Plant glycoside hydrolases involved in cell wall polysaccharide degradation

Plant Physiology and Biochemistry 44 (2006) 435–449 www.elsevier.com/locate/plaphy

Review

Plant glycoside hydrolases involved in cell wall polysaccharide degradation Z. Minic, L. Jouanin* Laboratoire de biologie cellulaire, Institut national de la recherche agronomique, route de Saint-Cyr, 78026 Versailles cedex, France Received 23 December 2005 Available online 07 September 2006

Abstract The cell wall plays a key role in controlling the size and shape of the plant cell during plant development and in the interactions of the plant with its environment. The cell wall structure is complex and contains various components such as polysaccharides, lignin and proteins whose composition and concentration change during plant development and growth. Many studies have revealed changes in cell walls which occur during cell division, expansion, and differentiation and in response to environmental stresses; i.e. pathogens or mechanical stress. Although many proteins and enzymes are necessary for the control of cell wall organization, little information is available concerning them. An important advance was made recently concerning cell wall organization as plant enzymes that belong to the superfamily of glycoside hydrolases and transglycosidases were identified and characterized; these enzymes are involved in the degradation of cell wall polysaccharides. Glycoside hydrolases have been characterized using molecular, genetic and biochemical approaches. Many genes encoding these enzymes have been identified and functional analysis of some of them has been performed. This review summarizes our current knowledge about plant glycoside hydrolases that participate in the degradation and reorganisation of cell wall polysaccharides in plants focussing particularly on those from Arabidopsis thaliana. © 2006 Elsevier Masson SAS. All rights reserved. Keywords: Arabidopsis; Cell wall; Polysaccharides; Glycoside hydrolases

1. Introduction The plant cell wall represents half of the organic carbon in the biosphere. Many organisms can efficiently degrade the cell wall and use by-products of its degradation for nutrition [1]. During plant life, the cell wall plays an important role in various physiological functions such as growth, intercellular communication, defense against pathogen attack, mechanical resistance, and interaction with the environment [2–5]. Abbreviations: D-Glc, D-glucose; D-Xyl, D-xylose; EG, Endoglucanase; EndoPG, endopolygalacturonase; ExoPG, exopolygalacturonase; GalA, α-DGalacturonide; HPLC, high performance liquid chromatography; HG, homogalacturonan; L-Ara, L-arabinose; pNP, p-nitrophenyl; OSX, oat spelt xylan; RAX, rye arabinoxylan; SBA, sugar beet arabinan; RG-I, rhamnogalacturonan I; RG-II, rhamnogalacturonan II; WAX, wheat arabinoxylan; X-Gal, 5-bromo-4-chloro-3-indoxyl-β-D-galactopyranoside; XET, xyloglucan endotransglucosylase; XTH, xyloglucan endotransglycosylase/hydrolase; XGA, xylogalacturonan. * Corresponding author. Tel.: +33 1 30 83 30 63; fax: +33 1 30 83 30 99. E-mail address: [email protected] (L. Jouanin). 0981-9428/$ - see front matter © 2006 Elsevier Masson SAS. All rights reserved. doi:10.1016/j.plaphy.2006.08.001

Plant cells undergo two types of cell wall deposition: the primary and secondary cell walls. The primary cell wall is synthesized during cell expansion at the first stages of development and is composed of cellulose, hemicelluloses, pectic polysaccharides and many proteins. The polysaccharide composition of the primary cell wall can vary and is designated as type I or II. Type I is the most common while type II is typical for grasses [6]. The non-cellulosic biopolymers of type I walls are xyloglucan and about 35% of the wall mass are pectins. Type I walls are found in all dicotyledons, the nongraminaceous monocotyledons, and gymnosperms [7]. Type II walls have a low pectin and xyloglucan content and a high arabinoxylan content [6]. Type II walls also contain mixed linked β-D-glucan and possess ester linked ferulic bridges in the xylan, which have not as yet been found in type I walls. The secondary cell wall is deposited in fully expanded and specialized cells (xylem and fibers). Compared to primary walls, secondary walls contain more cellulose with a higher degree of polymerization and crystallinity which confers desirable quality traits for wood product industries [8]. The thicken-

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ing of the secondary cell wall is mostly due to the deposition of polysaccharides (as cellulose in cotton fibers and xylans in wood) and accumulation of phenolic polymers such as lignins in xylem vessels or sclerenchyma fibers. Plant cell wall polysaccharides are the most abundant organic compounds found in nature. They make up 90% of the plant cell wall and can be divided into three groups: cellulose, hemicelluloses, and pectins and their composition varies from one plant species to another (dicot/monocot), but also in different tissues of the same plant [2,9,10]. Structural proteins and enzymes constitute the remaining 10% of the plant cell wall. They play a crucial role in cell wall structure and architecture, cell wall metabolism, cell enlargement, signalling, response to abiotic and biotic stresses, and many other physiological processes [2–7]. 2. Plant cell wall polysaccharides Cellulose represents the major cell wall polysaccharide and consists of a linear polymer of β-(1→4)-linked D-glucose residues forming (1→4)-β-D-Glucan [1]. In addition, cell walls may contain callose, a linear (1→3)-β-linked polymer of DGlc with occasional (1→6)-β-linked branches. Plant callose is produced in pollen and synthetised in response to wounding, and pathogen attack [11]. Angiosperm microsporocytes synthesize callose before meiosis [12]. Cellulose and callose are synthesized at the plasma membrane level and then deposited into the wall [11]. The cellulose polymer is present as microfibrils, and its main function is to ensure the rigidity of the plant cell wall [13]. The mixed (1→3)(1→4)-β-D-glucan is especially abundant in walls of the starchy endosperm. For example, it constitutes about 75% of cell walls in the starchy endosperm of barley grain whereas it is not very abundant in walls of other tissues [14–16]. (1→3)-β-D- and (1→4)-β-D Glycosyl bonds are not randomly distributed in the (1→3)(1→4)-β-D-glucan. Hydrolysis of the native polymer reveals that 70% of the polysaccharide consists of cellotriosyl units connected by single (1→3)-β- linkages [16]. (1→3)(1→4)-β-D-Glucan is tightly associated with cellulose and other noncellulosic polysaccharides in the primary walls of growing cells. This polymer is synthetised in the Golgi apparatus [16]. Hemicelluloses constitute up to 30-35% of the dry weight of the plant cell wall [9] and are mainly composed of xyloglucan and xylan with minor amounts of several mannose-containing polysaccharides [17, 18]. The hemicelluloses are synthesized in the Golgi apparatus and are then excreted into the wall. The major hemicellulose polymer in cereals and wood is xylan [19,20]. Xylan is composed of a backbone of β-(1→4)-linked Dxylopyranosyl units substituted with L-arabinofuranose, glucuronic acid, 4-O-methylglucuronic acid, and acetyl sidegroups to varying extent [19,20]. Xylans can form crosslinkages with lignin via α-L-arabinofuranosyl residues, which can be esterified with hydroxycinnamic acids, such as ferulic or p-coumaric acids [6]. Arabinoxylan has a β-(1→4)-D-xylopyranoside backbone carrying O-2, O-3 or O-2,3 α-L-arabino-

furanosyl substitutions with an overall L-Ara/D-Xyl ratio of around 0.7 [21]. An unusual feature of arabinoxylan from barley is the high proportion of xylose residues monosubstituted at O-2 [22]. Some of the arabinose substituents are esterified at C-5 with ferulic acid or p-coumaric acid resulting in the formation of intermolecular cross-links [23]. Xyloglucans as constituents of the primary cell wall, possess a β-(1→4)-linked D-glucosyl backbone substituted with xylosyl, galactosyl and L-fucosyl residues [24]. Xyloglucans interact with cellulose microfibrils by formation of hydrogen bonds [25], thus contributing to the structural integrity of the cellulose network [26]. Binding capacity between cellulose and xyloglucans depends on the number of side-chains branching fucose and galactose residues [24,25]. A minor amount of the hemicellulose is composed of galacto(gluco)mannan, which consists of a backbone of β(1→4)-linked D-mannose and D-glucose residues with Dgalactose side groups [27]. Galactoglucomannans are abundant polysaccharides in the thickened secondary walls of gymnosperms [28] and Arabidopsis [18]. Pectins, like hemicelluloses, are synthesized in the Golgi and excreted to the wall [29,30]. Pectins are heteropolysaccharides composed of homogalacturonan (HG), rhamnogalacturonan I (RG-I), substituted homogalacturonans named rhamnogalacturonan II (RG-II) and xylogalacturonan (XGA) [30]. Pectins interact in the wall in many ways. First, the helical chains of polygalacturonic acids can condense by crosslinking with Ca2+ to form junction zones, linking several chains together and forming a gel. Pectins may by crosslinked further via ester linkages with dihydroxycinnamic acids, such as diferulic acid, to form covalent bonds with other polymers [31]. Homogalacturonan consists of an α-(1→4)-linked Dgalacturonic acid backbone which is partially methylesterified. RG-I is a heteropolysaccharide consisting of the repeating disaccharide unit α-(1→2)-L-rhamnosyl α-(1→4)-GalA [31]. Numerous side chains can be attached to the rhamnosyl residues of the RG-I backbone. These chains include galactans (β1,4-linked) or arabinans (α-1,5-linked), both of which can be decorated further with arabinosyl residues or side chains. In certain pectins, ferulic acid can be present as terminal residues attached to O-5 of the arabinose residues or O-2 of the galactose residues. Xylogalacturonan (XGA) is a branched galacturonan with β-(1→3)-D-xylopyranoside chains [32,33]. The αD-Galacturonide (GalA) residues of XGA can be methylesterified like HG [32]. RG-II is a very complex pectic polysaccharide. RG-II is composed of at least 12 different glycosyl residues linked together by more than 20 different glycosidic linkages [34]. This polysaccharide exists in the primary wall as a dimer that is cross linked by a 1/2 borate-diol diester [35]. RG-II is extremely resistant to known microbial polysaccharide degrading enzymes [36]. At the present time little is known about the degradation of RGII by plant enzymes. These polysaccharides (hemicelluloses, pectins) and the aromatic lignin polymer interact with the cellulose fibrils to create a rigid structure strengthening the plant cell wall. These poly-

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mers also form covalent cross-links, which are thought to be involved in limiting cell growth and reducing cell wall biodegradability. Such covalent cross-links have been identified between plant cell wall polysaccharides and lignin [37,38], between arabinoxylan polymers [23], between pectin polymers [39], between pectins and xyloglucan [40], and between pectinxyloglucan-xylan complexes [41]. During plant growth, the size and morphology of cells change. A coordinated series of biochemical events occur resulting in the biosynthesis and degradation of cell wall components. Consequently, numerous enzymes must be implicated in these processes to allow the remodelling of cell wall structure by selective degradation of some polysaccharide components [2,9,42,43]. This review focuses on plant enzymes that participate in the modification of cell wall polysaccharides. 3. Enzymes involved in the degradation of cell wall polysaccharides The degradation of cell wall polysaccharides is dependent on the action of numerous enzymes. Fry [5] classified such enzymes into three groups: exopolysaccharidases, endopolysaccharidases and other hydrolases that do not belong to these two groups. Exopolysaccharidases attack poly- and oligosaccharides progressively from the nonreducing terminus, or substituted side groups, releasing monosaccharides and sometimes disaccharides. Endopolysaccharidases attack the polysaccharide backbone at any position. They have an immediate and large impact on the molecular weight of polysaccharides. Hydrolases from the third group can break some substituted noncarbohydrate groups linked to wall polysaccharides such as O-acetyl, O-methyl, O-feruloyl and others [5]. After biosynthesis and deposition of polysaccharides, these enzymes, located in the wall itself or in the plasma membrane, participate in the degradation of different cell wall polysaccharides. These modifications allow changes in the structure and composition of polysaccharides. In this manner, enzyme polysaccharidases participate in regulation of cell wall expansion and alteration. No data have been reported about the cellular localisations of these enzymes, other than those localized in the wall itself or in the plasma membrane, that may be directly involved in the degradation of cell wall polysaccharides except for a cytoplasmic xylanase from barley [44]. Plant cell wall polysaccharides are very heterogenous/complex polymers and consequently the spectra of activities of the glycoside hydrolases must be very diverse with many exo- and endo- polysaccharidases involved in their degradation. 4. Experimental approaches used for studying glycoside hydrolases The functions of plant cell wall glycoside hydrolases have been studied using different biochemical and molecular genetic approaches. The biochemical approach consists in the extraction and purification of enzymes from the cell wall and analysis of their enzymatic activity, molecular and catalytical prop-

