Category Archives: Biomass Recalcitrance

Molecular approaches for defining biomass-degrading communities

The challenge of studying the ecology of biomass-degrading communities has always been the ability to accurately determine a representative picture of the true diversity of the mi­crobial consortia, and the biochemical processes. Advances in molecular biology have led to development of culture-independent methods for describing microbial communities based on analysis of DNA extracted directly from natural populations thereby circumventing the need to isolate and culture bacteria for phylogenetic analysis. The recent surge of research in molecular microbial ecology has provided evidence for the existence of many novel types of microorganisms in the environment, in numbers, and varieties far greater than culti­vated in the laboratory, which probably comprise less than 1% of all microorganisms (33). Additional corroboration comes from estimates of DNA complexity and the discovery of many unique bacterial 16S rRNA gene sequences from numerous environmental sources (2, 35, 83). One approach to classifying complex communities is to use “marker” genes as phylogenetic anchors for identification of source microorganisms. Ribosomal RNAs are highly conserved and are commonly used to determine phylogenic relationships between organisms. Other marker genes can be used including recA and rpoB genes, however, the largest available database is for 16S rRNA. Another approach is to use gene-centric analysis of large datasets that focus on the identification and taxonomic characterization of genes important to the overall community function.

DNA-based fingerprinting techniques developed to characterize and compare whole genomes of organisms include amplified fragment length polymorphism (84), terminal restriction fragment length polymorphism (68), denaturing gradient gel electrophoresis (70), amplified rRNA gene restriction analysis (85), restriction landmark genome scan­ning (86), and automated ribosomal intergenic spacer analysis (87). Dominate members of biomass-degrading communities can be determined using methods such as terminal restriction fragment length polymorphism (T-RFLP) and denaturing gradient gel elec­trophoresis (DGGE).

Cell Wall Polysaccharide Synthesis

Debra Mohnen, Maor Bar-Peled and Chris Somerville

5.1 Introduction

One of the many challenges associated with trying to understand the synthesis, structure, and function of higher plant cell walls is the obvious chemical diversity between tissue types within a plant and between plant species. For instance, an analysis of the cell wall sugar composition of different tissues in Arabidopsis revealed that each of the tissue types analyzed had a strikingly different composition (1). In spite of this diversity, there is sub­stantial evidence in support of the hypothesis that most cells of all higher plants have the same six major types of polysaccharides: cellulose, xyloglucan, xylan, homogalacturonan, rhamnogalacturonan I, and rhamnogalacturonan II (2). According to this idea, variation in wall composition arises from variation in the amounts of the various polysaccharides and in variations of the structures of the various polysaccharides. Thus, in order to understand how cell walls are synthesized, much of the task may be condensed into understanding how these classes of polysaccharides are synthesized and deposited in walls. Additionally, several other polysaccharides are found in some species. These include xylogalacturonan, mannan, and mixed-linkage glucans. Lignin associated with secondary cell walls represents the other major component. Although cell wall proteoglycans are a quantitatively minor component, they may be relevant for understanding certain aspects of assembly. Similarly, callose plays an important role in synthesis of de novo cell walls or in specialized cell walls such as pollen tubes but is not usually a quantitatively significant component of most types of cell walls.

Plant walls are divided into two basic types, the primary and secondary wall (Figure 5.1). The primary wall is the first wall laid down in dividing and growing plant cells and it is the terminal wall in many cells in the soft parts of the plant, including the palisade cells in leaves and parenchyma cells present throughout the plant. The primary wall contains 80-90% polysaccharide and 10-20% protein. Cellulose, hemicellulose, and pectin are the main polysaccharide components in the primary wall. The most abundant hemicellulose in most primary walls is xyloglucan. However, in grasses and other commelinoid monocots glucuronoarabinoxylan is the major hemicellulose and during cell expansion in grasses, (3-1,3;p1,4-mixed linkage glucans are prevalent in the primary wall. Secondary walls are produced by specialized cells that serve a structural role such as fibers and xylem cells in vascular bundles. Secondary walls generally have less pectin, contain more cellulose and more of the hemicellulose (3-1,4-xylan (often as glucuronoxylan), and are often rigidified

