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Nucleotide sugars and glycosyltransferases for synthesis of cell wall matrix polysaccharides
Plant Physiol. Biochem., 2000, 38 (1/2), 69−80 / © 2000 Éditions scientifiques et médicales Elsevier SAS. All rights reserved S0981942800001674/REV
Nucleotide sugars and glycosyltransferases for synthesis of cell wall matrix polysaccharides David M. Gibeaut Department of Plant Science, University of Adelaide, Waite Campus, Glen Osmond SA 5064, Australia
(fax +61 8 8303 7109; e-mail [email protected]) (Received September 30, 1999; accepted October 19, 1999) Abstract — The current interest in cell wall biosynthesis is expanding because of the increasing evidence that the properties of the cell wall mediate cellular interactions during growth, development and differentiation. Much effort has been put forward to the identification of glycosyltransferases because of their obvious importance in polysaccharide synthesis. Enzymes involved in nucleotide sugar production and transport are also important because of the potential to manipulate the composition of cell walls through substrate level control. Molecular genetics have begun to uncover genes for important enzymes in polysaccharide biosynthesis including glycosyltransferases and enzymes of nucleotide sugar metabolism; but at this time, much is inferred from comparisons to bacteria, yeast and animal cells. This review examines the production and transport of nucleotide sugars, the protein structure of glycosyltransferases, and implications for the cellular mechanisms of cell wall biosynthesis. © 2000 Éditions scientifiques et médicales Elsevier SAS Glycosyltransferase / Golgi apparatus / hydrophobic cluster analysis / nucleotide sugar / nucleotide sugar transporter / polysaccharide / virus-induced gene silencing Ara, arabinose / BLAST, basic local alignment search tool / dTDP-, deoxythymidinediphospho- / Fuc, fucose / gal, galactose / Glc, glucose / HCA, hydrophobic cluster analysis / Man, mannose / Rhm, rhamnose / UDP-, uridinediphospho/ VIGS, virus-induced gene silencing / Xyl, xylose
1. INTRODUCTION Knowledge of cell wall metabolism is crucial to our understanding of plant development because few growth events occur without a change in the chemistry and structure of the cell wall [10]. In particular, the biosynthesis of matrix components during growth is critical for cellular differentiation and these biosynthetic processes are a principal function of the endomembrane system of plants [20]. Even cellulose deposition is influenced by the endomembrane system by the placement of the terminal complexes of cellulose synthesis [16], and in delivery of the matrix polysaccharides that must be co-deposited as cellulose is synthesized and the wall assembled. Other than the glycosyltransferase activities of cellulose and callose synthesis, the greatest number of the various glycosyltransferase activities are found in the Golgi apparatus. Knowledge of the topology of the enzymatic machinery, including the glycosyltransferases and the enzymes of nucleotide sugar conversion and transport, will increase our understanding of the requirements for biosynthesis of the cell wall. In this review, we describe the production and transport of nucleotide
sugars, the likely topology and protein structure of glycosyltransferases, and some ideas for ways to identify the genes of glycosyltransferases.
2. SUBSTRATES FOR MATRIX POLYSACCHARIDES: NUCLEOTIDE SUGARS Recent interest in the nucleotide sugar metabolism of plants has been spurred by the possibility that cell wall properties could be manipulated by altering the substrate availability for polysaccharide glycosyltransferases. This is shown, for example by the mur1 mutation of GDP-Man-4,6-dehydratase [4], the cloning and heterologous expression of UDP-Glc 4-epimerase [13], and by the possibility of manipulating the flux of substrate using UDP-Glc dehydrogenase [51]. Additionally, the discovery of specific transporters of nucleotide sugars across the Golgi membrane in animal and yeast cells has suggested important roles for these transport proteins in control of the glycan processing of glycoproteins and proteoglycans [25, 27]. However, in plants, there is little
Plant Physiol. Biochem., 0981-9428/00/1-2/© 2000 E´ditions scientifiques et médicales Elsevier SAS. All rights reserved
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comparable and convincing evidence for many aspects of nucleotide sugar metabolism including the genetic identity and location of the enzymes involved in nucleotide sugar activation, conversion, and transport steps. Yet to be demonstrated are the nucleotide sugar substrates for a number of polysaccharides.
2.1. Nucleotide sugar pools From radiolabeling studies using monosaccharides, sucrose or CO2, there is tempting speculation that separate pools of nucleotide sugars exist for the synthesis of the cell wall. When 14CO2 and photosynthesis are used as a means to pulse label tissues, cellulose labels more quickly than matrix components, whereas matrix components label more rapidly when 14 C-Glc is used [21]. The difference in labeling kinetics could arise by the relative activities of sucrose synthase and UDP-Glc pyrophosphorylase. The regulation of flux through these enzymes is the subject of much debate in terms of source-sink balance and starch synthesis or degradation, and will not be discussed here. Separate pools of nucleotide sugars can also arise from the recycling of cell wall polysaccharides. However, most importantly, separate pools may arise by the activity of a plasmamembrane-associated sucrose synthase, and by the regulation of the enzymes of nucleotide sugar conversion and transport. In cells that are growing rapidly and thus synthesizing cell wall material, including those undergoing secondary wall thickening such as cotton fibers, UDPGlc is derived by the activity of sucrose synthase from sucrose that is delivered symplastically [46]. Amor et al. [2] found that a form of sucrose synthase was associated with the plasmamembrane, and because UDP-Glc free in the cytosol was not an efficient substrate for the synthesis of cellulose, they postulated that sucrose synthase channeled UDP-Glc directly to cellulose synthase. On the other hand, a separate cytoplasmic pool of nucleotide sugars can arise from either sucrose synthase or nucleotide sugar pyrophosphorylases. Thus, two sources or pools of UDP-Glc, one for synthesis of cellulose and another for synthesis of matrix polysaccharide could arise within a cell. The ‘free cytoplasmic’ pool can be further segregated by the activities of enzymes involved in conversion of UDP-Glc to the other nucleotide sugars required for synthesis of the matrix polysaccharides. The location of the enzymes involved in the activation and conversion of nucleotide sugars, whether cytoplasmic (free or membrane-associated) or within the lumen of the Golgi apparatus, is not yet established for plants, and the summary of many reports has Plant Physiol. Biochem.
indicated both possibilities [18]. For example, it is possible that UDP-Glc pyrophosphorylase, an enzyme that produces UDP-Glc, may have a Golgi-located isozyme. Most activity is cytosolic [29], although putative N-glycosylation sites (as well as biochemical data) indicate the possibility of Golgi-associated isoforms [15]. However, these studies have only shown an association with the membranes and have not ascertained a lumenal or cytoplasmic orientation of the enzymes. To date, no studies in animal, yeast or plant cells have demonstrated a lumenal orientation. Instead, specific transporters of nucleotide sugars are well established to reside in the membranes of the Golgi apparatus and function in complex glycan and polysaccharide production in animal and yeast cells [1, 25]. Pyrophosphorylases have an important role in the production of nucleotide sugars for both the initial activation of sugars and for the scavenging or recycling of the sugars released from matrix polysaccharides during growth. Ara and Gal based polysaccharides, especially non-classical type II AGP, exhibit rapid turnover from the cell walls of growing tissues [19, 21, 24]. The (1→3),(1→4)-β-D-glucan of grasses [19], and galactans of fibrous tissues in flax stems [21] also show rapid turnover. Furthermore, terminal arabinosyl units such as those decorating glucuronoarabinoxylans are particularly susceptible to turnover after which the parent backbone becomes more tightly bound in the wall [19]. The sugars released from polysaccharides are absorbed by the cell and phosphorylated in the cytoplasm. Although a general C-1 hexokinase may operate on all available sugars including those scavenged or recycled during cell wall metabolism, C-1 kinases for specific sugars, especially Ara and Gal do exist [18], as do specific pyrophosphorylases for each sugar and nucleoside.
2.2. Nucleotide sugar conversions Regardless of how sugars become activated – either through the activities of pyrophosphorylases or sucrose synthase – the majority of the glycosyl units of cell wall polysaccharide pass through either UDP-Glc or GDP-Man. A possible exception is the source of Rhm as discussed below. Although many pathways for activation and conversion of nucleotide sugars have been demonstrated in plants [18], only a few dominate in the synthesis of cell wall polysaccharides as shown in figure 1. For example, metabolism of inositol can lead to the production of UDP-GlcA, UDP-Xyl and UDP-Ara in pollen tubes grown in media lacking metabolizable carbon, but growth of pollen tubes is only normal with added sucrose [48]. In addition, in
Matrix polysaccharides
Figure 1. Pathways for synthesis of the nucleotide sugar substrates of cell wall polysaccharides. Although other pathways to many of the substrates have been demonstrated in plants, the pathways and enzymes shown are proposed to dominate the flux of substrate for cell wall synthesis. Activation of Rhm as dTDP-Rhm has been demonstrated in plants, and is well established in bacteria. The actual substrate used in the glycosyltransferase activity for synthesis of rhamnogalacturonan has not been demonstrated.
radioactive labeling with [3H]-inositol, about 1 % of the label absorbed by squash hypocotyls was incorporated into the cell wall during a 9-h pulse; whereas, 40 % of the radioactivity from [14C]-Glc was incorporated [54]. Metabolism of inositol may be more important in the turnover of phytic acid in germinating legumes, in the intracellular signaling pathways of inositol-phosphates, and possibly in the turnover of the glycosylphosphotidylinositol membrane anchors of arabinogalactan-proteins [49, 55]. Once UDP-Glc or GDP-Man is formed, several enzymes act to provide most of the other nucleotide sugars required for synthesis of the cell wall. Control of
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the expression and activity of these enzymes may influence the amounts and characteristics of several matrix polysaccharides synthesized by cells. In the first steps of metabolism, UDP-Glc may be converted to UDP-Gal by the activity of UDP-Glc-4-epimerase, or to UDP-GlcA by UDP-Glc dehydrogenase. As part of the Leloir pathway of galactose metabolism, UDP-Glc-4epimerase has been extensively studied, and recently the gene has been identified in Arabidopsis [13]. Deficiencies in this enzyme activity lead to impaired growth and improper glycan processing in yeast. In plants, Gal-containing polysaccharides are abundant in stems and roots, and Gal is a critical component of lipids, especially in chloroplasts. Not surprisingly, the mRNA for UDP-Glc-4-epimerase is expressed in all plant organs, and highly expressed in stems and roots of Arabidopsis [13]. In amino acid sequence, this enzyme is remarkably similar in bacteria, yeast and animal, and conserved regions include NAD+ and UDP-hexose binding domains. No signal peptide is present. Manipulation of the expression of UDP-Glc-4-epimerase in plants could lead to metabolic deficiencies in other UDP-sugars by sequestration of the pool of uridinenucleosides. A similar situation is observed in grasses when Gal is infiltrated into tissues [7], because the recycling pathway leads to overproduction of UDP-Gal and starvation of UTP for production of the other UDP-sugars. Alternatively, the properties of galactanrich fibrous tissues, such as in flax stems, could be altered by enrichment of the cell wall galactan. Another enzyme, UDP-Glc dehydrogenase, catalyzes an important step in the synthesis of the uronosyl and pentosyl units of the cell wall. UDP-Glc dehydrogenase catalyzes the conversion of UDP-Glc to UDPGlcA requiring two NAD+. The location of this protein is cytosolic, and it probably forms a hexameric structure [50]. No signal sequence is present in the cDNA [51]; however, there is an indication that it is glycosylated [50]. In soybean and Arabidopsis, the gene is apparently present as a single copy [51]. Because the majority of the matrix polysaccharides are comprised of uronosyl and pentosyl units, regulation of UDP-Glc dehydrogenase activity could be a critical control point. It is in low concentration relative to other enzymes in the pathway, operates far from equilibrium, and is inhibited by UDP-Xyl, one of the products in the pathway [43, 51]. In addition, the expression of mRNA for UDP-Glc dehydrogenase was shown to be greatest in the growing portions of leaves and roots of soybean [51] and soybean nodules [50]. From the UDP-GlcA produced by the activity of UDP-Glc dehydrogenase, both UDP-GalA and UDPvol. 38 (1/2) 2000
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Xyl are formed. A UDP-GalA-C-4 epimerase, distinct from UDP-Glc-C-4 epimerase, forms UDP-GalA in a reversible reaction. The gene has recently been cloned from bacteria, and some of the enzymatic properties studied [34]. The enzyme binds two NAD+ and is strongly inhibited by UDP-Glc, UDP-Gal and UDPXyl. Several conserved amino acids important in binding of NAD+ and UDP-Glc were present, as well as the motif YXXXK, thought to be important for catalytic activity. The activities of UDP-GlcA decarboxylase and UDP-Xyl-4-epimerase sequentially produce UDP-Xyl and UDP-Ara [18]. These enzyme activities have been well studied, but information of their molecular genetics is lacking. Feingold and Avigad [18] also questioned whether UDP-Ara might be formed by the decarboxylation of UDP-GalA. To date no evidence for UDP-GalA decarboxylase in plants has been presented. The mannosyl units of the cell wall are derived from the conversion of the phosphorylated intermediates of sugar metabolism, although a specific C-1 kinase for Man is also present in plants [18]. A GDP-Man pyrophosphorylase catalyzes the activation of the mannosyl units used in cell wall and glycoprotein synthesis. A gene for GDP-Man pyrophosphorylase has recently been cloned in yeast [22]. A hydropathy plot of the predicted amino acid sequence indicates the protein is hydrophilic and lacks a signal peptide, suggesting the protein is a soluble cytoplasmic protein. The significance for the use of GDP instead of UDP sugars is unknown, but it is interesting that GDP-Glc, along with GDP-Man, are the preferred substrates in the synthesis of glucomannan [37]. Further metabolism of GDP-Man provides GDP-LFuc and probably GDP-L-Gal. Fucose-deficient mutants of Arabidopsis (mur1) produced by chemical mutagenesis [40, 41] are due to the mutation and loss of function of GDP-D-Man-4,6-dehydratase [4], the first of two enzymes involved in the formation of GDP-LFuc from GDP-D-Man. The following epimerization and reduction activities apparently are performed by one enzyme in animal cells [52]. Discovery of the fucose deficient mutants of Arabidopsis was the first demonstration that cell wall polysaccharide composition and properties could be addressed by a molecular genetic approach. The mur1 plants have a brittle phenotype in the stems and it was postulated that the lack of an α-L-Fuc unit on the trisaccharide branch of xyloglucan [56] disrupted the ability of xyloglucan to associate with cellulose [30]. Interestingly, α-L-Gal replaced about one-third of the α-L-Fuc found in wild type xyloglucan [56], but only a small proportion of the Plant Physiol. Biochem.