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erties, determination of post-translational modifications and structural characterization [5]. The biochemical strategy has also been used to determinate substrate specificities and their mode of action. Enzymatic activities of many exo glycoside hydrolases have generally been assayed using chromogenic or fluorogenic artificial substrates such as 4-nitrophenylglucosides or 4-methylumbelliferyl glucosides [5]. The genetic approach consists of identifying mutant plants presenting modifications in their polysaccharide structure [45,46]. Comparative molecular genetic studies have not been efficient because plant cell wall enzymes are functionally and structurally different from those of bacteria and yeast cell walls [47]. Screens to identify cell wall mutants were reviewed by Turner et al.. [48]. Most of the cell wall screens have used chemically mutagenized plant collections [45,49]. However, cloning of a gene from a mutated line is time-consuming and difficult. Screens based on promoter or gene trap strategies have allowed the isolation of mutants of interest by direct cloning of the mutated genes [50,51]. Microarray transcript profiling is a new technology for large-scale gene regulation studies in multicellular eukaryotes [52]. The complete Arabidopsis Affimetrix gene chips and the CATMA microarrays [53] contain most of the predicted genes in the Arabidopsis thaliana genome. The identification of genes coding for potential glycoside hydrolases specifically expressed in a tissue or under a particular condition could constitute another strategy for the identification of cell wall degrading enzymes [54,55]. The function of these enzymes could then be further studied using functional genetics and biochemical methodologies. Recently, progress has been made in proteomic approaches for extraction and identification of plant cell wall proteins [56– 59]. For instance, an efficient protocol was developed for the extraction of cell wall proteins from plant tissues based on vacuum-infiltration [58]. Using this protocol several families of glycoside hydrolases were identified. 5. The cell-wall degrading CAZymes in Arabidopsis The superfamily of glycoside hydrolases and transglycosidases is divided into numbered families based on amino acid sequence similarities [59,60]. Superfamilies of glycoside hydrolases and transglycosidases can be conveniently accessed through the carbohydrate-active enzyme web site CAZy (http:// afmb.cnrs-mrs.fr/CAZY/). For each family, this continuously updated database provides a list of identified members with links to their nucleotide sequence and most recent structural analysis. In addition, conserved catalytic residues are presented for each family. Arabidopsis thaliana with its complete genomic sequence data [61] is considered as the model plant for studying carbohydrate metabolism in plants. Arabidopsis contains over 730 open reading frames corresponding to two main classes of carbohydrate-active enzymes, glycoside hydrolases and transglycosidases [62]. The comparison of Arabidopsis and rice genomes shows variations in genome size and gene number [63]. The number of certain genes encoding glycoside hydrolases differs significantly between Arabidopsis and rice,

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particularly for the genes coding polygalacturonases. This observation suggests that the glycoside hydrolases participating in cell-wall dynamics differ in these plant species. This observation is in agreement with the different pectin composition found in the cell walls of these two species. The commelinoid monocots, such as rice, have a low content of pectin in contrast to Arabidopsis, a dicot which is rich in this constituent. The Arabidopsis glycoside hydrolases are thought to play various functions in plant defense, signalling, hydrolysis of glycoproteins and starch. Members of glycoside hydrolases of Arabidopsis families have been reported by Henrissat et al [62]. The CAZY (http://afmb.cnrs-mrs.fr/CAZY/) databases and TAIR (http://www.arabidopsis.org/info/genefamily/GlycosideHydrolase.html) have been used to try to select potential or experimentally determined enzyme activities of cell wall glycoside hydrolases in Arabidopsis. Enzyme activities involved in plant cell wall degradation were explored from previously known data [4,5]. In addition, the PSORT algorithm (http:// psort.nibb.ac.jp/form.html) was used to predict the cellular localization of selected sequences. A total of 200 genes in the Arabidopis genome that encode enzymes predicted to have a membrane or an extracellular localization are listed in Table 1. The selected enzymes belong to 13 different families. Endoand exo- glycoside hydrolases belong to 7 different families with family 28 containing both exo- and endo- glycoside hydrolases. Previous phylogenetic analyses demonstrated that this family contains three distinct clades. Clades A and B were suggested to have endo-polygalacturonase activities while members of clade C probably possess exopolygalacturonidase activity [64]. Enzymatic activities have never been determined for most of the proteins listed in Table 1. Very little information (existing references are indicated in Table 1) is available concerning the biochemical properties and mode of action of these proteins. Most of the characterized enzymes have been studied using genetic approaches and there may remain for this reason some ambiguity concerning their exact role. In addition, listed glycoside hydrolases may have other functions than hydrolysis of cell wall components such as defense against pathogen attack, hydrolysis of the sugar moieties of glycoproteins or glycolipids, storage of polysaccharides or of different metabolites present in cell wall. The last column of Table 1 indicates the number of ESTs (reflecting the amount of transcript) in the cDNA libraries (http:// flagdb-genoplante-info.infobiogen.fr). Only a few of the glycoside hydrolases have been shown to be expressed and generally they correspond to those already studied and reported in publications. The expression of others might be limited to particular tissues or specific conditions. 6. The cell-wall hydrolysing enzymes characterized in plants Putative functional properties of selected glycoside hydrolase families obtained by exploring available Arabidopsis sequences (Table 1) are based on published reports concerning functional properties of characterized glycoside hydrolases

from different organs of various plants (see below). This report describes glycoside hydrolases with known amino acid sequences and enzymatic properties that are, or could be, involved in modification and reorganization of plant cell wall polysaccharides. They are presented according to the polysaccharide they are hypothesized to degrade. 7. Degradation of Glucans and Callose Two classes of enzyme activities have been identified that are involved in the degradation of glucans and callose. Endoglucanases (EG) can hydrolyze glucans and callose to glucooligosaccharides. Exoglucanases degrade the oligosaccharides to D-glucose. The endoglucanases from plants show different substrate specificities and can be divided into three major groups: the (1→4)-β-D-glucanase (EC 3.2.1.74) or endocellulases, (1→3)-β-D-glucanase (EC 3.2.1.6) and (1→3,1→4)-β-D-glucanases (EC 3.2.1.73) [12,15]. A membrane bound endo-1,4-β-glucanase (cellulase) from Brassica napus showed higher substrate specificity for low substituted carboxymethyl-cellulose and amorphous cellulose but could hydrolyze crystalline cellulose, xyloglycan, xylan, (1→3)(1→4)-β-D-glucan and other polysaccharides and oligosaccharides [65]. This EG can degrade beta-glucans with pure β-(1→4)-linked backbone (cellulose) and is localized in the plasma membrane. It presents amino acid sequence similarity with a small gene family of three genes (KOR1, KOR2 and KOR3) in Arabidopsis [66,67] and is defined as an ortholog of korrigan. It was proposed that these membrane bound EG could play a role in cell wall biosynthesis. The involvement of KOR in plant cellulose biosynthesis has been suggested due to a lack of hypocotyl elongation and a reduction in cellulose content in the KOR mutant [66,68]. KOR enzymes are also thought to be involved in various developmental processes such as leaf abscission, fruit ripening [69] and cell expansion [66]. The membrane endo (1→4)-β-D-glucanases belongs to the family 9 glycoside hydrolases. Extracellular EG belong to the same family. Some extracellular EG were cloned and characterized in Arabidopsis [70], strawberry [71] and tomato [69]. They play specific roles in normal wall assembly and cell elongation, fruit ripening and floral abscission. The membrane and extracellular plant endo-glucosidases of family 9 have not been studied at the biochemical level. Only one report details the enzymatic specificity of a Brassica glucosidase ortholog of korrigan [65]. Family 17 of plant glycoside hydrolases contains distinct enzymes with high structural similarity: (1→3)-β-D-glucanases and (1→3,1→4)-β-D-glucanases [69]. The molecular studies suggested that these glucanases share a common ancestry. Moreover, it seems that (1→3,1→4)-β-D-glucanases diverged from (1→3)-β-D-glucanases during the appearance of the graminaceous monocotyledons [72]. (1→3)-β-D-Glucanases are abundant proteins found in all higher plants [73]. It has been proposed that (1→3)-β-D-glucanases are important for diverse physiological processes including pollen development, fertilization, mobilization of storage

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Table 1 List of the Arabidopsis thaliana glycoside hydrolases obtained by bioinformatic analyses which are potentially involved in cell wall polysaccharide degradation Genes Family 1 (total At1g02850 At1g52400 At1g61810 At1g60260 At1g60270 At1g60090 At1g66270 At1g66280 At1g75940 At2g44450 At2g44470 At2g44480 At3g03640 At3g21370 At3g18070 At3g18080 At3g60140 At3g62750 At4g22100 At4g27820 At4g27830 At5g24540 At5g24550 At5g54570 Family 3 (total At1g02640 At1g78060 At3g19620 At3g62710 At5g04885 At5g09730 At5g10560 At5g20940 At5g20950 At5g49360 At5g64570 Family 5 (total At1g13130 At2g20680 At3g10890 At3g10900 At3g26130 At3g30540 At4g28320 At5g01930 At5g16700 At5g17500 At5g66460 Family 9 (total At1g19940 At1g22880 At1g23210 At1g48930 At1g64390 At1g70710 At1g71380 At1g75680 At1g65610 At2g32990 At2g44540 At2g44560 At2g44570

Enzyme names 47: 24 outside or plasma membrane) beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase/beta-mannosidase beta-glucosidase/beta-mannosidase beta-glucosidase beta-glucosidase. beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase beta-glucosidase 15; 11 outsise or plasma membrane) beta-xylosidase beta-xylosidase beta-xylosidase beta-glucosidase beta-glucosidase beta-xylosidase beta-xylosidase beta-glucosidase beta-glucosidase α-arabinofuranosidase/β-xylosidase (XYL1) beta-xylosidase (XYL4) 13; 11 outside or plasma membrane) endo–glucanase/cellulase (1-4)-beta-mannan endohydrolase (1-4)-beta-mannan endohydrolase (1-4)-beta-mannan endohydrolase endo–glucanase/cellulase (1-4)-beta-mannan endohydrolase endo–glucanase/cellulase (1-4)-beta-mannan endohydrolase endo–glucanase/cellulase endo–glucanase/cellulase (1-4)-beta-mannan endohydrolase 25; 22 outside or plasma membrane) endo-beta-1,4-D-glucanase endo-beta-1,4-D-glucanase endo-beta-1,4-D-glucanase endo-beta-1,4-D-glucanase endo-beta-1,4-D-glucanase endo-beta-1,4-D-glucanase endo-beta-1,4-D-glucanase endo-beta-1,4-D-glucanase endo-beta-1,4-D-glucanase (KOR2) endo-beta-1,4-D-glucanase endo-beta-1,4-D-glucanase endo-beta-1,4-D-glucanase endo-beta-1,4-D-glucanase

E.C No.

Cell loc.