plasma membrane (cellulose) and in the Golgi (pectin and hemicellulose) by the action of glycosyltransferases that use nucleotide-sugar substrates. (b) Some cells (e. g., xylem and fiber cells) form secondary walls internal to the primary wall. Secondary walls have increased amounts of cel­lulose and hemicellulose, less pectin, and are often rigidified with lignin. Secondary walls in wood tissue consist of three layers (S1, S2, and S3) that differ in cellulose microfibril orientation and chemical compo­sition. (c) Secondary wall synthesis includes formation of cellulose microfibrils at the plasma membrane, hemicellulose within the Golgi followed by deposition in the wall, and lignin polymerization from mono — lignols within the wall matrix. (Figure with permission by Malcolm O’Neill, CCRC, University of Georgia)

by lignification. The synthesis of both the primary and secondary walls utilizes nucleotide — sugar substrates that are synthesized in the cytosol or in the lumen of the endoplasmic reticulum and the Golgi.

In the following overview, we have summarized the current status of knowledge about how the major classes of cell wall polymers are synthesized, modified, secreted, and assembled.

Because of the importance of Arabidopsis as an experimental model, much of the recent progress has been obtained using Arabidopsis. However, information from other species is included where it is available. The following description of cellulose synthesis is an update of a recent review (3).

Direct production of NDP-sugars

Some activated sugars such as CMP-KDO are exceptional since the free sugar, synthesized via intermediary metabolism products, is condensed directly with CTP without a prior phosphorylation of C1.

Ara-5-P + PEP ^ KDO-8-P ^ KDO ^ KDO + CTP ^ CMP-KDO

2.5.2 NDP-sugar Interconversion Pathway

The interconversion pathway ofnucleotide-sugars is a major pathway where specific enzymes convert preexisting NDP-sugars into different stereospecific NDP-sugars (Figure 5.4).


Figure 5.4 The metabolism of NDP-sugars in plants. Glucose, fructose, inositol, and sucrose are ma­jor sources of carbon that can be metabolized into NDP-sugars. A salvage pathway is defined as the ability of plant enzymes to recycle sugars that are released from glycoconjugates. The enzyme activity de­picted as numbers (italic) are UDP-glucose pyrophosphorylase (1), ADP-glucose pyrophosphorylase (2), sucrose synthase (SuSy, 3), UDP-glucose dehydrogenase (UGD, 4), UDP-apiose/xylose synthase (5), UDP — glucuronic acid decarboxylase (UXS, 6), UDP-glucuronic acid 4-epimerase (UGlcAE, 7), UDP-galactose 4-epimerase (UGE, 8), UDP-glucose 4,6 dehydratase (9), UDP-4-keto-6deoxyglucose-3′,5′-epimerase and 4′,6′-keto-reductase (NRSER, 10). Reaction 9 and reaction 10 are also carried out by a single polypep­tide (URS/MUM/RHM, 9.10), inositol oxygenase (MIOX, 11), glucuronic acid-1-P kinase (12), UDP — sugar pyrophosphorylase (Sloppy, 13), UDP 5′-diphospho-6-sulfoquinovose synthase (14), UDP-xylose epimerase (UXE, 15), UDP-arabinomutase (UAM, RGP, 38), L-arabinose-1-P kinase (Ara1, AraK, 16), D-galactose-1-P kinase (GalK, 17), D-galacturonic acid-1-P kinase (GalAk, 18), L-rhamnose-1-P kinase (RhaK, 19), UDP-rhamnose pyrophosphorylase (20), AMP sugar-1-P transferase or ADP-glucose phospho — rylase (21), glucoseamine-6-P acetyltransferase (GNA1,22), UDP-N-acetylglucoseamine pyrophosphory­lase (23), mannose-1-P pyrophosphorylase (24), GDP-Man 3′,5′ epimerase (GME, 25), GDP-L-galactose phosphorylase (39), L-galactose-1-P phosphatase (40), GDP-mannose 4,6 dehydratase (GMD, 26), GDP- 4-keto-6-deoxymannose-3,5-epimerase-4-reductase (GER1, FX, 27), fucose-1-P kinase (29), GDP-fucose pyrophosphorylase (30), mannose-6-P kinase (31), fructose-6-P kinase (32), glucose-6-P kinase (hexoki — nase, 33), phosphoglucose isomerase (PGI, 41), phosphomannose isomerase (PMI, 42), phosphomannose mutase (PMM, 43), phosphoglucose mutase (PGM, 44), KDO-8-P synthase (34), KDO-8-P phosphatase (35), CMP-KDP synthase (36).