fucosyl units of N-glycans of proteins were replaced with α-L-Gal [38]. Furthermore, it is worth noting that solanaceous xyloglucans do not possess the Fuc-GalXyl trisaccharide and that the lack of α-L-Fuc on essential proteins [38] could be an important part of the brittle phenotype. Other mutants with altered cell wall monosaccharide composition were also identified [40, 41], but it is notable that screening of this mutant population may have missed alterations in the uronic acids, hence much of the pectic polysaccharides, because the wall materials were not carboxylreduced [28] during the derivatization procedures. The nucleotide sugar used in the synthesis of the α-L-Rhm, containing rhamnogalacturonans I and II, has not been demonstrated, but has been variously reported as GDP-L-Rhm, UDP-D-Rhm, or UDP-LRhm due to the presence of these compounds in plants [18]. However, in bacteria dTDP-L-Rhm is the substrate used in the synthesis of the α-L-Rhm containing polysaccharides of the bacterial cell wall. The synthesis pathway of dTDP-L-Rhm begins with dTDPD-Glc pyrophosphorylase forming dTDP-D-Glc. This activity has been demonstrated in plants and suggested to be distinct from the activity of UDP-Glc pyrophosphorylase [18]. A specific nucleotide sugar 4,6dehydratase activity followed by epimerization and reduction, similar to that producing GDP-L-Fuc as described above, produces dTDP-L-Rhm. The set of enzymes involved in the metabolism of GDP-L-Fuc, and the dTDP-L-Rhm dehydratase in bacteria have been cloned and extensively studied [52]. Some bacteria can also produce polysaccharides comprised of D-Rhm, but the synthesis of this polysaccharide requires GDP-D-Rhm provided through the conversion of GDP-D-Man [44]. By analogy, because plant rhamnogalacturonan contains α-L-Rhm, plants may use dTDP-L-Rhm for the synthesis of pectic polysaccharides. The evolutionary origin of α-L-Rhm containing polysaccharides in plants could be interesting in regard to the symbiotic evolution of plants and bacteria, and in the nature and the function of Rhm containing pectic polysaccharides and Rhm-rich proteoglycans. A survey for similar genes and enzyme activities, as well as an evaluation of the glycosyltransferase activities in vitro from plant membranes would be prudent.
2.3. Nucleotide sugar transporters At least three possible scenarios exist for the delivery and utilization of nucleotide sugars by the plant Golgi apparatus as shown in figure 2. In the first, each nucleotide sugar is transported across the Golgi membrane and the active sites of Type II membrane
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glycosyltransferases (discussed below) are oriented toward the lumen. In the second, nucleotide sugars are delivered from the cytoplasmic side to the active site of an integral membrane glycosyltransferases and the growing glycan chain is extruded into the Golgi lumen. In the third, sugar, sugar phosphates or a nucleotide sugar are transported into the Golgi lumen where nucleotide sugar activation or conversions occur. The cytoplasmic location of pyrophosphorylases [18, 29], the existence of specific transporters for each nucleotide sugar species in animal and yeast cells [27], and the indication that UDP-Xyl [23] and UDP-Glc [33] are transported across the Golgi apparatus in plants would argue for a cytoplasmic location of nucleotide sugar conversion in plants. Except for the unlikely possibility that a second set of nucleotide sugar conversion enzymes are present in the lumen of the Golgi apparatus – a condition that would be unique in plants – specific transporters for each nucleotide sugar substrate must exist in the plant Golgi apparatus and endoplasmic reticulum, similar to the situation in animal and yeast cells. Furthermore, the demonstrated presence of nucleoside diphosphatase (NDPase) within the Golgi apparatus of plants and its likely function in polysaccharide synthesis [32] argues for a Golgi lumenal orientation of the active sites of the glycosyltransferases involved in polysaccharide synthesis. NDPase activity produces the nucleoside monophosphate from the nucleoside-diphosphate released by the activity of glycosyltransferases. The resulting nucleoside-monophosphates are removed from the lumen in an antiport process with the corresponding nucleotide sugar transporter [1, 25]. Nucleotide sugar transporters have been implicated as control points for the synthesis of extracellular proteoglycans in animal and yeast cells [27]. In rat hepatocytes, protein glycosylation can be affected by the availability of nucleotide sugars in the Golgi lumen [42]. Increased UDP-sugar concentrations allowed increased incorporation of N-acetylhexosamines at the expense of sialation because of impaired transport of CMP-N-acetylneuraminate. As another example, the loss of UDP-Gal transport in mutant MDCK cells results in a loss of the polysaccharide, keratin sulfate, without affecting galactosylation of the linkage regions of heparin and chondroitin sulfates [53]. These authors suggested that the Km values for the galactosyltransferases that synthesize heparin sulfate and chondroitin sulfate must be significantly lower than the Km value for keratin sulfate suggesting
Figure 2. Proposed scenarios for delivery of nucleotide sugar substrates to sites of polysaccharide synthesis in the Golgi apparatus. A, Specific transporters for each species of nucleotide sugar act in an antiporter process with the corresponding nucleoside monophosphate. Nucleotide sugars are concentrated in the lumen where Type II membrane protein glycosyltransferases produce polysaccharide. Nucleoside diphosphatase produces the monophosphate, releasing Pi, which is removed by a putative Pi transporter. B, Integral membrane protein glycosyltransferases, by analogy to cellulose synthase, would have catalytic sites oriented toward the cytosol and would not require transport of nucleotide sugar. C, Nucleotide sugars if produced in the lumen of the Golgi apparatus, would require transport of sugars, sugar phosphates, UDP-Glc and other required compounds for nucleotide sugar conversions. No evidence for this scenario is present in animal or yeast cells.
a regulation of glycosylation by nucleotide sugar levels through limitations in nucleotide sugar transporters. The location of the transporters, whether in the endoplasmic reticulum or the early or late elements of the Golgi apparatus is often correlated with the location of glycosyltransferase activity [11]. Several cDNA’s for nucleotide sugar transporters have been cloned [27]. The transporters are structurally related hydrophobic proteins with multiple transmembrane regions. Some possess putative leucine zippers that may facilitate protein-protein interactions. Furthermore, a regulation of the type of polysaccharide and possibly the degree of substitution could also be controlled by the relative Km values of glycosyltransferases and the transport of nucleotide sugars. Plants may also control polysaccharide synthesis by regulating nucleotide sugar concentration. Reported Kms for the glycosyltransferases forming the backbones of xyloglucan [33], (1→3),(1→4)-β-D-glucan [7] and galactomannan [39] are one to two orders of vol. 38 (1/2) 2000
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Table I. Properties of some glycosyltransferases involved in cell wall synthesis. Polysaccharide
Resulting linkage
Substrate
Retaining or inverting
Cellulose Callose (1→3),(1→4)-β-D-Glucan Xyloglucan
β-D-Glc-1→4-β-D-Glc β-D-Glc-1→3-β-D-Glc β-D-Glc-3-or 4-β-D-Glc β-D-Glc-1→4-β-D-Glc α-D-Xyl-1→6-β-D-Glc β-D-Gal-1→2-α-D-Xyl α-L-Fuc-1→2-β-D-Gal β-D-Man-1→4-β-D-Glc β-D-Glc-1→4-β-D-Man α-D-Gal-1→2-β-D-Man β-D-Xyl-1→4-β-D-Xyl α-D-GlcA-1→2-β-D-Xyl α-L-Ara-1→2-β-D-Xyl α-D-GalA-1→4-α-D-GalA α-L-Rhm-1→2-α-L-GalA α-D-GalA-1→4-α-D-Rhm
UDP-D-Glc UDP-D-Glc UDP-D-Glc UDP-D-Glc UDP-D-Xyl UDP-D-Gal GDP-L-Fuc GDP-D-Man GDP-D-Glc UDP-D-Gal UDP-D-Xyl UDP-D-GlcA UDP-L-Ara UDP-D-GalA ? UDP-D-GalA
inverting inverting inverting inverting retaining inverting inverting inverting inverting retaining inverting retaining retaining retaining retaining retaining
Galactoglucomannan
Glucuronoarabinoxylan
Homogalacturonan Rhamnogalacturonan
magnitude greater than those reported for the glycosyltransferases of side group additions such as the fucosyltransferase of xyloglucan [36] and the galactosyltransferases of xyloglucan [17] and galactomannan [39]. Although accuracy of the Km determinations are questionable because of disruption of the membrane environment [20], lower Km values of protein processing and side group adding glycosyltransferases would ensure their activity to proceed to completion at low substrate concentration while the higher Km of the glycosyltransferases of backbone polysaccharides could allow overall control of polysaccharide synthesis by substrate level. Furthermore, the ratio of Glc to Man in glucomannan [37], and the ratio of the tri- and tetrasaccharide repeating units of mixed linkage (1→3),(1→4)-β-D-glucan [7] can be manipulated in vitro by varying the concentration of substrates. However, these ratios do not vary in planta even when the substrate concentrations are manipulated. These observations indicate that plants may closely regulate the concentration of substrate within the Golgi lumen.
3. GLYCOSYLTRANSFERASES Until the recent biochemical and molecular genetic characterization of a fucosyltransferase [36] that is involved in the addition of a terminal α-L-fucosyl unit to a xyloglucan acceptor, a cDNA had not been matched to a suitably purified glycosyltransferase from plants. The central difficulty associated with the purification of glycosyltransferases from plants, in particular those forming the backbone, is the loss of activity when solubilized. However, even if the activPlant Physiol. Biochem.
Type of membrane protein integral integral ? ? ? ? Type II ? ? ? ? ? ? ? ? ?
ity can be solubilized, as is the general case for the Type II glycosyltransferases involved in side group formation, we must also have the proper acceptor glycans in a soluble form [17]. Additionally, for many polysaccharides, the substrates are not readily available. Fortunately, genetic approaches can be very useful to identify putative glycosyltransferases. The genes for many glycosyltransferases have been identified by mutation and complementation analysis in bacteria, yeast and animal cells. This information has formed the basis for classification of glycosyltransferases into a number of families based upon primary sequence. The classification has revealed that sequence related proteins share the same mechanisms of activity [9], which may be expected because the structure and function of a protein is intrinsically related to its primary sequence. The mechanisms referred to above are stereochemical descriptions of the donor and acceptor molecules. Glycosyltransferase activities either retain or invert the configuration of the anomeric carbon as it is transferred from the nucleotide α-sugar (β- in the case of GDP-Fuc) substrate [9]. These transfers can be either to the nonreducing end or to a side group position of a glycan. A family classification of glycosyltransferases can be found at http://afmb.cnrs-mrs.fr/∼pedro/CAZY/ db.html, and a summary of linkages, mechanisms and protein classifications of glycosyltransferases involved in cell wall biosynthesis is shown in table I. The above mentioned classifications distinguish between the three-dimensional folding of enzymes and their mechanisms of activity, but do not directly consider the transmembrane spanning regions of the
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Integral membrane glycosyltransferases such as cellulose synthase and callose synthase are known in bacteria, yeast and plants. These integral membrane glycosyltransferases possess several membrane spanning regions and both the NH2 and COOH termini are predicted to be oriented toward the cytosol. Evidence for the identity of cellulose synthase genes is convincing [12]. In addition, sequence similarity of two plant cDNA’s (accessions AAD25952, AAD30609) with those of yeast 1→3-β-D-glucan synthase genes [14, 26, 31] is quite strong. Importantly, this type of integral membrane glycosyltransferase has only been found in the plasmamembrane.
