC MODELS OF CELL-WALL DEVELOPMENT IN THE GRASSES................... 467

Xyloglucan ............................................... .......................................................................... Other Glycans .......................................... .......................................................................... Pectic Substances..................................... .......................................................................... Aromatic Substances................................ .......................................................................... Structural Proteins................................... .......................................................................... Other Cell-Wall Substances..................... .......................................................................... CELL-WALL COMPOSITION AS A TAXONOMIC CHARACTER IN THE MONOCOTYLEDONAE..................... ..........................................................................


In the early 1900s, Walter Norman Haworth and Edmund Langley Hirst, founding fathers of modern carbohydrate chemistry, began studies of the pentose constituents of cell walls of plants. Armed with only rudimentary analytical techniques, they and their colleagues defined the cell walls of esparto grass as composed largely of (14)--D-xylans (58). By 1970, gas chromatography­mass spectrometry (GC-MS) was employed routinely for unequivocal determination of linkage structure of complex cell-wall polysaccharides. Techniques such as 1H- and 13C-nuclear magnetic resonance (NMR) spectroscopy provided anomeric configurations, linkage structures, and some three-dimensional configurations. Sequence-dependent endoglycanases were used to cleave polysaccharides into oligosaccharides that could be completely sequenced. From characteristic repeating unit structures, the sequence and conformation of very large polymers were deduced (20). By such analyses, the major polysaccharides of the walls of a wide range of flowering plants were defined, and the first models of how cell walls are put together emerged. In subsequent years, the dynamic interactions of individual components were reflected in more current models of the architecture of the primary cell wall of flowering plants--a strong framework of cellulose microfibrils intertwined with xyloglucans that is embedded in a gel of uronic-acid-rich pectins and cross-linked with hydroxyproline-rich glycoproteins (20, 101). When the first conceptual models were proposed about twenty years ago, the differences in wall compositions between monocots and dicots were just beginning to be catalogued (33). Perhaps because of the socioeconomic importance of the cereals, the vast majority of the monocots studied were grasses. Whistler (164) described grasses as rich sources of xylan, and Aspinall (1), in a review of plant cross-linking glycans, noted the enrichment of xylans and mixed-linked glucans in grasses. Wilkie (165) offered the first comprehensive survey of the cross-linking glycans of grasses. More recent



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studies of the carbohydrate and aromatic components of cell walls from a broad spectrum of monocots have revealed that the Poales (family Poaceae, formerly the order Graminales, family Gramineae), their progenitors, and related taxonomic orders have primary cell walls completely different from those of other monocots (3, 20, 68). Dahlgren et al (32) proposed phyletical relationships between some two dozen orders of the Monocotyledonae on the basis of several anatomical features and chemical constituents. One of these features, the presence or absence of ferulic acid in the primary walls, is a major distinguishing feature of the Poales and related orders (56). Nonlignified cells of grasses are enriched in aromatic substances, and polymeric forms constitute a second architectural element. A third type of architectural element is structural protein. The primary walls of the Poales contain substantially less protein than other species, but several classes of proteins are found in elevated amounts in specific cell types during differentiation. Recent reviews (83, 147) note that the grass wall proteins bear reasonable homology to those representing major classes of structural proteins of nongramineous species. In an earlier review (20), structural models for two types of primary walls were provided: the Type I wall, composed of a cellulose-xyloglucan framework embedded in a pectin gel, and the Type II wall, the special wall of the Poales. This review focuses on the Type II wall of the Poales, its composition, architecture, biogenesis, and dynamics during growth.


Cellulose microfibrils in all flowering plants are composed of about three dozen linear chains of (14)--linked D-Glc condensed to form long paracrystalline arrays that spool around each cell (35). Although each chain may be only several thousand units long, they begin and end at different places within a microfibril and make very long microfibrils whose ends are rarely detected.


The linkage structure of the grass glucuronoarabinoxylans (GAX) has been known for much of the 20th century. The t--L-arabinofuranosyl units are attached primarily at the O-3 positions along the (14)--D-xylan backbone, and the t--D-glucuronic acids are attached to the O-2 positions (1, 165). The highly substituted GAXs of the barley aleurone and barley malts contain significant O-2- and doubly branched O-2, O-3-linked arabinosyl units in addition to the abundant O-3-linked units (5, 160). Arabinoxylans are widespread in the walls of all flowering plants, but in nongramineous species the polymer



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is of much lower abundance, and the -L-arabinosyl units are attached mostly at the O-2 rather than the O-3 of the xylosyl units (33). Whereas some of the xylem-rich straws may contain exclusively the 4-O-methyl derivative (1, 165), underivatized GlcA is the major acidic constituent of the maize primary wall GAXs (22). A highly substituted GAX (HS-GAX), with six of seven xylosyl units bearing appendant groups, is associated with the maximum growth rate of coleoptiles (22). Smith degradation indicated a unit structure in which the arabinosyl and GlcA units are added to the growing chain in a specific pattern during synthesis (22). Nishitani & Nevins (117) found a sequence-dependent xylanase that requires an appendant GlcA to cleave the neighboring (14)-D-xylosyl linkage. When maize GAXs are depleted of arabinosyl units by mild-acid hydrolysis, this endo--D-xylanase releases a homogeneous group of deca- or undecamers of glucuronoxylan. The molecular structure of the HS-GAX may be a unit structure with GlcA units added to alternate xylan heptamer units (Figure 1A). The arabinosyl units are hydrolyzed after incorporation into the wall to yield the broad range of grass GAXs (46). The xylosyl units are also substituted with acetyl groups at the O-2 and O-3 position (30); the acetyl content of some ryegrasses has been reported to be almost 10% of the dry mass of the wall (52).