Score

n.d. 3.2.1.21 n.d. n.d. n.d. n.d. 3.2.1.21 n.d. n.d. n.d. n.d. n.d. n.d n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d.

outside outside outside outside outside outside outside outside outside outside outside plasma mem outside outside outside outside plasma mem outside outside outside outside outside outside plasma mem

0.820 0.820 0.820 0.375 0.480 0.814 0.820 0.820 0.820 0.820 0.676 0.685 0.820 0.685 0.370 0.370 0.685 0.820 0.820 0.820 0.633 0.700 0.619 0.685

n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. 3.2.1.55/3.2.1.37 3.2.1.37

outside plasma mem outside outside plasma mem plasma mem plasma mem plasma mem plasma mem outside outside

0.820 0.460 0.666 0.820 0.640 0.460 0.460 0.460 0.460 0.820 0.820

n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d.

plasma mem outside outside outside outside outside plasma mem plasma mem outside outside outside

0.811 0.820 0.776 0.820 0.790 0.820 0.685 0.685 0.370 0.820 0.820

n.d. n.d. n.d. n.d. n.d. 3.2.1.4 n.d. n.d. 3.2.1.4 n.d. n.d. n.d. n.d.

outside outside outside outside outside outside outside outside plasma mem outside plasma mem outside outside

0.820 0.370 0.556 0.820 0.820 0.820 0.604 0.820 0.790 0.820 0.919 0.690 0.820

Ref.

[82,83]

[82,84]

[102] [102]

[102] [102] [107,108] [107]

EST 17 85 6 10 0 0 40 8 4 4 11 3 3 14 0 37 16 4 1 2 9 0 0 1 15 9 0 3 9 7 12 3 26 92 31 1 5 1 0 0 0 1 10 0 0 5

[70]

[67]

5 3 0 7 29 19 4 39 5 12 4 2 2 (continued)

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Table 1 (continued) Genes Enzyme names At3g43860 endo-beta-1,4-D-glucanase At4g09740 endo-beta-1,4-D-glucanase At4g11050 endo-beta-1,4-D-glucanase At4g23560 endo-beta-1,4-D-glucanase At4g38990 endo-beta-1,4-D-glucanase At4g39000 endo-beta-1,4-D-glucanase At4g39010 endo-beta-1,4-D-glucanase At4g24260 endo-beta-1,4-D-glucanase (KOR3) At5g49720 endo-beta-1,4-D-glucanase (KOR1) Family 10 (total 12; 5 outside) At2g14690 1,4-beta-xylan endohydrolase At4g33820 1,4-beta-xylan endohydrolase At4g33840 1,4-beta-xylan endohydrolase At4g33860 1,4-beta-xylan endohydrolase At4g38650 1,4-beta-xylan endohydrolase Family 16 (total 33; 30 outside or plasma membrane) At1g14720 xyloglucan endotransglycosylase (XTR2) At1g11545 xyloglucan endotransglycosylase At1g32170 xyloglucan endotransglycosylase At1g65310 xyloglucan endotransglycosylase At2g01850 xyloglucan endotransglycosylase (EXGT-A3) At2g06850 xyloglucan endotransglycosylase (EXGT-A1) At2g14620 xyloglucan endotransglycosylase At2g18800 xyloglucan endotransglycosylase At3g23730 xyloglucan endotransglycosylase At3g25050 xyloglucan endotransglycosylase At3g48580 xyloglucan endotransglycosylase At4g03210 xyloglucan endotransglycosylase At4g14130 xyloglucan endotransglycosylase (XTR7) At4g13080 xyloglucan endotransglycosylase At4g13090 xyloglucan endotransglycosylase At4g18990 xyloglucan endotransglycosylase At4g25810 xyloglucan endotransglycosylase (XTR6) At4g25820 xyloglucan endotransglycosylase (XTR9) At4g28850 xyloglucan endotransglycosylase At4g30270 xyloglucan endotransglycosylase(Meri-5) At4g30280 xyloglucan endotransglycosylase At4g30290 xyloglucan endotransglycosylase At4g37800 xyloglucan endotransglycosylase At5g13870 xyloglucan endotransglycosylase(EXGT-A4) At5g48070 xyloglucan endotransglycosylase At5g57530 xyloglucan endotransglycosylase At5g57540 xyloglucan endotransglycosylase At5g57550 xyloglucan endotransglycosylase (XTR3) At5g57560 xyloglucan endotransglycosylase (TCH4) At5g65730 xyloglucan endotransglycosylase Family 17 (total 49; 26 outside or plasma membrane) At1g30080 endo-1,3-beta-glucanase At1g32860 endo-1,3-beta-glucanase At1g33220 endo-1,3-beta-glucanase At1g66250 endo-1,3-beta-glucanase At1g77780 endo-1,3-beta-glucanase At1g77790 endo-1,3-beta-glucanase At2g01630 endo-1,3-beta-glucanase At2g39640 endo-1,3-beta-glucanase At3g04010 endo-1,3-beta-glucanase At3g07320 endo-1,3-beta-glucanase At3g13560 endo-1,3-beta-glucanase At3g23770 endo-1,3-beta-glucanase At3g24330 endo-1,3-beta-glucanase At3g46570 endo-1,3-beta-glucanase At3g55430 endo-1,3-beta-glucanase At3g55780 endo-1,3-beta-glucanase At3g57260 endo-1,3-beta-glucanase

E.C No. n.d. n.d. n.d. n.d. n.d. n.d. n.d. 3.2.1.4 3.2.1.4

Cell loc. plasma mem plasma mem outside outside outside outside outside plasma mem plasma mem

Score 0.811 0.685 0.820 0.719 0.820 0.820 0.662 0.790 0.790

Ref.

[67] [66]

EST 4 0 1 8 0 3 4 0 64

n.d. n.d. n.d. n.d. n.d.

outside outside outside outside outside

0.820 0.556 0.657 0.461 0.820

[105] [105] [105] [105] [105]

3 4 1 0 3

2.4.1.207 n.d. n.d. n.d. 2.4.1.207 2.4.1.207 n.d. n.d. n.d. n.d. n.d. n.d. 2.4.1.207 n.d. n.d. n.d. 2.4.1.207 2.4.1.207 n.d. 2.4.1.207 n.d. n.d. n.d. 2.4.1.207 n.d. n.d. n.d. 2.4.1.207 2.4.1.207 n.d.

outside outside outside outside outside outside outside outside outside outside outside outside outside outside outside outside outside outside outside outside outside outside outside outside plasma mem outside outside outside plasma mem outside

0.757 0.576 0.685 0.820 0.820 0.820 0.820 0.820 0.542 0.466 0.820 0.820 0.743 0.709 0.428 0.820 0.820 0.820 0.614 0.800 0.719 0.652 0.370 0.820 0.685 0.820 0.820 0.566 0.685 0.820

[86,132]

22 14 10 1 17 54 6 0 10 2 2 11 26 0 0 1 13 9 0 43 13 10 9 1 4 4 14 14 26 17

n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. 3.2.1.39

plasma mem plasma mem plasma mem plasma mem outside outside plasma mem outside plasma mem outside plasma mem outside plasma mem plasma mem outside plasma mem outside

0.685 0.919 0.919 0.640 0.370 0.820 0.919 0.820 0.919 0.728 0.919 0.700 0.919 0.460 0.820 0.460 0.709

[86,132] [86,90,133]

[86]

[86] [86,90] [82,86]

[86,132]

[86] [86,90,134]

[135]

8 5 0 0 1 0 10 1 0 11 7 1 0 0 42 0 22 (continued)

Z. Minic, L. Jouanin / Plant Physiology and Biochemistry 44 (2006) 435–449

441

Table 1 (continued) Genes Enzyme names At3g57270 endo-1,3-beta-glucanase At3g61810 endo-1,3-beta-glucanase At4g16260 endo-1,3-beta-glucanase At4g17180 endo-1,3-beta-glucanase At4g34480 endo-1,3-beta-glucanase At5g20340 endo-1,3-beta-glucanase At5g20560 endo-1,3-beta-glucanase At5g42720 endo-1,3-beta-glucanase At5g58090 endo-1,3-beta-glucanase Family 27 (total 4; 4 outside or plasma membrane) At3g26380 alpha-galactosidase At3g56310 alpha-galactosidase At5g08370 alpha-galactosidase At5g08380 alpha-galactosidase Family 28 (total 66; 46 outside or plasma membrane) At1g02790 exopolygalacturonase-1 At1g05650 pectinase At1g05660 pectinase At1g23460 pectinase At1g70500 pectinase At1g56710 pectinase At1g65570 pectinase At1g78400 pectinase At1g80170 pectinase At2g23900 pectinase At2g33160 pectinase At2g41850 endo-polygalacturonase At1g43090 pectinase At1g43100 pectinase At2g43860 pectinase At2g43870 pectinase At2g43890 pectinase At3g06770 pectinase At3g07820 pectinase At3g07830 pectinase At3g07840 pectinase At3g07850 exopolygalacturonase At3g07970 pectinase At3g14040 endo-polygalacturonase At3g15720 pectinase At3g16850 pectinase At3g42950 pectinase At3g48950 pectinase At3g57510 endo-polygalacturonase At3g57790 pectinase At3g59850 pectinase At3g61490 pectinase At3g62110 pectinase At4g01890 pectinase At4g18180 pectinase At4g23500 pectinase At4g23820 pectinase At4g33440 pectinase At4g35670 pectinase At5g14650 pectinase At5g17200 pectinase At5g27530 pectinase At5g39910 pectinase At5g41870 pectinase At5g44840 pectinase At5g48140 pectinase Family 31 (total 5; 4 outside) At1g68560 alpha-xylosidase (XYL1) At3g45940 alpha-xylosidase

E.C No. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d.

Cell loc. outside outside outside outside plasma mem plasma mem outside plasma mem plasma mem

Score 0.571 0.628 0.614 0.728 0.460 0.460 0.514 0.919 0.919

Ref.

n.d. n.d. n.d. n.d.

outside outside outside plasma mem

0.690 0.820 0.537 0.685

3.2.1.67 n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. 3.2.1.67 n.d. 3.2.1.15 n.d. n.d. n.d. n.d. 3.2.1.15 n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d. n.d.

outside plasma mem plasma mem outside outside outside plasma mem outside outside outside outside outside plasma mem plasma mem outside outside outside outside outside outside outside outside outside outside plasma mem outside plasma mem outside plasma mem outside outside outside outside outside outside outside outside plasma mem outside outside outside plasma mem outside outside outside outside

0.810 0.685 0.685 0.820 0.685 0.475 0.685 0.820 0.82 0.748 0.820 0.642 0.640 0.640 0.820 0.820 0.762 0.820 0.771 0.690 0.820 0.820 0.820 0.820 0.919 0.820 0.685 0.820 0.650 0.432 0.781 0.820 0.599 0.820 0.461 0.537 0.518 0.790 0.820 0.676 0.781 0.790 0.547 0.623 0.820 0.705

[136]

3.2.1.20 n.d.

outside outside

0.820 0.820

[88,102]

EST 1 0 52 3 1 0 15 11 17 9 30 7 8

[136] [136]

[137]

18 0 0 0 0 0 0 0 2 0 0 2 0 0 0 0 0 12 11 3 0 11 1 11 1 18 9 0 5 5 3 6 13 1 0 4 20 5 0 2 0 0 0 3 0 5 56 0 (continued)

