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3 Biosynthesis of pectinsDebra Mohnen

3.1 Introduction

Pectin is the most structurally complicated polysaccharide in the plant cell wall.Accordingly, the study of its synthesis is complex owing to the large numberof enzymes required. At least 12 activated sugar substrates are required forpectin synthesis. The known activated sugars are nucleotide-sugars, althoughthe possibility that lipid-linked sugars may be involved cannot be ruled out.The regulation of the synthesis of the activated sugar substrates is likely to beimportant in the overall regulation of pectin synthesis. Based on the knownstructure of pectin, at least 14 distinct enzyme activities are required to syn-thesize the activated sugar substrates and 58 distinct glycosyl-, methyl- andacetyltransferases are required to synthesize the complicated family of polymersknown as pectin. While progress has been made in characterizing some of thepectin biosynthetic enzymes in crude or partially purified plant homogenates,in no case has a single enzyme been completely characterized in regard toenzyme structure, subcellular location, protein sequence, gene identity andenzyme regulation. Several recent comprehensive reviews on pectin structure(Mohnen, 1999; Ridley et al., 2001), synthesis (Mohnen, 1999; Ridley et al.,2001) and function (Ridley et al., 2001; Willats et al., 2001a) and on nucleotide-sugar interconversion pathways (Reiter and Vanzin, 2001) have been publishedand should be consulted for detailed background information. In addition,several general reviews on cell wall synthesis (Gibeaut, 2000; Reid, 2000;Dhugga, 2001; Perrin et al., 2001) and on progress and strategies to identifywall biosynthetic genes (Keegstra and Raikhel, 2001; Perrin et al., 2001) andglycosyltransferases (Henrissat et al., 2001; Keegstra and Raikhel, 2001) haverecently been published. The goal of this review is to summarize our presentlevel of understanding of pectin synthesis.

The first part of the review outlines our understanding of the subcellularlocation of pectin synthesis. The nucleotide-sugars required for pectin synthesisare then introduced and progress towards understanding the mechanism, site(s)and regulation of their synthesis is reviewed. Finally, a list of the glycosyl-,methyl- and acetyltransferases required for pectin synthesis is provided andprogress towards identifying and characterizing these enzymes is summarized.It is hoped that this review will provide a foundation to facilitate the identificationof the pectin biosynthetic genes and the development of better molecular toolsto study pectin biosynthesis.

BIOSYNTHESIS OF PECTINS 53

3.2 What is the structure of newly synthesized pectin?

One of the challenges of studying pectin synthesis is that we do not know thestructure of de novo synthesized pectin. For example, the detailed structuralcharacterization of pectin isolated from the wall (ONeill et al., 1990; Mohnen,1999; Ridley et al., 2001) has led to the identification of the family of complexpolysaccharides known as homogalacturonan (HGA), rhamnogalacturonan I(RG-I) and the substituted galacturonans such as the ubiquitous rhamnogalac-turonan II (RG-II) (Ridley et al., 2001), and the less prevalent xylogalacturonan(Schols et al., 1990, 1995; Yu and Mort, 1996), and apiogalacturonan (Hartand Kindel, 1970; Watson and Orenstein, 1975). However, we do not yet knowwhether these polysaccharides are synthesized as one polymer or whether theyare synthesized as individual polymers that become interconnected during orfollowing their insertion into the wall (see Figure 3.1). Furthermore, we do notknow whether the structural differences found in each of the polysaccharidesisolated from the wall, such as variations in the degree and pattern of methyl-and acetyl-esterification of homogalacturonan (Willats et al., 2001b), arise inthe wall (i.e. in muro by wall-localized enzymes) or whether the differencesoccur, at least in part, during synthesis. This uncertainty makes it challenging topropose models for how pectin is synthesized and to predict which enzymes arerequired intracellularly for synthesis, rather than being required extracellularlyfor in muro modeling of pectin. For the purpose of this review, the workinghypothesis is that homogalacturonan can be synthesized as in independentpolymer. It is also proposed that substituted galacturonans such as ubiquitousRG-II (Ridley et al., 2001) and the other less prevalent xylogalacturonan (Scholset al., 1990, 1995; Yu and Mort, 1996) and apiogalacturonan (Ridley et al.,2001) can be synthesized as modified versions of homogalacturonan. Finally,it is hypothesized that RG-I is synthesized as an independent polymer thatmay, or may not, be covalently linked to homogalacturonan or to substitutedgalacturonans (see Figure 3.1). It must be stressed, however, that no unequivocalevidence is available to support the hypothesized independence of RG-I andHGA synthesis or of the dependence of RG-II synthesis on HGA. One ofthe goals, or outcomes, of current research in pectin synthesis should be theelucidation of how the three main pectic polymers, HGA, RG-I and RG-II, arelinked together during synthesis.

3.3 Subcellular location of pectin synthesis

Several lines of evidence indicate that pectin is synthesized in the Golgi andtransported to the wall via membrane-bound vesicles. The plant Golgi apparatusis a dynamic series of membrane-bound vesicles and stacks that function inthe biosynthesis pectins and hemicellulose, in the glycosylation of proteins

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and in the synthesis of lipids (Staehelin and Moore, 1995; Nebenfuhr andStaehelin, 2001). The Golgi vesicles move along actin filaments via myosinmotors (Nebenfuhr et al., 1999) and it is believed that this movement targetsthe transport of pectin and other macromolecules to the cell wall.

The autoradiographic identification of radiolabeled polysaccharides in Golgicisternae and their chase from the Golgi to the cell wall using non-radiolabeledprecursors (Northcote and Pickett-Heaps, 1966; Northcote, 1970) was amongthe first evidence that pectin is synthesized in the Golgi. For example, Golgi-enriched fractions isolated from cells grown in the presence of radiolabeledglucose contained radioactive galactose (Gal), arabinose (Ara) and galactosy-luronic acid (GalA, galacturonic acid), the major glycosyl residues found in thepectic polysaccharides (Stoddart and Northcote, 1967; Harris and Northcote,1971).

The identification of pectin-specific carbohydrate epitopes in the Golgi, viaimmunocytochemistry of thin cell sections using anti-pectin antibodies (Mooreet al., 1991; Staehelin and Moore, 1995; Willats et al., 2000), provides fur-ther evidence that pectins are synthesized in the Golgi apparatus (Staehelinand Moore, 1995) and suggests that specific pectic carbohydrate epitopes aresublocalized in the Golgi. Studies using antibodies reactive to HGA-like andthe RG-I-like epitopes suggest that the syntheses of homogalacturonan (HGA)and rhamnogalacturonan I (RG-I) begin in the cis-Golgi (Lynch and Staehelin,1992; Zhang and Staehelin, 1992; Staehelin and Moore, 1995) and continue intothe medial Golgi (Moore et al., 1991; Zhang and Staehelin, 1992; Staehelin andMoore, 1995) with more extensive branching taking place in the trans-Golgicisternae (Zhang and Staehelin, 1992; Staehelin and Moore, 1995). Further-more, studies using antibodies reactive against relatively unesterified HGA(JIM5 (VandenBosch et al., 1989; Knox et al., 1990); PGA/RG-I (Moore et al.,1986; Moore and Staehelin, 1988; Lynch and Staehelin, 1992); 2F4 (Linerset al., 1989)) and relatively esterified HGA (JIM7 (Knox et al., 1990)) sug-gest that HGA becomes methyl-esterified in the medial and trans-Golgi (Vianand Roland, 1991; Liners and Van Cutsem, 1992; Zhang and Staehelin, 1992;Sherrier andVandenBosch, 1994; Staehelin and Moore, 1995), and is transportedto the plasma membrane in vesicles as a highly methyl-esterified polymer thatis inserted into the wall (Carpita and Gibeaut, 1993; Liners et al., 1994; Dolanet al., 1997). The de-esterification of HGA by pectin methylesterases (Micheli,2001) in the wall or at cell plate (Dolan et al., 1997) produces more acidicHGA (Stoddart and Northcote, 1967; Shea et al., 1989; Liners and Van Cutsem,1992; Li et al., 1994; Marty et al., 1995). Such a spatial partitioning of HGAesterification and de-esterification is supported by the localization of esterifiedHGA throughout the cell wall (Fujiki et al., 1982; Knox et al., 1990; Vianand Roland, 1991; Liners and Van Cutsem, 1992; Liners et al., 1994; Sherrierand VandenBosch, 1994; Marty et al., 1995; Dolan et al., 1997; Willats et al.,2001a), the localization of relatively unesterified HGA in the middle lamella,

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58 PECTINS AND THEIR MANIPULATION

and the frequently observed absence of unesterified HGA epitopes in the trans-Golgi vesicles. In spite of such results, however, the fact that some cell typesshow a different localization of unesterified HGA, such as melon callus cells(Vian and Roland, 1991) that contain unesterified HGA in the trans-Golgi,suggests that HGA can be inserted into the wall in a relatively unesterifiedstate in some cells. This calls into question the general belief that HGA isnecessarily synthesized in a highly esterified form (Knox et al., 1990; Caseroand Knox, 1995). It is important to realize that specific pectic epitopes localizeto different Golgi compartments in different cell types (Knox et al., 1990;Lynch and Staehelin, 1992; Casero and Knox, 1995; Staehelin and Moore,1995), suggesting that pectin synthesis may differ in different cell types, indifferent species, at different points during development, or even at differentlocations in the same wall (Stacey et al., 1995; Willats et al., 1999; McCartneyet al., 2000; Orfila and Knox, 2000; Willats et al., 2000). However, since theabsence of a specific carbohydrate epitope may be due to masking of the epitoperather than to a lack of the synthesis of the epitope, it is necessary to confirmcarbohydrate-epitope immunocytochemistry results with localization studies onthe biosynthetic enzymes themselves in order to make conclusions regardingthe mechanism of biosynthesis.

The first enzymatic evidence for the location of a pectin biosynthetic glyco-syltransferase was the localization of HGA-galacturonosyltransferase (GalAT)to the Golgi and the demonstration that its catalytic site faces the Golgi lumen(Sterling et al., 2001). Most of the GalAT activity in pea (Pisum sativum) co-localizes in linear and discontinuous sucrose gradients with the Golgi markerlatent UDPase and is separated from the endoplasmic reticulum, mitochondriaand plasma membrane (Sterling et al., 2001). The catalytic site of GalAT wasshown to reside within the lumen of the Golgi, since GalAT activity was reducedby treatment with proteinase K only if Golgi membranes were first permeabi-lized with detergent (Sterling et al., 2001). The enzymes that methyl-esterifyHGA have also been localized to the Golgi, further confirming the location ofpectin synthesis. Tobacco HGA methyltransferase (HGA-MT) activity localizesto the Golgi and the enzymes catalytic site was shown to face the Golgi lumen(Goubet and Mohnen, 1999a). The localization of roughly half of the pectinmethyltransferase activity in flax to the Golgi (Vannier et al., 1992; Bourlardet al., 1997b) provides further evidence that HGA methyl-esterification occursin the Golgi.