Some of the types of modifications involved in the interconversion pathway are the iso­merization of l — and D-sugars, 4-epimerization, specific C-6 oxidation, decarboxylation, and the formation of NDP-deoxy sugar derivatives. Examples of the interconversion pathway are the conversion of UDP-GlcA to UDP-GalA and the conversion of UDP-GlcA to UDP-Xyl. The synthesis of each nucleotide-sugar is described in the following sections.

Molecular modeling

Molecular dynamics and molecular mechanics calculations have been used extensively to examine cellulose, often giving unexpected results. Because of inadequate information con­tained in fiber diffraction patterns noted above, all models for cellulose have been developed to some extent through modeling wherein constraints are imposed on the solution to com­plement inadequate data sets. The constraints most often used are dimensions of the unit cell together with assumptions regarding symmetry that allow the development of a recip­rocal space. These constraints are not unlike boundary conditions necessary for the solution of differential equations in the contexts of mathematical formulations of descriptions of specific physical phenomena.

The molecular dynamics (MD) simulations carried out recently by Matthews and cowork­ers represent the least constrained analyses (14). The only constraint used was an initial condition corresponding to the published structure of the Ip form. More recently, similar simulations were carried out with the structure of the Ia form as the initial condition and conclusions were essentially the same as those derived from simulation with the structure of the Ip form as the initial condition. During the course of the simulations, a number of structural fluctuations and changes occurred. Over the length of the simulations, aver­age unit cell dimensions shifted away from those reported on the basis of diffractometric measurements; here we use the term unit cell only to provide a basis for comparison with the published structures of the same forms. These dimensions varied with position in the aggregate relative to the surfaces and the chain termini. The results, averaged over these cellobiose units, are summarized in Figure 6.4 and compared with the crystallographic unit cell of the Ip form.

In the simulation, the aggregate was observed to undergo an expansion in which the value of lattice constant a increased from 7.784 to 8.470 A, while the b value decreased slightly from 8.201 to 8.112 A. The c value expanded significantly, from 10.380 to 10.512 A. In addition, the у angle decreased from 96.5° to almost orthogonal, у ~ 90°. The unit cell a-axis (corresponding to the distance between hydrogen bonded sheets) in this simulation differs considerably from that proposed in the crystallographic models. The terminology regarding lattice structures and unit cells is used here primarily to allow comparisons with the structures proposed on the basis of diffractometric measurements.

Another extremely significant change in the structure of the aggregate that occurred during the simulations is that many of the C6 primary alcohol groups underwent rotational transitions away from the conformations reported for the diffraction-based structure. This exocyclic group has three low-energy staggered conformations, which are named TG, GG, and GT, with the first letter in these labels specifying the position of the O6 atom as either trans or gauche with respect to the O5 atom, and the second letter specifies its relationship to the C4 atom (see Figure 6.5) (20). In both the Ia and Ip diffraction-based structures, all of these exocyclic groups are in the TG conformation. This is also one of the consequences of the constraints of symmetry.