Figure 3. Two types of membrane bound glycosyltransferases may be responsible for polysaccharide synthesis in the plant Golgi apparatus. Integral membrane protein glycosyltransferases such as cellulose synthase and callose synthase are found in the plasma membrane. The presence of integral membrane protein glycosyltransferases in the Golgi membrane for synthesis of polysaccharide backbones has not been demonstrated. Integral membrane proteins are characterized by multiple strands of amino acids spanning the membrane. If the catalytic domain were oriented toward the cytosol, as in cellulose synthase, then a pore would be required for channeling the polysaccharide into the lumen. Type II membrane protein glycosyltransferases have a well-established domain structure. A single strand of amino acids anchors the protein in the membrane. The cytosolic oriented NH2 tail and the lumenal stem region each contribute to the retention of the protein within the Golgi apparatus. The catalytic domain is globular and oriented toward the lumen.
glycosyltransferases. A distinction based upon the nature of the transmembrane regions is apparent and will be discussed below.
3.1. Integral and Type II integral membrane glycosyltransferases Glycosyltransferases involved in glycan and polysaccharide synthesis are integral membrane proteins that can be classified into two broad groups based upon the number of their membrane spanning regions and topology of the catalytic domains as shown in figure 3. The glycosyltransferases of eukaryotes comprise these two groups but prokaryotes have additional variety in the number and position of membrane spanning regions.
Type II membrane glycosyltransferases are well studied in animal and yeast cells for their significance in glycan processing and polysaccharide synthesis [11]. The recently identified fucosyltransferase from Arabidopsis is a predicted Type II membrane protein [36]. Type II membrane proteins are located in the Golgi apparatus. They are characterized by a short NH2 tail oriented toward the cytoplasm, a single membrane spanning region, and a variable length stem connecting with a globular catalytic region oriented toward the lumen. Type II glycosyltransferases are about 400 amino acids long, which is much smaller than the integral glycosyltransferases, such as cellulose synthase and callose synthase that are four to five times longer. Modification of the membrane anchor and stem regions of Type II glycosyltransferases can alter the resulting glycan products [11, 35] by placing the glycosyltransferase in a different sub-compartment of the Golgi apparatus. A variation of the commonly known Type II glycosyltransferases described above involves the cytoplasmic NH2 region of the protein as seen in the KRE6 gene product of yeast [45]. The KRE6 gene product is a Type II membrane protein located in the Golgi apparatus and is the catalytic peptide for synthesis of the 1→6-β-D-glucan of yeast cell walls. It is different from most Type II glycosyltransferases however in that the cytoplasmic NH2 tail is much larger. The cytoplasmic region of Kre6p is 252 amino acids, and contains several consensus sites for phosphorylation (RXXS/T where S or T is phosphorylated) that resemble those of PKC1, MAP kinase, cAMPdependent protein kinase and casein kinase. Several lines of evidence suggest the PKC1 pathway participates in the regulation of yeast cell wall synthesis. It is tempting to speculate that plants may also regulate the synthesis of matrix polysaccharides by phosphorylation-mediated pathways. vol. 38 (1/2) 2000
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3.2. Conserved amino acids of glycosyltransferases Sequence analysis of DNA or protein has helped to identify by similarity many hundreds of glycosyltransferases. For example, BLAST analysis has been used to identify members within the large families of cellulose synthases (http://cellwall.stanford.edu), fucosyltransferases [6, 36] and galactosyltransferases [5]. However, even within a family, there can be as little as 25 % similarity in primary sequence. The diversity of sequence is presumably a reflection of the different substrates, the linkages formed, and the acceptor recognition regions. Furthermore, convergent evolution has very likely occurred and this may obscure homologies when searching for sequence similarities. At present, there are insufficient examples from plants for comparisons to be made that will help identify many glycosyltransferases involved in cell wall synthesis because many activities are unique to plants and presumably, the genes for these glycosyltransferases will be highly divergent in sequence. The grouping of glycosyltransferases into 36 families – more are likely to be identified – is testament to their diversity [9]. However, within each family, similarity in threedimensional folding structure is expected. Thus, knowledge of the three-dimensional protein structure of a few glycosyltransferases can aid in identifying divergent glycosyltransferases. Underlying the threedimensional folding of a protein is the protein’s secondary structure. Hydrophobic cluster analysis (HCA) is an indicator of the secondary structure of a protein, and HCA has been used to help identify a limited number of conserved regions and amino acids that are related to active sites and acceptor binding regions. Clearly, the search for putative genes for glycosyltransferases would be greatly advanced if we could identify motifs that are conserved among all or within families of glycosyltransferases.
Glycosyltransferases most often have aspartate residues in the active sites. Three such conserved aspartate residues are found in the integral glycosyltransferases such as cellulose, and chitin synthases whereas only two are conserved in the Type II glycosyltransferases. These structures are clearly seen in the enzymes studied by Saxena et al. [47], the galactosyltransferases [5] and less clearly in the fucosyltransferases [6]. Comparing the integral and Type II glycosyltransferases, it appears that the integral glycosyltransferases have a catalytic region divided into two domains, A and B, whereas Type II enzymes have only one domain which appears equivalent to domain A. One of the aspartate residues in domain A is preceded by a vertical hydrophobic cluster. This motif is present in both integral and Type II glycosyltransferases. This motif can be found in proteins that are not glycosyltransferases, so it is not a unique signature of glycosyltransferases, but most if not all glycosyltransferases have it. Sometimes, this motif is difficult to see especially if a non-hydrophobic amino acid interrupts the vertical hydrophobic cluster. However, in addition to the motif, the second of the conserved aspartate regions should be looked for when searching the database for potential genes of glycosyltransferases. Sometimes the order of the conserved aspartate residues appears reversed, and the distance between can vary from 10 to well over 100 amino acids. Additionally, most of the conserved aspartate residues are found in a DD or DXD motif. Furthermore, a few conserved motifs are present within a group of glycosyltransferases that have the same substrate. In this respect, it is interesting that the xyloglucan fucosyltransferase [36] has a KPW motif that is seen in the galactosyltransferases [5]. Representations of topol-
Figure 4. Topology predictions (left panels) and HCA plots (right panels) of a bacterial cellulose synthase (AcscA, accession CAA38487), a plant cellulose synthase (GhCesA-1, accession AAB37766), a plant fucosyltransferase (XG-FT1, accession AAD41092), a yeast (1→6)-β-D-glucan glucosyltransferase (KRE6, accession P32468), and a plant putative glycosyltransferase (T7N9.14, accession AAB61490). The topology predictions were based upon a hidden Markov model and were drawn using the TMHMM program (http://www.cbs.dtu.dk/services/TMHMM-1.0/). The height of the vertical bars represents the probability of a transmembrane helix. Topology predictions are a useful indicator of integral membrane proteins (top two panels) and type II integral membrane proteins (bottom three panels). The HCA’s were performed using the drawhca program (http://www.lmcp.jussieu.fr/∼soyer/www-hca/hca-file.html). HCA involves drawing the amino acid sequence on a cylinder representing an α-helix. The cylinder is split lengthwise, flattened, and the sequences duplicated to restore the connectivity of the amino acids. The one-letter code is used for amino acids except for glycine (diamonds), proline (stars), serine (squares with dots), and threonine (squares). Acidic amino acids are shown in red, basic amino acids in blue, and hydrophobic amino acids in green. Contours are also drawn around the clusters of hydrophobic amino acids to aid in the visual analysis of their shape, length and distribution. The conserved QXXRW motif of cellulose synthases is shown over a gray background and conserved aspartate residues are circled. For XG-FT1, a KPW motif is also indicated over a gray background although this motif appears conserved in family B of galactosyltransferases [5]. Along with the identification of conserved amino acids using sequence alignment methods, HCA is a useful tool to uncover conserved motifs. The regions indicated by the yellow background were identified as described in the text; however, their usefulness in predicting the function of a gene product as a glycosyltransferase is highly speculative. More examples of HCA plots of glycosyltransferases can be found in references [5, 6, 47].
Plant Physiol. Biochem.
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ogy predictions and HCA’s of some glycosyltransferases are shown in figure 4. It should be emphasized that only a few glycosyltransferases have been studied in enough detail to identify the critical amino acids in active sites and nucleotide binding regions. The number and arrange-
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ment of putative catalytic sites and binding regions is still speculative for the majority. These issues, in particular the number and identity of active sites will be resolved with amino acid modification studies, cryo-electron microscopy and if possible with X-ray crystallographic studies of crystallized proteins.
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3.3. Identifying genes for glycosyltransferases The rapid pace of EST and genome sequencing means that many genes for glycosyltransferases are already available in the database. The difficulty is to identify putative genes and then demonstrate their function. This is especially problematic for the polysaccharide producing glycosyltransferases of plants because we do not have a similar means to produce and recover knock-out mutants in a way that has been so useful in other organisms (but see Fagard et al. in this issue). We may infer from the screening of mutant Arabidopsis [40, 41] that many loss of function mutations will not be recovered. This may also happen in anti-sense transformation strategies unless inducible promoters are used. A more rapid approach, and one that avoids some of the pitfalls of lethal mutations, is the use of virus-induced gene silencing [3]. To perform virus-induced gene silencing (VIGS), genes or gene fragments of interest are inserted into a potato X potexvirus (PVX), and RNA transcripts are used for infection of seedlings of Nicotiana benthamiana. Post-transcriptional gene silencing results in reduced mRNA levels of endogenously expressed plant genes that have a high level of sequence identity with the sequence carried by the virus. Furthermore, a length of only 300–500 nucleotides is sufficient to effect silencing. Thus, full-length cDNAs or genes are not required which is particularly advantageous for the ‘fast forward’ screening of candidate genes [3]. The major advantage of the VIGS system relates to the relative speed with which the role of a gene product can be identified, compared with antisense or co-suppression approaches in transgenic plants. In addition, VIGS can be used to knock out potentially lethal genes, because young seedlings are established before the gene is introduced via the viral RNA vector. We have used a homologous fragment of the cellulose synthase gene, GhCesA, and VIGS to demonstrate the knock-out of cellulose synthesis. The silencing of cellulose synthase resulted in dwarfism, and a disruption of leaf cell shape. Interestingly, there was compensation for the loss of cellulose by an increase of homogalacturonan with a low degree of carboxyl esterification [8]. With a suitable method for functional analysis now in hand, efficient selection of candidate genes can be increased by knowledge of the gene and protein structures of glycosyltransferases as described above. A combination of BLAST, topology prediction, and HCA can be used to identify protein sequences by the structural characteristics that are consistent with the large number of glycosyltransferases already identified. Plant Physiol. Biochem.
In addition to considerations of protein structure, identification of sequence motifs within a family of glycosyltransferases will narrow the field of candidates, and may be critical for selecting gene sequences.
4. CONCLUSIONS Within the lumen of the Golgi apparatus, many polysaccharides, proteoglycans and glycoproteins are synthesized and organized for secretion to the cell wall. The mechanisms for the location and organization of nucleotide sugar transporters and the glycosyltransferases that use the corresponding substrate are hot topics in animal and yeast research because of the potential to understand certain disease processes, and for the production of valuable proteins and biopolymers. Knowledge of these processes in plants may also generate the means to understand and modify plant development through alteration of the chemistry and structure of the cell wall. Even with the promise given by genetic analysis of the glycosyltransferases fast approaching as shown by the genetic descriptions of cellulose synthase [12] and fucosyltransferase [36], we realize there will also be much to be discovered in the cellular mechanics of cell wall biosynthesis. One important question yet to be answered is whether the backbone producing glycosyltransferases in the Golgi apparatus are integral or Type II membrane proteins. The answers will suggest much about the intracellular mechanisms of matrix polysaccharide synthesis.
Acknowledgments This work was supported by a grant from the Grains Research and Development Corporation of Australia (to Geoffrey B. Fincher).
REFERENCES [1] Abeijon C., Mandon E.C., Hirschberg C.B., Transporters of nucleotide sugars, nucleotide sulfate and ATP in the Golgi apparatus, Trends Biochem. Sci. 22 (1997) 203–207. [2] Amor Y., Haigler C.H., Johnson S., Wainscott M., Delmer D.P., A membrane-associated form of sucrose synthase and its potential role in synthesis of cellulose and callose in plants, Proc. Natl. Acad. Sci. USA 92 (1995) 9353–9357.
