Proteins are ubiquitous components of plant cell walls where they may account for as much as 10% of the dry weight of the wall. The wall proteins include enzymes, wall loos­ening proteins (expansins) (71), and signaling molecules (AGPs) (70) but the structural proteins are quantitatively the most important. These include the glycine-rich proteins (GRP) (72, 73), the hydroxyproline-rich glycoproteins (HRGP, extensins), the proline — and hydroxyproline-rich proteins (PRP), and non-extensin proteins (70, 74-77). The HRGP are glycoproteins bearing short arabinofuranoside side chains at the Hyp residues (70). They are highly elongated molecules with extended polyproline II helix conformations imparted by the runs of Hyp residues. The HRGP, PRP, and AGP families also contain members that are chimeras having, for example, an HRGP domain fused to a domain with AGP characteristics (70). The structural proteins of the walls are thought to form an independent wall matrix phase.

RG-II is the most structurally complicated polysaccharide in the cell wall. It is present in the walls of all plants and its structure is highly conserved. RG-II makes up approximately 4% of the cell wall in dicotyledonous plants and less than 1% of the wall in monocots (157). It contains 12 different types of glycosyl residues in at least 20 different linkages (159) including both methyl etherified (e. g., 2-0 Me-xylose and 2-O Me-fucose) and O — acetylated glycosyl residues (e. g., 3-O-, or 4-0-Ac-fucose). RG-II also contains unusual sugars such as aceric acid (3-C-carboxy-5-deoxy-L-xylose) (346), KDO (2-keto-3-deoxy-D- manno-octulopyranosylonicacid (345), andDHA (3-deoxy-D-lyxo-2-heptulopyranosylaric acid) (346). RG-II has a backbone of a-1,4-linked D-galactosyluronic acid with structurally complex side chains attached to C-2 and/or C-3 (2, 157, 159, 269, 345-352). RG-II in the wall exists largely complexed with borate as an RG-II dimer that is cross-linked by a borate diester (159,352-357). At least 24 transferase activities are expected to be required for RG-II synthesis. There have been very few systematic studies of RG-II synthesis, although progress is beginning to be made on several enzymes.

GDP-Man (485) is a major sugar donor that provides flux of sugar to the synthesis of glycoproteins, polysaccharides, and ascorbic acid in plants. GDP-Man is the precursor for GDP-Fuc and GDP-Gal. Guanosine 5′-diphosphate-mannose pyrophosphorylase (GDP — Man PPase) catalyzes the conversion of a-D-Man-1-P and GTP to GDP-Man and pyrophos­phate (486). The enzyme activity requires Mg2+ and the enzyme appears to be cytosolic. The pyrophosphorylase is well studied in various organisms and the crystal structure is known. In Arabidopsis, a wall mutant cytl (487), an ozone-sensitive mutant, and ascorbic acid mutant, vtcl, identified the same gene product At2g39770 as the locus responsible for the production of GDP-Man (488, 489). The mutant likely survives since another homol­ogous gene, At4g30570, may compensate for its activity. Although the enzymatic activity

of recombinant protein was not described, work in our laboratory confirmed that the en­coded gene At2g39770 has GDP-Man PPase activity and is very specific toward Man-1-P as a substrate (Echole and Bar-Peled, unpublished).

Although this chapter attempts to give justice later to what we now know about native lignin macromolecular configuration and modulation thereof, it must be robustly stressed that the analytical methodologies currently available for the study of polymeric lignins (structure, content, and composition) are woefully inadequate, as well as being unsatisfactory from any holistic quantification perspective. Indeed, such inherent shortcomings hold for all of the commonly used protocols — including the ones that we use, such as either NMR, thioaci — dolysis, alkaline nitrobenzene oxidation, or Klason analyses, to give just a few examples. For these reasons, it is important to appreciate and understand the very limited information that is being gained with all these approaches. As discussed below, often only a very small fraction (i. e., 20-40%) of the lignins can be accounted for in a chemically definitive and quantitative manner through degradative analyses.

Such massive deficits and/or discrepancies in the analytical procedures have, in one way or another, existed for nearly a century. For this reason, it is imperative that in the near future new approaches/technologies be urgently developed to determine, for example, the nature and frequency of all interunit linkages in lignin(s) — an essential requirement if we are to fully understand how lignins are, in fact, being formed in vivo.