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Table 1 (continued) Genes Enzyme names E.C No. Cell loc. Score Ref. EST At5g11720 alpha-glucosidase AGLU1 3.2.1.20 outside 0.805 [138] 11 At5g63840 alpha-glucosidase/xylosidase n.d. outside 0.685 17 Family 35 (total 18; 12 outside) At1g31740 beta-galactosidase n.d. outside 0.820 [102] 0 At1g72990 beta-galactosidase n.d. outside 0.820 [102] 8 At1g77410 beta-galactosidase n.d. outside 0.820 [102] 3 At2g16730 beta-galactosidase n.d. outside 0.820 [102] 3 At2g28470 beta-galactosidase n.d. outside 0.820 [102] 8 At2g32810 beta-galactosidase n.d. outside 0.820 [102] 12 At3g13750 beta-galactosidase n.d. outside 0.820 [102] 132 At3g52840 beta-galactosidase n.d. outside 0.571 [102] 16 At4g26140 beta-galactosidase n.d. outside 0.820 [102] 5 At5g56870 beta-galactosidase n.d. outside 0.820 [102] 26 At5g63800 beta-galactosidase n.d. outside 0.820 [102] 12 At5g63810 beta-galactosidase n.d. outside 0.819 [102] 6 Family 43 (total 2; 2 outside) At3g49880 1,4-beta-xylan endohydrolase n.d. plasma mem 0.640 4 At5g67540 1,4-beta-xylan endohydrolase n.d. plasma mem 0.790 10 Family 51 (total 2; 2 outside or plasma membrane) At3g10740 α-L-arabinofuranosidase/β-D-xylosidase 3.2.1.55/3.2.1.37 outside 0.752 [107,139] 30 (ARAf) At5g26120 alpha-L-arabinofuranosidase n.d. outside 0.561 [139] 10 Database accession numbers of the genes are shown. The main determined enzymatic activities reported for each family are indicated with corresponding references. Only the ones with EC numbers have been tested for activity. The number of expressed sequence tags (ESTs) reported in databases is given.

reserves in the endosperm, and cell division [73]. The (1→3)β-D-glucanases are believed to play a major role in plant protection against pathogen attack. These enzymes are classified among pathogenesis-related (PR) proteins [74]. Specific RNA accumulation after virus infection in Arabidopsis was observed for one glucanase [74]. (1→3)-β-D-glucanase is synthesized in microsporocytes before meiosis in angiosperms. The biological role of (1→3)-β-D-glucanase in relation to callose has not been clearly established but Worrall et al. [12] demonstrated that premature callose degradation in tobacco is sufficient to cause male sterility and suggested that callose is essential for the formation of a normal microspore cell wall. It has been established that class I (1→3)-β-D-glucanases in tobacco are involved in endosperm rupture by weakening the endosperm [75]. In contrast to (1→3)-β-D-glucanases, (1→3,1→4)-β-glucanases are involved in cell wall degradation in graminaecous monocotyledons and in elongation of vegetative tissues [3, 72]. These enzymes have been purified and characterized from germinated barley [76]. Members of this endoglucanase group can specifically hydrolyze xyloglucan and the mixedlink (1→3)(1→4)-β-D-glucan. Such enzymes are abundant in the growing cell walls of dicot and grass seedlings [3]. Cleavage of glucan polymers by endoglucanases into smaller fragments facilitates its complete hydrolysis to D-Glucose by Exoglucanases (EC 3.2.1.21). Plants possess a large number of exoglucanases (most of the 48 genes identified as putative beta-glucosidase genes in Arabidopsis thaliana do not have a known function). These enzymatic activities were detected in plants and classified in the families 1 or 3 of glycoside hydrolases [15]. Examination of substrate specificity reveals that the barley enzymes from family 1 exhibit preference for (1→4)-βD-oligoglucosides and the rate of hydrolysis increased with the

degree of polymerization of the substrate [77–79]. In contrast, the family 3 exoglucanases exhibits broad substrate specificity and can hydrolyze the non-reducing terminal glucosidic linkage of polymeric β-D-glucans and β-D-oligoglucosides containing (1→2)-, (1→3)-, (1→4)- or (1→6)-linkages and in some β-D-oligoxyloglucosides [79–81]. The broad specificity for glycosidic linkages suggests that these enzymes could be involved in diverse functions during plant development. Both families 1 and 3 of exoglucanases can hydrolyze aryl β-D-glucosides such as 4-NP-β-D-glucopyranoside. In addition, family 3 of exo-β-D-glucanases exhibits transglycosylase activities and generated tri- and tetrasaccharides from disaccharides [79–81]. One exo-β-D-glucanase which is associated with the external surface of the plasma membrane has been purified and characterized from maize seedlings [81]. Exoglucanases are involved in plant development processes but can also be induced in response to biotic and abiotic stress [77,82]. In Arabidopsis, genes encoding exo-β-D-glucanases (β-D-glucosidases) have been studied in the context of plant defense against insect attack [83] and during phosphate deprivation [84]. The precise role of exo-β-D-glucanases in cell wall degradation is not clear but these enzymes were shown to possess hydrolytic activity towards native cell wall substrates [85]. 8. Degradation of Xyloglucan Enzymes presumed to be involved in the modification of xyloglucan structure are xyloglucan specific endoglucanases including xyloglucan endo-transglycosylase/hydrolases (XTH)(2.4.1.207) [5,86], a xyloglucan-active β-D-galactosidase (3.2.1.23) [87], a xyloglucan specific α-L-fucosidase (3.2.1.51/3.2.1.63) and a xyloglucan oligosaccharide-specific α-D-xylosidase (3.2.1.37) [88].

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XTHs appear to play a major role in plant growth and development because they are probably involved in the construction and restructuration of the plant cell wall [5]. XTH enzymes exhibit either xyloglucan endotransglycosylation (XET) or xyloglucan hydrolysis (XEH) activities or both in vitro. The majority exhibit XET activity, some exhibit both XET and XEH activities and some exhibit only XEH activity [86]. Kinetic and enzymatic studies suggest that the XET reaction uses two substrates (donor and acceptor) involved in a ping-pong mechanism [5,89]. The enzyme cleaves a donor xyloglucan to release a smaller xyloglucan, with a reducing end in the first step, then transfers the remainder to the acceptor, an oligosaccharide. The properties and function of XTHs have been reviewed in several papers [5,86,90]. XTHs in plants are assigned to family 16 of glycoside hydrolases. The structure and organization of the XTH gene family were analyzed in Arabidopsis and 33 open reading frames (ORFs) potentially encoding XTH proteins that were divided into three major phylogenetic groups or subfamilies were identified [86]. In addition, it appears that β-D-glucosidases, exoglucanases from the family 3 glucoside hydrolases (see above for degradation of glucan), can also hydrolyse xyloglucan oligosaccharides. Purified β-D-glucosidases from the cotyledons of germinated nasturtium exhibited broad specificity for glycosidic linkage like other enzymes of the same family [91]. This enzyme has been shown to hydrolyse glucose disaccharides with different linkages at different rates (1→3), (1→4), (1→2) (1→6) and also exhibited transglycosylase activity [91,92]. β-D-Glucosidase from nasturtium shows strong sequence similarity with a β-D-glucan exo-hydrolase from barley [92]. The cotyledons of germinated nasturtium seeds were used for purification and characterization of a xyloglucan specific β-D-galactosidase [87]. This enzyme catalyzes removal of the terminal non-reducing β-D-galactopyranosyl residues from xyloglucans but cannot release the terminal non-reducing αD-galactopyranosyl residues from seed galactomannans. Other purified β-D-galactosidases have not been reported to hydrolyze xyloglucans, probably because their activity on xyloglucan has not been tested. Usually β-D-galactosidase activities are probed using synthetic substrates such as p-NP-galactopyranoside, 4-methylumbelliferyl-β-D-galactopyranoside, X-Gal or natural substrates such as (1→4)-β-D-galactan [93]. Generally, these enzyme activities belong to family 35 glycoside hydrolases. α-L-Fucosidase activity was detected in crude enzyme extracts of growing region of etiolated pea stems and in cotyledons of germinating nasturtium seedlings [94]. An α-L-fucosidase from pea epicotyls has been purified and characterized [95]. It hydrolyzed the terminal α-(1→2)-fucosidic linkage of oligosaccharides and did not cleave p-NP-α-L-fucoside. In addition, the enzyme did not release measurable amounts of fucosyl residues from large polysaccharides [96]. The amino acid sequence deduced from mRNA was not similar to other known α-L-fucosidases, but was strongly homologous to Kunitz-type trypsin inhibitors [96]. The cDNA clones encoding

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this protein were isolated and expressed in Escherichia coli, insect cells and Arabidopsis thaliana cells, and produced recombinant proteins which did not exhibit α-L-fucosidase activity [97]. Another α-L-fucosidase with similar substrate specificity was purified from cabbage (Brassica oleracea) leaves [98]. Two peptide sequences obtained for this protein were used to identify Arabidopsis genes coding for α-L-fucosidase. One gene identified was heterologously expressed in Pichia pastoris cells. This recombinant α-L-fucosidase was active against the oligosaccharides from xyloglucan XXFG as well as against commercial milk oligosaccharide 2′-fucosyl-lactitol but not against p-NP-α-L-fucopyranoside. Its amino acid sequence is similar to lipases. Another α-L-fucosidase, homologous to known α-L-fucosidases was expressed in Pichia pastoris cells. Its belongs to family 29 of glycoside hydrolases. This enzyme only possesses activity against 2′-fucosyl-lactitol [98]. Finally, α-D-xylosidase cleaves specifically the α-xylosyl residue attached to the glucose residue of the xyloglucanoligosaccharide [99–101]. Plant α-D-xylosidases do not hydrolyze the artificial substrate p-NP-α-D-xylopyranoside, but hydrolyze natural xyloglucan oligosaccharides. Two plant αD-xylosidases were sequenced, one from nasturtium [80] and another from Arabidopsis leaves [88]. Their amino acid sequences indicated that these plant α-D-xylosidases are members of family 31 glycoside hydrolases. The amino acid sequences exhibit significant homology with apoplastic α-Dglucosidases. α-D-Glucosidase activity from the nasturtium enzyme was demonstrated to degrade several α-(1→4)- and α-(1→6)-linked substrates [80] while Arabidopsis enzyme activity was demonstrated against p-NP-α-D-glucoside [88]. Recently, analyses of various Arabidopsis tissues using quantitative real-time reverse transcription-PCR allowed the identification of genes coding for enzymes involved in the degradation of xyloglucan oligosaccharides. In this study, genes coding for α-L-fucosidase (At1g67830), α-D-xylosidase (At1g68560), 12 putative β-D-galactosidases and four β-D-glucosidases were identified [102]. 9. Degradation of Xylan and Arabinoxylan Three classes of plant enzymes which degrade xylan were identified: endoxylanases (EC 3.2.1.8), β-D-xylosidases (EC 3.2.1.37) and α-L-arabinofuranosidases (3.2.1.55). Endoxylanases and β-D-xylosidases are enzymes responsible for cleavage of xylan backbone groups while α-L-arabinofuranosidase removes side-chain arabinose substituents from xylan or oligoxylan. Endo-β-1,4-xylanases hydrolyze the insoluble xylan backbone into shorter, soluble xylo-oligosaccharides, while βD-xylosidases hydrolyze xylo-oligosaccharides and xylobiose from their nonreducing ends to release D-xylose [103]. Endoxylanase activities were identified in several fruit and higher plants [44,104,105]. In barley two xylan endohydrolases have been purified, characterized and corresponding cDNAs and genes identified [44]. These enzymes belong to family 10 glycoside hydrolases. Four putative xylanase genes identified