The location of pectin synthesis in the Golgi leads to the question of where thenucleotide-sugar substrates are synthesized and of how the substrates gain accessto the enzyme. It has been proposed that the nucleotide-sugars required for pectinsynthesis are synthesized on the cytosolic side of the Golgi and transportedinto the Golgi lumen by specific nucleotide-sugar:nucleoside monophosphateantiporters (Sterling et al., 2001) (see Figure 3.2a).While this model is consistentwith the topology of some nucleotide-sugar biosynthetic enzymes in animals


(Berninsone and Hirschberg, 2000) and plants (Schroeder and Hagiwara, 1989;Munoz et al., 1996; Neckelmann and Orellana, 1998; Baldwin et al., 2001),there are also indications that some nucleotide biosynthesis enzymes, such asUDP-glucuronic acid decarboxylase (Hayashi et al., 1988; Kearns et al., 1993),may actually reside in the Golgi (see Figure 3.2b). Thus, until the subcellularlocation of the enzymes is confirmed experimentally, two models for the locationof the nucleotide-transforming enzymes must be considered (Figure 3.2). Forthose nucleotide-sugars that are synthesized in the cytosol, it has been shown inboth animals (Capasso and Hirschberg, 1984) and plants (Munoz et al., 1996;Wang et al., 1997; Neckelmann and Orellana, 1998; Baldwin et al., 2001) thattransport occurs via nucleotide-sugar:nucleoside monophosphate antiportersthat reside in the endoplasmic reticulum or Golgi membranes. As shown inFigure 3.2a, nucleotide-sugars that are synthesized on the cytosolic side of theGolgi are predicted to be transported into the Golgi lumen by specific nucleotide-sugar:nucleoside monophosphate antiporters. The channeling of nucleotide-sugars from the cytosol into the Golgi may be facilitated by nucleotide-sugarbinding proteins (Faik et al., 2000). Once the nucleotide-sugar is transportedinto the Golgi it is used as a substrate by a pectin biosynthetic glycosyltrans-ferase that transfers the glycosyl residue onto a growing polymer. The releasednucleoside diphosphate (NDP) is hydrolyzed by a Golgi-localized nucleotide-5-diphosphatase (NDPase) (Orellana et al., 1997) into NMP and inorganicphosphate. The nucleoside monophosphate is then transported out of the Golgiby a nucleotide-sugar:nucleoside monophosphate antiporter.

3.4 Synthesis of the nucleotide-sugar substratesrequired for pectin synthesis

It is commonly accepted that nucleotide-sugars are the immediate substrates forthe glycosyltransferases that drive pectin biosynthesis. Radiolabeled nucleotide-sugars have been used in vitro as substrates to assay a number of pectin biosyn-thetic glycosyltransferases including 1,4-galacturonosyltransferase (Sterlinget al., 2001; Takeuchi and Tsumuraya, 2001) and galactosyltransferase (Geshiet al., 2000; Peugnet et al., 2001). Figure 3.3 shows a summary of the nucleotide-sugar interconversion pathways that are believed or proposed to be the primarypathways for the synthesis of the activated sugars required for pectin synthe-sis (Feingold and Avigad, 1980; Feingold and Barber, 1990; Mohnen, 1999;Gibeaut, 2000; Reiter and Vanzin, 2001; Ridley et al., 2001). Although the inter-conversion pathways are likely the main pathways for the synthesis of thenucleotide-sugars, there is a set of alternative pathways for the synthesis of theNDP-sugars known as the salvage pathway. In the salvage pathwayl-Ara,d-Gal,d-Man (mannose),d-GalA,d-GlcA, l-Rha, l-Fuc (fucose),d-Glc (glucose) andd-Xyl (xylose) are recycled from the wall by conversion into sugar-1-phosphates

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by the action of C-1-Kinases (Hassid et al., 1959; Hassid, 1967; Feingold andAvigad, 1980; Feingold and Barber, 1990). The resulting sugar-1-phosphatesare transformed into nucleotide-sugars by pyrophosphorylases that transfer thenucleoside monophosphate (NMP) from a nucleoside triphosphate (NTP) ontothe phosphate of the sugar-1-phosphate with the release of pyrophosphate: sugar-1-P + NTP NDP-sugar + PPi (Hassid et al., 1959). The so-called salvagepathway is thought to function in the re-utilization of glycosyl residues followingturnover of the wall. The synthesis of UDP-GlcA from GlcA-1-P derived frommyoinositol (the myoinositol pathway) (Hassid, 1967; Feingold and Barber,1990) also uses, in part, some of the enzymes from the salvage pathway. Thegenes for some of the enzymes in the salvage pathway have been identified,including galactokinase (AGK1 and GAL1) (Kaplan et al., 1997; Sherson et al.,1999), arabinose kinase (ARA1) (Sherson et al., 1999), and GDP-d-mannosepyrophosphorylase (also referred to mannose-1-phosphate guanylyltransferase)(VTC1 and CYT1) (Keller et al., 1999; Lukowitz et al., 2001).

Although the amount of each type of glycosyl residue required for wallsynthesis will depend on the exact structure of pectin being synthesized inthat cell type at that specific point of development, a comparison of the glycosylresidue composition of pectins isolated from walls of sycamore cell suspensions(Eberhard et al., 1989), tobacco cell suspensions (Mohnen et al., 1996) andArabidopsis leaves (Zablackis et al., 1996) (see Table 3.1) gives an indicationof the relative proportion of the different glycosyl residues in pectin. The mostabundant glycosyl residue in pectin is d-GalA followed by, depending on thespecies and cell type, l-Ara, l-Rha, d-Gal, d-GlcA and d-Xyl. The otherglycosyl residues required for pectin synthesis (i.e. l-Fuc, l-Gal, d-Apiose,l-aceric acid, d-Kdo (ketodeoxymanno-octulopyranosylonic acid), and d-Dha(deoxylyxo-heptulopyranosylaric acid)) represent 1% or less of normalizedmol% of pectin (Ridley et al., 2001).

The nucleotide-sugars involved in pectin synthesis (Mohnen, 1999; Ridleyet al., 2001) and in general cell wall synthesis (Feingold and Avigad, 1980;Feingold and Barber, 1990; Gibeaut, 2000) and the molecular genetics of plantnucleotide-sugar interconversion pathways (Reiter and Vanzin, 2001) have beenreviewed. The reader is directed to these reviews for additional backgroundinformation. Here, only the salient features of the biosynthetic pathways willbe presented. Hexose phosphates made directly from carbohydrate products ofphotosynthesis or from transported sucrose or stored starch are the precursorsfor the nucleotide-sugars (see Figure 3.3). As mentioned above, while it has tra-ditionally been held that the nucleotide-sugars are synthesized on the cytosolicside of the Golgi, there is evidence that some nucleotide-sugars, such as UDP-Xyl, may at least in part be synthesized in the Golgi lumen (Hayashi et al.,1988). The nucleotide-sugars required for pectin synthesis will be described inthe order of the abundance of their respective glycosyl residues in pectin, asbased on the composition of the three different pectins shown in Table 3.1.


Table 3.1 Comparison of the glycosyl residue compositiona of pectic polysaccharidesb released fromtobacco, sycamore and Arabidopsis walls

Normalized mol%

Glycosyl residuea Sycamore suspensionc Tobacco suspensiond Arabidopsis leavese

Galacturonic acid 56.0 74.2 72.1Arabinose 15.2 2.6 6.6Rhamnose 7.4 6.4 9.8Galactose 11.6 1.4 6.5Glucuronic acid 5.0 7.5 0.3Xylose 1.5 0.5 2.0Fucose 1.2 1.0 0.7Unknownf 6.5 Mannose 1.8 0 0

aGlycosyl residues


and Mohnen, 1999) moiety. UDP-[14C]GalA has also been synthesized forin vitro use by the enzymatic oxidation of UDP-[14C]Gal to UDP-[14C]GalA(Rao and Mendicino, 1976; Kelleher and Bhavanandan, 1986; Basu et al., 2000)with a yield of>90% UDP-[14C]GalA using a simple polyethyleneimine (PEI-cellulose) column chromatography purification step (Basu et al., 2000). Wefind it necessary to purify the UDP-[14C]GalA synthesized from UDP-[14C]Galby high-pressure liquid chromatography (HPLC) to remove contaminants thatinhibit 1,4-galacturonosyltransferase activity. With the HPLC purification stepwe obtain 27% yield of UDP-[14C]GalA (J. Sterling and D. Mohnen, unpub-lished results). The UDP-GlcA 4-epimerase from plants has not been purifiedto homogeneity, nor has its gene been cloned. However, a gene (Cap1J) for abacterial UDP-GlcA 4-epimerase from Streptococcus pneumoniae type 1 hasbeen identified (Munos et al., 1999). The bacterial enzyme has a Km for UDP-GlcA of 240M, a pH optimum of 7.5, an equilibrium constant of 1.3 in thedirection of UDP-GalA, and Mr of 80,000. The bacterial enzyme appears torequire a tightly bound NAD+ for activity (Munos et al., 1999). Reiter andVanzin (2001) report that BLAST searches of the Arabidopsis genome yields sixpredicted coding regions with high degrees of sequence similarity to the Cap1Jgene from Streptococcus pneumoniae. Definitive evidence that these putativeUDP-GlcA epimerase genes encode functional UDP-GlcA 4-epimerase has notyet been reported.

3.4.2 Uridine diphosphate--l-arabinose (UDP-l-Ara)UDP-l-Ara is a substrate for the synthesis of RG-I and RG-II. UDP-l-Arais formed by the 4-epimerization of UDP-Xyl catalyzed by UDP-arabinose 4-epimerase (EC 5.1.3.5) (Feingold et al., 1960; Fan and Feingold, 1970; Feingoldand Avigad, 1980; Robertson et al., 1995). UDP-arabinose 4-epimerase activ-ity has been identified in particulate preparations from multiple plant species(Feingold and Avigad, 1980).

UDP-d-XylUDP-xylose 4-epimerase UDP-l-Ara (2)

A partially purified UDP-xylose 4-epimerase from wheat germ (Fan andFeingold, 1970) was shown to have a pH optimum of 8.0, an apparent Km of1.5 mM for UDP-Xyl and 0.5 mM for UDP-l-Ara, and an equilibrium constantof 0.8 in the direction of UDP-Ara (Fan and Feingold, 1970). A comparableequilibrium constant of 1.0 has been reported for the UDP-xylose 4-epimerasefrom mung bean (Feingold et al., 1960). If NAD+ is required for the reaction,it must be tightly bound to the enzyme (Feingold and Avigad, 1980). Thewheat germ UDP-xylose 4-epimerase (Fan and Feingold, 1970; Feingold andAvigad, 1980) has been used to synthesize non-radiolabeled and radiolabeledUDP--l-arabinopyranose (Pauly et al., 2000) for use for in vitro wall polymer


biosynthesis studies. The Ara in the synthesized UDP-Ara is in the pyranoseform, while the predominant form of arabinose is arabinofuranose in the wallpolysaccharides, proteoglycans, arabinans and arabinogalactan proteins (Hassidet al., 1959; Feingold and Avigad, 1980; Carpita, 1996). It is not known whenthe mutorotation occurs; however, it is believed to occur during polysaccha-ride biosynthesis, presumably by arabinosyltransferase(s) that catalyze ringrearrangement before formation of the glycosidic bond (Carpita, 1996). AnArabidopsis mutant (mur4) has been identified that has reduced membrane-bound UDP-xylose 4-epimerase activity in the leaves, cotyledons and flowers(Burget and Reiter, 1999).