The (1®3),(1®4)-b-D-glucans

Noncellulosic glucans are also found at certain stages in grass development, particularly in the seed brans. These unbranched "mixed-linked" glucans (D-glucans) contain both (13) and (14)-linkages in a ratio of about 1:2 to 1:3. A general sequence structure was unequivocally deduced with a sequence-dependent endoglycanase, a -D-glucanohydrolase from Bacillus subtilis that catalyzes the hydrolysis of a (14)--D-glucosyl linkage only if preceded by a (13)--D-linked glucosyl unit on the nonreducing side (151). With oat or barley endosperm walls, this enzyme hydrolyzes about 90% of the -D-glucan macromolecules into cellobiosyl- and cellotriosyl-(13)-D-Glc in a ratio of a little over 2:1 (Figure 1B) (151). The remainder of the polymer is made of small amounts of longer runs of the cellodextrin series inFigure 1 (A) (Feruloylated) glucuronoarabinoxylan (GAX). The highly substituted GAX contains -L-arabinose and -D-glucuronic acid units on six of every seven (14)--D-linked xylosyl units (22). Many of the arabinosyl units are cleaved from the GAX during assembly in the wall (46). Ferulic acid esters are attached at the O-5 position of a small portion of the arabinosyl units, and these esters may dimerize with other feruloyl groups in several ways to cross-link the GAX into a network (126). (B) Trimer and tetramer unit structure of the mixed-linkage (13),(14)--D-glucans. The Bacillus subtilis endo-glucanase cleaves (14)--D-glucosyl linkages just in front of (13) linkages (arrows) to yield cellobiosyl- and cellotriosyl-(13)--D-glucose oligomers in a ratio of about 2:1 (151).

Annu. Rev. Plant. Physiol. Plant. Mol. Biol. 1996.47:445-476. Downloaded from by University of Georgia on 04/26/06. For personal use only.




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terrupted by single (13)-linked units (80, 169) These longer runs of cellodextrin units are apparently uniformly distributed throughout the length of the polymer. An endo--D-glucanase from cell walls of developing maize seedlings, which can hydrolyze the -D-glucan macromolecule only at the cellodextrin-rich regions and not the tri- and tetramers, yields a homogeneous fraction of polymers about 50 sugar units long (57, 67). The distribution of sizes of the cellodextrin lengths larger than four are remarkably constant among the cereals (167).


Small amounts of xyloglucan (XyG) are found in the grasses, but hydrolysis with 4-glucanohydrolases does not yield the hepta- and nonasaccharides characteristic of the polymer in many Type I walls. The XyGs are enriched in grass meristematic cells before the onset of enhanced -D-glucan and GAX synthesis during elongation. Enzyme hydrolysis yields the diagnostic disaccharide isoprimeverose, the -D-Xyl-(16)-D-Glc, but the xylosyl units appear on isolated or two contiguously linked glucosyl units of the (14)linked glucan backbone instead of the regular blocks of three typically found in many flowering plants (78). An exception may be the XyG of rice endosperm walls. Shibuya & Misaki (145) found XyG fragments containing both two and three contiguous xylosyl side-groups and a possible attachment of t-Gal at the O-2 of some of the xylosyl units. The common Type I XyG trisaccharide, t-L-Fuc-(12)--D-Gal-(12)--D-Xyl-, is absent from the grass XyGs. Consistent with this lack of Fuc and Gal in grass XyG, monoclonal antibodies that recognize this feature of Type I XyGs fail to recognize grass XyG (123). Surprisingly, Fuc is assimilated by suspension cultures of fescue and added to the occasional Gal unit in grass XyG (103).

Other Glycans

Small amounts of glucomannan are also found tightly bound to the cellulose microfibrils. Urea is able to extract selectively a glucomannan from the walls of wheat and barley endosperm walls (5) and the maize coleoptile (15). Although grasses are generally devoid of Fuc, the root cap slime can be considerably enriched in this sugar (24). The maize slime is almost 20% Fuc, in mostly t- and 3-linkages, whereas the remainder is pectic-like (4).

Pectic Substances

Two fundamental constituents of all flowering plant pectins are polygalacturonic acid (PGA), which is a homopolymer of (14)--D-galactosyluronic acid (GalA), and rhamnogalacturonan I (RG I), which is a heteropolymer of repeating (12)--L-rhamnosyl-(14)--D-GalA disaccharide units (73). RG I is found in walls of somatic cells of both maize and rice (156). PGA and RG I are found in grasses but in much smaller amounts (17, 146).



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Like many flowering plants, grasses contain arabinans, galactans, and highly branched arabinogalactans (AGs) of various configurations and sizes, and they are attached to the O-4 of the rhamnosyl residues of RG (17, 146). The arabinans are mostly 5-linked arabinofuranosyl units but can be connected to one another at virtually every free position, the O-2, O-3, and the O-5, to form a diverse group of branched arabinans. The galactans and two classes of AGs are the major side-chains. One class of AGs is associated only with pectins and is composed of (14)--D-galactan chains with mostly t-arabinosyl, and sometimes t-galactosyl, units at the O-3 of the galactosyl residues of the backbone (3). In maize and rice pectins, the branched 5-linked arabinan and 4-galactan side-chains are attached to the O-4 of about two of every three rhamnosyl units (17, 146, 156). The neutral sugar side-chains of the grass RGs are notable only for their lack of Fuc (156). In walls from maize seedlings and rice endosperm, the chelator-soluble pectins are enriched in the HS-GAX as well as RG and arabinogalactan (protein) (AGP). Two fractions of grass pectins are resolved by ion-exchange chromatography (17, 146); one fraction is enriched in HS-GAX and an associated AGP, whereas the other contains mostly GAX and RG I (17). The second class of AGs constitutes a broad group of short (13)- and (16)--D-galactan chains connected to each other by (13,16)-linked branch point residues. Most of the remaining O-3 or O-6 galactosyl positions are filled with t-arabinosyl groups (2, 40). These AGs are also associated with proteins (AGPs) whose functions in intracellular, plasma membrane, and cell-wall locations are still unknown.

Aromatic Substances

A major feature of the Poales and their relatives is the enrichment of aromatic substances in nonlignified walls (56, 135). A large portion of the aromatic substances are esters of the hydroxycinnamates, ferulate, and p-coumarate (56). The GAXs are cross-linked in walls by both esterified and etherified hydroxycinnamates and by other phenolic substances (68, 138), and the etherified linkages represent complexes of polysaccharide and lignin (69). The ferulate and p-coumarate esters are attached to the O-5 of arabinosyl units of GAX (54, 79, 114). Markwalder & Neukom (100) suggested that neighboring feruloylated GAX chains are cross-linked by formation of 5,5-diferulate, and this was demonstrated directly in bamboo GAX by Ishii (72). The 5,5diferulate is only one of a series of dehydrodimers present in grasses (126). These esters are broken by dilute alkali, and release of ferulic acid and diferulic acid is coincident with the release of HS-GAX (16). Polysaccharides may also be cross-linked photochemically by [2 + 2]-homo- and -heterodimerization of the ferulate and p-coumarate esters to cyclobutane derivatives called truxillic and truxinic acids (55, 157).