While these limitations are, in large part, due to both the intractable and polymeric na­ture of native macromolecular lignins, there is yet another confounding feature. That is, with one exception (77), most of the research carried out on lignification (and its modu­lation) to date has not been directed toward any systematic and holistic determination of trends in the lignin-forming processes. Furthermore, there are increasing numbers of re­ports of lignin contents and compositions that are unreliable, and which presumably reflect a lack of scientific rigor and knowledge about lignin/cell-wall chemistries. We would plead that reliable/rigorous chemical and biochemical analyses must be an expectation for lignin investigations as they are for all other fields of (plant) chemistry/phytochemistry.

An additional complicating feature in the analysis of native lignin macromolecular con­figuration is that of the large variations in H, G, and S monomeric compositions and lignin contents between and within plant species. Quite remarkably, no comprehensive attempts to correlate such variability in lignin content/composition with cell wall anatomy have been reported. Yet, this would appear to be useful in terms of potentially identifying simpli — fying/unifying features in lignin macromolecular configuration, e. g., by correlating lignin amounts/composition with overall dimensions of the vascular apparatus and distinct cell types present in different plant lines, such as in the syringyl-rich fiber cell walls, for example.

We initially considered it instructive to determine how control over monolignol radical coupling might be effectuated, particularly given that this was a factor largely not considered as being possible during the 1950s/1960s. On the other hand, this was a relevant issue with the

large numbers of structurally related lignans now known. In this regard, many different forms of specific coupling have been reported in isolated metabolites (e. g., containing specifically — linked 8-1′, 8-5′, 8- O-4′, 5-5′, 3- O-4′, 7-1′, 8-7′, 1-5′, and 2- O-3′ interunit linkages) depending upon the metabolite and/or plant species in question (321, 322). This indicated, at the very least, that a mechanism of regiospecific coupling control had evolved in planta for phenolic coupling. Additionally, large numbers of lignan metabolites are optically active suggesting, in turn, that stereoselective coupling might also be occurring in many instances. For the purposes of this discussion, however, there are two pertinent examples of potential control over phenoxy radical-radical coupling. These include formation of the 8-8′-linked (+)-pinoresinol (69a, Figure 7.19A) in Forsythia species (323-327), and that of (meso)

8-8r-linked nor dihydroguaiaretic acid (84, Figure 7.20C) in the creosote bush (Larrea tridentata) (328, 329), respectively.

Work was thus undertaken to initially establish how the lignan (+)-pinoresinol (69a) was formed, this having been an earlier unresolved scientific interest of Holgar Erdtman. Using Forsythia suspensa as a biological partner, we were able to demonstrate that preferential stereoselective coupling of two Е-coniferyl alcohol (3) moieties occurred to afford (+)- pinoresinol (69a) when incubated with crude “cell wall’Vinsoluble preparations (323). Sub­sequent purification of the various proteinaceous components in this crude preparation ul­timately afforded (as estimated by SDS-PAGE) an ~ 110 kDa one-electron oxidase (a laccase) and an ~26 kDa protein, with the latter protein lacking monolignol oxidizing capacity (324). The laccase alone in vitro generated coniferyl alcohol radicals which underwent the well — known nonspecific coupling to afford the corresponding racemic (±)-dihydrodiconiferyl alcohols (68a/b), (±)-pinoresinols (69a/b), and (±)-threo/erythro guaiacylglycerol 8-0-4

coniferyl alcohol ethers (71a/b) (Figure 7.19A). On the other hand, when the 26 kDa pro­tein, which existed as an ~50-52 kDa dimer, was added to the assay mixture, stereoselective coupling occurred instead to afford (+)-pinoresinol (69a) (Figure 7.19A).

Interestingly, in addition to the laccase, other one-electron oxidases (peroxidase) and one-electron oxidizing agents were able to effectuate stereoselective coupling in the pres­ence of this protein (324). Given this striking ability to dictate the outcome of stereoselective coupling with the (+)-pinoresinol-forming protein, we coined the term dirigent protein (DP) from the Latin: dirigere, to guide or to align (324). The corresponding gene was next obtained, with this encoding a protein of ~18 kDa. The discrepancy in the molecular size (~26 versus ~18 kDa) was due to posttranslational (glycosylation) modification (325); functionally competent recombinant protein was also obtained when expressed in a “glyco­sylated” form using insect (Spodoptera frugiperda) cell cultures (325).