Matrix polysaccharides
[3] Baulcombe D.C., Fast forward genetics based on virus-induced gene silencing, Curr. Opin. Plant Biol. 2 (1999) 109–113. [4] Bonin C.P., Potter I., Vanzin G.F., Reiter W.D., The MUR1 gene of Arabidopsis thaliana encodes an isoform of GDP-D-mannose-4,6-dehydratase, catalyzing the first step in the de novo synthesis of GDP-Lfucose, Proc. Natl. Acad. Sci. USA 94 (1997) 2085–2090. [5] Breton C., Bettler E., Joziasse D.H., Geremia R.A., Imberty A., Sequence-function relationships of prokaryotic and eukaryotic galactosyltransferases, J. Biochem. 123 (1998) 1000–1009. [6] Breton C., Oriol R., Imberty A., Conserved structural features in eukaryotic and prokaryotic fucosyltransferases, Glycobiology 8 (1998) 87–94. [7] Buckeridge M.S., Vergara C.E., Carpita N.C., The mechanism of synthesis of a mixed-linkage (1→3),(1→4) β-D-glucan in maize (Zea mays L.). Evidence for multiple sites of glucosyl transfer in the synthase complex, Plant Physiol. 120 (1999) 1–12. [8] Burton R.A., Gibeaut D.M., Bacic A., Findlay K., Roberts K., Hamilton A., Baulcombe D.C., Fincher G.B., Virus-induced silencing of a plant cellulose synthase gene, Plant Cell (1999) (in press). [9] Campbell J.A., Davies G.J., Bulone V., Henrissat B., A classification of nucleotide-diphospho-sugar glycosyltransferases based on amino acid sequence similarities, Biochem. J. 326 (1997) 929–942. [10] Carpita N.C., Gibeaut D.M., Structural models of primary cell walls in flowering plants: consistancy of molecular structure with the physical properties of the walls during growth, Plant J. 3 (1993) 1–30. [11] Colley K.J., Golgi localization of glycosyltransferases – more questions than answers, Glycobiology 7 (1997) 1–13. [12] Delmer D.P., Cellulose biosynthesis: exciting times for a difficult field of study, Annu. Rev. Plant Physiol. Plant Mol. Biol. 50 (1999) 245–276. [13] Dörmann P., Benning C., Functional expression of uridine 5’-diphospho-glucose 4-epimerase (EC 5.1.3.2) from Arabidopsis thaliana in Saccharomyces cerevisiae and Escherichia coli, Arch. Biochem. Biophys. 327 (1996) 27–34. [14] Douglas C.M., Foor F., Marrinan J.A., Morin N., Nielsen J.B., Dahl A.M., Mazur P., Baginsky W., Li W., El-Sherbeini M., Clemas J.A., Mandala S.M., Frommer B.R., Kurtz M.B., The Saccharomyces cerevisiae FKS1 (ETG1) gene encodes an integral membrane protein which is a subunit of 1,3-β-D-glucan synthase, Proc. Natl. Acad. Sci. USA 91 (1994) 12907–12911. [15] Eimert K., Villand P., Kilian A., Klecczkowski L.A., Cloning and characterization of several cDNAs for UDP-glucose pyrophosphorylase from barley (Hordeum vulgare) tissues, Gene 170 (1996) 227–232. [16] Emons A.M.C., Mulder B.M., The making of the architecture of the plant cell wall: how cells exploit geometry, Proc. Natl. Acad. Sci. USA 95 (1998) 7215–7219.
79
[17] Faïk A., Chileshe C., Sterling J., Maclachlan G., Xyloglucan galactosyl- and fucosyltransferase activities from pea epicotyl microsomes, Plant Physiol. 114 (1997) 245–254. [18] Feingold D.S., Avigad G., Sugar nucleotide transformations in plants, in: Preiss J. (Ed.), The Biochemistry of Plants. Carbohydrates: Structure and Function, vol. 3, Academic Press, 1980, pp. 101–170. [19] Gibeaut D.M., Carpita N.C., Tracing the biosynthesis of the cell wall in intact cells and plants. Selective turnover and alteration of soluble and cell wall polysaccharides in grasses, Plant Physiol. 97 (1991) 551–561. [20] Gibeaut D.M., Carpita N.C., Biosynthesis of plant cell wall polysaccharides, FASEB J. 8 (1994) 904–915. [21] Gorshkova T.A., Chemikosova S.B., Lozovaya V.V., Carpita N.C., Turnover of galactans and other cell wall polysaccharides during development of flax plants, Plant Physiol. 114 (1997) 723–729. [22] Hashimoto H., Sakakibara A., Yamasaki M., Yoda K., Saccharomyces cerevisiae VIG9 encodes GDPmannose pyrophosphorylase, which is essential for protein glycosylation, J. Biol. Chem. 272 (1997) 16308–16314. [23] Hayashi T., Koyama T., Matsuda K., Formation of UDP-xylose and xyloglucan in soybean Golgi membranes, Plant Physiol. 87 (1988) 341–345. [24] Hirano Y., Tsumuraya Y., Hashimoto Y., Characterization of spinach leaf α-L-arabinofuranosidases and β-galactosidases and their synergistic action on an endogenous arabinogalactan-protein, Physiol. Plant. 92 (1994) 286–296. [25] Hirschberg C.B., Transporters of nuleotide sugars, nucleotide sulfate and ATP in the Golgi apparatus membrane – where next?, Glycobiology 7 (1997) 169–171. [26] Inoue S.B., Takewaki N., Takasuka T., Mio T., Adachi M., Fujii Y., Miyamoto C., Arisawa M., Furuichi Y., Watanabe T., Characterization and gene cloning of 1,3-β-D-glucan synthase from Saccharomyces cerevisiae, Eur. J. Biochem. 231 (1995) 845–854. [27] Kawakita M., Ishida N., Miura N., Sun-Wada G.H., Yoshioka S., Nucleotide sugar transporters: elucidation of their molecular identity and its implication for future studies, J. Biochem. 123 (1998) 777–785. [28] Kim J.B., Carpita N.C., Changes in esterification of the uronic acid groups of cell wall polysaccharides during elongation of maize coleoptiles, Plant Physiol. 98 (1992) 646–653. [29] Klecczkowski L.A., Glucose activation and metabolism through UDP-glucose pyrophosphorylase in plants, Phytochemistry 37 (1994) 1507–1515. [30] Levy S., Maclachlan G., Staehelin L.A., Xyloglucan sidechains modulate binding to cellulose during in vitro binding assays as predicted by conformational dynamics simulations, Plant J. 11 (1997) 373–386. vol. 38 (1/2) 2000
80
D.M. Gibeaut
[31] Mazur P., Morin N., Baginsky W., El-Serbeini M., Clemas J.A., Nielsen J.B., Foor F., Differential expression and function of two homologous subunits of yeast 1,3-β-D-glucan synthase, Mol. Cell. Biol. 15 (1995) 5671–5681. [32] Mitsui T., Honma M., Kondo T., Hashimoto N., Kimura S., Igaue I., Structure and function of the Golgi complex in rice cells II. Purification and characterization of Golgi membrane-bound nucleoside diphosphatase, Plant Physiol. 106 (1994) 119–125. [33] Muñoz P., Norambuena L., Orellana A., Evidence for a UDP-glucose transporter in Golgi apparatus-derived vesicles from pea and its possible role in polysaccharide biosynthesis, Plant Physiol. 112 (1996) 1585–1594. [34] Muñoz R., López R., de Frutos M., Garcia E., First molecular characterization of a uridine diphosphate galacturonate 4-epimerase: an enzyme required for capsular biosynthesis in Streptococcus pneumoniae type 1, Mol. Microbiol. 31 (1999) 703–713. [35] Paulson J.C., Colley K.J., Glycosyltransferases. Structure, localization, and control of cell type-specific glycosylation, J. Biol. Chem. 264 (1989) 17615–17618. [36] Perrin R.M., DeRocher A.E., Bar-Peled M., Zeng W., Norambuena L., Orellana A., Raikhel N.V., Keegstra K., Xyloglucan fucosyltransferase, an enzyme involved in plant cell wall biosynthesis, Science 284 (1999) 1976–1979. [37] Piro G., Dalessandro G., Northcote D.H., Glucomannan synthesis in pea epicotyls: the mannose and glucose transferases, Planta 190 (1993) 206–220. [38] Rayon C., Cabanes-Macheteau M., Loutelier-Bourhis C., Salliot-Maire I., Lemoine J., Reiter W-D., Lerouge P., Faye L., Characterization of N-glycans from Arabidopsis. Application to a fucose-deficient mutant, Plant Physiol. 119 (1999) 725–733. [39] Reid J.S.G., Edwards M., Gidley M.J., Clark A.H., Enzyme specificity in galactomannan biosynthesis, Planta 195 (1995) 489–495. [40] Reiter W.D., Chapple C., Somerville C.R., Altered growth and cell walls in a fucose-deficient mutant of Arabidopsis, Science 261 (1993) 1032–1035. [41] Reiter W.D., Chapple C., Somerville C.R., Mutants of Arabidopsis thaliana with altered cell wall polysaccharide composition, Plant J. 12 (1997) 335–345. [42] Pels Rijcken W.R., Overdijk B., Van Den Eijnden D.H., Ferwerda W., The effect of increasing nucleotide-sugar concentrations on the incorporation of sugars into glycoconjugates in rat hepatocytes, Biochem. J. 305 (1995) 865–870. [43] Robertson D., McCormack B.A., Bolwell G.P., Cell wall polysaccharide biosynthesis and related metabolism in elicitor-stressed cells of French bean (Phaseolus vulgaris L.), Biochem. J. 306 (1995) 745–750. [44] Rocchetta H.L., Pacan J.C., Lam J.S., Synthesis of the A-band polysaccharide sugar D-rhamnose requires RMD and WBPW – identification of multiple ALGA homologues, WPBW and ORF488, in Pseudomonas aeruginosa, Mol. Microbiol. 29 (1998) 1419–1434. Plant Physiol. Biochem.
[45] Roemer T., Paravicini G., Payton M.A., Bussey H., Characterization of the yeast (1-6)-β-glucan biosynthetic components, Kre6p and Skn1p, and genetic interactions between the PKC1 pathway and extracellular matrix assembly, J. Cell Biol. 127 (1994) 567–579. [46] Ruan Y.L., Chourey P.S., Delmer D.P., Perez-Grau L., The differential expression of sucrose synthase in relation to diverse patterns of carbon partitioning in developing cotten seed, Plant Physiol. 115 (1997) 375–385. [47] Saxena I.M., Brown Jr R.M., Fevre M., Geremia R.A., Henrissat B., Multidomain architecture of β-glycosyl transferases: implications for mechanism of action, J. Bacteriol. 177 (1995) 1419–1424. [48] Schlüpmann H., Bacic A., Read S.M., Uridine diphosphate glucose metabolism and callose synthesis in cultured pollen tubes of Nicotiana alata Link et Otto, Plant Physiol. 105 (1994) 659–670. [49] Schultz C., Gilson P., Oxley D., Youl J., Bacic A., GPI-anchors on arabinogalactan-proteins: implications for signalling in plants, Trends Plant Sci. 3 (1998) 426–431. [50] Stewart D.C., Copeland L., Uridine 5’-diphosphateglucose dehydrogenase from soybean nodules, Plant Physiol. 116 (1998) 349–355. [51] Tenhaken R., Thulke O., Cloning of an enzyme that synthesizes a key nucleotide-sugar precursor of hemicellulose biosynthesis from soybean: UDP-glucose dehydrogenase, Plant Physiol. 112 (1996) 1127–1134. [52] Tonetti M., Sturla L., Bisso A., Zanardi D., Benatti U., De Flora A., The metabolism of 6-deoxyhexoses in bacterial and animal cells, Biochimie 80 (1998) 923–931. [53] Toma L., Pinhal M.A.S., Dietrich C.P., Nader H.B., Hirschberg C.B., Transport of UDP-galactose into the Golgi lumen regulates the biosynthesis of proteoglycans, J. Biol. Chem. 271 (1996) 3897–3901. [54] Wakabayashi K., Sakurai N., Kuraishi S., Effects of ABA on synthesis of cell-wall polysaccharides in segments of etiolated squash hypocotyl. I. Changes in incorporation of glucose and myo-inositol into cellwall components, Plant Cell Physiol. 30 (1989) 99–105. [55] Youl J.J., Bacic A., Oxley D., Arabinogalactanproteins from Nicotiana alata and Pyrus communis contain glycosylphosphatidylinositol membrane anchors, Proc. Natl. Acad. Sci. USA 95 (1998) 7921–7926. [56] Zablackis E., York W.S., Pauly M., Hantus S., Reiter W.D., Chapple C.C.S., Albersheim P., Darvill A., Substitution of L-fucose by L-galactose in cell walls of Arabidopsis mur1, Science 272 (1996) 1808–1810.