More complex interactions between aromatic substances and polysaccharides involve ester-ether interactions, and the ether bonds are not broken by dilute alkali (70). Scalbert et al (138) suggested that such ester-ether interactions form bridges between polysaccharides and lignin via phenol addition to quinone methide lignin intermediates in several ways. Ferulic acid is the principal component in the ester-ether linkage of carbohydrate and lignin (69, 91, 92). Ferulates are also incorporated into lignins via radical mechanisms to form not only the -ether linkages but other structural forms that cannot be released by any solvolytic method (125). p-Coumaric and ferulic acids form single esters, but only rarely does the p-coumaric acid form an ester-ether bridge (91, 92). The p-coumaric acid is more heavily associated with lignin, particularly later in cell-wall development, and attached at the -positions of the lignin side-chains (124). The principal monomers of grass lignin are coniferyl and sinapyl alcohols, with some p-OH coumaryl alcohol. The latter is often overestimated because the nitrobenzene products from p-coumaric acid are attributed to p-OHcinnamyl units of lignin. Syringyl lignin increases in proportion relative to guaiacyl and p-hydroxyphenyl lignins during maturation of some grasses. The formation of the hydroxycinnamate esters of GAX most likely occurs cytosolically, presumably in the Golgi apparatus (115). The cinnamyl alcohol precursors of lignin are also synthesized cytosolically, and several of the key enzymes in the biosynthetic pathway have been characterized in grasses. In addition to phenylalanine ammonia lyase, grasses also possess a tyrosine ammonia lyase that forms p-coumaric acid from tyrosine (116). Other hydroxycinnamates are made through oxygenase and methyl transferase reactions. These include trans-cinnamate-4-monooxygenase, which forms p-coumarate and caffeate from trans-cinnamic acid (121); caffeate-O-methyl transferase, which catalyzes the conversion of caffeate to ferulate (41); and cinnamyl alcohol dehydrogenase, which catalyzes the conversion of the cinnamyl aldehydes to p-coumaryl, coniferyl, and sinapyl alcohols, the direct precursors of lignin (120). The formation of cinnamyl alcohol glucosides may be important in transport, because the unglycosylated precursors are not very soluble in water. Cleavage of the glucosides by cell-wall -D-glycosidases may generate the active precursors in muro (163). The UDP-Glc:coniferyl alcohol transferases have been described in many nongramineous species (163). The mechanism of the polymerization of the cinnamyl alcohols into lignin is not completely established. For many years, the reactions were thought to be catalyzed solely by peroxidases, but recent evidence has implicated laccase as a participating enzyme (163). Use of 13C-labeling techniques has permitted a convenient monitoring of the synthesis of phenylpropanoid synthesis and poly merization (96, 124). A role of ferulates and diferulates in nucleation of lignification has been suggested (125).

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Structural Proteins

In Type I walls, the cell-wall carbohydrates of fully elongated and differentiating cells are cross-linked with structural proteins to produce an inextensible structure. In the primary walls of grasses, this function is carried out largely by phenolic substances. Homologs to several known classes of structural proteins are synthesized in the grasses and are in much larger amounts in specific cell types (83, 147). In nongramineous plants, the hydroxyproline-rich glycoproteins (HRGPs)--the extensins--are ubiquitous. These rod-shaped proteins owe their structure to two features: a polyproline II helix due to glycosylated Ser(Hyp)4 repeats and a Tyr-X-Tyr-Lys motif that is the site of formation of a stabilizing intramolecular isodityrosine linkage (83). The grass homolog is a threonine-hydroxyproline-rich glycoprotein (THRGP) found in maize (63, 82, 152), Sorghum (127), and rice (12). In the maize and Sorghum HRGPs, only one "signature" Ser-(Hyp)4 sequence remains near the carboxy terminus, whereas a majority of the repeats contain a Lys substitution for a Hyp (84, 127). A rice HRGP contains no Ser-Hyp4 repeats and a repeating ProPro-Thr-Tyr-Lys-Pro in place of Pro-Pro-Thr-Tyr-Thr-Pro of the maize and Sorghum proteins (12). This substitution is predicted to interrupt the polyproline II helix, and the increased molecular flexibility should prevent maintenance of a rod-like structure (84). The periderm, a firm, suberized structure, is quite enriched in the THRGP, which supports the idea of a special structural role for these proteins (42, 63). The THRGP is enriched in protoxylem and metaxylem and in the longitudinal radial walls of the epidermis (148). A unique, extensin-like protein with Ser-(Hyp)4 repeats has also been described in the maize pollen grain (27, 134). Glycine-rich proteins (GRPs) are encoded by a large family of genes and perform a broad spectrum of cytosolic and cell-wall functions (147). GRPs homologous to vascular cell-wall proteins from nongramineous species have been found only in rice (95) and barley (133). Many other GRPs of grasses do not have signal peptides (147), which indicates a possible cytosolic function. Another member of a large gene family, a proline-rich protein (PRP), was also described in maize (76). This protein has repeated Pro-Pro-Tyr-Val and Pro-Pro-Thr-Pro-Arg-Pro-Ser sequences at the N-terminal domain and a cysteine-rich C-terminal domain, features similar to some PRPs from nongramineous species. The protein portions of AGPs are also from a diverse family of genes encoding Hyp-rich and Hyp-poor polypeptides (40). Ryegrass AGP-peptides are Ala- and Ser-rich and contain Ala-Hyp repeats (51), a motif that is also found in the maize histidine-rich protein (81). van Holst & Fincher (158) showed that the Lolium AGP formed a polyproline II helix, but considering the broad diversity of AGP protein structures, this may or may not turn out to