The biochemical mode of action of the (+)-pinoresinol forming DP has since been the subject of our recent work (326, 327), where it is considered to function by trapping monolignol (radicals). Figure 7.19B thus depicts our current understanding of the kinetic and “rate-limiting” processes presumed to be involved in stereoselective coupling, versus that of nonspecific coupling leading to racemic dimers (327). Importantly, it was demonstrated that proteins had indeed evolved monolignol (radical) binding capacity, and the ability to engender specific coupling modes, in vascular plants.

However, we considered the (+)-pinoresinol forming DP discovery as a special example of control over monolignol radical-radical coupling, i. e., whereby the monolignol (radical) was bound to the active site of each monomer in the DP dimer. In this way, the (+)- pinoresinol-forming DP dimer was able to orientate both coniferyl alcohol radicals in such a way as to where only (+)-pinoresinol (69a) formation could occur (Figure 7.19A).

More recent work has established that dirigent proteins and their homologues are found throughout the vascular plant kingdom (330-332). However, they appear to be restricted to land plants, suggesting they obtained their function(s) during the transition of plants to land. Dirigent proteins are generally also present in large multigene families with varying levels of homology (e. g., from 99.5 to 12.5% identity) (331, 332), with most biochem- ical/physiological functions of their homologues currently unknown. As a beginning to define their functions, we have investigated their expression profiles, using a GUS-reporter system linked to each putative DP promoter of two dirigent multigene families and/or ho­mologues in both western red cedar (Thuja plicata) (333) and Arabidopsis (334) (Kim etal., manuscript in preparation). [Western red cedar was of particular interest since it accumu­lates the (+)-pinoresinol-derived metabolite, plicatic acid (74, Figure 7.19C), and several of its DPs have the capacity for (+)-pinoresinol (69a) formation (330).] Determination of their expression profiles was carried out for both families; this appears to be a useful approach to begin to establish the biochemical/physiological functions of the various homologues.

There have also been a number of other studies which provisionally suggest DP control over other coupling modes. For example, using Eucommia ulmoides “insoluble residues” Lourith et al. (335, 336) reported diastereoselective homo — and hetero-coupling of sinapyl (5) and coniferyl (3) alcohol moieties in vitro, without addition of any cofactor, to afford the 8-0-4′ lignans 75-78 (Figure 7.20A) of differing levels of erythro/threo ratios and optical activities (335, 336). The proteins involved in such coupling now need to be purified, and their encoding genes cloned, in order to more fully characterize the basis of such transformations.

Another intriguing report ofpresumed dirigent protein involvement is in the stereoselec­tive coupling of two molecules of the terpenoid, hemigossypol (79), to afford (+)-gossypol (80a) in cotton (Gossypium hirsutum L. var. marie galante) flower petals (Figure 7.20B) (337). Interestingly, different Gossypium varieties accumulate varying levels of (+)- and ( — )-gossypols (80a and 80b), with both having equal toxicity toward insects and pathogens. However, cotton seeds generally cannot be used in animal feed because of the toxicity of (-)-gossypol (80b). Breeding has thus been used to select varieties accumulating the (+)-, but not the ( —)-, antipode of gossypol (80). Research has also recently been conducted to

understand the biochemical basis of (+)-gossypol (80a) formation in cotton flowers (337). In the presence of a one-electron oxidase (peroxidase, laccase), coupling of hemigossypol (79) only afforded racemic gossypol (80a/b), whereas when a presumed dirigent protein — again lacking oxidative capacity was added — stereoselective coupling occurred to essen­tially only give (+)-gossypol (80a). Thus, it would provisionally appear that DP control over radical-radical coupling can involve metabolically quite distinct plant product classes.