3.4.3 Uridine diphosphate--l-rhamnose (UDP-l-Rha)l-Rhamnose is a component of RG-I and RG-II. Plants such as tobacco (Barber,1963), mung bean (Barber, 1962), Silene dioica (Kamsteeg et al., 1978),pummelo (Citrus maxima) (Bar-Peled et al., 1991) and Chlorella pyrenoidasa(Barber and Chang, 1967) can convert UDP-d-Glc to UDP-l-rhamnose in anNADH-dependent reaction (reviewed in Feingold and Avigad, 1980; Feingoldand Barber, 1990). UDP-4-keto-6-deoxy-d-Glc is an intermediate in the con-version (Barber, 1963; Barber and Chang, 1967; Kamsteeg et al., 1978). UDP-Rha has been shown to be a substrate in plants for the rhamnosylation ofsecondary metabolites such as flavonoid-glycosides (Bar-Peled et al., 1993)and it is assumed that UDP-l-Rha is the nucleotide-sugar substrate for thesynthesis of RG-I and RG-II. However, it should be noted that it has not yetbeen experimentally confirmed that UDP-Rha is the substrate for RG-I and RG-II synthesis. A biosynthetic scheme for the synthesis of UDP-Rha in plants hasbeen proposed (Kamsteeg et al., 1978; Feingold and Avigad, 1980; Mohnen,1999) based on the rfb genes-encoded pathway for the synthesis of dTDP-rhamnose from dTDP-glucose in bacteria (Stevenson et al., 1994). The proposedpathway is shown in equation 3.

UDP-d-Glc

UDP-glucose4,6-dehydratase [UDP-4-keto-6-deoxy-Glc]

UDP-4-keto-l-rhamnose3,5-epimerase

[UDP-4-keto-6-deoxy-l-mannose]UDP-4-ketorhamnose

reductaseUDP-l-Rha (3)

Based on the bacterial pathway, the proposed pathway for UDP-Rha synthesisin plants begins with the conversion of UDP-d-Glc to UDP-4-keto-6-deoxy-Glccatalyzed by UDP-glucose 4,6-dehydratase (EC 4.2.1.76). The UDP-4-keto-6-deoxy-Glc is subsequently epimerized to UDP-4-keto-6-deoxy-l-mannose


by UDP-4-keto-l-rhamnose 3,5-epimerase. Finally, the UDP-4-keto-6-deoxy-l-mannose is reduced to UDP-l-rhamnose by UDP-4-ketorhamnose reductase.None of these enzymes has been purified to homogeneity or cloned in plants. It isalso not clear how many enzymes would encode the required enzyme activities.Blast searches of the Arabidopsis genome database using E. coli dTDP-l-rhamnose biosynthetic genes (E. coli has unique genes for each of the threerequired enzymatic activities for dTDP-rhamnose synthesis) have identifiedthree Arabidopsis genes with significant sequence similarity to dTDP-d-glucose4,6-dehydratase (Reiter and Vanzin, 2001). Based on the primary sequence ofthese genes, it has been proposed that they may each encode all three enzymaticactivities required for UDP-Rha synthesis (Reiter and Vanzin, 2001). Blastsearchers have also identified other Arabidopsis genes with sequence similarityto individual E. coli genes that encode only one of the enzymatic activitiesrequired for dTDP-Rha synthesis (see Reiter and Vanzin, 2001). However,direct proof that any of these Arabidopsis putative UDP-Rha biosynthetic genesactually encodes a UDP-Rha biosynthetic protein has not yet been reported.

3.4.4 Uridine diphosphate--d-galactose (UDP-Gal)UDP-Gal is a substrate for the synthesis of RG-I and RG-II. UDP-Gal isformed from UDP-Glc by a 4-epimerization catalyzed by UDP-Glc 4-epimerase(EC 5.1.3.2) (Fan and Feingold, 1969; Feingold and Avigad, 1980). The reac-tion mechanism includes an enzyme-bound UDP-4-keto-hexose intermediate(Maxwell, 1957; Maitra and Ankel, 1971; Wee and Frey, 2001) which bindsthe enzyme approximately 100 times more tightly than UDP-Glc (Feingold andAvigad, 1980; Wee and Frey, 2001).

UDP-GlcUDP-Glc 4-epimerase UDP-Gal (4)

The structure of the enzyme from E. coli has been determined by X-ray crys-tallography (Bauer et al., 1992). The bacterial enzyme comprises two identical39.5 kDa subunits (Wilson and Hogness, 1969; Bauer et al., 1992), each of whichbinds a NAD+ cofactor (Bauer et al., 1992). Each subunit folds into a distinctN-terminal domain, primarily responsible for NAD+/NADH positioning, thathas a seven-stranded parallel -pleated sheet flanked on either side by -helices.The small C-terminal motif is responsible for binding the UDP-sugar (Thodenet al., 1996a, 1996b; Thoden and Holden, 1998). The active site is locatedbetween the two domains. The size of the enzyme varies in different species, asdoes the tightness by which the enzyme binds NAD+. For example, the bovineenzyme is a monomer of 40 kDa that requires exogenous NAD+ for activitywhile the UDP-d-Glc 4-epimerase from Candida pseudotropicalis is made upof two identical 60 kDa subunits each of which contains one tightly boundNAD+ (Maxwell, 1957; Geren and Ebner, 1977; Feingold and Avigad, 1980).


The UDP-Glc 4-epimerase from leaves of Vicia faba is a soluble cytoplasmicprotein with a pH optimum of 8.8 and an apparent Km for UDP-Gal of 95M(Konigs and Heinz, 1974). A UDP-Glc 4-epimerase purified from wheat germextract has Mr of 100 000 and requires NAD+ for activity (Fan and Feingold,1969; Feingold and Avigad, 1980). The tightness of binding of NAD+ to plantUDP-Glc 4-epimerase appears to be species-specific (Feingold and Avigad,1980) and both soluble and membrane-bound activities have been recoveredin plants (Feingold and Avigad, 1980). An Arabidopsis gene for UDP-Glc 4-epimerase (UGE1) has been cloned and expressed in E. coli (Dormann andBenning, 1996). The Arabidopsis expressed protein encodes a 39 kDa proteinwith a broad pH optimum from 7.0 to 9.55 and an apparentKm for UDP-Glc of110M (Dormann and Benning, 1996). A cDNA encoding a predicted 39 kDaputative UDP-Glc 4-epimerase from pea (Pisum sativum) that has 92% sequencehomology to the Arabidopsis gene has also been cloned (Lake et al., 1998)although no characteristics of the enzyme have been reported. Two cDNAs thatencode UDP-Glc epimerases of 39.3 and 38.4 kDa from developing seeds ofguar (Cyamopsis tetragonoloba) endosperm have also been reported (Joersboet al., 1999). Analysis of the Arabidopsis genome has led to the identificationof four coding regions with significant sequence identity to UGE1 (Reiter andVanzin, 2001), two of which (UGE2 and UGE3) encode functional UDP-Glcepimerases (Reiter and Vanzin, 2001).

3.4.5 Uridine diphosphate--d-glucuronic acid (UDP-GlcA)UDP-d-Glucuronic acid is the likely substrate for the incorporation of GlcA intoRG-II and into some side branches of RG-I. UDP-GlcA is produced either bythe oxidation of UDP-Glc catalyzed by UDP-Glc 6-dehydrogenase (Feingoldet al., 1960; Feingold and Avigad, 1980) or by the uridylation of Glc-1-P viathe myoinositol pathway (Feingold and Avigad, 1980).

UDP-d-Glc UDP-glucose dehydrogenaseUDP-d-GlcA (5)

UDP-Glc 6-dehydrogenase (EC 1.1.1.22) catalyzes the 4-electron oxidationof UDP-Glc at C-6 and the reduction of two moles of NAD+ (Feingold andAvigad, 1980; Feingold and Franzen, 1981). The reaction is ordered: beginningwith binding of UDP-Glc, followed by binding of NAD+ (Feingold and Avigad,1980; Hempel et al., 1994; Campbell et al., 2000), reduction of the first boundNAD+ and release of the first NADH. This is followed by binding of the secondNAD+, reduction and release of the second NADH and finally release of theUDP-GlcA (Feingold and Avigad, 1980; Campbell et al., 1997). Bovine UDP-Glc 6-dehydrogenase consists of six 52 kDa subunits (Zalitis and Feingold,1969; Gainey et al., 1972; Franzen et al., 1978; Feingold and Avigad, 1980;Jaenicke et al., 1986; Hempel et al., 1994) with one mole of substrate bound per


two moles of enzyme (i.e. half-of-the-site behavior) (Franzen et al., 1978;Hempel et al., 1994). In contrast, UDP-Glc 6-dehydrogenase from E. coliconsists of two identical subunits of 50 kDa each (Schiller et al., 1976; Feingoldand Franzen, 1981). The X-ray crystal structure of UDP-glucose dehydroge-nase from the bacteria Streptococcus pyogenes has been solved (Campbellet al., 2000). The S. pyogenes UDP-Glc dehydrogenase appears to exist aseither a monomer or dimer in solution. Each monomer consists of two discrete/ domains connected by a long -helix. Each domain consists of a core-sheet sandwiched between -helices. The N-terminal domain contains a six-stranded parallel -sheet that binds NAD+ (Campbell et al., 2000). UDP-Glc 6-dehydrogenases have been purified 1000-fold from pea ((Stromingerand Mapson, 1957), 12-fold from germinating lily (Lilium longiflorum) pollen(Davies and Dickinson, 1972), 62-fold from soybean nodules (Stewart andCopeland, 1998), and 341-fold from elicitor-treated French bean (Phaseolusvulgaris L) cell suspensions (Robertson et al., 1996). The apparent Km valuesfor UDP-Glc for the different enzymes were 70M, 300M, 50M, and5.5 mM, respectively, and the apparent Km values for NAD+ were 115M,400M, 120M and 20M. The soybean enzyme has a pH optimum of 8.4,a native molecular mass of 272 kDa, and a subunit molecular mass of 47 kDa,suggesting that the enzyme functions as a hexamer (Stewart and Copeland,1998). All known eukaryotic UDP-Glc 6-dehydrogenases are cooperativelyinhibited by UDP-Xyl, suggesting a feedback inhibition of the enzyme byUDP-Xyl (Feingold and Avigad, 1980; Campbell et al., 1997). A cDNA clonefor UDP-Glc 6-dehydrogenase from soybean that is highly homologous tothe cloned bovine UDP-GlcDH gene (Hempel et al., 1994) encodes a proteinwith a predicted molecular mass of 52.9 kDa (Tenhaken and Thulke, 1996).The soybean gene has a conserved NAD+-binding site motif and contains thecatalytic Cys residue (Hempel et al., 1994; Tenhaken and Thulke, 1996). AnArabidopsis gene (UGD) encoding UDP-Glc dehydrogenase has been identifiedand its gene expression has been studied using -glucuronidase and green fluo-rescent protein reporter constructs (Seitz et al., 2000). Three additional putativeUDP-Glc dehydrogenases have been identified in Arabidopsis via sequenceanalysis (Reiter and Vanzin, 2001). The four genes share 8393% sequenceidentity (Seitz et al., 2000). The protein with UDP-Glc 6-dehydrogenase activitythat was purified from French bean (Robertson et al., 1996) does not sharethe characteristics of the cloned UDP-GlcDH from soybean (Strominger andMapson, 1957) or Arabidopsis (Seitz et al., 2000). The putative UDP-GlcDHfrom French bean (Robertson et al., 1996) has a molecular mass of 40 kDa, a highapparent Km of 5.5 mM for UDP-Glc, co-purifies with alcohol dehydrogenaseactivity and is preferentially located in cells that make secondary walls. It isunclear whether the 40 kDa protein from French bean represents a bona fidemultifunctional UDP-GlcDH preferentially expressed during secondary wallsynthesis (Robertson et al., 1996), or whether it is an alcohol dehydrogenase


that can oxidize UDP-Glc in vitro but plays little or no role in the formation ofUDP-GlcA in planta.