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in the Arabidopsis genome belong to family 10 glycoside hydrolases [105]. In addition, in the Arabidopsis genome two putative xylanases were identified that belong to family 43 of glycoside hydrolases. However, the enzymatic properties of these enzymes have not yet been demonstrated. β-D-xylosidase is a key enzyme for the complete degradation of xylan. One β-D-xylosidase from young barley seedlings (Hordeum vulgare) and one from Arabidopsis thaliana stems were purified, sequenced and biochemically characterized [106,107]. The characterized β-D-xylosidases released Dxylose from various natural cell wall polysaccharides and oligosaccharides or synthetic substrates. These two enzymes belong to family 3 of glycoside hydrolases. Two bifunctional α-L-arabinofuranosidase/β-D-xylosidases belonging to family 3 of glycoside hydrolases were identified from barley seedlings and Arabidopsis stems [106,107]. Bifunctional α-L-arabinofuranosidase/β-D-xylosidases released D-xylose less efficiently than β-D-xylosidase from oat spelt xylan (OSX), oligoxylan, arabinoxylans, and synthetic substrate pNP-β-xyloside. In contrast, this enzyme released Larabinose more efficiently than β-D-xylosidase from arabinan (SBA), arabinoxylan (WAX and RAX), oligoarabinoxylans (obtained from WAX and RAX) and synthetic substrate pNPα-L-arabinofuranoside. An Arabidopsis bifunctional α-L-arabinofuranosidase/β-D-xylosidase has been proposed to be involved in secondary cell wall metabolism during plant development [108]. An α-L-arabinofuranosidase was purified from stem tissues of Arabidopsis and shown to belong to glycoside hydrolase family 51. This enzyme shows similar hydrolytic properties to bifunctional α-L-arabinofuranosidase/β-D-xylosidases from family 3 glycoside hydrolase when tested on various natural and synthetic substrates [107]. In addition, two α-L-arabinofuranosidases from family 51, that remove α-L-arabinofuranosyl residues from the arabinoxylan polymer, have been purified, sequenced and characterized from barley seedlings [106,109]. These enzymes have a relatively broad substrate specificity but prefer arabinoxylan as a substrate [109]. 10. Degradation of the galactomannan The galactomannans in the endosperm cell walls of tomato and lettuce are hydrolysed during seed germination. This degradation depends of endo-β−mannanases (EC 3.2.1.78), βD-mannosidases (EC 3.2.1.25) and α-D-galactosidases (3.2.122) [17]. Endo-β−mannanases, hydrolyze the mannan backbone releasing manno-oligosaccharides which are hydrolyzed further by β-D-mannosidases and α-D-galactosidases to remove the galactose unit on side-chains. An endo-β-mannanase was partially purified then sequenced from coffee seeds [110]. This coffee cDNA expressed in Escherichia coli produced an enzyme that exhibited endo-βmannanase activity using bean galactomannan. Mannobiose and mannotriose were released by this endo-β−mannanase predominantly from mannan [111]. Characterized plant endo-

β−mannanases are members of the family 5 glycoside hydrolases. The activity of β-D-mannosidases from many plant sources has been investigated. Nonetheless, only recently a β-D-Mannosidase from tomato seeds was purified, characterized and sequenced [111]. This enzyme was able to hydrolyse synthetic pNP-β-D-manopyranoside and belongs to family 1 glycoside hydrolases. Complete degradation of galactomannan depends on the action of α-D-galactosidase. The α-D-galactosidase from rice cell cultures was purified to homogeneity, cloned and expressed in E. coli [112]. This rice α-D-galactosidase hydrolyzes melibiose, raffinose, stachyose and galactomannooligosaccharide. These enzymes belong to family 27 glycoside hydrolases. 11. Degradation of Homogalacturonan Three classes of enzymes involved in the degradation of homogalacturonan (HG) are known: endopolygalacturonases (EC 3.2.1.15), exopolygalacturonases (EC 3.2.1.67) and pectin methyl esterases (EC 3.2.2.11) [29,113]. The backbone of the HG can be hydrolyzed by endo-polygalacturonases and terminally cleaved by exo-polygalacturonases. It has been suggested that these two enzymes play an important role in the degradation of acidic polymers in cell walls and allow cellcell adhesion [114]. Pectin methyl esterase catalyzes the hydrolysis of methyl ester groups on cell wall pectins. Modification of the cell wall by endopolygalacturonases (EndoPGs) has been studied intensively in the context of fruit ripening and softening [115]. EndoPGs have been purified and characterized from Capsicum [115] and tomato [116]. The EndoPG from avocado was sensitive to methylation of the substrate polyuronide. In addition, a highly methylesterified polyuronide was resistant to the action of endoPG [116]. The role of exopolygalacturonase (ExoPG) has been proposed for various physiological functions such as plant growth and development, fruit ripening, plant microorganism symbiosis, defense and production of potential elicitors [117]. ExoPG purified from carrot was not active when expressed in Escherichia coli [117]. This exo-PGase showed 49.3% sequence identity with the ENOD8 gene product of alfalfa (Medicago sativa). These two classes of enzymes (endopolygalacturonase and exopolygalacturonase involved in the degradation of HG) belong to family 28 glycoside hydrolases. Pectin methyl esterases have been implicated in a number of processes including cell growth, fruit ripening, abscission and senescence, pathogenesis, cambial cell differentiation, dormancy termination, germination and stem elongation [118]. Biochemical and physiological studies and possible functions of these enzymes were reviewed recently [118]. 12. Degradation of RG-I So far, only two enzyme activities are known to participate in hydrolysis of RG-I: bifunctional α-L-arabinosidases/β-xylo-

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sidases (EC.3.2.1.55/3.2.1.37) and β-D-galactosidase (3.2.1.23), that can be involved in degradation of arabinan and galactan, respectively [107,119–121]. Bifunctional α-L-arabinosidases/β-xylosidases from families 3 and 51 isolated from Arabidopsis stems are able to release Larabinose from arabinan [107]. β-D-galactosidases have been purified from various plants [5,119] and are associated with ripening processes, pectin solubility, cell wall porosity, cell wall rigidity and other processes required for cell wall growth and development [120]. Purified β-D-galactosidases were shown to participate in the removal of galactose from galactans and other cell wall components [93,119,120]. A purified β-Dgalactosidase from the cotyledons of germinated Lupinus angustifolius L. seeds was demonstrated to degrade galactooligosaccharides [122]. An exo-(1→4)-β-D-galactanase was purified, characterized and sequenced from ripe tomato fruit. This enzyme was specific for (1→4)-β-D-galactan indicating that the enzyme was an exo-(1→4)-β-D-galactanase [123]. Purified β-D-galactosidase from Cicer arietinum was able to degrade the galactan side chain of cell walls during the cell wall loosening that occurs prior to cell elongation [124]. The β-D-galactosidase from ripe carambola fruit was purified and characterized [125]. This enzyme was able to hydrolyse β(1→4)- and a mixture of β(1→3)- and β(1→6)-galactan. β-D-Galactosidases might be involved in degradation of glycoproteins. Recently, an β-D-galactosidase was purified and characterized in Raphanus sativus [126]. This enzyme was able to hydrolyse both β(1→3)- and β(1→6) galactosyl residues from the carbohydrate chains of arabinogalactan proteins (AGPs). These purified and characterized β-D-galactosidases belongs to family 35 glycoside hydrolases. 13. Degradation of other polysaccharides Plant enzymes that participate in hydrolysis of xylogalacturonan (XGA) and RG-II have not been identified. With regards to XGA degradation, it can be supposed that β-D-xylosidases from family 3 glycoside hydrolases could participate in its side chain debranching. Likewise, it is to be expected that many enzymes participate in the hydrolysis of RG-II because of its complex structure, composed of more than 20 different glycosidic linkages [34]. Some enzymatic activities such as a β-Dglucuronidase from rye [127] and an α-L-arabinopyranosidase from Arabidopsis (our unpublished data) have been detected but their amino acid sequences remain to be identified. 14. Concluding remarks and perspectives Recently many studies have reported the identification and characterization of plant enzymes that play a role in the reorganization of the cell wall. In this review, we present the large number of plant glycoside hydrolases that may be involved in the modification of cell wall polysaccharides. These selected glycoside hydrolases were chosen from published reports and by exploring available sequence information from the Arabidopsis genome [61]. Based on the derived amino acid

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sequences, the gene products have been assigned to different families of glycoside hydrolases [59]. The enzyme families identified cover most of the activities that are necessary for the modification and reorganization of the various plant cell wall polysaccharides. However for many of these enzymes, the detailed analysis of their physiological roles remain to be performed. This is particularly true for the enzymes that might be involved in the reorganization of RG-I and RG-II. Different strategies (biochemical, genetic, proteomic and micro-array approaches) need to be used for their identification. Proteomic and cDNA microarrays analyses of different plant organs at various stages of development can be used to evaluate eventual enzyme candidates for further investigation [86]. A genetic approach using knockout mutants could be of interest although the presence of large multigene families for each enzyme-type may be a problem. Information concerning the pattern of gene expression should help in choosing the candidate gene and may indicate phenotypes to be tested. Purification of the enzymes or heterologous expression will no doult be important for the determination of substrate specificity. However, the absence of commercially available, appropriate, natural or synthetic substrates (example: α-D-glucuronidase) could be a problem. In addition, some glycoside hydrolases might have different enzymatic activities when tested with natural oligo- and/ or polysaccharides than with synthetic substrates [4]. For example, some plant α-D-xylosidases act on the oligosaccharide xyloglucan to remove a single xylose residue from the reducing terminus, but do not hydrolyse the synthetic substrate 4-NP-α-D-xylopyranoside [100,101]. Enzymatic assays using synthetic substrates are very easy to perform. In contrast, enzymatic assays using natural substrates are very difficult since they involve chromatographic steps (HPLC). New and easy ways to perform biochemical assays with adequate substrates are necessary for the identification of novel enzymatic activities. A major advance would be if appropriate substrates could be used to visualized in vivo specific enzymatic activities. In addition, different glycoside linkages are observed in plant cell wall polysaccharides. For instance, L-arabinose can be substituted with α-(1→2), α-(1→3) and α-(1→5) glycoside linkages in xylan, arabinoxylan and arabinan polysaccharides. Similarly, D-galactose, D-glucose, D-xylose can have different glycoside linkages. For most of the characterized glycoside hydrolases, their enzymatic specificity using various polysaccharides and oligosaccharides representing the possible linkages remains to be established. Furthermore, the activity of endopolysaccharidases strongly depends on the componants of the backbone chain [124] and this specificity is not clearly determined for some of them (e.g. xylanase). Modification and re-structuration of cell wall polysaccharides might be regulated according to tissue and development stages and the expression profiles of glycoside hydrolases need to be determined. In addition, the enzyme activities must be coordinated for the correct development of the plant [2,9,10,42,43]. It will be of interest to compare the mechanisms of plant cell wall degradation with those occurring in microorganisms. Many microorganisms degrade plant polysaccharides

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using the synergistic and cooperative action of several glycoside hydrolases which cleave different linkages [128,129,130]. For example, the degradation of xylan by hydrolytic enzymes from Aspergillus involves a main-chain-cleaving enzyme and one or more accessory enzymes [129]. It is possible that similar synergistic action occurs in plant cell wall modification [131]. In addition, according to the plant species and the abundance of hemicelluloses, the network could be different [63]. The understanding of the mode of action of plant hydrolytic enzymes will have uses in practical applications. Plant cell wall polysaccharides are abundant and their biodegradability is an important potential source for the synthesis of new compounds. Plant enzymes able to degrade polysaccharides could be used in industrial processes such as the biobleaching of pulp in the paper industry, clarification of fruit juices, preparation of animal feed, the cosmetic and pharmaceutical industries, fabrication of sugar beet syrup, caramel and fruit flavours. Industrial processes use glycoside hydrolase enzymes originating from microorganisms or fungi, but enzymes of plant origin could be useful in these industrial applications in the future. Currently, the industrial use of plant enzymes is limited by their availability and price. In addition, in contrast to plant enzymes, microbial industrial enzymes are characterized by pH and thermal stability. For these reasons plant enzymes have not yet replaced microbial enzymes in industrial applications. However, it is possible that with appropriate genetic engineering and heterologous expression of mutated proteins, enzymes of plant origin could be produced in microorganisms on an industrial scale.