Other forms of control over regiospecific radical-radical coupling have also been noted for formation of the 8-8r-linked lignans, such as in the creosote bush (Larrea tridentata). The latter accumulates (meso)-nordihydroguaiaretic acid (NDGA, 84) and other 8-8r-linked lignans. Interestingly, while the presumed precursor of NDGA, p-anol (81) can potentially undergo various forms of coupling at different sites on the molecules, such as at the 4- O, C-5, C-1, and C-8 positions, only regiospecific coupling at 8-8r occurs (see Figure 7.20C). This is again indicative of, at the minimum, regiospecific control over phenolic radical — radical coupling (328, 329). Other examples of regiospecific coupling are also apparently found in ellagitannin metabolism, such as in the intermolecular coupling of pentagalloyl glucose (85) moieties to afford tellimagrandin II (86) and subsequent intramolecular cou­pling of the latter to generate cornusiin E (87) (Figure 7.21) (338-340). While these are presumably considered to be “laccase-like” protein mediated (338-340), it will be impor­tant to establish how their biochemical mechanisms differ — if they do — from the dirigent

protein-mediated coupling. Work is currently underway to explore this possibility. In any event, biochemical mechanisms have been preliminarily described for proteinaceous control over both stereoselective and regiospecific coupling.

7.7.3.3.1 PROTEIN VERSUS NON-PROTEIN DIRECTED NATIVE LIGNIN

MACROMOLECULAR CONFIGURATION AND THE QUESTION OF RACEMISATION IN LIGNIN STRUCTURE The preceding sections have emphasized some of the difficulties in establishing native lignin macromolecular configuration — beginning with the technological limitations experienced in lignin analyses more than five decades ago — to the present date. In addition, verification of the long-known racemic nature of lignins and of lignin subunit fragments apparently con­vinced some researchers more recently that lignin formation must indeed occur randomly in vivo (175,341). However, the presence of racemic substructures does not eliminate proteina­ceous control over lignin macromolecular configuration as described below. Additionally, the notion of random coupling has in turn led to other suggestions — but again without rigorous proof — that various cell wall constituents, such as hemicelluloses, cellulose, have important roles in determining and/or establishing lignin configuration in vivo. Several of the hemicelluloses may, however, indeed be involved in forming lignin-carbohydrate bonds, e. g., via reaction with intermediate quinone methides.

However, such considerations fully ignore both the presence and the roles of cell wall pro­teins, the vast majority of whose functions remain currently unknown (342). In this regard, all of the advances made to the current day in the study of phenylpropanoid metabolism (and the effects of its modulation) resulted solely from the study of the proteins, enzymes, and genes involved. Thus, given the paucity in our knowledge of cell wall biochemistry (and of the proteins involved), we consider it inopportune not to systematically examine their roles in controlling native lignin macromolecular configuration. Indeed, this is a more likely and presumably more productive direction than the study of non-proteinaceous components, such as the effects of cellulose, hemicelluloses, etc.

In this regard, various proteins have been considered for their potential roles in establish­ing lignin macromolecular configuration in vivo. For example, using polyclonal antibodies raised against the (+)-pinoresinol-forming dirigent protein, our preliminary analyses in­dicated that their epitopes could be detected in the cell wall areas (e. g., S1 sublayers and cell corners) where lignification was initiated (284). This was considered as being due to recognition of the monolignol (radical) binding motif(s). This, in turn, led to our proposal that — in contrast to stereoselective coupling — there were arrays of dirigent (monolignol) radical binding sites in those subcellular regions, thereby providing the basis for forming a predetermined — albeit racemic — lignin structure (or structures) (29, 31, 284, 285).

Other studies have also implicated various other proteins (e. g., proline-rich proteins, PRP) as potential “lignin scaffolds” in the cell wall, based on co-localization ofPRP epitopes and lignins in developing cell walls of maize coleoptiles (343) and in secondary cell walls of soybean (Glycine max) differentiating protoxylem elements (344), albeit without any precise indication as to what this meant either biochemically or in terms of how they influ­ence lignin macromolecular configuration. Before investigating the involvement of any one of these possibilities, we considered it useful to begin to develop more robust approaches to: probe native lignin structure in vivo (type and frequency of interunit linkages); identify conditions for obtaining native lignin facsimiles through in vitro assays, as well as to identify the biochemical (structural motifs) in dirigent proteins that are required for monolignol (radical) substrate binding (work in progress).