Nucleotide sugars and glycosyltransferases for synthesis of cell wall matrix polysaccharides David M. Gibeaut Department of Plant Science, University of Adelaide, Waite Campus, Glen Osmond SA 5064, Australia
(fax +61 8 8303 7109; e-mail [email protected]) (Received September 30, 1999; accepted October 19, 1999) Abstract — The current interest in cell wall biosynthesis is expanding because of the increasing evidence that the properties of the cell wall mediate cellular interactions during growth, development and differentiation. Much effort has been put forward to the identification of glycosyltransferases because of their obvious importance in polysaccharide synthesis. Enzymes involved in nucleotide sugar production and transport are also important because of the potential to manipulate the composition of cell walls through substrate level control. Molecular genetics have begun to uncover genes for important enzymes in polysaccharide biosynthesis including glycosyltransferases and enzymes of nucleotide sugar metabolism; but at this time, much is inferred from comparisons to bacteria, yeast and animal cells. This review examines the production and transport of nucleotide sugars, the protein structure of glycosyltransferases, and implications for the cellular mechanisms of cell wall biosynthesis. © 2000 Éditions scientifiques et médicales Elsevier SAS Glycosyltransferase / Golgi apparatus / hydrophobic cluster analysis / nucleotide sugar / nucleotide sugar transporter / polysaccharide / virus-induced gene silencing Ara, arabinose / BLAST, basic local alignment search tool / dTDP-, deoxythymidinediphospho- / Fuc, fucose / gal, galactose / Glc, glucose / HCA, hydrophobic cluster analysis / Man, mannose / Rhm, rhamnose / UDP-, uridinediphospho/ VIGS, virus-induced gene silencing / Xyl, xylose
1. INTRODUCTION Knowledge of cell wall metabolism is crucial to our understanding of plant development because few growth events occur without a change in the chemistry and structure of the cell wall [10]. In particular, the biosynthesis of matrix components during growth is critical for cellular differentiation and these biosynthetic processes are a principal function of the endomembrane system of plants [20]. Even cellulose deposition is influenced by the endomembrane system by the placement of the terminal complexes of cellulose synthesis [16], and in delivery of the matrix polysaccharides that must be co-deposited as cellulose is synthesized and the wall assembled. Other than the glycosyltransferase activities of cellulose and callose synthesis, the greatest number of the various glycosyltransferase activities are found in the Golgi apparatus. Knowledge of the topology of the enzymatic machinery, including the glycosyltransferases and the enzymes of nucleotide sugar conversion and transport, will increase our understanding of the requirements for biosynthesis of the cell wall. In this review, we describe the production and transport of nucleotide
sugars, the likely topology and protein structure of glycosyltransferases, and some ideas for ways to identify the genes of glycosyltransferases.
2. SUBSTRATES FOR MATRIX POLYSACCHARIDES: NUCLEOTIDE SUGARS Recent interest in the nucleotide sugar metabolism of plants has been spurred by the possibility that cell wall properties could be manipulated by altering the substrate availability for polysaccharide glycosyltransferases. This is shown, for example by the mur1 mutation of GDP-Man-4,6-dehydratase [4], the cloning and heterologous expression of UDP-Glc 4-epimerase [13], and by the possibility of manipulating the flux of substrate using UDP-Glc dehydrogenase [51]. Additionally, the discovery of specific transporters of nucleotide sugars across the Golgi membrane in animal and yeast cells has suggested important roles for these transport proteins in control of the glycan processing of glycoproteins and proteoglycans [25, 27]. However, in plants, there is little
Plant Physiol. Biochem., 0981-9428/00/1-2/© 2000 E´ditions scientifiques et médicales Elsevier SAS. All rights reserved
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comparable and convincing evidence for many aspects of nucleotide sugar metabolism including the genetic identity and location of the enzymes involved in nucleotide sugar activation, conversion, and transport steps. Yet to be demonstrated are the nucleotide sugar substrates for a number of polysaccharides.
2.1. Nucleotide sugar pools From radiolabeling studies using monosaccharides, sucrose or CO2, there is tempting speculation that separate pools of nucleotide sugars exist for the synthesis of the cell wall. When 14CO2 and photosynthesis are used as a means to pulse label tissues, cellulose labels more quickly than matrix components, whereas matrix components label more rapidly when 14 C-Glc is used [21]. The difference in labeling kinetics could arise by the relative activities of sucrose synthase and UDP-Glc pyrophosphorylase. The regulation of flux through these enzymes is the subject of much debate in terms of source-sink balance and starch synthesis or degradation, and will not be discussed here. Separate pools of nucleotide sugars can also arise from the recycling of cell wall polysaccharides. However, most importantly, separate pools may arise by the activity of a plasmamembrane-associated sucrose synthase, and by the regulation of the enzymes of nucleotide sugar conversion and transport. In cells that are growing rapidly and thus synthesizing cell wall material, including those undergoing secondary wall thickening such as cotton fibers, UDPGlc is derived by the activity of sucrose synthase from sucrose that is delivered symplastically [46]. Amor et al. [2] found that a form of sucrose synthase was associated with the plasmamembrane, and because UDP-Glc free in the cytosol was not an efficient substrate for the synthesis of cellulose, they postulated that sucrose synthase channeled UDP-Glc directly to cellulose synthase. On the other hand, a separate cytoplasmic pool of nucleotide sugars can arise from either sucrose synthase or nucleotide sugar pyrophosphorylases. Thus, two sources or pools of UDP-Glc, one for synthesis of cellulose and another for synthesis of matrix polysaccharide could arise within a cell. The ‘free cytoplasmic’ pool can be further segregated by the activities of enzymes involved in conversion of UDP-Glc to the other nucleotide sugars required for synthesis of the matrix polysaccharides. The location of the enzymes involved in the activation and conversion of nucleotide sugars, whether cytoplasmic (free or membrane-associated) or within the lumen of the Golgi apparatus, is not yet established for plants, and the summary of many reports has Plant Physiol. Biochem.
indicated both possibilities [18]. For example, it is possible that UDP-Glc pyrophosphorylase, an enzyme that produces UDP-Glc, may have a Golgi-located isozyme. Most activity is cytosolic [29], although putative N-glycosylation sites (as well as biochemical data) indicate the possibility of Golgi-associated isoforms [15]. However, these studies have only shown an association with the membranes and have not ascertained a lumenal or cytoplasmic orientation of the enzymes. To date, no studies in animal, yeast or plant cells have demonstrated a lumenal orientation. Instead, specific transporters of nucleotide sugars are well established to reside in the membranes of the Golgi apparatus and function in complex glycan and polysaccharide production in animal and yeast cells [1, 25]. Pyrophosphorylases have an important role in the production of nucleotide sugars for both the initial activation of sugars and for the scavenging or recycling of the sugars released from matrix polysaccharides during growth. Ara and Gal based polysaccharides, especially non-classical type II AGP, exhibit rapid turnover from the cell walls of growing tissues [19, 21, 24]. The (1→3),(1→4)-β-D-glucan of grasses [19], and galactans of fibrous tissues in flax stems [21] also show rapid turnover. Furthermore, terminal arabinosyl units such as those decorating glucuronoarabinoxylans are particularly susceptible to turnover after which the parent backbone becomes more tightly bound in the wall [19]. The sugars released from polysaccharides are absorbed by the cell and phosphorylated in the cytoplasm. Although a general C-1 hexokinase may operate on all available sugars including those scavenged or recycled during cell wall metabolism, C-1 kinases for specific sugars, especially Ara and Gal do exist [18], as do specific pyrophosphorylases for each sugar and nucleoside.
2.2. Nucleotide sugar conversions Regardless of how sugars become activated – either through the activities of pyrophosphorylases or sucrose synthase – the majority of the glycosyl units of cell wall polysaccharide pass through either UDP-Glc or GDP-Man. A possible exception is the source of Rhm as discussed below. Although many pathways for activation and conversion of nucleotide sugars have been demonstrated in plants [18], only a few dominate in the synthesis of cell wall polysaccharides as shown in figure 1. For example, metabolism of inositol can lead to the production of UDP-GlcA, UDP-Xyl and UDP-Ara in pollen tubes grown in media lacking metabolizable carbon, but growth of pollen tubes is only normal with added sucrose [48]. In addition, in
Matrix polysaccharides
Figure 1. Pathways for synthesis of the nucleotide sugar substrates of cell wall polysaccharides. Although other pathways to many of the substrates have been demonstrated in plants, the pathways and enzymes shown are proposed to dominate the flux of substrate for cell wall synthesis. Activation of Rhm as dTDP-Rhm has been demonstrated in plants, and is well established in bacteria. The actual substrate used in the glycosyltransferase activity for synthesis of rhamnogalacturonan has not been demonstrated.
radioactive labeling with [3H]-inositol, about 1 % of the label absorbed by squash hypocotyls was incorporated into the cell wall during a 9-h pulse; whereas, 40 % of the radioactivity from [14C]-Glc was incorporated [54]. Metabolism of inositol may be more important in the turnover of phytic acid in germinating legumes, in the intracellular signaling pathways of inositol-phosphates, and possibly in the turnover of the glycosylphosphotidylinositol membrane anchors of arabinogalactan-proteins [49, 55]. Once UDP-Glc or GDP-Man is formed, several enzymes act to provide most of the other nucleotide sugars required for synthesis of the cell wall. Control of
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the expression and activity of these enzymes may influence the amounts and characteristics of several matrix polysaccharides synthesized by cells. In the first steps of metabolism, UDP-Glc may be converted to UDP-Gal by the activity of UDP-Glc-4-epimerase, or to UDP-GlcA by UDP-Glc dehydrogenase. As part of the Leloir pathway of galactose metabolism, UDP-Glc-4epimerase has been extensively studied, and recently the gene has been identified in Arabidopsis [13]. Deficiencies in this enzyme activity lead to impaired growth and improper glycan processing in yeast. In plants, Gal-containing polysaccharides are abundant in stems and roots, and Gal is a critical component of lipids, especially in chloroplasts. Not surprisingly, the mRNA for UDP-Glc-4-epimerase is expressed in all plant organs, and highly expressed in stems and roots of Arabidopsis [13]. In amino acid sequence, this enzyme is remarkably similar in bacteria, yeast and animal, and conserved regions include NAD+ and UDP-hexose binding domains. No signal peptide is present. Manipulation of the expression of UDP-Glc-4-epimerase in plants could lead to metabolic deficiencies in other UDP-sugars by sequestration of the pool of uridinenucleosides. A similar situation is observed in grasses when Gal is infiltrated into tissues [7], because the recycling pathway leads to overproduction of UDP-Gal and starvation of UTP for production of the other UDP-sugars. Alternatively, the properties of galactanrich fibrous tissues, such as in flax stems, could be altered by enrichment of the cell wall galactan. Another enzyme, UDP-Glc dehydrogenase, catalyzes an important step in the synthesis of the uronosyl and pentosyl units of the cell wall. UDP-Glc dehydrogenase catalyzes the conversion of UDP-Glc to UDPGlcA requiring two NAD+. The location of this protein is cytosolic, and it probably forms a hexameric structure [50]. No signal sequence is present in the cDNA [51]; however, there is an indication that it is glycosylated [50]. In soybean and Arabidopsis, the gene is apparently present as a single copy [51]. Because the majority of the matrix polysaccharides are comprised of uronosyl and pentosyl units, regulation of UDP-Glc dehydrogenase activity could be a critical control point. It is in low concentration relative to other enzymes in the pathway, operates far from equilibrium, and is inhibited by UDP-Xyl, one of the products in the pathway [43, 51]. In addition, the expression of mRNA for UDP-Glc dehydrogenase was shown to be greatest in the growing portions of leaves and roots of soybean [51] and soybean nodules [50]. From the UDP-GlcA produced by the activity of UDP-Glc dehydrogenase, both UDP-GalA and UDPvol. 38 (1/2) 2000
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Xyl are formed. A UDP-GalA-C-4 epimerase, distinct from UDP-Glc-C-4 epimerase, forms UDP-GalA in a reversible reaction. The gene has recently been cloned from bacteria, and some of the enzymatic properties studied [34]. The enzyme binds two NAD+ and is strongly inhibited by UDP-Glc, UDP-Gal and UDPXyl. Several conserved amino acids important in binding of NAD+ and UDP-Glc were present, as well as the motif YXXXK, thought to be important for catalytic activity. The activities of UDP-GlcA decarboxylase and UDP-Xyl-4-epimerase sequentially produce UDP-Xyl and UDP-Ara [18]. These enzyme activities have been well studied, but information of their molecular genetics is lacking. Feingold and Avigad [18] also questioned whether UDP-Ara might be formed by the decarboxylation of UDP-GalA. To date no evidence for UDP-GalA decarboxylase in plants has been presented. The mannosyl units of the cell wall are derived from the conversion of the phosphorylated intermediates of sugar metabolism, although a specific C-1 kinase for Man is also present in plants [18]. A GDP-Man pyrophosphorylase catalyzes the activation of the mannosyl units used in cell wall and glycoprotein synthesis. A gene for GDP-Man pyrophosphorylase has recently been cloned in yeast [22]. A hydropathy plot of the predicted amino acid sequence indicates the protein is hydrophilic and lacks a signal peptide, suggesting the protein is a soluble cytoplasmic protein. The significance for the use of GDP instead of UDP sugars is unknown, but it is interesting that GDP-Glc, along with GDP-Man, are the preferred substrates in the synthesis of glucomannan [37]. Further metabolism of GDP-Man provides GDP-LFuc and probably GDP-L-Gal. Fucose-deficient mutants of Arabidopsis (mur1) produced by chemical mutagenesis [40, 41] are due to the mutation and loss of function of GDP-D-Man-4,6-dehydratase [4], the first of two enzymes involved in the formation of GDP-LFuc from GDP-D-Man. The following epimerization and reduction activities apparently are performed by one enzyme in animal cells [52]. Discovery of the fucose deficient mutants of Arabidopsis was the first demonstration that cell wall polysaccharide composition and properties could be addressed by a molecular genetic approach. The mur1 plants have a brittle phenotype in the stems and it was postulated that the lack of an α-L-Fuc unit on the trisaccharide branch of xyloglucan [56] disrupted the ability of xyloglucan to associate with cellulose [30]. Interestingly, α-L-Gal replaced about one-third of the α-L-Fuc found in wild type xyloglucan [56], but only a small proportion of the Plant Physiol. Biochem.