[11] [12]

[13]

[14]

[15]

[16]

[17]

[18]

[19] [20] [21]

[22]

[23]

Acknowledgements

[24]

The authors wish to thank Deborah Goffner and Helen North for their critical review of the manuscript.

[25]

References [1]

[2] [3] [4] [5] [6] [7]

[8]

[9] [10]

G.P. Hazlewood, H.J. Gilbert, Structure and function analysis of Pseudomonas plant cell wall hydrolases, Prog. Nucleic Acid Res. Mol. Biol. 61 (1998) 211–241. D.J. Cosgrove, Assembly and enlargement of the primary cell wall in plants, Annu. Rev. Cell Dev. Biol. 13 (1997) 171–201. D.J. Cosgrove, Enzymes and other agents that enhance cell wall extensibility, Annu. Rev. Plant Physiol. Plant Mol. Biol. 50 (1999) 391–417. S.C. Fry, Polysaccharide-modifying enzymes in the plant cell wall, Annu. Rev. Plant Physiol. Plant Mol. Biol. 46 (1995) 497–520. S.C. Fry, Primary cell wall metabolism: tracking the careers of wall polymers in living plant cells, New Phytol. 161 (2004) 641–675. N.C. Carpita, Structure and biogenesis of the cell walls of grasses, Annu. Rev. Plant Physiol. Plant Mol. Biol. 47 (1996) 445–476. N.C. Carpita, D.M. Gibeaut, Structural models of primary cell walls in flowering plants: consistency of molecular structure with the physical properties of the walls during growth, Plant J. 3 (1993) 1–30. E.J. Mellerowicz, M. Baucher, B. Sundberg, W. Boerjan, Unraveling cell wall formation in the woody dicot stem, Plant Mol. Biol. 47 (2001) 239–274. A. Heredia, A. Jimenez, R. Guillen, Composition of plant cell walls, Z. Lebensm. Unters. Forsch. 200 (1995) 24–31. Z.A. Popper, S.C. Fry, Primary cell wall composition of bryophytes and charophytes, Ann. Bot. (Lond.) 91 (2003) 1–12.

[26]

[27] [28] [29]

[30]

[31] [32]

[33]

[34]

L. Ostergaard, M. Petersen, O. Mattsson, J. Mundy, An Arabidopsis callose synthase, Plant Mol. Biol. 49 (2002) 559–566. D. Worrall, D.L. Hird, R. Hodge, W. Paul, J. Draper, R. Scott, Premature dissolution of the microsporocyte callose wall causes male sterility in transgenic tobacco, Plant Cell 4 (1992) 759–771. M.S. Doblin, I. Kurek, D. Jacob-Wilk, D.P. Delmer, Cellulose biosynthesis in plants: from genes to rosettes, Plant Cell Physiol. 43 (2002) 1407–1420. G.B. Fincher, Cell wall metabolism in barley, in: P.R. Shewry (Ed.), Barley: Genetics, Biochemistry, Molecular Biology and Biotechnology, CAB International, UK, 1992, pp. 413–437. M. Hrmova, G.B. Fincher, Structure-function relationships of beta-Dglucan endo- and exohydrolases from higher plants, Plant Mol. Biol. 47 (2001) 73–91. M.S. Buckeridge, C. Rayon, B. Urbanowicz, M.A.S. Tine, N.C. Carpita, Mixed linkage (1→3),(1→4)-ß-D-glucans of grasses, Cereal Chem. 81 (2004) 115–127. J.D. Bewley, R.A. Burton, Y. Morohashi, G.B. Fincher, Molecular cloning of a cDNA encoding a (1→4)-beta-mannan endohydrolase from the seeds of germinated tomato (Lycopersicon esculentum), Planta 203 (1997) 454–459. M.G. Handford, T.C. Baldwin, F. Goubet, T.A. Prime, J. Miles, X. Yu, P. Dupree, Localisation and characterisation of cell wall mannan polysaccharides in Arabidopsis thaliana, Planta 218 (2003) 27–36. R.A. Prade, Xylanases: from biology to biotechnology, Biotechnol. Genet. Eng. Rev. 13 (1996) 101–131. N. Kulkarni, A. Shendye, M. Rao, Molecular and biotechnological aspect of xylanases, FEMS Microbiol. Rev. 23 (1999) 411–456. M.S. Izydorczyk, C.G. Biliaderis, Cereal arabinoxylans: advances in structure and physicochemical properties, Carbohydr. Polym. 28 (1995) 33–48. R.J. Viëtor, S.A.G.F. Angelino, A.G.J. Voragen, Structural features of arabonoxylans from barley and malt cell wall material, J. Cereal Sci. 15 (1992) 213–222. K. Iiyama, T. Lam, B.A. Stone, Covalent Cross-Links in the Cell Wall, Plant Physiol. 104 (1994) 315–320. M. Madson, C. Dunand, X. Li, R. Verma, G.F. Vanzin, J. Caplan, D.A. Shoue, N.C. Carpita, W.D. Reiter, The MUR3 gene of Arabidopsis encodes a xyloglucan galactosyltransferase that is evolutionarily related to animal exostosins, Plant Cell 15 (2003) 1662–1670. D.U. Lima, M.S. Buckeridge, Interaction between cellulose and storage xyloglucans: the influence of the degree of galactosylation, Carbohydr. Polym. 46 (2001) 157–163. M.C. McCann, K. Roberts, Architecture of the primary cell wall, in: C.W. Lloyd (Ed.), The cytoskeletal basis of plant growth and form, Academic Press, Inc, New York, N.Y, 1991, pp. 109–129. P.M. Dey, Biochemistry of plant galactomannans, Adv. Carbohydr. Chem. Biochem. 35 (1978) 3411–3476. P. Capek, J. Alfoldi, D. Liskova, An acetylated galactoglucomannan from Picea abies L, Karst. Carbohydr. Res. 337 (2002) 1033–1037. W.G. Willats, L. McCartney, W. Mackie, J.P. Knox, Pectin: cell biology and prospects for functional analysis, Plant Mol. Biol. 47 (2001) 9– 27. J.P. Vincken, H.A. Schols, R.J. Oomen, M.C. McCann, P. Ulvskov, A.G. Voragen, R.G. Visser, If homogalacturonan were a side chain of rhamnogalacturonan I. Implications for cell wall architecture, Plant Physiol. 132 (2003) 1781–1789. M.C. Jarvis, Structure and properties of pectin gels in plant cell walls, Plant Cell Environ. 7 (1984) 153–164. H.A. Schols, E. Vierhuis, E.J. Bakx, A.G. Voragen, Different populations of pectic hairy regions occur in apple cell walls, Carbohydr. Res. 275 (1995) 343–360. J. Visser, A.G.J. Voragen, Progress in Biotechnology 14: Pectins and Pectinases. Proceedings of an International Symposium, Wageningen. Elsevier, Amsterdam. Science BV, Amsterdam, 1996 [990 pp]. M.A. O’Neill, T. Ishii, P. Albersheim, A.G. Darvill, Rhamnogalacturonan II: structure and function of a borate cross-linked cell wall pectic polysaccharide, Annu. Rev. Plant Biol. 55 (2004) 109–139.

Z. Minic, L. Jouanin / Plant Physiology and Biochemistry 44 (2006) 435–449 [35]

[36] [37]

[38]

[39]

[40]

[41]

[42]

[43] [44]

[45]

[46] [47]

[48] [49]

[50]

[51] [52]

[53]

[54]

M. Kobayashi, H. Nakagawa, T. Asaka, T. Matoh, Boraterhamnogalacturonan II bonding reinforced by Ca2+ retains pectic polysaccharides in higher-plant cell walls, Plant Physiol. 119 (1999) 199– 203. H. Höfte, Plant biology. A baroque residue in red wine, Science 294 (2001) 795–797. T. Bach Tuyet Lam, K. Iiyama, B.A. Stone, Cinnamic acid bridges between cell wall polymers in wheat and phalaris internodes, Phytochemistry 31 (1992) 1179–1183. T. Imamura, K.M. Watanabe, T. Koshijima, Ester linkages between lignin and glucuronic acid in lignin-carbohydrate complexes from Fagus crenata, Phytochemistry 37 (1994) 1165–1173. A. Oosterveld, J.H. Grabber, G. Beldman, A.G.J. Voragen, Formation of ferulic acid dehydrodimers through oxidative cross-linking of sugar beet pectin, Carbohydr. Res. 300 (1997) 179–189. J.E. Thompson, S.E. Fry, Evidence for covalent linkage between xyloglucan and acidic pectins in suspension-cultured rose cells, Planta 211 (2000) 275–286. A. Femenia, N.M. Rigby, R.R. Selvendran, K.W. Waldron, Investigation of the occurrence of pectic-xylan-xyloglucan complexes in the cell walls of cauliflower stem tissues, Carbohydr. Polym. 39 (1999) 151– 164. T. Stolle-Smits, J.G. Beekhuizen, M.T. Kok, M. Pijnenburg, K. Recourt, J. Derksen, A.G. Voragen, Changes in cell wall polysaccharides of green bean pods during development, Plant Physiol. 121 (1999) 363–372. W.D. Reiter, Biosynthesis and properties of the plant cell wall, Curr. Opin. Plant Biol. 5 (2002) 536–542. M.P. Caspers, F. Lok, K.M. Sinjorgo, M.J. van Zeijl, K.A. Nielsen, V. Cameron-Mills, Synthesis, processing and export of cytoplasmic endobeta-1,4-xylanase from barley aleurone during germination, Plant J. 26 (2001) 191–204. W.D. Reiter, C. Chapple, C.R. Somerville, Mutants of Arabidopsis thaliana with altered cell wall polysaccharide composition, Plant J. 12 (1997) 335–345. M. Fagard, H. Hofte, S. Vernhettes, Cell wall mutants, Plant Physiol. Biochem. 38 (2000) 15–25. T. Girke, J. Lauricha, H. Tran, K. Keegstra, N. Raikhel, The Cell Wall Navigator database. A systems-based approach to organism-unrestricted mining of protein families involved in cell wall metabolism, Plant Physiol. 136 (2004) 3003–3008. S.R. Turner, N.G. Taylor, L. Jones, Mutations of the secondary cell wall, Plant Mol. Biol. 47 (2001) 209–219. S.R. Turner, C.R. Somerville, Collapsed xylem phenotype of Arabidopsis identifies mutants deficient in cellulose deposition in the secondary cell wall, Plant Cell 9 (1997) 689–701. V. Sundaresan, P. Springer, T. Volpe, S. Haward, J.D. Jones, C. Dean, H. Ma, R. Martienssen, Patterns of gene action in plant development revealed by enhancer trap and gene trap transposable elements, Genes Dev. 9 (1995) 1797–1810. P.J. Krysan, J.C. Young, M.R. Sussman, T-DNA as an insertional mutagen in Arabidopsis, Plant Cell 11 (1999) 2283–2290. P. Hilson, J. Allemeersch, T. Altmann, S. Aubourg, A. Avon, J. Beynon, R.P. Bhalerao, F. Bitton, M. Caboche, B. Cannoot, V. Chardakov, C. Cognet-Holliger, V. Colot, M. Crowe, C. Darimont, S. Durinck, H. Eickhoff, A.F. de Longevialle, E.E. Farmer, M. Grant, M.T. Kuiper, H. Lehrach, C. Leon, A. Leyva, J. Lundeberg, C. Lurin, Y. Moreau, W. Nietfeld, J. Paz-Ares, P. Reymond, et al., Versatile gene-specific sequence tags for Arabidopsis functional genomics: transcript profiling and reverse genetics applications, Genome Res. 14 (2004) 2176–2189. M.L. Crowe, C. Serizet, V. Thareau, S. Aubourg, P. Rouze, P. Hilson, J. Beynon, P. Weisbeek, P. van Hummelen, P. Reymond, J. Paz-Ares, W. Nietfeld, M. Trick, CATMA: a complete Arabidopsis GST database, Nucleic Acids Res. 31 (2003) 156–158. D.M. Brown, L.A.H. Zeef, J. Ellis, R. Goodacre, S.R. Turner, Identification of novel genes in Arabidopsis involved in secondary cell wall formation using expression profiling and reverse, Plant Cell 17 (2005) 2281–2295.