Proponents of the random coupling/combinatorial biochemistry model — affording 1066 (per 100-mer) isomers (196) alone for monolignols 1-, 3-, and 5-derived structures — have given additional speculation as to why it is their consideration that no proteinaceous control over lignification is in effect. These include: (i) the presumed absence of optical activity in lignins; (ii) that for proteinaceous control, there would be a requirement for complementary chains of proteins in both d and l configurations (of their amino acids) for monolignol (radical) binding leading to racemic lignins; and (iii) that presumed lignin subunits, such as the ~1% or so of secoisolariciresinol components considered part of gymnosperm lignin, are formed nonenzymatically.

In this regard, in 1999, Ralph et al. (341) endeavored to demonstrate that the pres­ence of racemic lignin fragments eliminated the possibility of proteinaceous control over macromolecular lignin configuration. This was an unexpected interpretation, given that provisional reasoning had been provided beforehand (31) and later (52) to rationalize the hitherto well-known absence of lignin optical activity. Nevertheless, presumed 8-5′, 8-1′, 8­8′, and 8- O-4′ lignin fragments were isolated from pine sapwood, using the reductive DFRC method (341), which converts benzylic hydroxyl groups of various lignin/lignan-derived entities to their corresponding methylenic functionalities. Analyses of these products estab­lished that two (8-5′ and 8-1′-derived) reduction products were, as expected, racemic; on the other hand, the reduced derivatives obtained from the presumed 8-8′ pinoresinol (69) and the 8-O-4′ (71) substructures could not be resolved into specific enantiomeric forms. Interestingly, the 8-8′-linked product was not pinoresinol (69) per se, but was instead sec­oisolariciresinol (89) (tetraacetate): as indicated above, the latter 89 is currently considered by some researchers as being a minor (~1%) component of gymnosperm (spruce) lignin (345). [Indeed, the abundance of this presumed structure was estimated to be ~1.0 unit per 100 C9 units in spruce lignin, and 1.0-1.5 units per 100 C9 units in kraft and kraft pulp residual lignin based on quantitative 13C NMR and HSQC NMR analyses (345).]

Formation of secoisolariciresinol (89) from pinoresinol (69) (Figure 7.22A), however, has been extensively investigated in this laboratory, this resulting from the action of ei­ther enantiospecific pinoresinol-lariciresinol reductases (PLRs) or with PLRs that do not display strict enantiomeric preferences (346-348). Moreover, the X-ray crystal structure of PLR has been obtained (348), a catalytic mechanism proposed (348), and work has been completed to identify the changes in PLR substrate-binding pockets for differing enan- tiospecificities/preferences (Kim et al., manuscript in preparation). It is now quite well established that enantiospecific differences and/or slight enantiomeric preferences involve, as anticipated, changes in the nature of the substrate-binding pocket(s). That is, this does not require an interconversion of the d — and L-configurations of the amino acids present in the protein/enzyme structure, as proposed by Ralph and Brunow (349). In a somewhat analogous manner, the 7-O-4′ reduction of (±)-dehydrodiconiferyl alcohols (68a/b) by phenylcoumaran benzylic ether reductase (PCBER) results from binding either enantiomer in its large substrate-binding pocket (348), i. e., again without any necessity to have d and l forms of the amino acids in the enzymes involved.

Interestingly, Holmgren etal. (350) also attempted to demonstrate the reductive conver­sion of pinoresinol (69) into secoisolariciresinol (89) upon incubation of coniferyl alcohol (3) with horseradish peroxidase/H2O2 in presence of NADH, i. e., in the absence of any

(PLR) protein. As expected, no such reduction occurred in the absence of functional PLR; indeed, such experiments are generally carried out as controls in our enzyme assays. Thus, given that biochemical mechanisms for the reduction of both (+)- and ( — )-pinoresinols (69a and 69b) have been described (347), it may be more instructive to examine whether such processes are involved — even to a small amount — in either the lignification process, or whether they result from an infusion of heartwood-forming components, including small amounts of (±)-secoisolariciresinols (89).