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be a general feature. The soluble AGPs bind specifically to the Yariv reagent containing phloroglucinol--D-linked Glc units (75). In addition to the arabinosylated (13,16)--D-galactan structure (40), small amounts of other neutral and acidic sugars and sugar linkages are found that may give diversity in the carbohydrate domain. The carbohydrates are attached to Hyp residues in some AGPs (2), but it is uncertain whether Ser and Thr residues are the site of attachment in other classes of AGPs. AGPs are found in numerous cellular locations. They are the major polysaccharide in secretory vesicles (46), and distinct AGPs are associated with both the plasma membrane and cell-wall compartments in nongramineous species. Their functions in any location are still speculative. Nonstructural proteins reside in the grass cell wall. Many are hydrolases, transferases, esterases, peroxidases, and several other enzymes that function in the modification of cell-wall polymers at the various stages of development and differentiation (23). In addition, thionins, which are toxic to fungal pathogens, accumulate in the walls of many grasses (7), and grasses also accumulate pathogenesis-related proteins similar to those found in nongramineous species.

Other Cell-Wall Substances

Silica is particularly abundant in the walls of grasses, mostly as inclusion bodies in the epidermis, periderm, and other specialized cells of the root, rhizome, and aerial shoots (119). Little has been reported on any chemical interaction with other cell-wall constituents. Silica binds uronic acid residues of animal extracellular polysaccharide and peptidoglycans (142), which is an indication of possible complementary interactions with PGA or GAX. A group of related calcium-binding glycoproteins was discovered in specialized silica deposition vesicles involved in the synthesis of the silica-rich walls of marine diatoms (88). Similar to all flowering plants, grasses possess cutin, suberin, and waxes in specialized cells (3). Osmiophilic granules, apparently derived from the Golgi apparatus, appear in the outer epidermal layers during rapid growth in deep-water rice (89). These granules may carry precursors and enzymes responsible for the synthesis of the cuticle and secretion of waxes (61).


In a hallmark paper, Chase et al (26) examined the phylogenetics of flowering plants by nucleotide sequence variability of a ribulose bisphosphate carboxylase gene rbcL. Duvall et al (36) elaborated on these data in their reevaluation of the Monocotyledonae. From these extensive studies, two major



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divisions among the Monocotyledonae have been proposed: the "commelinoids," a large clade that includes the grasses, sedges, rushes, palms, and gingers, and the "noncommelinoids," a more basal group of aroids, alismatids, and lilioids (135). Harris & Hartley's (56) detection of the green fluorescence enriched in nonlignified cells is confined, almost without exception, to the commelinoids (Figure 2). Ruddall & Cadick (135) surveyed a broader range of monocots for wall fluorescence, including some of questionable lineage, and established that several genera that had previously been classified as lilioids are actually commelinoids. This classification is largely consistent with the system of Dahlgren et al (32), who showed that several anatomical, developmental, and biochemical traits distinguish the commelinoids from the noncommelinoids. Among these characteristics, the Poales are noted for a starchy endosperm; the presence of silica bodies in the epidermis, pericycle, and other specialized cells; and the absence of calcium oxalate raphide crystals (Figure 2). Silica bodies are found in most commelinoids, with the exceptions of many Commelinales and more basal species. Apart from a few Liliiflorae, silica bodies are absent from the noncommelinoids. Whereas the Poales, Cyperales, Commelinales, and most Zingiberales lack oxalate raphides, the crystals are detected in the Arecales and more primitive commelinoids (32). Unfortunately, cell-wall polysaccharide composition has not been used extensively enough to confirm some of the phylogenetic relationships, but the data available indicate at least two possible transitional stages in the development of the novel wall structure in the Poales. The first transition is the replacement of XyG with GAX as the principal cross-linking glycan in walls of meristematic cells (Figure 2b). Ferulate, which produces an alkali-induced green fluorescence, and p-coumarate are esterified specifically to the O-5 position of the arabinofuranosyl units of GAX (54). Because of this chemical feature, all commelinoids should have GAX as a major cell-wall polymer. Consistent with this prediction, GAX is the major cell-wall polysaccharide in the few nongramineous commelinoids that have been examined, including the Cyperales (10), Zingiberales (3, 165), and Bromeliales (149). NoncomFigure 2 Comparison of anatomical and chemical features of the commelinoid and noncommelinoid Monocotyledonae (after 32). (A) The presence of silica bodies (r) is associated with many of the commelinoids, whereas the presence of calcium oxalate raphide crystals (·) is associated with the noncommelinoids (32). (B) Detection of UV-fluorescence (í) in nonlignified cells is found in most commelinoids and progenitors, whereas there is an absence of fluorescence (ì) in the walls of all noncommelinoids (56, 135). The glucuronoarabinoxylan (t) is a major cross-linking glycan in many of the commelinoids (3, 10, 149, 165), whereas xyloglucan (s) predominates in the few noncommelinoids that have been examined (3, 128). There is a trend toward low pectin content (|) in the advanced commelinoids, although some have intermediate amounts (z), and the noncommelinoids are rich in pectin (z) (74). The mixed-linkage -D-glucan is present (r) only in the Poales and absent (·) from all other commelinoid and noncommelinoid species that have been examined (153).

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Annu. Rev. Plant. Physiol. Plant. Mol. Biol. 1996.47:445-476. Downloaded from by University of Georgia on 04/26/06. For personal use only.

melinoid monocots have XyG- and pectin-rich walls typical of the Type I walls (74, 128). The transition from XyG to GAX also represents an exchange of an acidic polymer for a neutral one, and the tight association of HS-GAX with the small amount of pectic fraction indicates that GAX may have replaced part of the role of the pectins in providing an anionically charged matrix. Jarvis et al (74) found that the content of the pectins among the commelinoids was consistently well below that of noncommelinoids and dicots (Figure 2b). The second transitional stage is marked by the appearance of the (13),(14)--D-glucan (Figure 2). This special glucan is found exclusively in the Poales (149, 153).