3.4.6 Uridine diphosphate--d-xylose (UDP-Xyl)UDP-Xyl is the expected substrate for the synthesis of xylogalacturonan andRG-II. UDP-Xyl is produced by the decarboxylation of UDP-GlcA catalyzedby UDP-GlcA carboxylase (EC 4.1.1.35) (Feingold et al., 1960; Feingold andAvigad, 1980; Hayashi et al., 1988; Hannapel, 1991).

UDP-d-GlcA UDP-GlcA decarboxylaseUDP-d-Xyl (6)UDP-GlcA decarboxylase contains a tightly-bound NAD+ and catalyzes a

reaction that proceeds via a UDP-4-keto-hexose intermediate (Feingold andAvigad, 1980). Partially purified UDP-GlcA decarboxylase from wheat germ(John et al., 1977) has a pH optimum of 7.0 and consists of two 210 kDaisoenzymes that do not require exogenous NAD+ for activity. Both isozymes areactivated by low (


species can convert GDP-d-Man to GDP-l-Fuc (Liao and Barber, 1971). Thesynthesis of GDP-l-Fuc using enzyme preparations from Phaseolus vulgarisrequires NADPH or NADH, occurs at a pH optimum of 6.97.8, has an apparentKm for GDP-d-Man of 160M (Liao and Barber, 1971) and an apparentmolecular mass of 120 kDa (Liao and Barber, 1972). The reaction proceedsvia the C-4 oxidation and C-6 reduction of GDP-Man catalyzed by GDP-d-Man 4,6-dehydratase (EC 4.2.1.47) (Liao and Barber, 1971; Feingold andAvigad, 1980). The product formed, GDP-4-keto-6-deoxy-d-mannose, appearsto tightly bind a GDP-4-keto-6-deoxy-d-Man 3,5-epimerase, which convertsit to a GDP-4-keto-6-deoxy-l-galactose intermediate (Feingold and Avigad,1980; Bonin et al., 1997) that is then reduced by a GDP-4-keto-l-fucose reduc-tase activity to yield GDP-l-fucose (Feingold and Avigad, 1980). Although itwas previously thought that the final two enzyme activities might reside onseparate proteins, it has been shown in humans (Sullivan et al., 1998) andin transgenic Saccharomyces cerevisiae harboring the E. coli genes (Mattilaet al., 2000) that a single enzyme, GDP-keto-6-deoxymannose 3,5-epimerase-4-reductase, catalyzes both the epimerization and reduction steps. The firstenzyme in the pathway, GDP-d-Man-4,6-dehydratase, is inhibited by GDP-fucose (Sullivan et al., 1998; Kornfeld and Ginsburg, 1966), indicating feedbackinhibition.

GDP-d-Man GDP-d-Man 4,6-dehydratase GDP-4-keto-6-deoxy-d-mannoseGDP-keto-6-deoxymannose3,5-epimerase-4-reductase GDP-Fuc (7)

The Arabidopsis mutant mur1 is defective in the synthesis of l-fucose inthe aerial parts of the plant (Reiter et al., 1993). The MUR1 gene encodesa 41.9 kDa GDP-d-mannose 4,6-dehydratase (Bonin et al., 1997). A secondArabidopsis gene, GMD1, encodes a second GDP-d-mannose 4,6-dehydratasethat is highly expressed in roots (Bonin et al., 1997). The Arabidopsis gene GER1encodes a GDP-keto-6-deoxymannose 3,5-epimerase-4-reductase (Bonin andReiter, 2000). A second putative GDP-keto-6-deoxymannose 3,5-epimerase-4-reductase gene, GER2, with 88% amino acid identity to GER1 has been identi-fied by DNA sequence analysis (Reiter and Vanzin, 2001). The dwarf phenotypeassociated with mur1-1 and mur1-2 was shown to be due to a substitution of l-galactose for the l-fucose and 2-O-methyl-l-galactose for 2-O-methyl-l-fucosein RG-II. This change in RG-II structure results in a reduction in the amountof borate crosslinked RG-II dimer (ONeill et al., 2001) that is present in thewalls and leads to the dwarfism. This result provides unequivocal proof that thepectic polysaccharide RG-II is essential for normal plant growth (ONeill et al.,2001).


3.4.8 Uridine diphosphate--d-apiose (UDP-apiose)UDP-apiose is a substrate for the synthesis of the species-specific substitutedgalacturonan apiogalacturonan that is found in some aquatic monocotyledonousplants such as Spirodela polyrrhiza (Watson and Orenstein, 1975) and Lemnaminor (Hart and Kindel, 1970). Apiogalacturonan is a homogalacturonan inwhich d-apiose or apiobiose (d-Apif-1,3-d-apiose) are attached to O-2 or O-3 of HGA. UDP-apiose is also the likely substrate for the synthesis of RG-II(ONeill et al., 2001; Ridley et al., 2001). UDP-apiose is formed by a decarboxy-lation and rearrangement of UDP-GlcA catalyzed by a NAD+-dependent UDP-apiose/UDP-Xyl synthase (Wellmann and Grisebach, 1971; Baron et al., 1973;Kindel and Watson, 1973; Watson and Orenstein, 1975; Matern and Grisebach,1977; Feingold and Barber, 1990). UDP-Xyl is also a product of the in vitroenzymatic reaction (Matern and Grisebach, 1977) with UDP-apiose:UDP-Xylratios of 1.4 reported (Matern and Grisebach, 1977). The UDP-apiose synthaseand UDP-Xyl synthase activities could not be separated in a 1400-fold purifiedprotein preparation, leading to the suggestion that a single multifunctionalprotein is responsible for both activities (Wellmann and Grisebach, 1971; Maternand Grisebach, 1977). However, it has more recently been suggested that xylosemay be an artificial product recovered in in vitro reactions (Gardiner et al., 1980)and the name UDP-apiose synthase has been used for the enzyme (Gardineret al., 1980). It is believed that UDP-apiose synthesis occurs via the formationof an l-threo-4-pentosulose intermediate, common to both UDP-apiose andUDP-Xyl formation, followed by ring contraction and epimerization (Maternand Grisebach, 1977).

UDP-d-GlcA UDP-apiose synthaseUDP-d-apioseUDP-Xyl (possible in vitro side product?) (8)

Partially purified UDP-apiose/UDP-Xyl synthase from Lemna minor hasoptimum activity at 1 mM NAD+ and a pH of 8.08.3 (Kindel et al., 1971).Partially purified UDP-apiose/UDP-Xyl synthase from parsley (Matern andGrisebach, 1977) is composed of an 86 kDa protein consisting of two identical44 kDa subunits and a 65 kDa protein consisting of two identical 34 kDa subunits(Matern and Grisebach, 1977). The 86 kDa protein contains all the enzymeactivity, binds 0.5 mol of UDP-GlcA per mol of protein and, in the presenceof UDP-GlcA, binds 0.5 mol NAD+ per mol of catalytic protein (Matern andGrisebach, 1977). The 65 kDa protein is enzymatically inactive but was reportedto be required for stability of the 86 kDa protein (Matern and Grisebach, 1977).UDP-d-glucose and UDP-methyl-d-GlcA are competitive inhibitors of UDP-apiose/UDP-Xyl synthase (Gebb et al., 1975).


3.4.9 Uridine diphosphate--l-galactose (GDP-Gal)GDP-l-Gal is a possible substrate for the l-Gal in RG-II and for the l-Gal thatis substituted for the l-Fuc (Zablackis et al., 1996) in xyloglucan synthesized inArabidopsis mur1 mutants. Mur1 mutants have a mutation in GDP-d-mannose4,6-dehydratase and, thus, have reduced amounts of l-Fuc in the aerial portionsof the plant that is replaced with l-Gal (Zablackis et al., 1996). It has beenproposed that GDP-d-Man is converted to GDP-l-Gal by GDP-d-mannose 3,5-epimerase (Feingold and Avigad, 1980).

GDP-d-ManGDP-d-mannose 3,5-epimerase GDP-l-Gal (9)

The reversible 3,5-epimerization of GDP-Man has been reported usingextracts from Chlorella pyrenoidosa (Hebda et al., 1979; Hebda and Barber,1978). The partially purified Chlorella GDP-d-mannose 3,5-epimerase had amolecular mass of 100 kDa, a broad pH optimum centering at 8.1, an appar-ent Km of 96M for GDP-d-Man and 97M for GDP-l-Gal, and anequilibrium constant of 2.9 in the direction of GDP-Man (Hebda et al.,1979).

3.4.10 XXX-Kdo, XXX-Dha and XXX-aceric acidThe identity and biosynthetic pathways for the activated glycosyl donors of theKdo, Dha and aceric acid in RG-II have not been experimentally established inplants. In contrast, a great deal of information is available regarding the synthe-sis in bacteria of cytidine 5-monophosphate-3-deoxy-d-manno-octulosonate(CMP-Kdo), the activated donor of the Kdo found in lipopolysaccharides andother extracellular bacterial polysaccharides (Unger, 1981; Raetz, 1990; Pazzaniet al., 1993; Baasov and Kohen, 1995; Rosenow et al., 1995; Jelakovic et al.,1996; Jelakovic and Schulz, 2001). Assuming that plants use CMP-Kdo tosynthesize RG-II, and that they synthesize CMP-Kdo using a similar pathway asbacteria, the following pathway is proposed. d-Ribulose 5-phosphate is isomer-ized to d-arabinose 5-phosphate by d-arabinose-5-phosphate isomerase (Unger,1981). Thed-arabinose 5-phosphate is condensed with phosphoenolpyruvate byKdo-8-phosphate synthetase (2-dehydro-3-deoxyphosphooctonate aldolase) toform Kdo 8-phosphate (2-dehydro-3-deoxy-d-octonate 8-phosphate) (Unger,1981; Doong et al., 1991). The 8-phosphate is removed from Kdo 8-phosphateby Kdo-8-phosphate phosphatase to produce Kdo and inorganic phosphate(Unger, 1981). Finally, CMP-Kdo and pyrophosphate are formed from cytidine5-triphosphate (CTP) and Kdo by CMP-Kdo synthetase (Unger, 1981). Kdo-8-phosphate synthetase has been identified in multiple plant species and hasbeen partially purified from spinach (Doong et al., 1991). Kdo-8-phosphate


synthetase has a pH optimum of 6.2, an apparent Km of 270M for arabinose5-phosphate and an apparent Km of 35M for phosphoenolpyruvate (Doonget al., 1991). A cDNA from pea has been identified that encodes a 31.7 kDafunctional Kdo-8-phosphate synthetase when expressed in E. coli (Brabetz et al.,2000). The expressed pea enzyme has a pH optimum of 6.1.