Similar to the simulations carried out in vacuum (14), ab initio MD simulations of xy­lose degradation in water started when a protonated (3-D-xylose was positioned in a unit cell surrounded by 32 water molecules (43). During the course of our entire simulation (~5 ps), the initiation proton attached to the -OH groups on the sugar ring was ob­served to be transferred back to the surrounding water molecules. This transfer is rapid and occurs in less than 100 fs for all of the hydroxyl groups on the xylose ring. Once the proton was transferred to the neighboring water molecule, it is quickly transferred to other water molecules and away from the sugar molecule. This result shows that proto­nation is probably the rate-limiting step in sugar degradation under acidic media because our earlier simulations in vacuum demonstrate that protonated (3-D-xylose molecules de­compose rapidly. Figure 9.4 shows the proton transfer from xylose C2-OH to the sol­vent water molecules from our simulations. The simulations started with a protonated xylose molecule surrounded by 32 water molecules. After 34 fs, a neighboring water molecule forms a bond with the protonated hydroxyl group (C2-OH+-OH2) as shown in the figure. After 62 fs, the proton from the C2-OH transfers to a water molecule, forming an H3O+. After the proton was transferred from the xylose hydroxyl group to the water molecule, it was quickly transferred to other water molecules and away from the xylose molecule due to the strong hydrogen bonding interactions between the water molecules and the high proton mobility. It appears that protonation is a slow rate-limiting step.

In order to test the notion that protonation is the rate-limiting step, reaction barriers have been estimated using the hybrid density functional B3LYP (18). A single water molecule has a lower proton affinity (44), PA = 165 kcal mol-1, than xylose (Table 9.1 PA = 186.7-191.3 kcal mol-1), but the MD simulations appear to indicate that a proton will be transferred from xylose to a neighboring water cluster. CBS-QB3 calculations (18) show that for water clusters, the proton affinity increases with cluster size due to the increased stability of the hydronium ion. These calculations show that for a four-molecule cluster, the proton affinity has increased to 220.2 kcal mol-1. This and the MD simulations suggest that bulk water will have a higher proton affinity than xylose. The high proton affinity of bulk water relative to xylose must then add energy to the intrinsic energy barrier for the dehydration of xylose. This could explain the discrepancy between the energy barriers calculated in vacuum shown inTable9.1 (~16kcalmol-1 for the formation of furfural) and the experimentally observed barriers in solution, about 32 kcal mol-1. This was investigated further with static electronic structure calculations using B3LYP. Energy barriers for the initial step of xylose conversion to furfural were calculated in the presence of water clusters. With water clusters present, the barrier for xylose dehydration increased (18) to around 30 kcal mol-1, which is consistent with experimental values. This suggests that the experimentally measured barrier for xylose dehydration contains an energy contribution due to transferring the proton from the solvent to the substrate.

12.3.1 Rumen fungi

Rumen anaerobic fungi, which are related to chytrids (66), have been shown to produce highly active cellulase systems (67) and have been the source of recombinant enzymes with high specific activities that have attracted interest for a variety of biotechnological applications [e. g. (68, 69)]. As in cellulolytic bacteria, individual enzyme structures show multidomain organization (70). Furthermore, manypolysaccharidases from anaerobic fungi exhibit a proposed 40 amino acid docking domain that maybe present in one, two, or three copies (71). This domain is cysteine-rich and shows no sequence homology with bacterial dockerins, but has been reported in enzymes from a range of anaerobic fungi including Orpinomyces, Piromyces, and Neocallimastix frontalis (72). There is evidence in Piromyces equi for an interaction between this docking domain and a scaffolding protein of 97 kDa (73). Recent evidence in Neocallimastix frontalis, however, also suggests that removal of the docking domain influences the activity and temperature optimum of the adjacent catalytic domains (74). The nature of the docking domain interaction with the putative scaffolding protein remains to be established.

The close sequence relationships between catalytic domains from cellulolytic anaerobic fungi and bacteria suggest that horizontal gene transfer between these two groups has played an important role in the evolution of their cellulase systems (75).

Extracellular enzymes catalyze the initial rate-limiting step of decomposition and are the primary means by which microbes degrade complex biomass into smaller molecules that can be assimilated. Therefore, it is reasonable to assume that determining the amount and type of hydrolytic activities associated with biomass can be an indicator of the hydrolytic po­tential of a given microbial community. Defining biomass degrading communities through the application of traditional biochemistry tools combined with new proteomic technolo­gies are advancing our understanding of how microorganisms attack biomass as well as our understanding of natural enzyme diversity. Proteomic analysis can be used to identify and validate hydrolytic enzyme targets and profile protein expression patterns in complex communities (71). By using two-dimensional PAGE it is possible to fingerprint the secreted proteins of cellulose-degrading microorganisms and obtain sufficient sequence informa­tion for cloning. Surveying the proteome of natural microbial communities can lead to the discovery and analysis of more diverse hydrolytic enzymes that can break down cellu­lose, hemicellulose, and lignin. New classes of ligninases and hemicellulases will likely be identified, their mechanisms of action understood, and their performance refined to allow introduction of enzymatic pretreatment that will free cellulose microfibrils for enzymatic saccharification (breakdown to sugars).