Biochemical studies have provided a good catalog of the wall constituents, but until recently we could only estimate, by virtue of the physical and chemical properties of the polymers in solution and solid-state, how the constituents were assembled, arranged, and cross-linked into a functional matrix. Cryopreservation techniques for electron microscopy have helped in visualizing the fine structure of the wall (102), and antibody and enzyme probes for specific cell-wall epitopes reveal the organization of certain polymers within domains of a single cell wall (87, 136). Fourier transform infrared and FTRaman spectroscopy are used to detect specific chemical bonds and their orientation within the underivatized cell walls of grasses (143). In dividing cells, the microfibrils are wound around each cell randomly, and this pattern continues throughout isodiametric expansion. When elongation begins, microfibrils are wound transversely or in a shallow helix around the longitudinal axis. Microfibrils of nongramineous walls are 5­15 nm wide and are spaced 20­40 nm apart (102). Preliminary measurements of mesophyll and epidermal cells of the maize coleoptile showed spacings that were slightly smaller than these dimensions (132), but comparisons of the spacings in growing and nongrowing walls with different architectures still need more thorough evaluation (MC McCann, personal communication). McCann & Roberts (101) suggested that the XyGs not only tether the microfibrils but also establish the spacings between the microfibrils. The ability of the GAX to self­hydrogen bond might result in altered spacings. Wilkie's review (165) of the gramineous xylans summarized broad variation among cereals in the Ara:Xyl ratios, an estimate of the degree of substitution of the xylan with arabinosyl side-groups. The Ara:Xyl ratio decreases in leaves and stems throughout the development of the oat plant (129), and the number of Ara units along the xylan chain varies markedly during coleoptile elongation--from GAXs whose Xyl units are nearly all branched to those with <10% of the xylosyl units bearing side-groups (14). The side-groups greatly affect the ability of the GAXs to bind to one another and to cellulose.



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Like XyG, the unbranched (14)-linked xylans hydrogen bond to cellulose and to one another, whereas the attachment of arabinosyl and GlcA sidegroups to the internal O-2 and O-3 secondary alcohols of the xylan backbone hinder the formation of hydrogen bonds (13). Because the Ara units markedly alter water solubility and the ability to hydrogen bond, the GAXs may exist in at least two distinct domains within the matrix: an HS-GAX domain that is a structural continuum with the pectin matrix and a relatively unbranched xylan domain that forms tight bonds to the cellulose microfibrils (13, 15). The distribution of methyl esterified and unesterified PGAs in Type II cell walls was detected by antibodies against these polymers (87). The grasses display a marked developmental preference for accumulating methyl esterified or unesterified pectins in specific cell types. Grasses have mostly esterified PGAs in vascular tissues, whereas walls of the cortical and parenchyma cells have mostly unesterified polymers (87). Neither antibody recognizes PGAs in walls of the root epidermis or root cap cells. As estimated by infrared spectroscopy, the esterified PGAs of grass walls represent a sizeable portion of the total pectin (111) and are easily distinguished by other spectral features from nongramineous species (143). Whereas pectins of the Type II wall are comprised of both PGA and RG, HS-GAX is also a major component closely associated with pectins, particularly in a fraction containing an AGP (17). The Type II cell-wall model has cellulose microfibrils bound and interlaced with unbranched GAXs (Figure 3a). Additional GAXs with varying degrees of branching may have functionally replaced the predominant pectic substances in the Type I cell wall, and the spacing of the appendant arabinosyl and GlcA units could determine porosity and surface charge. Given that the spacing between cellulose microfibrils measured tangentially is the same as the spacing radially, the lamellate structure is only about four to eight strata thick (Figure 3a).

Figure 3 Architecture of the Type II cell wall of the Poaceae. (A) Representation of four strata of microfibrils and associated polysaccharides and aromatic substances just after cell division. The microfibrils are interlocked by glucurononarabinoxylans. Unlike xyloglucans, the xylans are secreted in a form highly substituted with arabinosyl units that prevent hydrogen bonding. The units are cleaved from a portion of the GAX to yield runs of xylan that can bind on either face to cellulose or to one another (13, 20). Porosity of the GAX domain could be determined by the extent of removal of the appendant units. Some highly substituted GAX remains intercalated in the small amount of pectins that also are found in the primary wall (17). A portion of the noncellulosic polymers are cross-linked to the microfibrils by alkali-resistant phenolic linkages. (B) The expanding Type II cell wall. Absent from the developing wall of dividing cells, the -D-glucan is synthesized specifically during cell elongation and is the major cross-linking glycan. Long runs of (14)-linked glucan could bind to cellulose or to other glucan. Although some tissues accumulate structural proteins, the fundamental cross-linking in the primary wall of recently enlarged cells is by esterified and etherified phenolic compounds that lock the wall into place and halt further stretching of the microfibrils.

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Several structural and architectural changes accompany elongation of grass cell walls. Some of the changes involve the synthesis and turnover of developmental-stage-specific cross-linking glycans that are mechanical determinants of growth, whereas subtle alterations in pectin composition and orientation reflect enzymic or physiological control of growth. Undoubtedly, all of these changes reflect either directly or indirectly the biochemical and physical events of wall loosening, expansion, and retightening. During isodiametric expansion of meristematic cells, the major noncellulosic polysaccharides are GAX and the pectic polysaccharides (14). Liquid cultures of somatic cells, in which little cell elongation is observed, have cell walls that are essentially devoid of -D-glucan (21). In dividing and elongating cells, highly branched GAXs are abundant, whereas after elongation and differentiation more and more unbranched GAX accumulates (14, 15, 46). Turnover of Ara in coleoptiles has been observed in vitro with isolated coleoptile walls and wall fragments (34, 60) and in vivo by kinetic analysis of the interconversion of labeled Ara into the Xyl moieties of GAX by continuous recycling through the nucleotide-sugar pools (46). Interactions between the HS-GAX, PGA, and RG may function as a control mechanism of wall metabolism. The PGAs of maize pectins contain nonmethyl esters, whose formation and disappearance are coincident with the most rapid rate of cell elongation (85) and the accumulation of the HS-GAX (14). Maize GalAs are esterified in a greater proportion than can be accounted for by methyl esters alone, and formation of these nonmethyl esters is associated with covalent attachment to the primary wall (85). Brown & Fry (9) discovered an unusual hydrophobic ester of GalA, but they did not identify the alcohol moiety. These nonmethyl esters of PGAs may have escaped detection by both kinds of pectin antibodies (87). Schopfer (141) found Yariv-positive AGPs located at the inner surface of the growth-controlling epidermal wall of the maize coleoptile, a position that suggests an involvement in cell expansion. Schindler et al (140) tested this hypothesis with three monoclonal antibodies directed against different AGP epitopes. Their finding of AGPs in the membranes of developing sclerenchyma and tracheids supports the idea that the AGPs are developmental markers, but the inability of auxin to modulate the amount of soluble AGPs in nonvascular tissues or epidermis is evidence for a lack of involvement of these peptidoglycans in wall loosening (140). The pectin domain may exert growth-controlling functions, but the interaction of the cross-linking glycans and cellulose is the tensile force-bearing structure that must be loosened. Taiz (154) reviewed much of the early literature on enzymatic activities associated with elongation in grasses and dicots. Subsequent studies focused on the specific polysaccharides in the walls and