Figure 6.4 Left: the cellulose Ip crystal unit cell determined by fiber diffraction; right: the trajectory — averaged unit cell for the simulation of the diagonal crystal. Hydrogen atoms are omitted for clarity, and positions obtained by symmetry operations are transparent. (Reproduced in color as Plate 7.)

In the Ip diffraction-based structure, all these exocyclic groups are in the TG confor­mation. In this conformation, the exocyclic hydroxyl groups can hydrogen-bond along the chain or to adjacent chains in the same layer, but no hydrogen bonds between layers can occur. For those layers of the aggregate made up of the origin chains, there was little change in structure in the MD simulation from that of the diffraction structure, and the hydro­gen bonding pattern remained the same. This result is remarkably similar to the reported


Figure 6.5 Nomenclature of primary alcohol conformation. The dihedral angle measured by C4-C5-C6- O6 is shown. (Reproduced in color as Plate 8.)

image082 image083 image084

image085Hydrogen bonding in

Figure 6.6 Single frames from the center chain layers illustrating three different hydrogen bond patterns. Left: the pattern is similar to the predominant pattern from the crystal structure, but the rotation to GG makes the HO2-O6 hydrogen bond across the glycosidic linkage impossible;center: the hydrogen bond pattern is very similar to the less occupied pattern from the crystal structure;right: hydrogen bonds from HO6 in a center chain to O2 in an origin layer chain, which is not shown for clarity. (Reproduced in color as Plate 9.) crystallographic hydrogen bond network in origin chains, where the O2 hydroxyl group was refined to just one of the two possible hydrogen bond positions (8). However, in the MD simulations, in every other layer in the interior of the aggregate, made up of the center chains in the diffraction-based structure, this primary alcohol group rotated from the starting TG conformation to the GG position. In this GG conformation, three rapidly interchanging hydrogen bond patterns were possible, as shown in Figure 6.6.

One of these patterns allowed hydrogen bonding between layers, which was not possible when all the hydroxymethyl groups were in the TG conformation. On the surfaces, where the anhydroglucose monomers were in direct contact with water, the hydrogen bonds to the freely diffusing water molecules helped to introduce considerable disorder into these primary alcohol conformations and promoted frequent transitions, but the interior portions of the aggregate developed two distinct patterns of hydroxymethyl conformations between the center and origin layers. Primary alcohol groups in surface chains alternate between facing toward the interior and facing the solvent, and the conformation of these surface groups corresponds to the local environment. Several NMR studies have determined that the conformations of surface cellulose chains are different from the interior, and as in the new simulation, contain both GG and GT rotamers (20-23). The presence of two rotamers is also consistent with the Raman spectra of Ia and Ip to be discussed further in a following section. In the spectra of both Ia and Ip the scissors vibration of the methylene group on C6 results in two bands in the region above 1450 cm-1; the methylene scissors vibration is


Figure 6.7 Views of the central portion of one center chain and one origin chain from the middle of the diagonal crystal, illustrating the inter-plane hydrogen bonds which can occur after the center chain primary alcohol groups rotate to the GG conformation. Hydrogen bonds between layers are indicated with dashed lines. (Reproduced in color as Plate 10.)

the only one that gives rise to a band above 1450 cm-1. Thus, the occurrence of two bands is consistent with the presence of two rotamers.

In the GG conformation, the primary alcohol groups are essentially perpendicular to the average planes of the anhydroglucose rings and as a result are pointing up and down toward the origin chains of the layers above and below. In this conformation, the exocyclic groups can make good O6-O2 hydrogen bonds between layers. Since under normal conditions cellulose exhibits no tendency for layers to slip relative to one another, the existence of such stabilizing hydrogen bonds may not seem so implausible. However, in this conformation steric clashes between the center chain primary alcohol groups and the origin layers above and below force the center chains to tilt significantly with respect to the plane of their own layer. Such a tilt was also found by Heiner and Teleman (24).