The traditional biochemical approach to understanding microbial biomass utilization is through kinetic assays designed to quantitatively determine the presence or absence of hydrolytic enzymes. Hydrolytic enzymes can be detected using either natural substrates such as cellulose or xylan by measuring sugar release or by using synthetic substrates that contain easily detected chromophores such as p-nitrophenol or 4-methyl umbelliferone (69). The detection of 4-methyl umbelliferone can be used as a sensitive, quantitative assay for endoglucanse or other enzymes that cleave substrates linked to 4-methyl umbelliferone (72). However, how tightly the enzymes are bound to the substrate or are associated with the microorganism is a limitation of direct enzymatic assays. A range of soil enzyme assays was developed by Lynch and coworkers as alternatives to population measurements (46). These included assays for determining chitobiosidase, N-acetyl glucosaminidase, p-glucosidase, p-galactosidase, acid phosphatase, alkaline phosphatase, phosphodiesterase, aryl sulfatase, and urease activities from small soil samples. Soil enzyme activities, therefore, can index changes in the microbial functioning in soil, and there is ample evidence in the literature of the importance of glycosyl hydrolases, and proteases to the soil’s performance (69, 73-77).

Monitoring many proteins simultaneously in a complex system can be best accom­plished for many hundreds of protein species across a large number of samples using mod­ern technology for two-dimensional differential gel electrophoresis in combination with sophisticated statistical algorithms for data analysis. Two-dimensional gel electrophoresis is capable of resolving several hundreds to several thousands of proteins on a single gel (78). This method utilizes independent properties of proteins (i. e., isoelectric point and molecular mass) to resolve proteins present in a biological extract in two dimensions. Methods for two­dimensional differential gel electrophoresis have been greatly improved over the last several years to enable quantitative analysis of relative protein abundance among a set of samples. The first major improvement involves derivatization of proteins in samples with spectrally distinct, covalently coupled, charge — and mass-balanced fluorescent dyes (79). These highly fluorescent tags allow for extremely sensitive detection limits [i. e., 1 ng; (80)] and a broad linear response range [i. e., 3-4 orders of magnitude; (81)]. Derivatization of two different protein extracts with spectrally distinct fluorescent tags enables multiplexing of two un­known samples on gels, which eliminates any question about which protein in one sample is co-migrating with what spot on another gel. A second major improvement of the ex­perimental design for two-dimensional differential gel electrophoresis is the inclusion of an internal standard labeled with a third, spectroscopically resolvable, fluorescent dye (82). This enables normalization of the data from a series of gels, which permits statistically valid comparison of protein amounts across a series of gels. Because of this improved experimen­tal design, it is now possible to detect changes in absolute protein abundance on the order of 10% with 95% confidence.

The degree of recalcitrance of cell wall polysaccharides to depolymerization by enzymes, or other hydrolytic reagents, depends on the ability of the enzymes to access their substrates in the walls. This is manifest at two levels. The first relates to the surface area ofthe wall exposed to the hydrolytic agent: the greater the extent of comminution of the feedstock, the greater the surface area exposed as the cellular organization of the plant material is disrupted (99). The second level of constraint is manifest in the organization of the polysaccharides in the
cell walls and in particular to the extent that lignin deposition and its covalent cross-linking to other wall polymers limits enzymatic degradation as discussed in Sections 4.3 & 4.4.