fucosyl units of N-glycans of proteins were replaced with α-L-Gal [38]. Furthermore, it is worth noting that solanaceous xyloglucans do not possess the Fuc-GalXyl trisaccharide and that the lack of α-L-Fuc on essential proteins [38] could be an important part of the brittle phenotype. Other mutants with altered cell wall monosaccharide composition were also identified [40, 41], but it is notable that screening of this mutant population may have missed alterations in the uronic acids, hence much of the pectic polysaccharides, because the wall materials were not carboxylreduced [28] during the derivatization procedures. The nucleotide sugar used in the synthesis of the α-L-Rhm, containing rhamnogalacturonans I and II, has not been demonstrated, but has been variously reported as GDP-L-Rhm, UDP-D-Rhm, or UDP-LRhm due to the presence of these compounds in plants [18]. However, in bacteria dTDP-L-Rhm is the substrate used in the synthesis of the α-L-Rhm containing polysaccharides of the bacterial cell wall. The synthesis pathway of dTDP-L-Rhm begins with dTDPD-Glc pyrophosphorylase forming dTDP-D-Glc. This activity has been demonstrated in plants and suggested to be distinct from the activity of UDP-Glc pyrophosphorylase [18]. A specific nucleotide sugar 4,6dehydratase activity followed by epimerization and reduction, similar to that producing GDP-L-Fuc as described above, produces dTDP-L-Rhm. The set of enzymes involved in the metabolism of GDP-L-Fuc, and the dTDP-L-Rhm dehydratase in bacteria have been cloned and extensively studied [52]. Some bacteria can also produce polysaccharides comprised of D-Rhm, but the synthesis of this polysaccharide requires GDP-D-Rhm provided through the conversion of GDP-D-Man [44]. By analogy, because plant rhamnogalacturonan contains α-L-Rhm, plants may use dTDP-L-Rhm for the synthesis of pectic polysaccharides. The evolutionary origin of α-L-Rhm containing polysaccharides in plants could be interesting in regard to the symbiotic evolution of plants and bacteria, and in the nature and the function of Rhm containing pectic polysaccharides and Rhm-rich proteoglycans. A survey for similar genes and enzyme activities, as well as an evaluation of the glycosyltransferase activities in vitro from plant membranes would be prudent.
2.3. Nucleotide sugar transporters At least three possible scenarios exist for the delivery and utilization of nucleotide sugars by the plant Golgi apparatus as shown in figure 2. In the first, each nucleotide sugar is transported across the Golgi membrane and the active sites of Type II membrane
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glycosyltransferases (discussed below) are oriented toward the lumen. In the second, nucleotide sugars are delivered from the cytoplasmic side to the active site of an integral membrane glycosyltransferases and the growing glycan chain is extruded into the Golgi lumen. In the third, sugar, sugar phosphates or a nucleotide sugar are transported into the Golgi lumen where nucleotide sugar activation or conversions occur. The cytoplasmic location of pyrophosphorylases [18, 29], the existence of specific transporters for each nucleotide sugar species in animal and yeast cells [27], and the indication that UDP-Xyl [23] and UDP-Glc [33] are transported across the Golgi apparatus in plants would argue for a cytoplasmic location of nucleotide sugar conversion in plants. Except for the unlikely possibility that a second set of nucleotide sugar conversion enzymes are present in the lumen of the Golgi apparatus – a condition that would be unique in plants – specific transporters for each nucleotide sugar substrate must exist in the plant Golgi apparatus and endoplasmic reticulum, similar to the situation in animal and yeast cells. Furthermore, the demonstrated presence of nucleoside diphosphatase (NDPase) within the Golgi apparatus of plants and its likely function in polysaccharide synthesis [32] argues for a Golgi lumenal orientation of the active sites of the glycosyltransferases involved in polysaccharide synthesis. NDPase activity produces the nucleoside monophosphate from the nucleoside-diphosphate released by the activity of glycosyltransferases. The resulting nucleoside-monophosphates are removed from the lumen in an antiport process with the corresponding nucleotide sugar transporter [1, 25]. Nucleotide sugar transporters have been implicated as control points for the synthesis of extracellular proteoglycans in animal and yeast cells [27]. In rat hepatocytes, protein glycosylation can be affected by the availability of nucleotide sugars in the Golgi lumen [42]. Increased UDP-sugar concentrations allowed increased incorporation of N-acetylhexosamines at the expense of sialation because of impaired transport of CMP-N-acetylneuraminate. As another example, the loss of UDP-Gal transport in mutant MDCK cells results in a loss of the polysaccharide, keratin sulfate, without affecting galactosylation of the linkage regions of heparin and chondroitin sulfates [53]. These authors suggested that the Km values for the galactosyltransferases that synthesize heparin sulfate and chondroitin sulfate must be significantly lower than the Km value for keratin sulfate suggesting
Figure 2. Proposed scenarios for delivery of nucleotide sugar substrates to sites of polysaccharide synthesis in the Golgi apparatus. A, Specific transporters for each species of nucleotide sugar act in an antiporter process with the corresponding nucleoside monophosphate. Nucleotide sugars are concentrated in the lumen where Type II membrane protein glycosyltransferases produce polysaccharide. Nucleoside diphosphatase produces the monophosphate, releasing Pi, which is removed by a putative Pi transporter. B, Integral membrane protein glycosyltransferases, by analogy to cellulose synthase, would have catalytic sites oriented toward the cytosol and would not require transport of nucleotide sugar. C, Nucleotide sugars if produced in the lumen of the Golgi apparatus, would require transport of sugars, sugar phosphates, UDP-Glc and other required compounds for nucleotide sugar conversions. No evidence for this scenario is present in animal or yeast cells.
a regulation of glycosylation by nucleotide sugar levels through limitations in nucleotide sugar transporters. The location of the transporters, whether in the endoplasmic reticulum or the early or late elements of the Golgi apparatus is often correlated with the location of glycosyltransferase activity [11]. Several cDNA’s for nucleotide sugar transporters have been cloned [27]. The transporters are structurally related hydrophobic proteins with multiple transmembrane regions. Some possess putative leucine zippers that may facilitate protein-protein interactions. Furthermore, a regulation of the type of polysaccharide and possibly the degree of substitution could also be controlled by the relative Km values of glycosyltransferases and the transport of nucleotide sugars. Plants may also control polysaccharide synthesis by regulating nucleotide sugar concentration. Reported Kms for the glycosyltransferases forming the backbones of xyloglucan [33], (1→3),(1→4)-β-D-glucan [7] and galactomannan [39] are one to two orders of vol. 38 (1/2) 2000
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Table I. Properties of some glycosyltransferases involved in cell wall synthesis. Polysaccharide
Resulting linkage
Substrate
Retaining or inverting
Cellulose Callose (1→3),(1→4)-β-D-Glucan Xyloglucan
β-D-Glc-1→4-β-D-Glc β-D-Glc-1→3-β-D-Glc β-D-Glc-3-or 4-β-D-Glc β-D-Glc-1→4-β-D-Glc α-D-Xyl-1→6-β-D-Glc β-D-Gal-1→2-α-D-Xyl α-L-Fuc-1→2-β-D-Gal β-D-Man-1→4-β-D-Glc β-D-Glc-1→4-β-D-Man α-D-Gal-1→2-β-D-Man β-D-Xyl-1→4-β-D-Xyl α-D-GlcA-1→2-β-D-Xyl α-L-Ara-1→2-β-D-Xyl α-D-GalA-1→4-α-D-GalA α-L-Rhm-1→2-α-L-GalA α-D-GalA-1→4-α-D-Rhm
UDP-D-Glc UDP-D-Glc UDP-D-Glc UDP-D-Glc UDP-D-Xyl UDP-D-Gal GDP-L-Fuc GDP-D-Man GDP-D-Glc UDP-D-Gal UDP-D-Xyl UDP-D-GlcA UDP-L-Ara UDP-D-GalA ? UDP-D-GalA
inverting inverting inverting inverting retaining inverting inverting inverting inverting retaining inverting retaining retaining retaining retaining retaining
Galactoglucomannan
Glucuronoarabinoxylan
Homogalacturonan Rhamnogalacturonan
magnitude greater than those reported for the glycosyltransferases of side group additions such as the fucosyltransferase of xyloglucan [36] and the galactosyltransferases of xyloglucan [17] and galactomannan [39]. Although accuracy of the Km determinations are questionable because of disruption of the membrane environment [20], lower Km values of protein processing and side group adding glycosyltransferases would ensure their activity to proceed to completion at low substrate concentration while the higher Km of the glycosyltransferases of backbone polysaccharides could allow overall control of polysaccharide synthesis by substrate level. Furthermore, the ratio of Glc to Man in glucomannan [37], and the ratio of the tri- and tetrasaccharide repeating units of mixed linkage (1→3),(1→4)-β-D-glucan [7] can be manipulated in vitro by varying the concentration of substrates. However, these ratios do not vary in planta even when the substrate concentrations are manipulated. These observations indicate that plants may closely regulate the concentration of substrate within the Golgi lumen.
3. GLYCOSYLTRANSFERASES Until the recent biochemical and molecular genetic characterization of a fucosyltransferase [36] that is involved in the addition of a terminal α-L-fucosyl unit to a xyloglucan acceptor, a cDNA had not been matched to a suitably purified glycosyltransferase from plants. The central difficulty associated with the purification of glycosyltransferases from plants, in particular those forming the backbone, is the loss of activity when solubilized. However, even if the activPlant Physiol. Biochem.
Type of membrane protein integral integral ? ? ? ? Type II ? ? ? ? ? ? ? ? ?
ity can be solubilized, as is the general case for the Type II glycosyltransferases involved in side group formation, we must also have the proper acceptor glycans in a soluble form [17]. Additionally, for many polysaccharides, the substrates are not readily available. Fortunately, genetic approaches can be very useful to identify putative glycosyltransferases. The genes for many glycosyltransferases have been identified by mutation and complementation analysis in bacteria, yeast and animal cells. This information has formed the basis for classification of glycosyltransferases into a number of families based upon primary sequence. The classification has revealed that sequence related proteins share the same mechanisms of activity [9], which may be expected because the structure and function of a protein is intrinsically related to its primary sequence. The mechanisms referred to above are stereochemical descriptions of the donor and acceptor molecules. Glycosyltransferase activities either retain or invert the configuration of the anomeric carbon as it is transferred from the nucleotide α-sugar (β- in the case of GDP-Fuc) substrate [9]. These transfers can be either to the nonreducing end or to a side group position of a glycan. A family classification of glycosyltransferases can be found at http://afmb.cnrs-mrs.fr/∼pedro/CAZY/ db.html, and a summary of linkages, mechanisms and protein classifications of glycosyltransferases involved in cell wall biosynthesis is shown in table I. The above mentioned classifications distinguish between the three-dimensional folding of enzymes and their mechanisms of activity, but do not directly consider the transmembrane spanning regions of the
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Integral membrane glycosyltransferases such as cellulose synthase and callose synthase are known in bacteria, yeast and plants. These integral membrane glycosyltransferases possess several membrane spanning regions and both the NH2 and COOH termini are predicted to be oriented toward the cytosol. Evidence for the identity of cellulose synthase genes is convincing [12]. In addition, sequence similarity of two plant cDNA’s (accessions AAD25952, AAD30609) with those of yeast 1→3-β-D-glucan synthase genes [14, 26, 31] is quite strong. Importantly, this type of integral membrane glycosyltransferase has only been found in the plasmamembrane.