The identity and the biosynthetic pathway in plants for the activated glycosyldonor for Dha (3-deoxy-d-lyxo-2-heptulosaric acid) is not known. However,it has been proposed that 3-deoxy-d-arabino-heptulosonate 7-phosphate syn-thase could catalyze the condensation of phosphoenolpyruvate with threoseto generate a precursor of Dha (Doong et al., 1992). A cytosolic form of 3-deoxy-d-arabino-heptulosonate 7-phosphate synthase with a wide substratespecificity has been identified in plants (Doong et al., 1992). cDNAs encodingplastidic 3-deoxy-d-arabino-heptulosonate 7-phosphate synthases involved inthe shikimate pathway leading to aromatic secondary metabolism have beenidentified from potato (Solanum tuberosum L.) (Dyer et al., 1990) and otherplant species (Herrmann, 1995). An alternative route for the synthesis of theactivated donor of Dha might be the interconversion of CMP-Kdo to CMP-Dha through oxidation and decarboxylation reactions. There is no informationavailable regarding the nature or mode of synthesis of the activated donor foraceric acid.

3.5 Glycosyltransferases involved in pectin biosynthesis

Elucidating the mechanism by which polysaccharides are synthesized is a chal-lenging endeavor. Polymer synthesis involves at least three stages: initiation,elongation and termination. While nothing is known about the initiation andtermination of pectin synthesis, considerable effort has been directed towardsidentifying and characterizing glycosyltransferases that presumably catalyze theelongation of pectin. Such glycosyltransferases transfer a glycosyl residue froman activated sugar donor (e.g. a nucleotide-sugar) either onto endogenous pecticacceptors in the Golgi or onto exogenous oligosaccharide or polysaccharidepectic acceptors in in vitro reactions using permeabilized Golgi or microsomesor detergent-solubilized enzymes. Although it has recently been proposed to beuseful to make a distinction between the terms glycosyltransferase and glycansynthase (Perrin et al., 2001) when discussing polysaccharide synthesis, itappears premature at this time to make this distinction for the pectin biosyntheticenzymes. Glycan synthases have been defined as enzymes that synthesize thebackbone of a polysaccharide (i.e. the 1,4-linked d-galactosyluronic acid backbone of HGA or the alternating [-d-GalA-1,2--l-Rha-1,4] backbone ofRG-I). It is often presumed that such enzymes should act processively, thatis, catalyze multiple rounds of catalysis before releasing their oligosaccha-ride/polysaccharide acceptor. It is likely that such processivity, if it occurs


for pectin synthesis, could require protein complexes. Such complexes may,or may not, remain intact in the homogenates and fractions used for studyingpectin synthesis. Since both RG-I and the substituted regions of HGA (e.g.RG-II) are highly branched, it is not clear that it would be more favorable foroverall biosynthetic rates and structure fidelity for pectin synthesis to occurprocessively or distributively. No evidence for in vitro processivity of pectinsynthesis has yet been reported. For these reasons all the pectin biosyntheticenzyme activities identified to date are defined in this review as glycosyltrans-ferases. If these glycosyltransferases are eventually shown to act processivelywhen associated with other polypeptides in complexes, then it is recommendedthat the appropriate glycan synthase name be used for the holoenzyme complex.Alternatively, if the glycosyltransferases identified to date are eventually shownto act processively under specific reaction conditions (e.g. in the presence ofspecific cofactors or substrates), then it is recommended that these enzymesbe named their corresponding glycan synthase, as put forward by Perrin et al.(2001).

As mentioned above, it is not known whether all of the pectic polysaccharidesare synthesized as a single polysaccharide or whether separate populations ofpolymers are synthesized. It is likely that the substituted galacturonan namedRG-II is synthesized in the same polymer that contains contiguous regionsof HGA, since RG-II isolated from cell walls is extended on both ends byregions of HGA. It is also possible that some HGA is synthesized withoutsubstituted HGA regions. It is less clear whether the syntheses of HGA and RG-Ioccur on the same polysaccharide chain or whether they occur as independentbiosynthetic events. Work from the groups of Voragen (Schols et al., 1995)and Mort (Yu and Mort, 1996), suggesting that RG-I is covalently attached toxylogalacturonan-like regions, indicates that RG-I and HGA may exist as asingle polymer. In the following discussion, the enzymes activities required forthe synthesis of pectin will be divided into those required for HGA synthesis,substituted galacturoronan synthesis (including RG-II and xylogalacturonan),and RG-I synthesis.

3.5.1 Synthesis of homogalacturonanHomogalacturonan (HGA) is a partially methyl-esterified and acetylated homo-polymer of 1,4-linked d-galactosyluronic acid (Ridley et al., 2001). It isunclear how much the degree of polymerization (DP) of HGA varies withinpectin; however, a DP range of 72100 has been reported (Thibault et al.,1993). As shown in Table 3.2, the synthesis of HGA requires at least onehomogalacturonan1,4-galacturonosyltransferase (HGA-GalAT) (also referredas polygalacturonate: 1,4-galacturonosyltranferase), a homogalacturonan-methyltransferase (HGA-MT) (also referred to as pectin methyltransferase),and a homogalacturonan 3-O-acetyltransferase (HGA-AT).


Table 3.2 Glycosyltransferase activities required for HGA biosynthesisa

Enzymeb

Type of transferase Acceptor substrate Enzyme activity Reference for structureGlycosyl-d-GalAT GalA1,4-GalA 1,4-GalAT ONeill et al. (1990)Methyl-HGA methyltransferase GalA1,4-GalA(n) HGA-MT Goubet et al. (1998);(HGA-MT) Vannier et al. (1992);

Kauss et al. (1967)Acetyl-HGA: GalA 3-O- GalA1,4-GalA(n) HGA-AT Ishii (1997); De Vries et al.acetyltransferase (1986); Ishii (1995);(HGA-AT) Rombouts and Thibault (1986)aAdapted from Ridley et al. (2001).bAll sugars are d sugars and have pyranose rings unless otherwise indicated. Glycosyltranferases add tothe glycosyl residue on the left of the indicated acceptor.

3.5.1.1 Homogalacturonan galacturonosyltransferase (HGA-GalAT)Membrane-bound 1,4-galacturonosyltransferase (GalAT) activity has beenidentified and partially characterized in mung bean (Villemez et al., 1966;Kauss and Swanson, 1969), tomato (Lin et al., 1966), turnip (Lin et al., 1966),and sycamore (Bolwell et al., 1985), tobacco suspension (Doong et al., 1995),radish roots (Mohnen et al., 1999), enriched Golgi from pea (Sterling et al.,2001), Azuki bean (Vigna angularis) (Takeuchi and Tsumuraya, 2001) andArabidopsis (Sterling and Mohnen, unpublished results) (see Table 3.3). TheGalAT from pea has been localized to the Golgi (Sterling et al., 2001) withits catalytic site facing the lumenal side of the Golgi, (Sterling et al., 2001).These results provide the first direct enzymatic evidence that the synthesisof HGA occurs in the Golgi. In in vitro reactions, GalAT adds [14C]GalAfrom UDP-[14C]GalA (Liljebjelke et al., 1995) onto endogenous acceptors inmicrosomal membrane preparations to produce radiolabeled products of largemolecular mass (i.e. 105 kDa in tobacco microsomal membranes (Doonget al., 1995) and 500 kDa in pea Golgi (Sterling et al., 2001)). The cleav-age of up to 89% of the radiolabeled product into GalA, digalacturonic acid(diGalA) and trigalacturonic acid (triGalA) following exhaustive hydrolysiswith a purified endopolygalacturonase confirmed that the product synthesizedby tobacco GalAT was largely HGA. The product produced in vitro in tobaccomicrosomes is 50% esterified (Doong et al., 1995), while the product pro-duced in pea Golgi did not appear to be esterified (Sterling et al., 2001).These results suggest that the degree of methyl-esterification of newly synthe-sized HGA may be species-specific and that methyl-esterification occurs afterthe synthesis of at least a short stretch of HGA. GalAT activity in detergent-permeabilized microsomes from etiolated azuki bean seedlings adds [14C]GalA


Table 3.3 Comparison of catalytic constants and pH optimum of HGA 1,4-galacturonosyl-transferasesa,b

ApparentKm for Vmax

UDP-GalA pH (pmol mg1Enzymeb Plant source (M) optimum min1) ReferenceGalATa Mung bean 1.7 6.0 4700 Villemez et al. (1966)GalAT Mung bean n.d. n.d. n.d. Crombie and Reid (2001)GalAT Pea n.d.e 6.0 n.d. Cumming and Brett (1986)GalAT Pea n.d. n.d. n.d. Sterling et al. (2001)GalAT Sycamore 770 n.d. ? Bolwell et al. (1985)GalAT Tobacco 8.9 7.8 150 Doong et al. (1995)GalAT (sol)c Tobacco 37 6.37.8 290 Doong and Mohnen (1998)GalAT (per)d Azuki bean 140 6.87.8 2700 Takeuchi and Tsumuraya

(2001)aAdapted from Mohnen (1999).bUnless indicated, all enzymes are measured in particulate preparations.c(sol): detergent-solubilized enzyme.d(per): detergent-permeabilized enzyme.en.d.: not determined.

from UDP-[14C]GalA onto acid-soluble polygalacturonate (PGA) exogenousacceptors (Takeuchi and Tsumuraya, 2001). Treatment of the radiolabeled prod-uct with a purified fungal endopolygalacturonase yielded GalA and diGalA,confirming that the activity identified was GalAT. The azuki bean enzymehas a broad pH range of 6.87.8 and a surprisingly high specific activity of13002000 pmol mg1 min1, especially considering the large amount (3.14.1 nmol mg1 min1) of polygalacturonase activity that was also present in themicrosomal preparations. As with the product made by tobacco, no evidencefor the processive transfer of galactosyluronic acid residues onto the acceptorwas obtained (see below).

GalAT can be solubilized from membranes with detergent (Doong andMohnen, 1998). Solubilized GalAT adds GalA onto the nonreducing end(Scheller et al., 1999) of exogenous HGA acceptors of degrees of polymeri-zation 10 (Doong et al., 1995). The bulk of the HGA elongated in vitro bysolubilized GalAT from tobacco membranes (Doong and Mohnen, 1998) anddetergent-permeabilized Golgi from pea (Sterling et al., 2001) is elongated bya single GalA residue. These results suggest that solubilized GalAT in vitroacts nonprocessively, (i.e. distributively). The lack of in vitro processivity ofthe solubilized GalAT may indicate that the enzyme does not synthesize HGAin a processive manner in vivo, or it may be an artifact due to the dissociationof a required biosynthetic complex or cofactor(s)/substrate(s) during solubi-lization of the enzyme. Attempts to recover processive in vitro GalAT activityby using alternative pectic acceptors or high concentrations (up to 10 mM) of


UDP-GalA were not successful (H.F. Quigley and D. Mohnen, unpublishedresults; Ridley et al., 2001). There is also no evidence that the inclusion of themethyl donor S-adenosylmethionine (Takeuchi and Tsumuraya, 2001; Kaussand Swanson, 1969; Doong et al., 1995) and/or the acetyl donor acetyl-CoApromotes the processivity of GalAT (H.F. Quigley and D. Mohnen, unpublishedresults; Ridley et al., 2001). Thus, the question of whether or not GalAT in vivois processive remains to be resolved. The gene for HGA-GalAT has not yetbeen identified, although efforts to purify the enzyme are on-going in severallaboratories.