Probably the most significant change that occurred during the simulations was that the aggregate quickly developed a small right-hand twist during the heating and equilibration interval and the twist remained relatively stable throughout the rest of the simulation. Figure 6.7 illustrates this twist, with the middle hydrogen-bonded sheet shown in detail. In this figure, the average twist angle for each successive glycosidic linkage is shown. These angles are defined as the dihedral angle for the four C1 carbon atoms illustrated as joined by the dark lines in the figure. Although this angle varies considerably near the non-reducing end, apparently because of edge effects, the twist in the middle of the chain is fairly constant at around 1.4—1.7° per linkage, with an overall twist for this short oligosaccharide segment of almost 9.9° calculated from the first and last rows (which includes considerable irregularity due to the highly frayed structure of the non-reducing ends).

Imposition of the constraint of the symmetry of space group P21 confines cellulose chains to an exact twofold helix, and this constraint can be satisfied by many combinations of torsion angles across the glycosidic linkage (reported either as = H1-C1-O-C4/ and ф H = C1-O-

C4/-H4/ or as фо = O5-C1-O-C4/ and фC = C1-O-C4/-C5/). However, the line connecting twofold helical structures for cellobiose in ф, ф space does not coincide with a free energy minimum (25, 26). Cellulose oligomers in solution are extended, but do not have a flat ribbon structure (27, 28). The preference of cellulose chains to adopt conformations away from a twofold helix is frustrated in a crystalline state by packing and hydrogen bonding requirements. Equilibrium organization of the aggregate has each of the individual interior chains departing slightly from the flat starting structure, on average forming a right-handed helix. The helix of each chain corresponds to the overall twist of fiber in a manner similar to the twist seen in protein p-sheets (29, 30).

In addition to the findings of the simulation studies, it is helpful to consider the source of the helical patterns at the level of the individual monomers in cellulose; it is now accepted that anhydrocellobiose is the repeat unit of structure in cellulose as it implicitly defines the glucosidic linkage as well. The results of the earliest conformational energy mappings available (31,32) show that two energy minima associated with variations in dihedral angles of glycosidic linkage correspond to relatively small left — and right-handed departures from glycosidic linkage conformations consistent with twofold helical symmetry. More recent all-atom conformational energy maps for cellobiose exhibit the same qualitative topology (25). Local minima also represent values of dihedral angles very similar to those reported for cellobiose and methyl p-cellobioside on the basis of crystallographic analyses (9, 10). The relationship between different conformations is represented in Figure 6.8, which was adapted (6) from a diagram first presented by Reese and Skerrett (31).

Figure 6.8 is a ф /ф map presenting different categories of information concerning con­formation of the anhydrocellobiose unit as a function of the values of two dihedral angles


Figure 6.8 ф/ф map adapted from Ref. (9). (———— ) Loci of structures with constant anhydroglucose repeat

periods as noted in Angstroms. (…) Loci of structures of constant intramolecular bond O5-O31 distances.

(—- ) Contours of potential energy minima based on non-bonded interactions in cellobiose. J — cellobiose;

W — p-methylcellobioside;n = 2, the twofold helix line;n = 3, the threefold helix line;(R) right handed; (L) left handed. The Meyer and Misch structure is at ф = 180, ф = 0.

about bonds in the glycosidic linkage. ф is defined as the dihedral angle about the bond between C4 and the glycosidic linkage oxygen and ф as the dihedral angle about the bond between C1 and the glycosidic linkage oxygen. The parallel lines indicated by n = 3(L), 2, and 3(R) represent values of dihedral angles consistent with a left-handed threefold helical conformation, a twofold helical conformation, and a right-handed threefold helical confor­mation, respectively. A twofold helical conformation inherently does not have a handedness to it. Dashed contours represent conformations that have indicated repeat period per anhy — droglucose unit; the innermost represents a period of 5.25 A corresponding to 10.5 A per anhydrocellobiose unit. Two dotted lines indicate conformations corresponding to values of 2.5 and 2.8 A for the distance between the two oxygen atoms anchoring the intramolecular hydrogen bond between the C3 hydroxyl group of one anhydroglucose unit and the ring oxygen of the adjacent unit; values bracket the range wherein hydrogen bonds are regarded as strong.