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how they might be modified to permit extension. The synthesis of polysaccharides specific to cell expansion is a special feature of the grasses. Some arabinans, particularly the 5-linked arabinans, are found in the walls of dividing cells but are no longer made during cell expansion (14). Instead, the -Dglucans are now synthesized along with GAX (Figure 3b). The -D-glucans are substrate for two enzymes also enriched in growing seedlings: endo--Dglucanase, which requires at least four contiguous (14)--D-Glc units for activity (57), and an exo-glucanase, which yields Glc from the oligosaccharides produced by the endo--D-glucanase (67). These types of glycanohydrolases are thought to be secreted to or activated at specific sites within the cell wall to cleave tension-bearing -D-glucans that interlace the cellulose microfibrils (154). The -D-glucan is enriched in the outer walls of the epidermal cells (108, 137), the organ that is likely to be the layer that controls elongation rate (89, 90). The appearance of -D-glucan during cell expansion, the association of hydrolysis of -D-glucan in muro, and acceleration of its hydrolysis rates by growth regulators all implicate direct physical involvement of the polymer in extension growth. There is some skepticism about this hypothesis because of a lack of correlation between growth and loss of polysaccharide. -D-glucan is a stage-specific polymer that accumulates only transiently during expansion and disappears when growth is completed (14, 98). Growth in vivo is accompanied by a net accumulation of the -D-glucan that masks an extremely rapid turnover, and only in excised grass tissues not provided with sugar is the loss of glucan accelerated during elongation (154). Intact grass coleoptiles and deep-water rice accumulate -D-glucan during the elongation phase of growth (14, 98). In intact plants, the remaining glucan is gradually lost from the wall once growth and synthesis of new -D-glucan ceases. Hence, the synthesis and hydrolysis of the -D-glucan are in dynamic equilibrium. The correlation factor for -D-glucan and growth is not the relative abundance but the rate of turnover, i.e. the balance between synthesis and degradation. Because of the efficiency of sugar recycling mechanisms, turnover in vivo can be measured only indirectly (46). Enzyme- and substrate-specific antibodies offer an alternative approach to test the role of -D-glucan hydrolysis in wall expansion. Antibodies against exo- and endoglucanases purified from the walls of growing seedlings partially reverse auxin-induced growth (71). Antibodies against the -D-glucan itself can also inhibit growth (64, 66). Hoson et al (65) found that antibodies against the XyG nonasaccharide were able to inhibit growth in legumes but not grasses, which is an apparent demonstration of the specificity of the antibody interaction with a specific hydrolase. New hypotheses have been suggested for how stress relaxation of the wall may result in cell expansion without glucan hydrolysis (44, 105). Two differ-

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ent kinds of wall proteins are implicated in expansion in XyG-cross-linked Type I walls without net depolymerization of a cross-linking glycan. A XyG endotransglycosylase (XET) catalyzes transglycosylation rather than hydrolysis of XyG. In this reaction, one chain of XyG is cleaved but reattached to the nonreducing terminus of another XyG chain (44). By such a mechanism, the microfibrils could undergo a transient slippage without compromising tensile strength. It also provides a mechanism whereby new linkages between polymers can occur when juxtaposed after microfibril slippage. Grass root tips also exhibit substantial XET activity, and this activity is correlated with extension growth (122, 170). A wheat XET gene shows better than 70% sequence homology with several dicot XETs (118). These findings are surprising because XyGs are neither a major structural polymer of the grasses nor possess a unit structure that would be expected to produce effective oligomeric substrates for XET in vivo. No polysaccharide other than XyG has been shown to be substrate for XET. Cosgrove and his colleagues (106, 107) discovered two proteins, called expansins, that catalyze wall extension in vitro without detectable hydrolysis or transglycosylation. These proteins induce extension of paper in stress relaxation assays, which indicates that they probably catalyze breakage of hydrogen bonds (104). Homologous grass expansins could disrupt the tethering of cellulose by -D-glucans and xylans in the Type II walls. Expansins induce extension in heat-killed abraded coleoptile sections (31), and oat expansin induces extension of cucumber walls and is similar in activity to the cucumber enzymes (97). Although the oat expansin is abundant in sections of the coleoptile exhibiting substantial growth, expansin is still present in cells that are not growing, which suggests that other mechanisms modify the wall to make it insensitive to expansin action. The substrate(s) for expansin in grasses are not yet established, but expansin binding to cellulose is enhanced by coating the alkali-extracted walls or fibrous cellulose with -D-glucan (105). A 40kDa glycoprotein was localized specifically in the walls of elongating cells of several gramineous and nongramineous plant species (130). The protein is found in the walls of coleoptile tissues and is induced in expanding leaves in deep-water rice. Although the appearance of this cell-wall protein is tightly coupled with elongation, the function is unknown. To summarize, enzymes and proteins implicated directly and indirectly in wall extension and growth include those involved in pectin assembly and gel formation, the turnover of AGPs, the assembly of GAX, the breaking of hydrogen bonds between cross-linking glycans, and the turnover of the -D-glucans. Once the genes of these enzymes are cloned, molecular experiments could be used to assess the specific role of each of these proteins in growth processes.