3.5.1.2 HGA methyltransferase (HGA-MT)The methyl-esterification of HGA at the C-6 carboxyl group is catalyzed byHGA methyltransferase (HGA-MT). Although the enzyme has been referredto as pectin methyltransferase, the term HGA-MT is preferred to distinguishHGA-MT from the enzymes that methylate RG-I or RG-II. HGA-MT hasbeen identified in microsomal membranes from mung bean (Kauss et al., 1967,1969; Crombie and Reid, 1998), flax (Vannier et al., 1992; Schaumann et al.,1993), tobacco (Goubet et al., 1998), and soybean (Ishikawa et al., 2000) (seeTable 3.4).

Membrane-bound HGA-MTs from flax (Bruyant-Vannier et al., 1996;Bourlard et al., 1997) and tobacco (Goubet and Mohnen, 1999b) have been sol-ubilized using detergent. Two apparent HGA-MT isozymes, PMT5 and PMT7,from flax have been reported with pH optima of 5.0 and 6.5, respectively(Bourlard et al., 2001). Efforts to purify these apparent isozymes resultedin the identification of a small additional polypeptide with HGA-MT activ-ity designated PMT18. The isoelectric points and apparent molecular massesof PMT5, PMT 7 and PMT18 were 5.86.5, 8.79.2 and 4.04.5, and 40,110 and 18 kDa, respectively (Bourlard et al., 2001). It is proposed that the18 kDa protein is a subunit of the 40 and 110 kDa proteins, based on theappearance of the larger proteins when PMT18 is rechromatographed by size-exclusion chromatography and by the appearance of an 18 kDa band whenPMT5 and PMT7 are separated by SDS-polyacrylamide gel electrophoresis(Bourlard et al., 2001). Furthermore, photoaffinity labeling of PMT5 and PMT7with [3H]S-adenosylmethionine yielded a single labeled band at 18 kDa. Noinformation on the gene encoding HGA-MT is available.

HGA-MT is localized to the Golgi (Vannier et al., 1992; Bourlard et al.,1997b; Baydoun et al., 1999; Goubet and Mohnen, 1999a) with its catalytic sitefacing the Golgi lumen (Goubet and Mohnen, 1999a). Available biochemicalevidence suggests that at least a small stretch of HGA is synthesized priorto its methylation by HGA-MT in the Golgi. This conclusion is based onstudies with intact membranes which show that UDP-GalA stimulates HGA-MTactivity (Kauss and Swanson, 1969; Goubet et al., 1998) and from studies withdetergent-permeabilized membranes and solubilized HGA-MT which show that


Table 3.4 Comparison of catalytic constants and pH optimum of HGA methyltransferasesa

Apparent ApparentKm for molecular

Plant SAMd pH massEnzymeb,c source (M) optimum V emax (kDa) ReferenceHGA-MTb Mung 59 6.67.0 2.7f Kauss and Hassid (1967);

bean Kauss et al. (1969);Kauss et al. (1967)

PMT Flax 1030 6.8 n.d.h Schaumann et al. (1993);Vannier et al. (1992)

PMT (sol)g Flax 0.5 7.1i or n.d. Bourlard et al. (1997a,b);5.5j Bruyant-Vannier et al.

(1996)HGA-MT Tobacco 38 7.8 49 Goubet et al. (1998)HGA-MT (sol) Tobacco 18 7.8 7.3 Goubet and Mohnen

(1999b)PMT-MT Soybean 230 6.8 1360 Ishikawa et al. (2000)PMT-MT5k Flax n.d. 5.0 n.d. 40 Bourlard et al. (2001)PMT-MT7k Flax n.d. 6.5 n.d. 110 Bourlard et al. (2001)PMT-MT18k Flax n.d. n.d. n.d. 18 Bourlard et al. (2001)aAdapted from Ridley et al. (2001).bUnless indicated, all enzymes are measured in particulate preparations.cAll enzymes are thought to be HGA-MT. However, since some authors use the previous name (pectinmethyltransferase) it is included for clarity.dApparent Km for S-adenosylmethionine.eVmax in pmol min1 mg1 protein.fVmax is calculated from data in Kauss et al. (1969).g(sol): detergent-solubilized enzyme.hn.d.: not determined.iFrom Bruyant-Vannier et al. (1996).jFrom Bourlard et al. (1997a,b).kPurified enzymes.

polygalacturonic or pectin are acceptors for HGA-MT in vitro. Some of theHGA-MTs in detergent-permeabilized membranes from flax and soybean showa preference for partially esterified pectin (Bourlard et al., 1997b; Ishikawaet al., 2000; Bourlard et al., 2001) over polygalacturonic acid. These resultssuggest that multiple HGA-MTs may exist that differ in their specificity forHGA of differing degrees of methylation. Such HGA-MTs may be preferentiallyinvolved in the initial methylation of HGA or in the methylation of more highlyesterified HGA.

3.5.1.3 HGA acetyltransferase (HGA-AT)The GalA residues in HGA may, depending upon the species, be partially O-acetylated at C-2 or C-3 (Ishii, 1995; Ishii, 1997). Pectin O-acetyltransferaseactivity has been identified in microsomes from suspension-cultured potatocells (Pauly and Scheller, 2000). The incubation of potato microsomes with


[14C]acetyl-CoA yielded a salt/ethanol precipitable product from which approxi-mately 8% of the radioactivity could be solubilized by treatment with endopoly-galacturonase and pectin methylesterase. These results suggest that 8% of theradiolabeled acetate was transferred either onto HGA or onto solubilized RG-II or RG-I fragments. It remains to be shown whether the described activ-ity represents HGA-AT or an enzyme that acetylates one of the other pecticpolysaccharides that may be covalently linked to HGA.

3.5.2 Synthesis of substituted homogalacturonansThere have been no reports of a systematic effort to study the glycosyltrans-ferases required for the synthesis of RG-II, the most structurally complicatedpolysaccharide in the cell wall. As shown in Table 3.5, at least 24 transferaseactivities are likely required to synthesize RG-II. It is possible that the apio-syltransferase identified in the studies of apiogalacturonan synthesis in Lemnais related to, or is the same apiosyltransferase(s) as that required for RG-IIsynthesis (see below), however, this remains to be shown.3.5.2.1 Rhamnogalacturonan-II methyltransferase (RG-II-MT)The GalA in the HGA backbone of RG-II may be partially methyl-esterified. Adetergent-solubilized pectin methyltransferase activity from suspension-cultured flax cells was identified that could transfer methyl groups fromS-adenosylmethionine onto RG-II isolated from wine (Bourlard et al., 1997a).The addition of RG-II to enzyme reactions gave a 7-fold stimulation of methyl-transferase activity above the level obtained in the absence of exogenous accep-tor. The radiolabeled product had a size similar to RG-II monomers and RG-IIdimers (Bourlard et al., 1997; Ridley et al., 2001). It has not yet been shownwhere in RG-II the methyl group was added. The identified methylation couldrepresent methyl-esterification of the HGA backbone of RG-II. Alternatively,it could represent methyl-etherification of RG-II since RG-II contains methylgroups on non-galacturonic glycosyl residues (e.g. 2-O-methylxylose and 2-O-methylfucose (Darvill et al., 1978; ONeill et al., 1996)) of side chain residues.More research is required to determine the exact location of the methylationand of the identity of the potentially novel enzyme activity reported.

3.5.2.2 Apiogalacturonan apiosyltransferaseThe substituted galacturonan known as apiogalacturonan is produced in someaquatic monocotyledonous plants (Hart and Kindel, 1970;Watson and Orenstein,1975). Apiogalacturonan has apiose or apiobiose (d-Apif-1,3-d-apiose)attached to O-2 or O-3 of HGA (Hart and Kindel, 1970; Watson and Orenstein,1975). Table 3.6 shows some of the transferase activities likely to be requiredto synthesize apiogalacturonan. The anomeric configuration of the glycosidiclinkage of apiose to HGA may be in the configuration (Watson and Orenstein,1975). It is not known how apiogalacturonan is related to RG-II, which has


Table 3.5 Glycosyltransferase activities likely to be required for RG-II biosynthesisa,b

EnzymedType of RG-IIglycosyl side Parent Acceptor Enzyme Reference fortransferase chainc polymer substrate activity structured-GalAT HGA/RG-II4 GalA1,4-GalA 1,4-GalAT ONeill et al. (1990)d-GalAT A RG-II l-Rha1,3-Apif 1,2-GalAT ONeill et al. (1997);

Carpita and Gibeaut(1993)

d-GalAT A RG-II l-Rha1,3-Apif 1,3-GalAT ONeill et al. (1997);Carpita and Gibeaut(1993)

l-RhaT A,B RG-II Apif1,2-GalA 1,3-l-RhaT ONeill et al. (1997);Carpita and Gibeaut(1993)

l-RhaT C RG-II Kdo2,3-GalA 1,5-l-RhaT ONeill et al.(2001, 1997);Carpita and Gibeaut(1993)

l-RhaT B RG-II l-Ara1,4-Gal 1,2-l-RhaT ONeill et al. (1997);Carpita and Gibeaut(1993)

l-RhaT B2 RG-II l-Ara1,4-Gal 1,3-l-RhaTl-GalT B RG-II GlcA1,4-Fuc 1,2-l-GalT ONeill et al. (1997);


d-GalT B RG-II l-AcefA1,3-Rha 1,2-GalT Vidal et al. (2000);ONeill et al. (1997);Carpita and Gibeaut(1993)

l-AraT D RG-II Dha2,3-GalA 1,5-l-Araf T ONeill et al. (2001,1997); Carpita andGibeaut (1993)

l-AraT B RG-II Gal1,2-l-AcefA 1,4-l-ArapT Vidal et al. (2000,1997); Carpita andGibeaut (1993)

l-AraT B2 RG-II l-Rha1,2-l-Ara 1,2-l-Araf T ONeill et al. (1997);Carpita and Gibeaut(1993)

l-FucT A RG-II l-Rha1,3-Apif 1,4-l-FucT ONeill et al. (1997);Carpita and Gibeaut(1993)

l-FucT B RG-II Gal1,2-l-AceAf 1,2-l-FucT Vidal et al. (2000);ONeill et al. (1997);Carpita and Gibeaut(1993)

d-Apif T A,B RG-II GalA1,4-GalA 1,2-Apif T ONeill et al. (1997);Carpita and Gibeaut(1993)