[55]

[56]

[57]

[58]

[59] [60] [61] [62]

[63]

[64]

[65]

[66]

[67]

[68]

[69]

[70]

[71]

[72] [73] [74]

447

S. Persson, H. Wei, J. Milne, G.P. Page, C.R. Somerville, Identification of genes required for cellulose synthesis by regression analysis of public microarray data sets, Proc. Natl. Acad. Sci. USA 105 (2005) 8633– 8638. S. Chivasa, B.K. Ndimba, W.J. Simon, D. Robertson, X.L. Yu, J.P. Knox, P. Bolwell, A.R. Slabas, Proteomic analysis of the Arabidopsis thaliana cell wall, Electrophoresis 23 (2002) 1754–1765. G. Borderies, E. Jamet, C. Lafitte, M. Rossignol, A. Jauneau, G. Boudart, B. Monsarrat, M.T. Esquerre-Tugaye, A. Boudet, R. Pont-Lezica, Proteomics of loosely bound cell wall proteins of Arabidopsis thaliana cell suspension cultures: a critical analysis, Electrophoresis 24 (2004) 3421–3432. G. Boudart, E. Jamet, M. Rossignol, C. Lafitte, G. Borderies, A. Jauneau, M.T. Esquerre-Tugaye, R. Pont-Lezica, Cell wall proteins in apoplastic fluids of Arabidopsis thaliana rosettes: Identification by mass spectrometry and bioinformatics, Proteomics 5 (2004) 212–221. B. Henrissat, A classification of glycoside hydrolases based on amino acid sequence similarities, Biochem. J. 280 (1991) 309–316. B. Henrissat, Glycosidase families, Biochem. Soc. Trans. 26 (1998) 153–156. Arabidopsis Genome Initiative, Analysis of the genome sequence of the flowering plant Arabidopsis thaliana, Nature 408 (2000) 796–815. B. Henrissat, P.M. Coutinho, G.J. Davies, A census of carbohydrateactive enzymes in the genome of Arabidopsis thaliana, Plant Mol. Biol. 47 (2001) 55–72. R. Yokoyama, K. Nishitani, Genomic basis for cell-wall diversity in plants. A comparative approach to gene families in rice and Arabidopsis, Plant Cell Physiol. 45 (2004) 1111–1121. M. Torki, P. Mandaron, R. Mache, D. Falconet, Characterization of a ubiquitous expressed gene family encoding polygalacturonase in Arabidopsis thaliana, Gene 242 (2000) 427–436. M. Molhoj, P. Ulvskov, F. Dal Degan, Characterization of a functional soluble form of a Brassica napus membrane-anchored endo-1,4-betaglucanase heterologously expressed in Pichia pastoris, Plant Physiol. 127 (2001) 674–684. F. Nicol, I. His, A. Jauneau, S. Vernhettes, H. Canut, H. Hofte, A plasma membrane-bound putative endo-1,4-beta-D-glucanase is required for normal wall assembly and cell elongation in Arabidopsis, EMBO J. 17 (1998) 5563–5576. M. Molhoj, B. Jorgensen, P. Ulvskov, B. Borkhardt, Two Arabidopsis thaliana genes, KOR2 and KOR3, which encode membraneanchored endo-1,4-beta-D-glucanases, are differentially expressed in developing leaf trichomes and their support cells, Plant Mol. Biol. 46 (2001) 263–275. D.R. Lane, A. Wiedemeier, L. Peng, H. Hofte, S. Vernhettes, T. Desprez, C.H. Hocart, R.J. Birch, T.I. Baskin, J.E. Burn, T. Arioli, A.S. Betzner, R.E. Williamson, Temperature-sensitive alleles of RSW2 link the KORRIGAN endo-1,4-beta-glucanase to cellulose synthesis and cytokinesis in Arabidopsis, Plant Physiol. 126 (2001) 278–288. C.C. Lashbrook, C. Gonzalez-Bosch, A.B. Bennett, Two divergent endo-beta-1,4-glucanase genes exhibit overlapping expression in ripening fruit and abscising flowers, Plant Cell 6 (1994) 1485–1493. Z. Shani, M. Dekel, G. Tsabary, O. Shoseyov, Cloning and characterization of elongation specific endo-1,4-beta-glucanase (cel1) from Arabidopsis thaliana, Plant Mol. Biol. 34 (1997) 837–842. S. Spolaore, L. Trainotti, A. Pavanello, G. Casadoro, Isolation and promoter analysis of two genes encoding different endo-beta-1,4-glucanases in the non-climacteric strawberry, J. Exp. Bot. 54 (2003) 271– 277. P.B. Hoj, G.B. Fincher, Molecular evolution of plant beta-glucan endohydrolases, Plant J. 7 (1995) 367–379. B.A. Stone, A.E. Clarke, Chemistry and biology of (1,3)-β-glucans, La Trobe University Press, Victoria, Australia, 1992. G.L. Bucher, C. Tarina, M. Heinlein, F. Di Serio, F. Meins Jr., V.A. Iglesias, Local expression of enzymatically active class I beta-1,3-glucanase enhances symptoms of TMV infection in tobacco, Plant J. 28 (2001) 361–369.

448 [75]

[76]

[77]

[78]

[79]

[80]

[81]

[82]

[83]

[84]

[85]

[86]

[87]

[88]

[89]

[90]

[91]

[92]

Z. Minic, L. Jouanin / Plant Physiology and Biochemistry 44 (2006) 435–449 G. Leubner-Metzger, F. Meins Jr., Sense transformation reveals a novel role for class I ß-1,3-glucanase in tobacco seed germination, Plant J. 23 (2000) 215–221. R.J. Stewart, J.N. Varghese, T.P. Garrett, P.B. Hoj, G.B. Fincher, Mutant barley (1→3,1→4)-beta-glucan endohydrolases with enhanced thermostability, Protein Eng. 14 (2001) 245–253. R. Leah, J. Kigel, I. Svendsen, J. Mundy, Biochemical and molecular characterization of a barley seed beta-glucosidase, J. Biol. Chem. 270 (1995) 15789–15797. M. Hrmova, A.J. Harvey, J. Wang, N.J. Shirley, G.P. Jones, B.A. Stone, P.B. Hoj, G.B. Fincher, Barley beta-D-glucan exohydrolases with beta-D-glucosidase activity. Purification, characterization, and determination of primary structure from a cDNA clone, J. Biol. Chem. 271 (1996) 5277–5286. M. Hrmova, E.A. MacGregor, P. Biely, R.J. Stewart, G.B. Fincher, Substrate binding and catalytic mechanism of a barley beta-D-glucosidase/(1,4)-beta-D-glucan exohydrolase, J. Biol. Chem. 273 (1998) 11134–11143. H.J. Crombie, S. Chengappa, J.S. Reid, A xyloglucan oligosaccharideactive, transglycosylating beta-D-glucosidase from the cotyledons of nasturtium (Tropaeolum majus L) seedlings-purification, properties and characterization of a cDNA clone, Plant J. 15 (1998) 27–38. J.B. Kim, A.T. Olek, N.C. Carpita, Cell wall and membrane-associated exo-beta-D-glucanases from developing maize seedlings, Plant Physiol. 123 (2000) 471–486. Z. Xu, L. Escamilla-Trevino, L. Zeng, M. Lalgondar, D. Bevan, B. Winkel, A. Mohamed, C.L. Cheng, M.C. Shih, J. Poulton, A. Esen, Functional genomic analysis of Arabidopsis thaliana glycoside hydrolase family 1, Plant Mol. Biol. 55 (2004) 343–367. H.U. Stotz, B.R. Pittendrigh, J. Kroymann, K. Weniger, J. Fritsche, A. Bauke, T. Mitchell-Olds, Induced plant defense responses against chewing insects. Ethylene signaling reduces resistance of Arabidopsis against Egyptian cotton worm but not diamondback moth, Plant Physiol. 124 (2000) 1007–1018. M.A. Malboobi, D.D. Lefebvre, A phosphate-starvation inducible betaglucosidase gene (psr3.2) isolated from Arabidopsis thaliana is a member of a distinct subfamily of the BGA family, Plant Mol. Biol. 34 (1997) 57–68. R. Opassiri, Y. Hua, O. Wara-Aswapati, T. Akiyama, J. Svasti, A. Esen, J.R. Ketudat Cairns, Beta-glucosidase, exo-beta-glucanase and pyridoxine transglucosylase activities of rice BGlu1, Biochem. J. 379 (2004) 125–131. J.K. Rose, J. Braan, S.C. Fry, K. Nishitani, The XTH family of enzymes involved in xyloglucan endotransglucosylation and endohydrolysis: current perspectives and a new unifying nomenclature, Plant Cell Physiol. 43 (2002) 1421–1435. M. Edwards, Y.J. Bowman, I.C. Dea, J.S. Reid, A beta-D-galactosidase from nasturtium (Tropaeolum majus L.) cotyledons. Purification, properties, and demonstration that xyloglucan is the natural substrate, J. Biol. Chem. 263 (1988) 4333–4337. J. Sampedro, C. Sieiro, G. Revilla, T. Gonzalez-Villa, I. Zarra, Cloning and expression pattern of a gene encoding an alpha-xylosidase active against xyloglucan oligosaccharides from Arabidopsis, Plant Physiol. 126 (2001) 910–920. R. Baran, Z. Sulova, E. Stratilova, V. Farkas, Ping-pong character of nasturtium-seed xyloglucan endotransglycosylase (XET) reaction, Gen. Physiol. Biophys. 19 (2000) 427–440. P. Campbell, J. Braam, Xyloglucan endotransglycosylases: diversity of genes, enzymes and potential wall-modifying functions, Trends Plant Sci. 4 (1999) 361–366. H.J. Crombie, S. Chengappa, C. Jarman, C. Sidebottom, J.S. Reid, Molecular characterisation of a xyloglucan oligosaccharide-acting alpha-D-xylosidase from nasturtium (Tropaeolum majus L.) cotyledons that resembles plant ’apoplastic’ alpha-D-glucosidases, Planta 214 (2002) 406–413. M. Hrmova, R. De Gori, B.J. Smith, J.K. Fairweather, H. Driguez, J.N. Varghese, G.B. Fincher, Structural basis for broad substrate specificity in higher plant β-D-glucohydrolases, Plant Cell 14 (2002) 1033–1052.