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Both esterified and etherified cinnamic acid constituents accumulate at rates inversely correlated with elongation rates (16, 110, 155). The cessation of growth is correlated with the appearance of cinnamyl alcohol dehydrogenase (112, 137) and specific peroxidases (86, 99) required for cinnamate synthesis and polymerization in the grasses. The accumulation of the phenolic esters and ethers is an important event not only for locking the cells into their final shape but also for providing tensile and compression strength. Barley cells in liquid culture grown in the presence of dichlorobenzonitrile compensate for the lack of cellulose by increasing cross-linking of polymers by phenolic compounds, an alteration that increases the tensile strength of the wall despite cellulose levels only 30% of normal (144).


The chemical structure and architecture of the cell walls of grasses change markedly during the stages of cytokinesis, elongation, and maturation. Like all flowering plants, the developing cell plate--or phragmoplast--is derived from vesicles of the Golgi apparatus and grows outward until it fuses with the existing primary wall (150). As in animal cells, the plant Golgi apparatus is a factory for the synthesis, processing, and targeting of glycoproteins (150). The Golgi apparatus has been established by autoradiography and immunocytochemistry as the site of synthesis and export of all the cross-linking glycans and pectins. Cellulose and callose, an unbranched (13)-linked glucan, are the only polymers known to be made at the plasma membrane surface in any plant (35). Cellulose synthesis in higher plants is likely to occur at six-membered hexagonal terminal complexes called rosettes. The first rosettes observed in freeze fracture replicas of plasma membranes were those of the maize root (113). The quest for cellulose synthesis in vitro has been a long one. Recent attempts to identify polypeptides involved in the synthesis have focused on design of photoaffinity and other labels and the use of product entrapment (35, 48, 68). Meikle et al (109) precipitated callose synthase from Lolium membranes with monoclonal antibodies and detected several affinity-labeled polypeptides that were similar to those labeled in nongramineous species. The identity and function of these proteins are unknown. Pulse-labeling studies with intact seedlings and excised tissues have revealed many dynamic features of polysaccharide synthesis and turnover in vivo. In excised maize coleoptiles, Ara is incorporated into UDP-Ara and UDP-Xyl, but not into any hexose, whereas Xyl is incorporated into the hexose and pentose nucleotide sugars with distribution similar to that of Glc (19). These data reflect differing strategies that plants possess in managing the sugars hydrolyzed from polymers during growth. Plants possess C-1 ki-



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nases for Ara, Gal, Man, GalA, GlcA, Rha, and Fuc. Subsequently, NDP pyrophosphorylases catalyze the formation of the UDP or GDP sugars (37). All of these sugars are synthesized de novo from UDP-Glc, or in some instances from GDP-Glc, via nucleotide-sugar interconversion pathways. The existence of these salvage pathways is considered evidence that turnover of polysaccharides involves hydrolysis of the sugars from specific polysaccharides or proteins (46). Whereas stress relaxation of the wall is the physical event, synthesis is the biological event (132); both are vital and, to an extent, biochemically integrated during sustained growth. Even though substantial growth can occur without detectable secretion of new polysaccharide, inhibition by monensin or brefeldin A of secretory vesicle transport to the plasma membrane quickly stops growth (139).

Biosynthesis of (1®3),(1®4)-b-D-Glucan

We know very little about how cell-wall polysaccharides are organized for secretion and then modified for assembly into a functional matrix, but we are beginning to learn some of the features of the polymerization from studies of synthesis in vitro. Some special features of polysaccharide synthesis in the grasses are noteworthy. The grass -D-glucan differs from other (14)--Dglucans and callose because the (14)- and (13)--D-glucosyl linkages are ordered within the polymer. Synthesis of authentic -D-glucan in vitro is determined by hydrolysis of the radioactive products formed with Golgi membranes and UDP-Glc with the sequence-dependent B. subtilis endoglucanase, which yields diagnostic cellobiosyl- and cellotriosyl-(13)--D-Glc. These oligomers are separated by HPLC and assayed for incorporation of radioactivity (47). Synthesis of -D-glucan with cellular membranes was shown by comparable techniques (59, 109). UDP-Glc is substrate and Mg2+ or Mn2+ is required as cofactor. Improvement in the recovery assay of polymerase activity was provided by a flotation centrifugation method for isolating Golgi membranes and by improved procedures for separating and identifying, by GLC mass spectrometry and GLC gas proportional counting, diagnostic linkage derivatives of the products (45). The combination of gel permeation chromatography, linkage analysis, and enzymic digestion confirmed that entire tri- and tetrasaccharide units were synthesized and that the macromolecular -D-glucan synthesized in vitro in the Golgi apparatus was identical to the cell-wall polysaccharide (47). Unlike the Golgi apparatus from plants with Type I walls, the maize Golgi membranes also possess callose synthases. Because the Golgi synthase activity was stimulated only twofold by CaCl2 as compared with sevenfold in plasma membrane, the activity could not be attributed solely to contamination with plasma membrane (49). Disruption of membrane integrity with detergents and ionophores abolished -D-glucan synthase activity but increased



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the synthesis of callose. The Golgi-specific callose synthase may represent the default synthase, and similar to that of the plasma membrane, its activity is stable to solubilization with digitonin or CHAPS. Meikle et al (109) have demonstrated that a polypeptide transiently associated with the synthesis of -D-glucan synthesizes only (13)-linked glucosyl units when isolated with detergents. After removal of inhibitors of activity with a Ca2+-precipitation technique, microfibrillar callose was synthesized in copious amounts by a Lolium callose synthase enriched by product entrapment (11). Like no other polysaccharide from higher plants, the grass -D-glucan contains an ordered arrangement of cellulosic (14)--D-linkages and callosic (13)--D-linkages. Because of the kinds of linkages formed by the -D-glucan synthase, the divalent cation requirements, the substrate and apparent Km, and the appearance of callose synthase upon damage of Golgi membranes, the grass synthase genes may have evolved from ancestral cellulose synthase genes.