Figure 3. Two types of membrane bound glycosyltransferases may be responsible for polysaccharide synthesis in the plant Golgi apparatus. Integral membrane protein glycosyltransferases such as cellulose synthase and callose synthase are found in the plasma membrane. The presence of integral membrane protein glycosyltransferases in the Golgi membrane for synthesis of polysaccharide backbones has not been demonstrated. Integral membrane proteins are characterized by multiple strands of amino acids spanning the membrane. If the catalytic domain were oriented toward the cytosol, as in cellulose synthase, then a pore would be required for channeling the polysaccharide into the lumen. Type II membrane protein glycosyltransferases have a well-established domain structure. A single strand of amino acids anchors the protein in the membrane. The cytosolic oriented NH2 tail and the lumenal stem region each contribute to the retention of the protein within the Golgi apparatus. The catalytic domain is globular and oriented toward the lumen.
glycosyltransferases. A distinction based upon the nature of the transmembrane regions is apparent and will be discussed below.
3.1. Integral and Type II integral membrane glycosyltransferases Glycosyltransferases involved in glycan and polysaccharide synthesis are integral membrane proteins that can be classified into two broad groups based upon the number of their membrane spanning regions and topology of the catalytic domains as shown in figure 3. The glycosyltransferases of eukaryotes comprise these two groups but prokaryotes have additional variety in the number and position of membrane spanning regions.
Type II membrane glycosyltransferases are well studied in animal and yeast cells for their significance in glycan processing and polysaccharide synthesis [11]. The recently identified fucosyltransferase from Arabidopsis is a predicted Type II membrane protein [36]. Type II membrane proteins are located in the Golgi apparatus. They are characterized by a short NH2 tail oriented toward the cytoplasm, a single membrane spanning region, and a variable length stem connecting with a globular catalytic region oriented toward the lumen. Type II glycosyltransferases are about 400 amino acids long, which is much smaller than the integral glycosyltransferases, such as cellulose synthase and callose synthase that are four to five times longer. Modification of the membrane anchor and stem regions of Type II glycosyltransferases can alter the resulting glycan products [11, 35] by placing the glycosyltransferase in a different sub-compartment of the Golgi apparatus. A variation of the commonly known Type II glycosyltransferases described above involves the cytoplasmic NH2 region of the protein as seen in the KRE6 gene product of yeast [45]. The KRE6 gene product is a Type II membrane protein located in the Golgi apparatus and is the catalytic peptide for synthesis of the 1→6-β-D-glucan of yeast cell walls. It is different from most Type II glycosyltransferases however in that the cytoplasmic NH2 tail is much larger. The cytoplasmic region of Kre6p is 252 amino acids, and contains several consensus sites for phosphorylation (RXXS/T where S or T is phosphorylated) that resemble those of PKC1, MAP kinase, cAMPdependent protein kinase and casein kinase. Several lines of evidence suggest the PKC1 pathway participates in the regulation of yeast cell wall synthesis. It is tempting to speculate that plants may also regulate the synthesis of matrix polysaccharides by phosphorylation-mediated pathways. vol. 38 (1/2) 2000
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3.2. Conserved amino acids of glycosyltransferases Sequence analysis of DNA or protein has helped to identify by similarity many hundreds of glycosyltransferases. For example, BLAST analysis has been used to identify members within the large families of cellulose synthases (http://cellwall.stanford.edu), fucosyltransferases [6, 36] and galactosyltransferases [5]. However, even within a family, there can be as little as 25 % similarity in primary sequence. The diversity of sequence is presumably a reflection of the different substrates, the linkages formed, and the acceptor recognition regions. Furthermore, convergent evolution has very likely occurred and this may obscure homologies when searching for sequence similarities. At present, there are insufficient examples from plants for comparisons to be made that will help identify many glycosyltransferases involved in cell wall synthesis because many activities are unique to plants and presumably, the genes for these glycosyltransferases will be highly divergent in sequence. The grouping of glycosyltransferases into 36 families – more are likely to be identified – is testament to their diversity [9]. However, within each family, similarity in threedimensional folding structure is expected. Thus, knowledge of the three-dimensional protein structure of a few glycosyltransferases can aid in identifying divergent glycosyltransferases. Underlying the threedimensional folding of a protein is the protein’s secondary structure. Hydrophobic cluster analysis (HCA) is an indicator of the secondary structure of a protein, and HCA has been used to help identify a limited number of conserved regions and amino acids that are related to active sites and acceptor binding regions. Clearly, the search for putative genes for glycosyltransferases would be greatly advanced if we could identify motifs that are conserved among all or within families of glycosyltransferases.
Glycosyltransferases most often have aspartate residues in the active sites. Three such conserved aspartate residues are found in the integral glycosyltransferases such as cellulose, and chitin synthases whereas only two are conserved in the Type II glycosyltransferases. These structures are clearly seen in the enzymes studied by Saxena et al. [47], the galactosyltransferases [5] and less clearly in the fucosyltransferases [6]. Comparing the integral and Type II glycosyltransferases, it appears that the integral glycosyltransferases have a catalytic region divided into two domains, A and B, whereas Type II enzymes have only one domain which appears equivalent to domain A. One of the aspartate residues in domain A is preceded by a vertical hydrophobic cluster. This motif is present in both integral and Type II glycosyltransferases. This motif can be found in proteins that are not glycosyltransferases, so it is not a unique signature of glycosyltransferases, but most if not all glycosyltransferases have it. Sometimes, this motif is difficult to see especially if a non-hydrophobic amino acid interrupts the vertical hydrophobic cluster. However, in addition to the motif, the second of the conserved aspartate regions should be looked for when searching the database for potential genes of glycosyltransferases. Sometimes the order of the conserved aspartate residues appears reversed, and the distance between can vary from 10 to well over 100 amino acids. Additionally, most of the conserved aspartate residues are found in a DD or DXD motif. Furthermore, a few conserved motifs are present within a group of glycosyltransferases that have the same substrate. In this respect, it is interesting that the xyloglucan fucosyltransferase [36] has a KPW motif that is seen in the galactosyltransferases [5]. Representations of topol-
Figure 4. Topology predictions (left panels) and HCA plots (right panels) of a bacterial cellulose synthase (AcscA, accession CAA38487), a plant cellulose synthase (GhCesA-1, accession AAB37766), a plant fucosyltransferase (XG-FT1, accession AAD41092), a yeast (1→6)-β-D-glucan glucosyltransferase (KRE6, accession P32468), and a plant putative glycosyltransferase (T7N9.14, accession AAB61490). The topology predictions were based upon a hidden Markov model and were drawn using the TMHMM program (http://www.cbs.dtu.dk/services/TMHMM-1.0/). The height of the vertical bars represents the probability of a transmembrane helix. Topology predictions are a useful indicator of integral membrane proteins (top two panels) and type II integral membrane proteins (bottom three panels). The HCA’s were performed using the drawhca program (http://www.lmcp.jussieu.fr/∼soyer/www-hca/hca-file.html). HCA involves drawing the amino acid sequence on a cylinder representing an α-helix. The cylinder is split lengthwise, flattened, and the sequences duplicated to restore the connectivity of the amino acids. The one-letter code is used for amino acids except for glycine (diamonds), proline (stars), serine (squares with dots), and threonine (squares). Acidic amino acids are shown in red, basic amino acids in blue, and hydrophobic amino acids in green. Contours are also drawn around the clusters of hydrophobic amino acids to aid in the visual analysis of their shape, length and distribution. The conserved QXXRW motif of cellulose synthases is shown over a gray background and conserved aspartate residues are circled. For XG-FT1, a KPW motif is also indicated over a gray background although this motif appears conserved in family B of galactosyltransferases [5]. Along with the identification of conserved amino acids using sequence alignment methods, HCA is a useful tool to uncover conserved motifs. The regions indicated by the yellow background were identified as described in the text; however, their usefulness in predicting the function of a gene product as a glycosyltransferase is highly speculative. More examples of HCA plots of glycosyltransferases can be found in references [5, 6, 47].
Plant Physiol. Biochem.
Matrix polysaccharides
ogy predictions and HCA’s of some glycosyltransferases are shown in figure 4. It should be emphasized that only a few glycosyltransferases have been studied in enough detail to identify the critical amino acids in active sites and nucleotide binding regions. The number and arrange-
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ment of putative catalytic sites and binding regions is still speculative for the majority. These issues, in particular the number and identity of active sites will be resolved with amino acid modification studies, cryo-electron microscopy and if possible with X-ray crystallographic studies of crystallized proteins.
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3.3. Identifying genes for glycosyltransferases The rapid pace of EST and genome sequencing means that many genes for glycosyltransferases are already available in the database. The difficulty is to identify putative genes and then demonstrate their function. This is especially problematic for the polysaccharide producing glycosyltransferases of plants because we do not have a similar means to produce and recover knock-out mutants in a way that has been so useful in other organisms (but see Fagard et al. in this issue). We may infer from the screening of mutant Arabidopsis [40, 41] that many loss of function mutations will not be recovered. This may also happen in anti-sense transformation strategies unless inducible promoters are used. A more rapid approach, and one that avoids some of the pitfalls of lethal mutations, is the use of virus-induced gene silencing [3]. To perform virus-induced gene silencing (VIGS), genes or gene fragments of interest are inserted into a potato X potexvirus (PVX), and RNA transcripts are used for infection of seedlings of Nicotiana benthamiana. Post-transcriptional gene silencing results in reduced mRNA levels of endogenously expressed plant genes that have a high level of sequence identity with the sequence carried by the virus. Furthermore, a length of only 300–500 nucleotides is sufficient to effect silencing. Thus, full-length cDNAs or genes are not required which is particularly advantageous for the ‘fast forward’ screening of candidate genes [3]. The major advantage of the VIGS system relates to the relative speed with which the role of a gene product can be identified, compared with antisense or co-suppression approaches in transgenic plants. In addition, VIGS can be used to knock out potentially lethal genes, because young seedlings are established before the gene is introduced via the viral RNA vector. We have used a homologous fragment of the cellulose synthase gene, GhCesA, and VIGS to demonstrate the knock-out of cellulose synthesis. The silencing of cellulose synthase resulted in dwarfism, and a disruption of leaf cell shape. Interestingly, there was compensation for the loss of cellulose by an increase of homogalacturonan with a low degree of carboxyl esterification [8]. With a suitable method for functional analysis now in hand, efficient selection of candidate genes can be increased by knowledge of the gene and protein structures of glycosyltransferases as described above. A combination of BLAST, topology prediction, and HCA can be used to identify protein sequences by the structural characteristics that are consistent with the large number of glycosyltransferases already identified. Plant Physiol. Biochem.
In addition to considerations of protein structure, identification of sequence motifs within a family of glycosyltransferases will narrow the field of candidates, and may be critical for selecting gene sequences.
4. CONCLUSIONS Within the lumen of the Golgi apparatus, many polysaccharides, proteoglycans and glycoproteins are synthesized and organized for secretion to the cell wall. The mechanisms for the location and organization of nucleotide sugar transporters and the glycosyltransferases that use the corresponding substrate are hot topics in animal and yeast research because of the potential to understand certain disease processes, and for the production of valuable proteins and biopolymers. Knowledge of these processes in plants may also generate the means to understand and modify plant development through alteration of the chemistry and structure of the cell wall. Even with the promise given by genetic analysis of the glycosyltransferases fast approaching as shown by the genetic descriptions of cellulose synthase [12] and fucosyltransferase [36], we realize there will also be much to be discovered in the cellular mechanics of cell wall biosynthesis. One important question yet to be answered is whether the backbone producing glycosyltransferases in the Golgi apparatus are integral or Type II membrane proteins. The answers will suggest much about the intracellular mechanisms of matrix polysaccharide synthesis.
Acknowledgments This work was supported by a grant from the Grains Research and Development Corporation of Australia (to Geoffrey B. Fincher).
REFERENCES [1] Abeijon C., Mandon E.C., Hirschberg C.B., Transporters of nucleotide sugars, nucleotide sulfate and ATP in the Golgi apparatus, Trends Biochem. Sci. 22 (1997) 203–207. [2] Amor Y., Haigler C.H., Johnson S., Wainscott M., Delmer D.P., A membrane-associated form of sucrose synthase and its potential role in synthesis of cellulose and callose in plants, Proc. Natl. Acad. Sci. USA 92 (1995) 9353–9357.