Information is available from studies on forages, particularly grasses and legumes, for ruminant animals on the relative degradabilities (digestibilities) of the walls of different cell types by mixtures of fungal enzymes and by rumen fluid (5, 170-174). In general, non — lignified walls are highly degradable, whereas lignified walls are much less degradable. For example, many of the parenchyma cells in stems and leaves, including mesophyll cells, have thin, non-lignified, primary walls that are highly degradable. Even moderate amounts of diferulate cross-linking between GAXs in non-lignified grass walls apparently do not im­pede wall degradation (175). Thus, an approximate index of the suitability of a plant as a feedstock would be the relative proportion of cells in the vegetative tissue of the mature plant that have lignified walls. The relative proportions of different cell types in a range of C3 and C4 grass species have been reviewed by Buxton and Redfearn (172). For exam­ple, in leaf blades of switchgrass (Panicum virgatum), a warm-season C4 species, 28.9% of the cross-section area is occupied by bundle sheath cells that have moderately thick and weakly lignified walls, 35.7% by mesophyll cells with thin, non-lignified walls, and 2.9% by sclerenchyma fibers with thick, lignified walls (176). Furthermore, there maybe important genetic variations in the relative proportions of the different cell types among various clones of the same species (177) that may explain variations among the clones in their total cell wall degradabilities by polysaccharide degrading enzymes, e. g., in smooth bromegrass (Bromus inermis), a cool-season C3 species (177). The degradability of the cell walls of internodes from 100 lines of the temperate grass (Phalaris aquatica) using fungal enzymes was found to be markedly dependent on the line ranging from 22 to 38% (178).In investigations of the genetic background to digestibility of smooth bromegrass genotypes using fungal enzymes (179, 180), it was found that in high digestibility selections, variation in digestibility was due largely to variation in lignin concentration, whereas in low digestibility selections the influence of lignin concentration was dramatically reduced and esterified FA concentration showed a strong negative relationship with lignin concentration and appeared to be more associated with cell wall digestibility than was lignin.

During the development of forage plant organs, such as flowering stems, the overall degradability of the walls falls because some of the cell types develop lignified secondary walls (181, 182). Although this occurs in most sclerenchyma fibers and xylem tracheary elements, it occurs in only some of the parenchyma cells. Environmental conditions during growth such as temperature extremes, water deficit, nutrient stress, and shade also impact on cell wall composition and digestibility (183).

Interestingly, there are also differences between grasses and legumes in the degradabilities of their lignified walls. For example in legume stems, the lignified walls of the sclerenchyma fibers in the xylem are not degraded by rumen fluid, but in grass stems, the lignified walls of sclerenchyma fibers and parenchyma cells ingrasses show significant degradability (5). However, this degradation affects only the secondary walls and not the underlying more heavily lignified primary wall and middle lamella. The cleavage by feruloyl esterases (184) of ester linkages between heteroxylans and ferulic acid linked to lignin, which occur in walls of grasses but not legumes, maybe responsible for these differences.

Studies using rumen fluid and mixtures offungal enzymes have also shown that suberized layers and cuticles on cell walls in the periderm ofwoody species and epidermal cells of grasses are also barriers to the access of polysaccharide hydrolases to their substrates in the wall and thus hinder their depolymerization (5, 170).

It was exactly 101 years ago, while working on fermentation of sugars by yeast, that Harden and Young (408) first reported the chemistry and metabolic roles of sugar-phosphates. Later, different types of sugar kinases from yeast, muscle, and plant sources were identified that were able to convert sugar (monosaccharide) and ATP to the phosphorylated-sugar esters. The seminal work by Cori et al. in 1939 (409) proved, for the first time, the role of sugar-phosphates in the synthesis of polysaccharides (i. e., glycogen). Since the discovery of UDP-glucose in yeast (410), which was followed by the isolation of other NDP-sugars in yeast, plants, bacteria, algae, and humans, it has become apparent that NDP-sugars are the prime sugar-substrates used in the biosynthesis of glycans. The biosynthesis of activated- sugars is achieved in three general ways.

2.5.1 Sugar kinase — pyrophosphorylase pathway to synthesize NDP-sugars

Some sugars are first converted directly, or in a series of enzymatic steps, to phospho-1- sugars in the presence of ATP.

Sugar + ATP ^ Sugar-1-P + ADP

Following the phosphorylation of the anomeric center, enzymes known as pyrophos — phorylases transfer a nucleotide-monophosphate from NTP to the sugar-1-P to form the NDP-sugar. The synthesis of UDP-Glc and GDP-Man are examples of this type of synthesis.