Table 3.5 (continued)Enzymed

Type of RG-IIglycosyl side Parent Acceptor Enzyme Reference fortransferase chainc polymer substrate activity structured-XylT A RG-II l-Fuc1,4-l-Rha 1,3-XylT ONeill et al. (1997);


d-GlcAT A RG-II l-Fuc1,4-l-Rha 1,4-GlcAT ONeill et al. (1997);Carpita and Gibeaut(1993)

d-KdoT C RG-II GalA1,4-GalA 2,3-KdoT ONeill et al. (2001,1997); Carpita andGibeaut (1993)

d-DhaT D RG-II GalA1,4-GalA 2,3-DhaT ONeill et al. (2001,1997); Carpita andGibeaut (1993)

l-AcefA B RG-II l-Rha1,3-Apif 1,3-AceAf T Vidal et al. (2000,1997); Carpita andGibeaut (1993)

Methyl-RG-II: RG-II d-Xyl1,3-l-Fuc ONeill et al. (1997);xylose 2-O- Carpita and Gibeautmethyltransferase (1993)RG-II:fucose 2-O- RG-II l-Fuc1,2-d-Gal ONeill et al. (1997);methyltransferase Carpita and Gibeaut

(1993)Acetyl- RG-II l-Fuc1,2-d-Gal ONeill et al. (1997);RG-II:fucose Carpita and Gibeautacetyltransferase (1993)RG-II:aceric RG-II l-AcefA1,3-l-Rha Vidal et al. (2000,acid 3-O- 1997); Carpita andacetyltransferase Gibeaut (1993)aAdapted from Ridley et al. (2001).bThis list of glycosyltransferase activites is based on the most extended structure of RG-II (Ridley et al.,2001). Note that terminal Rhap3Arap- in side chain B and the terminal Araf2Rhap- in sidechain B are not present in all RG-II preparations (ONeill et al., 2001).cThe side chain of RG-II that contains the specified glycosyl residue. See Ridley et al. (2001), ONeillet al. (2001) for side chain structure.dAll sugars are d sugars and have pyranose rings unless otherwise indicated. Glycosyltranferases add tothe glycosyl residue on the left of the indicated acceptor.

two of its four side branches attached to an HGA backbone by a Apif linkedto the O-2 of HGA (Ridley et al., 2001). It is also not known whether theapiosyltransferases that synthesize RG-II are the same as those involved inapiogalacturonan synthesis since no studies specifically directed at the 1,2-apiosyltransferase involved in RG-II synthesis have been reported. The in vivo


Table 3.6 Some of the glycosyltransferase activities likely to be required for apiogalacturonanbiosynthesis

Enzymea,b

Type of transferase Acceptor substrate Enzyme activity Reference for structured-GalAT GalA1,4-GalA 1,4-GalAT ONeill et al. (1990)d-Apif T GalA1,4-GalA 1,2-Apif T Hart and Kindel (1970);

Watson and Orenstein (1975)d-Apif T GalA1,4-GalA 1,3-Apif T Hart and Kindel (1970);

Watson and Orenstein (1975)d-Apif T Apif 1,2-GalA 1,3-Apif T Hart and Kindel (1970);

Watson and Orenstein (1975)d-Apif T Apif 1,3-GalA 1,3-Apif T Hart and Kindel (1970);

Watson and Orenstein (1975)aAll sugars are d sugars and have pyranose rings unless otherwise indicated.bGlycosyltranferases add to the glycosyl residue on the left of the indicated acceptor.

synthesis of apiogalacturonan, however, has been studied in vegetative frondsof Spirodela polyrrhiza (Longland et al., 1989) and a d-apiosyltransferase hasbeen identified and characterized in cell-free particulate preparations from duck-weed (Lemna minor) (Pan and Kindel, 1977). The apiosyltransferase transferred[14C]apiose from UDP-[14C]apiose onto endogenous acceptors in particulatemembrane preparations from Lemna. The enzyme has a pH optimum of 5.7 andan apparentKm for UDP-apiose of 4.9M (Pan and Kindel, 1977). Interestingly,the rate of apiosyltransferase activity was increased twofold by the inclusion ofUDP-GalA in the reaction (Pan and Kindel, 1977) and the product synthesizedin the presence of UDP-GalA bound more tightly to anion exchange resinthan the product synthesized without UDP-GalA (Mascaro and Kindel, 1977).These results suggest either that the apiosyltransferase transfers apiose ontoa growing HGA chain or that the amount of endogenous HGA substrate waslimiting. The product was solubilized in 1% ammonium oxalate, as expected forapiogalacturonans isolated from Lemna wall (Mascaro and Kindel, 1977) andthe product was fragmented by treatment with a fungal pectinase as expectedfor apiogalacturonan. The fact that acid hydrolysis of the 14C-labeled productyielded [14C]apiose and [14C]apiobiose (Mascaro and Kindel, 1977) confirmedthat an apiogalacturonan apiosyltransferase was identified. There are no reportsof the purification of the enzyme.

3.5.2.3 Xylogalacturonan xylosyltransferaseXylogalacturonan is a region of HGA in which some of the GalA residuesare substituted at O-3 with -d-xylose (see (Schols et al., 1995). Table 3.7shows some of the transferases likely to be required for xylogalacturonansynthesis. Xylosyltransferase activity was identified during studies of apio-galacturonan synthesis (Mascaro and Kindel, 1977; Pan and Kindel, 1977).


Table 3.7 Glycosyltransferase activities likely required to synthesize the substituted galacturonan knownas xylogalacturonan

Enzymea

Type of transferase Acceptor substrate Enzyme activity Reference for structured-GalAT GalA1,4-GalA 1,4-GalAT ONeill et al. (1990)d-XylT GalA1,4-GalA 1,3-XylT Aspinall (1980); Yu and

Mort (1996); Kikuchi et al.(1996); Schols et al. (1995)

aAll sugars are d sugars and have pyranose rings unless otherwise indicated. Glycosyltranferases add tothe glycosyl residue on the left of the indicated acceptor.

The product produced was not characterized in detail; however, at least some ofthe radioactive xylose appeared to be incorporated into apiogalacturonan and/orHGA. Thus, the enzyme may have been a xylogalacturonan xylosyltransferase.There have been no reports of the identification of the 1,3-xylosyltransferasethat transfers xylose from UDP-Xyl onto the l-Fuc1,4-l-rhamnosyl portion ofthe side branch of RG-II (Ridley et al., 2001).

3.5.2.4 Other glycosyltransferasesThere are no reports of targeted studies for the glycosyltransferases that insertfucose, Kdo, Dha or aceric acid into pectins. However, success in identifyingfucosyltransferase genes involved in the synthesis of hemicellulose could beuseful in the study of pectin synthesis. Fucose is a component of RG-II and RG-I.No fucosyltransferase involved in the synthesis of either of these pectic poly-mers has knowingly been studied. However, an Arabidopsis gene for an 1,2-fucosyltranferase that fucosylates a side branch in the hemicellulose xyloglucanhas been described (Perrin et al., 1999) that has 3573.8% amino acid sequenceidentify to 10 other putative fucosyltransferase genes in Arabidopsis (Perrinet al., 2001). The possibility that one or more of these genes may encode fuco-syltransferase(s) involved in RG-I or RG-II synthesis remains to be investigated.

3.5.3 Synthesis of rhamnogalacturonan I (RG-I)RG-I is a family of polysaccharides with an alternating [4)--d-GalpA-(12)--l-Rhap-(1] backbone in which roughly 2080% of the rhamnosesare substituted by arabinans, galactans or arabinogalactans (Carpita and Gibeaut,1993; Mohnen, 1999; Ridley et al., 2001). Table 3.8 shows a list of all the likelyenzyme activities required to synthesize all the known structures of RG-I, basedon the available structural information. There is considerable evidence fromimmunocytochemistry studies of plant tissues using antibodies against specificcarbohydrate epitopes found in RG-I (Willats et al., 2001a) that supports theview that the precise structure of the side chains of RG-I varies in a cell type- and


Table 3.8 Glycosyltransferases required for RG-I biosynthesisa

EnzymebType of Parent Reference forglycosyltransferase polymer Acceptor substrate Enzyme activity structured-GalAT RG-I l-Rha1,4-GalA 1,2-GalAT ONeill et al. (1990);

Eda et al. (1986); Lauet al. (1985)

d-GalAT RG-I/HGAc GalA1,2-l-Rha 1,4-GalAT l-RhaT RG-I GalA1,2-l-Rha 1,4-l-RhaT ONeill et al. (1990);

Eda et al. (1986); Lauet al. (1985)

l-RhaT HGA/RG-Ic GalA1,4-GalA 1,4-l-RhaT d-GalT RG-I l-Rha1,4-GalA 1,4-GalT ONeill et al. (1990);

Lau et al. (1987)d-GalT RG-I Gal1,4-Rha 1,4-GalT ONeill et al. (1990);

Lau et al. (1987)d-GalT RG-I Gal1,4-Gal 1,4-GalT Morita (1965a,b);

Aspinall et al. (1967);Stephen (1983);Aspinall (1980);ONeill et al. (1990);Lau et al. (1987)

d-GalT RG-I Gal1,4-Gal 1,6-GalT ONeill et al. (1990);Lau et al. (1987)

d-GalT RG-I/AGPd Gal1,3-Gal 1,3-GalT Carpita and Gibeaut(1993)

d-GalT RG-I/AGPd Gal1,3-Gal 1,6-GalT Carpita and Gibeaut(1993)

d-GalT RG-I/AGPd Gal1,6-Gal1,3-Gal 1,6-GalT Carpita and Gibeaut(1993)

d-GalT RG-I l-Araf-1,4-Gal 1,5-GalT Huisman et al. (2001a)l-AraT RG-I Gal1,4-Rha 1,3-l-Araf T ONeill et al. (1990);

Lau et al. (1987)l-AraT RG-I l-Araf1,3-Gal 1,2-l-Araf T ONeill et al. (1990);

Lau et al. (1987)l-AraT RG-I l-Araf1,2-Ara 1,5-l-Araf T ONeill et al. (1990);

Lau et al. (1987)l-AraT RG-I l-Rha1,4-GalA 1,4-Araf T Lau et al. (1987)l-AraT RG-I l-Araf1,5-Ara 1,5-l-Araf T Carpita and Gibeaut

(1993)l-AraT RG-I l-Araf1,5-Ara 1,2-l-Araf T Carpita and Gibeaut



(1993)l-AraT RG-I Gal1,4-Gal 1,3-l-Araf T Morita (1965a,b);

Aspinall et al. (1967);Stephen (1983);Aspinall (1980);


Table 3.8 (continued)Enzymeb

Type of Parent Reference forglycosyltransferase polymer Acceptor substrate Enzyme activity structure

ONeill et al. (1990);Carpita and Gibeaut(1993)

l-AraT RG-I l-Araf-1,3-Gal 1,5-l-Araf T Morita (1965a,b);Aspinall et al. (1967);Stephen (1983);Aspinall (1980)

l-AraT RG-I/AGPd Gal1,6-Gal 1,3-l-Araf T Carpita and Gibeaut(1993)

l-AraT RG-I/AGPd Gal1,6-Gal 1,6-l-Araf T Carpita and Gibeaut(1993)

l-AraT RG-I Gal1,4-Gal 1,4-l-ArapT Huisman et al. (2001a)l-FucT RG-I Gal1,4-Gal 1,2-l-Fucf T ONeill et al. (1990);

Lau et al. (1987)d-GlcAT RG-I Gal. . . 1,6-GlcAT An et al. (1994)d-GlcAT RG-I Gal. . . 1,4-GlcAT An et al. (1994)Methyl-RG-I:GlcA 4-O- RG-I GlcA1,6-Gal An et al. (1994)methyltransferaseAcetyl- RG-I GalA1,2-l-Rha1,4(n) Lerouge et al. (1993);RG-I:GalA Ishii (1997);3-O/2-O-acetyl- ONeill et al. (1990);transferase Bacic et al. (1988);

Komalavilas andMort (1989)

aAdapted from Ridley et al. (2001).bAll sugars are d sugars and have pyranose rings unless otherwise indicated. Glycosyltranferases add tothe glycosyl residue on the left of the indicated acceptor.cEnzyme that may be required to make an HGA/RG-I junction.dEnzyme activity would also be required to synthesize arabinogalactan proteins (AGPs) (see Gasparet al., 2001).

development-specific manner. Thus, it not expected that all of the biosyntheticenzyme activities shown in Table 3.8 will necessarily be expressed in any givencell type synthesizing a specific RG-I structure.