[93]

[94]

[95]

[96]

[97]

[98]

[99]

[100]

[101]

[102]

[103] [104]

[105]

[106]

[107]

[108]

[109]

[110]

[111]

D.L. Smith, J.A. Abbott, K.C. Gross, Down-regulation of tomato betagalactosidase 4 results in decreased fruit softening, Plant Physiol. 129 (2002) 1755–1762. V. Farkas, R. Hanna, G. Maclachlan, Xyloglucan oligosaccharide α-Lfucosidase activity from growing pea stems and germinating nasturtium seeds, Phytochemistry 30 (1991) 3203–3207. C. Augur, N. Benhamou, A. Darvill, P. Albersheim, Purification, characterization, and cell wall localization of an alpha-fucosidase that inactivates a xyloglucan oligosaccharid, Plant J. 3 (1993) 415–426. C. Augur, V. Stiefel, A. Darvill, P. Albersheim, P. Puigdomenech, Molecular cloning and pattern of expression of an alpha-L-fucosidase gene from pea seedlings, J. Biol. Chem. 270 (1995) 24839–24843. T. Tarrago, I. Martinez, M. Torrent, A. Codina, E. Giralt, P. Puigdomenech, D. Ludevid, The fuc1 gene product (20 kDa FUC1) of Pisum sativum has no alpha-L-fucosidase activity, Plant Mol. Biol. 51 (2003) 877–884. F. de la Torre, J. Sampedro, I. Zarra, G. Revilla, AtFXG1, an Arabidopsis gene encoding alpha-L-fucosidase active against fucosylated xyloglucan oligosaccharides, Plant Physiol. 128 (2002) 247–255. R.A. O’Neill, P. Albersheim, A.G. Darvill, Purification and characterization of a xyloglucan oligosaccharide-specific xylosidase from pea seedlings, J. Biol. Chem. 264 (1989) 20430–20437. C. Fanutti, M.J. Gidley, J.S.G. Reid, A xyloglucan-oligosaccharide-specific α-D-xylosidase or exo-oligoxyloglucan-α-xylohydrolase from germinated nasturtium (Tropaeolum majus L.) seeds, Planta 184 (1991) 137–147. S. Chengappa, C. Jarman, C. Fanutti, J.S.G. Reid, Xyloglucan oligosaccharide-specific α-D-xylosidase: molecular mode of action and cloning of a cDNA from germinated nasturtium (Tropeaolum majus L.) seeds, J. Cell. Biochem. Suppl. 17A (1993) 27. N. Iglesias, J.A. Abelenda, M. Rodino, J. Sampedro, G. Revilla, I. Zarra, Apoplastic glycosidases active against xyloglucan oligosaccharides of Arabidopsis thaliana, Plant Cell Physiol. 47 (2006) 55–63. A. Sunna, G. Antranikian, Xylanolytic enzymes from fungi and bacteria, Crit. Rev. Biotechnol. 17 (1997) 39–67. S.S.H. Wu, D.F. Suen, H.C. Chang, A.H.C. Huang, Maize tapetum xylanase is synthesized as a precursor, processed and activated by a serine protease, and deposited on the pollen, J. Biol. Chem. 277 (2002) 49055–49064. M. Suzuki, A. Kato, N. Nagata, Y. Komeda, A xylanase, AtXyn1, is predominantly expressed in vascular bundles, and four putative xylanase genes were identified in the Arabidopsis genome, Plant Cell Physiol. 43 (2002) 759–767. R.C. Lee, R.A. Burton, M. Hrmova, G.B. Fincher, Barley arabinoxylan arabinofuranosidases: purification, characterization and determination of primary structures from cDNA clones, Biochem. J. 356 (2001) 181– 189. Z. Minic, C. Rihouey, C.-T. Do, P. Lerouge, L. Jouanin, Purification and characterization of enzymes exhibiting beta-D-xylosidase activities in stem tissues of Arabidopsis, Plant Physiol. 135 (2004) 867–878. T. Goujon, Z. Minic, A. El Amrani, O. Lerouxel, E. Aletti, C. Lapierre, J.P. Joseleau, L. Jouanin, AtBXL1, a novel higher plant (Arabidopsis thaliana) putative beta-xylosidase gene, is involved in secondary cell wall metabolism and plant development, Plant J. 33 (2003) 677– 690. H. Ferré, A. Broberg, J.O. Duus, K.K. Thomsen, A novel type of arabinoxylan arabinofuranohydrolase isolated from germinated barley: analysis of substrate preference and specificity by nano-probe NMR, Eur. J. Biochem. 267 (2000) 6633–6641. P. Marraccini, W.J. Rogers, C. Allard, M.L. Andre, V. Caillet, N. Lacoste, F. Lausanne, S. Michaux, Molecular and biochemical characterization of endo-beta-mannanases from germinating coffee (Coffea arabica) grains, Planta 213 (2001) 296–308. B. Mo, J.D. Bewley, Beta-mannosidase (EC 3.2.1.25) activity during and following germination of tomato (Lycopersicon esculentum Mill.) seeds. Purification, cloning and characterization, Planta 215 (2002) 141–152.

Z. Minic, L. Jouanin / Plant Physiology and Biochemistry 44 (2006) 435–449 [112] W.D. Kim, O. Kobayashi, S. Kaneko, Y. Sakakibara, G.G. Park, I. Kusakabe, H. Tanaka, H. Kobayashi, alpha-Galactosidase from cultured rice (Oryza sativa L. var. Nipponbare) cells, Phytochemistry 61 (2002) 621–630. [113] G.B. Seymour, K.C. Gross, Cell wall disassembly and fruit softening, Postharvest News and Information 7 (1996) 5N–52N. [114] R.G. Atkinson, R. Schroder, I.C. Hallet, D. Cohen, E.A. Macrae, Overexpression of polygalacturonase in transgenic apple trees leads to a range of novel phenotypes involving changes in cell adhesion, Plant Physiol. 129 (2002) 122–133. [115] G.U. Rao, I. Paran, Polygalacturonase: a candidate gene for the soft flesh and deciduous fruit mutation in Capsicum, Plant Mol. Biol. 51 (2003) 135–141. [116] K. Wakabayashi, T. Hoson, D.J. Huber, Methyl de-esterification as a major factor regulating the extent of pectin depolymerization during fruit ripening: a comparison of the action of avocado (Persea americana) and tomato (Lycopersicon esculentum) polygalacturonases, J. Plant Physiol. 160 (2003) 667–673. [117] R. Tanaka, M. Ikeda, K. Funatsuki, H. Yukioka, Y. Hashimoto, S. Fujimoto, M. Takata, K. Katoh, H. Konno, Molecular cloning and cytochemical analysis of exopolygalacturonase from carrot, Planta 215 (2002) 735–744. [118] F. Micheli, Pectin methylesterases: cell wall enzymes with important roles in plant physiology, Trends Plant Sci. 6 (2001) 414–419. [119] S.C. Li, J.W. Han, K.C. Chen, C.S. Chen, Purification and characterization of isoforms of β-galactosidases in mung bean seedlings, Phytochemistry 57 (2001) 349–359. [120] R. Esteban, B. Dopico, F.J. Munoz, S. Romo, I. Martin, E. Labrador, Cloning of a Cicer arietinum beta-galactosidase with pectin-degrading function, Plant Cell Physiol. 44 (2003) 718–725. [121] R.C. Lee, M. Hrmova, R.A. Burton, J. Lahnstein, G.B. Fincher, Bifunctional family 3 glycoside hydrolases from barley with α-L-arabinofuranosidase and β-D-xylosidase activity, J. Biol. Chem. 278 (2003) 5377– 5387. [122] M.S. Buckeridge, J.S. Reid, Purification and properties of a novel betagalactosidase or exo-(1→4)-beta-D-galactanase from the cotyledons of germinated Lupinus angustifolius L. seeds, Planta 192 (1994) 502–511. [123] A.T. Carey, K. Holt, S. Picard, R. Wilde, G.A. Tucker, C.R. Bird, W. Schuch, G.B. Seymour, Tomato exo-(1→4)-beta-D-galactanase. Isolation, changes during ripening in normal and mutant tomato fruit, and characterization of a related cDNA clone, Plant Physiol. 108 (1995) 1099–1107. [124] I. Martin, B. Dopico, F.J. Munoz, R. Esteban, R.J. Oomen, A. Driouich, J.P. Vincken, R. Visser, E. Labrador, In vivo expression of a Cicer arietinum beta-galactosidase in potato tubers leads to a reduction of the galactan side-chains in cell wall pectin, Plant Cell Physiol. 46 (2005) 1613–1622. [125] S. Balasubramaniam, H.C. Lee, H. Lazan, R. Othman, Z.M. Ali, Purification and properties of a beta-galactosidase from carambola fruit with

[126]

[127]

[128]

[129]

[130]

[131]

[132]

[133]

[134]

[135]

[136]

[137]

[138]

[139]

449

significant activity towards cell wall polysaccharides, Phytochemistry 66 (2005) 153–163. T. Kotake, S. Dina, T. Konishi, S. Kaneko, K. Igarashi, M. Samejima, Y. Watanabe, K. Kimura, Y. Tsumuraya, Molecular cloning of a βgalactosidase from radish that specifically hydrolyzes β-(1→3)- and β(1→6)-galactosyl residues of Arabinogalactan protein, Plant Physiol. 138 (2005) 1563–1576. S. Anhalt, G. Weissenböck, Subcellular localization of luteolin glucuronidases and related enzymes in rye mesophyll, Planta 187 (1992) 83– 88. R.P. de Vries, J. Visser, Aspergillus enzymes involved in degradation of plant cell wall polysaccharides, Microbiol. Mol. Biol. Rev. 65 (2001) 497–522. M. Tuncer, A.S. Ball, Co-operative actions and degradation analysis of purified xylan-degrading enzymes from Thermomonospora fusca BD25 on oat-spelt xylan, J. Appl. Microbiol. 94 (2003) 1030–1035. A.K. Rahman, N. Sugitani, M. Hatsu, K. Takamizawa, A role of xylanase, α-L-arabinofuranosidase, and xylanase in xylan degradation, Can. J. Microbiol. 49 (2003) 58–64. J.K.C. Rose, M. Saladie, C. Catala, The plot thickens: new perspectives of primary cell wall modification, Curr. Opin. Plant Biol. 7 (2004) 296– 301. T. Akamatsu, Y. Hanzawa, Y. Ohtake, T. Takahashi, K. Nishitani, Y. Komeda, Expression of endoxyloglucan transferase genes in acaulis mutants of Arabidopsis, Plant Physiol. 121 (1999) 715–722. Y. Hanzawa, T. Takahashi, Y. Komeda, ACL5: an Arabidopsis gene required for internodal elongation after flowering, Plant J. 12 (1997) 863–874. N.M. Steele, Z. Sulova, P. Campbell, J. Braam, V. Farkas, S.C. Fry, Ten isoenzymes of xyloglucan endotransglycosylase from plant cell walls select and cleave the donor substrate stochastically, Biochem. J. 355 (2001) 671–679. X. Dong, M. Mindrinos, K.R. Davis, F.M. Ausubel, Induction of Arabidopsis defense genes by virulent and avirulent Pseudomonas syringae strains and by a cloned avirulence gene, Plant Cell 3 (1991) 61–72. M. Torki, P. Mandaron, F. Thomas, F. Quigley, R. Mache, D. Falconet, Differential expression of a polygalacturonase gene family in Arabidopsis thaliana, Mol. Gen. Genet. 261 (1999) 948–952. L. Sander, R. Child, P. Ulvskov, M. Albrechtsen, B. Borkhardt, Analysis of a dehiscence zone endo-polygalacturonase in oilseed rape (Brassica napus) and Arabidopsis thaliana: evidence for roles in cell separation in dehiscence and abscission zones, and in stylar tissues during pollen tube growth, Plant Mol. Biol. 46 (2001) 469–479. J.D. Monroe, C.M. Gough, L.E. Chandler, C.M. Loch, J.E. Ferrante, P.W. Wright, Structure, properties, and tissue localization of apoplastic alpha-glucosidase in crucifers, Plant Physiol. 119 (1999) 385–397. L.M. Fulton, C.S. Cobbett, Two alpha-L-arabinofuranosidase genes in Arabidopsis thaliana are differentially expressed during vegetative growth and flower development, J. Exp. Bot. 54 (2003) 2467–2477.