Matrix polysaccharides
[3] Baulcombe D.C., Fast forward genetics based on virus-induced gene silencing, Curr. Opin. Plant Biol. 2 (1999) 109–113. [4] Bonin C.P., Potter I., Vanzin G.F., Reiter W.D., The MUR1 gene of Arabidopsis thaliana encodes an isoform of GDP-D-mannose-4,6-dehydratase, catalyzing the first step in the de novo synthesis of GDP-Lfucose, Proc. Natl. Acad. Sci. USA 94 (1997) 2085–2090. [5] Breton C., Bettler E., Joziasse D.H., Geremia R.A., Imberty A., Sequence-function relationships of prokaryotic and eukaryotic galactosyltransferases, J. Biochem. 123 (1998) 1000–1009. [6] Breton C., Oriol R., Imberty A., Conserved structural features in eukaryotic and prokaryotic fucosyltransferases, Glycobiology 8 (1998) 87–94. [7] Buckeridge M.S., Vergara C.E., Carpita N.C., The mechanism of synthesis of a mixed-linkage (1→3),(1→4) β-D-glucan in maize (Zea mays L.). Evidence for multiple sites of glucosyl transfer in the synthase complex, Plant Physiol. 120 (1999) 1–12. [8] Burton R.A., Gibeaut D.M., Bacic A., Findlay K., Roberts K., Hamilton A., Baulcombe D.C., Fincher G.B., Virus-induced silencing of a plant cellulose synthase gene, Plant Cell (1999) (in press). [9] Campbell J.A., Davies G.J., Bulone V., Henrissat B., A classification of nucleotide-diphospho-sugar glycosyltransferases based on amino acid sequence similarities, Biochem. J. 326 (1997) 929–942. [10] Carpita N.C., Gibeaut D.M., Structural models of primary cell walls in flowering plants: consistancy of molecular structure with the physical properties of the walls during growth, Plant J. 3 (1993) 1–30. [11] Colley K.J., Golgi localization of glycosyltransferases – more questions than answers, Glycobiology 7 (1997) 1–13. [12] Delmer D.P., Cellulose biosynthesis: exciting times for a difficult field of study, Annu. Rev. Plant Physiol. Plant Mol. Biol. 50 (1999) 245–276. [13] Dörmann P., Benning C., Functional expression of uridine 5’-diphospho-glucose 4-epimerase (EC 5.1.3.2) from Arabidopsis thaliana in Saccharomyces cerevisiae and Escherichia coli, Arch. Biochem. Biophys. 327 (1996) 27–34. [14] Douglas C.M., Foor F., Marrinan J.A., Morin N., Nielsen J.B., Dahl A.M., Mazur P., Baginsky W., Li W., El-Sherbeini M., Clemas J.A., Mandala S.M., Frommer B.R., Kurtz M.B., The Saccharomyces cerevisiae FKS1 (ETG1) gene encodes an integral membrane protein which is a subunit of 1,3-β-D-glucan synthase, Proc. Natl. Acad. Sci. USA 91 (1994) 12907–12911. [15] Eimert K., Villand P., Kilian A., Klecczkowski L.A., Cloning and characterization of several cDNAs for UDP-glucose pyrophosphorylase from barley (Hordeum vulgare) tissues, Gene 170 (1996) 227–232. [16] Emons A.M.C., Mulder B.M., The making of the architecture of the plant cell wall: how cells exploit geometry, Proc. Natl. Acad. Sci. USA 95 (1998) 7215–7219.
79
[17] Faïk A., Chileshe C., Sterling J., Maclachlan G., Xyloglucan galactosyl- and fucosyltransferase activities from pea epicotyl microsomes, Plant Physiol. 114 (1997) 245–254. [18] Feingold D.S., Avigad G., Sugar nucleotide transformations in plants, in: Preiss J. (Ed.), The Biochemistry of Plants. Carbohydrates: Structure and Function, vol. 3, Academic Press, 1980, pp. 101–170. [19] Gibeaut D.M., Carpita N.C., Tracing the biosynthesis of the cell wall in intact cells and plants. Selective turnover and alteration of soluble and cell wall polysaccharides in grasses, Plant Physiol. 97 (1991) 551–561. [20] Gibeaut D.M., Carpita N.C., Biosynthesis of plant cell wall polysaccharides, FASEB J. 8 (1994) 904–915. [21] Gorshkova T.A., Chemikosova S.B., Lozovaya V.V., Carpita N.C., Turnover of galactans and other cell wall polysaccharides during development of flax plants, Plant Physiol. 114 (1997) 723–729. [22] Hashimoto H., Sakakibara A., Yamasaki M., Yoda K., Saccharomyces cerevisiae VIG9 encodes GDPmannose pyrophosphorylase, which is essential for protein glycosylation, J. Biol. Chem. 272 (1997) 16308–16314. [23] Hayashi T., Koyama T., Matsuda K., Formation of UDP-xylose and xyloglucan in soybean Golgi membranes, Plant Physiol. 87 (1988) 341–345. [24] Hirano Y., Tsumuraya Y., Hashimoto Y., Characterization of spinach leaf α-L-arabinofuranosidases and β-galactosidases and their synergistic action on an endogenous arabinogalactan-protein, Physiol. Plant. 92 (1994) 286–296. [25] Hirschberg C.B., Transporters of nuleotide sugars, nucleotide sulfate and ATP in the Golgi apparatus membrane – where next?, Glycobiology 7 (1997) 169–171. [26] Inoue S.B., Takewaki N., Takasuka T., Mio T., Adachi M., Fujii Y., Miyamoto C., Arisawa M., Furuichi Y., Watanabe T., Characterization and gene cloning of 1,3-β-D-glucan synthase from Saccharomyces cerevisiae, Eur. J. Biochem. 231 (1995) 845–854. [27] Kawakita M., Ishida N., Miura N., Sun-Wada G.H., Yoshioka S., Nucleotide sugar transporters: elucidation of their molecular identity and its implication for future studies, J. Biochem. 123 (1998) 777–785. [28] Kim J.B., Carpita N.C., Changes in esterification of the uronic acid groups of cell wall polysaccharides during elongation of maize coleoptiles, Plant Physiol. 98 (1992) 646–653. [29] Klecczkowski L.A., Glucose activation and metabolism through UDP-glucose pyrophosphorylase in plants, Phytochemistry 37 (1994) 1507–1515. [30] Levy S., Maclachlan G., Staehelin L.A., Xyloglucan sidechains modulate binding to cellulose during in vitro binding assays as predicted by conformational dynamics simulations, Plant J. 11 (1997) 373–386. vol. 38 (1/2) 2000
80
D.M. Gibeaut
[31] Mazur P., Morin N., Baginsky W., El-Serbeini M., Clemas J.A., Nielsen J.B., Foor F., Differential expression and function of two homologous subunits of yeast 1,3-β-D-glucan synthase, Mol. Cell. Biol. 15 (1995) 5671–5681. [32] Mitsui T., Honma M., Kondo T., Hashimoto N., Kimura S., Igaue I., Structure and function of the Golgi complex in rice cells II. Purification and characterization of Golgi membrane-bound nucleoside diphosphatase, Plant Physiol. 106 (1994) 119–125. [33] Muñoz P., Norambuena L., Orellana A., Evidence for a UDP-glucose transporter in Golgi apparatus-derived vesicles from pea and its possible role in polysaccharide biosynthesis, Plant Physiol. 112 (1996) 1585–1594. [34] Muñoz R., López R., de Frutos M., Garcia E., First molecular characterization of a uridine diphosphate galacturonate 4-epimerase: an enzyme required for capsular biosynthesis in Streptococcus pneumoniae type 1, Mol. Microbiol. 31 (1999) 703–713. [35] Paulson J.C., Colley K.J., Glycosyltransferases. Structure, localization, and control of cell type-specific glycosylation, J. Biol. Chem. 264 (1989) 17615–17618. [36] Perrin R.M., DeRocher A.E., Bar-Peled M., Zeng W., Norambuena L., Orellana A., Raikhel N.V., Keegstra K., Xyloglucan fucosyltransferase, an enzyme involved in plant cell wall biosynthesis, Science 284 (1999) 1976–1979. [37] Piro G., Dalessandro G., Northcote D.H., Glucomannan synthesis in pea epicotyls: the mannose and glucose transferases, Planta 190 (1993) 206–220. [38] Rayon C., Cabanes-Macheteau M., Loutelier-Bourhis C., Salliot-Maire I., Lemoine J., Reiter W-D., Lerouge P., Faye L., Characterization of N-glycans from Arabidopsis. Application to a fucose-deficient mutant, Plant Physiol. 119 (1999) 725–733. [39] Reid J.S.G., Edwards M., Gidley M.J., Clark A.H., Enzyme specificity in galactomannan biosynthesis, Planta 195 (1995) 489–495. [40] Reiter W.D., Chapple C., Somerville C.R., Altered growth and cell walls in a fucose-deficient mutant of Arabidopsis, Science 261 (1993) 1032–1035. [41] Reiter W.D., Chapple C., Somerville C.R., Mutants of Arabidopsis thaliana with altered cell wall polysaccharide composition, Plant J. 12 (1997) 335–345. [42] Pels Rijcken W.R., Overdijk B., Van Den Eijnden D.H., Ferwerda W., The effect of increasing nucleotide-sugar concentrations on the incorporation of sugars into glycoconjugates in rat hepatocytes, Biochem. J. 305 (1995) 865–870. [43] Robertson D., McCormack B.A., Bolwell G.P., Cell wall polysaccharide biosynthesis and related metabolism in elicitor-stressed cells of French bean (Phaseolus vulgaris L.), Biochem. J. 306 (1995) 745–750. [44] Rocchetta H.L., Pacan J.C., Lam J.S., Synthesis of the A-band polysaccharide sugar D-rhamnose requires RMD and WBPW – identification of multiple ALGA homologues, WPBW and ORF488, in Pseudomonas aeruginosa, Mol. Microbiol. 29 (1998) 1419–1434. Plant Physiol. Biochem.
[45] Roemer T., Paravicini G., Payton M.A., Bussey H., Characterization of the yeast (1-6)-β-glucan biosynthetic components, Kre6p and Skn1p, and genetic interactions between the PKC1 pathway and extracellular matrix assembly, J. Cell Biol. 127 (1994) 567–579. [46] Ruan Y.L., Chourey P.S., Delmer D.P., Perez-Grau L., The differential expression of sucrose synthase in relation to diverse patterns of carbon partitioning in developing cotten seed, Plant Physiol. 115 (1997) 375–385. [47] Saxena I.M., Brown Jr R.M., Fevre M., Geremia R.A., Henrissat B., Multidomain architecture of β-glycosyl transferases: implications for mechanism of action, J. Bacteriol. 177 (1995) 1419–1424. [48] Schlüpmann H., Bacic A., Read S.M., Uridine diphosphate glucose metabolism and callose synthesis in cultured pollen tubes of Nicotiana alata Link et Otto, Plant Physiol. 105 (1994) 659–670. [49] Schultz C., Gilson P., Oxley D., Youl J., Bacic A., GPI-anchors on arabinogalactan-proteins: implications for signalling in plants, Trends Plant Sci. 3 (1998) 426–431. [50] Stewart D.C., Copeland L., Uridine 5’-diphosphateglucose dehydrogenase from soybean nodules, Plant Physiol. 116 (1998) 349–355. [51] Tenhaken R., Thulke O., Cloning of an enzyme that synthesizes a key nucleotide-sugar precursor of hemicellulose biosynthesis from soybean: UDP-glucose dehydrogenase, Plant Physiol. 112 (1996) 1127–1134. [52] Tonetti M., Sturla L., Bisso A., Zanardi D., Benatti U., De Flora A., The metabolism of 6-deoxyhexoses in bacterial and animal cells, Biochimie 80 (1998) 923–931. [53] Toma L., Pinhal M.A.S., Dietrich C.P., Nader H.B., Hirschberg C.B., Transport of UDP-galactose into the Golgi lumen regulates the biosynthesis of proteoglycans, J. Biol. Chem. 271 (1996) 3897–3901. [54] Wakabayashi K., Sakurai N., Kuraishi S., Effects of ABA on synthesis of cell-wall polysaccharides in segments of etiolated squash hypocotyl. I. Changes in incorporation of glucose and myo-inositol into cellwall components, Plant Cell Physiol. 30 (1989) 99–105. [55] Youl J.J., Bacic A., Oxley D., Arabinogalactanproteins from Nicotiana alata and Pyrus communis contain glycosylphosphatidylinositol membrane anchors, Proc. Natl. Acad. Sci. USA 95 (1998) 7921–7926. [56] Zablackis E., York W.S., Pauly M., Hantus S., Reiter W.D., Chapple C.C.S., Albersheim P., Darvill A., Substitution of L-fucose by L-galactose in cell walls of Arabidopsis mur1, Science 272 (1996) 1808–1810.