3.5.3.1 RG-I galactosyltransferase (GalT)The synthesis of the family of polysaccharides known as RG-I requires at leasteight different galactosyltransferase (GalT) activities to synthesize the diverseside chain linkages found in the RG-I structures identified to date (see Table 3.8).Probable 1,4-GalT and 1,3-GalT activities were identified in early studies ofmicrosomal preparations from mung bean (McNab et al., 1968; Panayotatos and


Villemez, 1973).A more recent study confirmed that a 1,4-galactosyltranferaseactivity with a pH of optimum of 6.5 was present in mung bean microsomesbased on sensitivity of the product to digestion with endo-1,4-galactanase(Brickell and Reid, 1996).

Galactosyltransferases have also been identified in particulate homogenates(Goubet and Morvan, 1993; Goubet and Morvan, 1994) and solubilized enzyme(Goubet, 1994) from flax (Linum usitatissimum L.). Flax GalTs solubilized frommicrosomes with detergent transferred [3H]Gal from UDP-[3H]Gal onto exoge-nous RG-I-enriched and pectic 1,4-galactan acceptors (Peugnet et al., 2001)to yield a radiolabeled product of high molecular mass. Interestingly, the pHoptimum for transfer onto lupin pectic 1,4-galactan (i.e. pH 6.5) was differentfrom the pH optimum for transfer of Gal onto an endopolygalacturonase-treatedRG-I-enriched fractions from flax (i.e. two optima: pH 6.5 and 8.0) (Peugnetet al., 2001). Cleavage of the large radiolabeled product with either rhamno-galacturonan I hydrolase (RGaseA) or with rhamnogalacturonan I lyase (RGaseB) resulted in a fragmentation of the radiolabeled product into lower molecularmass material that, at least in part, eluted as expected for small oligomers (i.e.DP 36) (Peugnet et al., 2001). This result confirmed that the GalTs added Galonto RG-I (Peugnet et al., 2001). The fragmentation of at least a portion of theradiolabeled product with 1,4-endogalactanase further demonstrated that atleast some of the GalT activity represented 1,4-galactosyltransferase (Peugnetet al., 2001). The GalT at pH 8.0 has an apparent Km of 460M for UDP-Gal.The characteristics of the GalT at pH 8.0 are consistent with an enzyme thatadds galactose onto short galactan side branches of RG-I.

An RG-I:1,4-galactosyltransferase was identified and partially character-ized from potato suspension cultured cells (Geshi et al., 2000). The potatoGalT transfers [14C]Gal from UDP-[14C]Gal onto endogenous acceptor(s) inthe membranes to produce a>500 kDa product. The product can be fragmentedby endo-1,4-galactanase into [14C]Gal and [14C]galactobiose. Treatment of theintact radiolabeled product with rhamnogalacturonase A, an endohydrolase thatcleaves the glycosidic linkage between the GalA and the Rha in the RG-I back-bone (Azadi et al., 1995), yielded radiolabeled fragments between 50 kDa and180 kDa in size (Geshi et al., 2000). The subsequent treatment of the [14C]Gal-labeled fragments with endo-1,4-galactanase yielded [14C]Gal and slightlylarger fragments that themselves could be cleaved to [14C]Gal by a purified -galactosidase. These results suggest that potato microsomal membranes containenzymes that both initiate and elongate 1,4-galactan side chains of RG-I. Thepotato 1,4-GalT has a pH optimum of 6.06.5.

A fenugreek gene for an -d-1,6-galactosyltransferase involved in the syn-thesis of the hemicellulose galactomannan (Edwards et al., 1999) has beenidentified. Eight Arabidopsis genes with sequence similarity exist in Arabidopsis(Perrin et al., 2001) and the possibility that one or more of these putativegalactosyltransferases is involved in pectin synthesis remains to be explored.


d-Galactose is a component of both RG-II and RG-I, while l-galactose is foundonly in RG-II. No information is available on the enzyme that adds l-Gal ontoRG-II.

3.5.3.2 RG-I arabinosyltransferaseRG-I and RG-II containl-arabinose in multiple linkages (seeTables 3.5 and 3.8).Most of the arabinose is in the furanose ring form, although it has recently beenreported that a terminal arabinose exists in the pyranose form in some RG-I sidechains (Huisman et al., 2001a). Arabinosyltransferase activity has been iden-tified in microsomes from mung bean (Phaseolus aureus) shoots (Odzuck andKauss, 1972) and from bean (Phaseolus vulgaris) hypocotyl and callus (Bolwelland Northcote, 1981). Definitive evidence that theseAraT activities are involvedin pectin synthesis was not demonstrated (see Mohnen, 1999, for review).The arabinosyltransferase activity in bean hypocotyl and callus was primarilyassociated with enriched Golgi, and to a lesser extent with enriched endoplasmicreticulum (Bolwell and Northcote, 1983). The difficulty of studying arabinosyl-transferases that specifically synthesize pectin, as opposed to other hemicellu-losic polysaccharides or arabinogalactan proteins has been discussed (Nunanand Scheller, 2001). An approach that entails the use of detergent-solubilizedmicrosomes and specific pectic oligo/polysaccharide acceptors has been reportedto yield arabinosylation of pectic acceptors (Nunan and Scheller, 2001).

3.5.3.3 RG-I methyltransferase (RG-I-MT)A detergent-solubilized pectin methyltransferase (PMT) from flax has beenreported to use an RG-I-enriched fraction as an exogenous acceptor (Bourlardet al., 1997a). Specifically, pectin methyltransferase activity was stimulatedin the presence of an enriched RG-I fraction 1.5-fold to 1.7-fold above levelsrecovered using endogenous acceptor. The resulting radiolabeled product had asize similar to RG-I. It was not shown, however, where in RG-I the methylationoccurred. Thus, it is not clear whether the methylation occurred on GalA inthe RG-I backbone, or whether it occurred on possible HGA tails that maybe covalently linked to RG-I. Also, it was not shown whether some of themethylation may have occurred on a non-galacturonic substituent in RG-I suchas methylation at the 4-position of glucuronic acid in the side branches of RG-I(An et al., 1994). The location on the polymer of the methylation in the RG-I-enriched fraction and, thus, the identity of the potentially novel enzyme activity,remains to be determined.

3.5.3.4 RG-I acetyltransferase (RG-I-AT)The GalA residues in the alternating [4)--d-GalpA-(12)--l-Rhap-(1]backbone of RG-I may be acetylated on C-2 and/or C-3 (Komalavilas and


Mort, 1989). Microsomes from suspension-cultured potato cells (Pauly andScheller, 2000) contain an RG-I acetyltransferase that transfers [14C]acetatefrom [14C]acetyl-CoA onto RG-I yielding a >500 kDa radiolabeled product(Pauly and Scheller, 2000). The release of [14C]acetate following incubation ofthe radiolabeled product with a purified rhamnogalacturonan O-acetyl esterase,and the fragmentation of the product by rhamnogalacturonan lyase (RGase B)confirmed that the enzyme was an RG-I acetyltransferase (Pauly and Scheller,2000). The RG-I acetyltransferase has an apparentKm for acetyl-CoA of 35M,an apparent Vmax of 54 pmol min1 mg1 protein and a pH optimum of 7.0, with80% of activity recovered at pH 6.58.0.

3.6 Future directions and resources for studying pectin biosynthesis

The partial characterization and, in some cases, purification of selected pectinbiosynthetic genes has provided a core of biochemical information on theenzymes. However, to significantly increase our understanding of when, whereand how the enzymes interact to produce pectin, it is essential that the genesfor the biosynthetic enzymes be identified (see Henrissat et al., 2001; Keegstraand Raikhel, 2001; Perrin et al., 2001; Reiter and Vanzin, 2001). The identifi-cation of the genes would provide primary structure information and allow thegeneration of antibodies against the biosynthetic enzymes that could be usedfor analysis of enzyme localization and provide a means to identify members ofthe expected biosynthetic protein complexes. The genes could also be used toexpress the enzymes for use in detailed kinetic, structure and function studies.The manipulation of the genes in transgenic plants should allow hypothesesregarding pectin structurefunction in the plant to be tested and would facilitatethe elucidation of how the individual biosynthetic enzymes interact to synthesizepectin.

The identification of the pectin biosynthetic genes will likely occur using mul-tiple strategies including enzyme purification, mutant identification and charac-terization, and DNA sequence/motif similarity computer searches. In all cases,however, the definitive identification of any pectin biosynthetic gene will requireproof of enzyme activity. Thus, continued progress in identifying and makingavailable the required nucleotide-sugar and oligo/polysaccharide substrates forthe biosynthetic enzymes is essential. The National Science Foundation-fundedPlant Cell Wall Biosynthesis Research Coordination Network, also referred toas WallBioNet (http://xyloglucan.prl.msu.edu), serves as an information centerand resource for researchers studying plant cell wall biosynthesis. It providesa central location for updated information on progress in, and available toolsfor, studying wall biosynthesis research. It also partially supports the synthesisof rare and needed substrates and acceptors for wall biosynthesis that are madeavailable to the scientific community.


Acknowledgments

I thank my colleagues at the CCRC for their helpful discussions. This effortwas supported in part by NSF grant No. MCB-0090281, NRI competitiveUSDA award 2001-35318-11111 and DOE-funded center grant DE-FG05-93-ER20097.

References

An, J., ONeill, M.A., Albersheim, P. and Darvill, A.G. (1994) Isolation and structural characterizationof -d-glucosyluronic acid and 4-O-methyl -d-glucosyluronic acid-containing oligosaccharidesfrom the cell-wall pectic polysaccharide, rhamnogalacturonan I. Carbohydr. Res., 252, 235243.

Ankel, H. and Tischer, R. (1969) UDP-d-Glucuronate 4-epimerase in blue-green algae. Biochim. Biophys.Acta, 178, 415419.

Aspinall, G.O. (1980) Chemistry of cell wall polysaccharides, in The Biochemistry of Plants, vol. 3 (ed.J. Preiss), Academic Press, New York, pp. 473500.

Aspinall, G.O., Begbie, R., Hamilton,A. and Whyte, J.N.C. (1967) Polysaccharides of soy-beans. Part III.Extraction and fractionation of polysaccharides from cotyledon meal. J. C

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