Biosynthesis of Other Cell-Wall Polysaccharides

Other cell-wall polysaccharides are made in the Golgi apparatus, and some of these activities have also been studied in vitro. Incorporation of Ara from UDP-Ara and Xyl from UDP-Xyl into ethanol-insoluble products thought to represent arabinan and xylan, respectively, have been demonstrated in membrane preparations from grass tissues (162). For complex xylans such as GAX, complementary additions of UDP-Ara and UDP-Xyl should result in branched polymers containing increasing amounts of 2,4- and 3,4-linked xylosyl units and a corresponding amount of nonreducing terminal arabinofuranosyl units. Xylans and glucuronoxylans can be made in vitro, although the radioactive products have not been characterized as extensively. One of the more intriguing questions is how arabinofuranosyl units are made. L-Arabinose is in the furanose ring conformation in most plant polymers containing this sugar, including GAX, 5-linked arabinans, AGP, and extensin, whereas UDP-Ara is exclusively in the pyranose form (43). Arabinosyl transferase must be distinct from other glycosyl transferases in its ability to permit ring rearrangement before addition of the sugar to the polymer.


The work of Reiter et al (131) and Chapple et al (25) illustrates how cell-wall mutants in Arabidopsis can be used to understand polysaccharide and lignin biosynthetic pathways and functions. A similar genetic model is needed for the grasses. There are many genera within the Poales that could potentially serve as pure genetic models. For example, Eragrostis and Chloris are small annual grasses with short life cycles, are 2N, and possess genome sizes only



two to three times that of Arabidopsis (6). The genome sizes of most of the cereal crop species are enormous by comparison with other flowering plants, which makes them rather unwieldy systems to generate and screen mutants for specific traits and to isolate genes by chromosome walking. Rice possesses the smallest genome of the major cereal crops but is difficult to manage in large numbers within a greenhouse environment. These problems are offset by genetic maps well-populated with markers, and current map-based cloning strategies make it possible to move at a reasonable pace from a phenotypic mutation to a cloned gene. Although not a single synthase involved in cell-wall polysaccharide synthesis has been purified and its gene knowingly cloned, many of the enzymes that function in the depolymerization of the cell wall are well characterized. In grasses, three separate enzyme activities are associated with -D-glucan metabolism. They are encoded by genes within several related gene families, and many representative members of the families have been sequenced (62). The activities include (13)--D-glucanases, which are present in the developing grains, persist throughout germination, and are homologous with the pathogenesis-related glucanases (38); the (13,14)--D-glucanases, which appear exclusively in the grains during germination and depolymerize the cereal -D-glucan (39); and the (14)--D-glucanases found in developing seedlings (57). Two exoglucanases have also been described: a 60-kDa enzyme that appears in the grains (94) and a 72-kDa enzyme that is associated with the walls of elongating cells (71). These enzymes hydrolyze nonreducing t-Glc(13)- or t-Glc-(14)-linked units from the oligosaccharides produced by endoglucanases. The cereal (13,14)--D-glucanases, similar to the B. subtilis endoglucanase, cleave a (14)-linkage only if the adjacent Glc on the nonreducing side is (13)-linked (151). This property makes the grass -D-glucan a good substrate for the enzyme, but the enzyme is unable to hydrolyze glucans composed of solely (13)- or (14)--D-glucosyl units. It is interesting to note that the barley (13,14)--D-glucanase genes are homologous to the (13)--D-glucanase genes but are completely unrelated to bacterial enzymes with the same carbohydrate linkage specificity (39, 171). The barley glucanase has been crystallized and its three-dimensional structure determined (28, 159). From mapping of the substrate-binding cleft, strategies to alter substrate specificity and other properties of the enzyme by site-directed mutagenesis are possible (62). Many of the genes that encode enzymes in the synthetic pathways of the lignin precursors have been cloned. Research is now focused on characterizing the regulatory elements as a strategy to modify lignin biosynthesis (163). Several "brown-midrib" (bmr) mutants of maize, Sorghum, and millet have impaired ability to synthesize lignin, and, in some instances, this factor in-

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creases digestibility (29). The maize bmr3 corresponds to an impaired gene encoding caffeate O-methyl transferase (161), an enzyme activity shown to be reduced in maize and millet bmr mutants (29, 53). Because lignin is a compression-strength component of the xylem and lignin-like substances can form a line of defense against pathogens, a simple reduction of phenylpropanoid and lignin content in plants is probably not a viable strategy to increase digestibility. Lignin content and structure are not the only factors in digestibility of the grasses. The extensive networks of ester- and ether-linked hydroxycinnamic acids in nonlignified cells and the pectin and other polysaccharide interactions with the phenylpropanoid network undoubtedly are contributing factors in decreased digestibility (77). The starchy endosperm, a special trait of the Poales, is the fundamental reason that cereals are of central importance in human nutrition (93). The world harvests over one billion tons of cereal grains annually. Rice and wheat alone provide at least half of the calories that humans ingest. The cell walls of the grasses also figure heavily in the nutrition equation, from the -D-glucans that constitute up to 70% of the endosperm walls, to the vast amounts of xylan- and cellulose-rich walls that are consumed by grazing animals. The -Dglucans can be both a benefit and a problem. As a benefit, they are the wall constituents implicated in the ability of barley and oat brans to reduce serum cholesterol in hypercholesterolemic individuals (8) and to modulate glucoregulation in diabetics (166). The composition of the endosperm cell walls in flours is a contributing factor to bread quality (50). Incomplete hydrolysis of the viscous -D-glucans during the brewing process is a major production problem and contributes to "hazing" of beers upon storage (168). Lignin, esterified and etherified aromatic substances, and other chemical modifications of the primary wall polysaccharides greatly reduce the nutritional quality of grasses for ruminants (77). Considering the social and economic impact of the cereals, it is not surprising that there is a considerable knowledge base in the genetics of the cereal crops. This knowledge base could be more widely used to provide information on the structure, biogenesis, and turnover of the special cell wall of the grasses and in the process reveal ways that the cell wall can be modified for agronomic benefit. ACKNOWLEDGMENTS I thank Tony Bacic, University of Melbourne; Larry Dunkle, Purdue University; Maureen McCann, John Innes Centre; and John Ralph, University of Wisconsin­Madison for valuable discussions and their critical review of this manuscript. I thank Debby Sherman, Purdue University Electron Microscopy Facility, for her help in the digital imaging of the figures. My work on cellwall biogenesis in the grasses has been supported largely by grant DE-FG0288ER13903 of the Division of Biological Energy Sciences, US Department



of Energy. This is publication number 14,910 of the Purdue University Agriculture Experiment Station.

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