The two domains defined by solid lines on either side of the twofold helix line (n = 2) represent the potential energy minima calculated by Reese and Skerrett for different confor­mations of cellobiose (31). Finally, the points marked by J and W represent the structure of cellobiose determined by Chu and Jeffrey (9) and that of methyl p cellobioside determined by Ham and Williams (10). The key point to be kept in mind with this diagram is that structures along the twofold helix line and with a repeat period of 10.3 A per anhydrocel­lobiose unit possess an unacceptable degree of overlap between the van der Waals radii of the hydrogen atoms on either side of the glycosidic linkage.

Figure 6.8 shows that the structure of glycosidic linkage in cellulose is not likely to coin­cide with the line representing twofold helical structures. Rather, it is likely to be on either side of the twofold helix line as are the structures of cellobiose determined by Chu and Jeffrey, designated (J) and of p-methylcellobioside determined by Ham and Williams des­ignated (W). However, because of the repeat distance per anhydroglucose unit, one would expect the glycosidic linkages in cellulose to be much closer to the twofold line than are the two dimeric structures. On the other hand, the SS 13C NMR spectra show a splitting of the resonances at C1 and C4. Thus, it seems plausible that values of the glycosidic di­hedrals in the cellulose chain might alternate between a small left-handed departure and a slightly larger right-handed departure from the twofold helix line. The net effect would be a slow, long-period, right-handed helical structure. Such an alternating pattern was ob­served in the stable equilibrium structure at the conclusion of the MD simulation (14). This pattern demonstrates that the long-period helical twist is a consequence of impor­tant characteristics of glycosidic linkages in cellulose rather than an artifact of a complex simulation.

From a broader perspective, a very important result of molecular modeling validates the approach represented by the theoretical model used. The finding that the cellulose aggregate is stable reflects that cellulose is insoluble in water beyond the octamer. The stability of the aggregate at equilibrium is not the result of any constraints or boundary conditions imposed on the solution of the equilibrium structure, but rather evidences that the molecular modeling has captured some essential distinctive properties of cellulose. Indeed, the measure of its true approximation of the nature of cellulose is that the insolubility is predicted for chains that are 12 anhydroglucose units in length. Furthermore, the results of the modeling are consistent with microscopic observations of long-period helical structures, and they explain the structure oftheHCH scissors vibration bands in the Raman spectra of Ia and Ip.

The results are also consistent with the effects of small-diameter fibrillation to be discussed further below.

It is important at this point to return to Figure 6.3, panel B and consider its implications. The application of a long period of 1200 nm to all of the fibrils is intended to allow comparison of the effects of lateral dimensions on the twisting. A period of 1200 nm is evidently too short a period for fibrils of Valonia and Halocynthia, considering that such periods are rarely observed in electron microscopy. On the basis of the observation of a long period of 1200 nm for Micrasterias, we anticipate that the period for a 20 by 20 nm fibril is likely to be 2500 nm or more. That dimension is 2.5 ^m and would be well beyond the field size in a high-magnification electron micrograph.

Another important point most obvious for the 20 by 20 nm fibril in panel B is that the center chain remains linear though it will be twisted by 90°. The other chains, however, develop some curvature so that the corner chains are obviously quite curved, and curvature increases with distance from the center. Thus, the resistance of the aggregate to inherent tendencies of the cellulose molecules to acquire a helical orientation increases with lateral dimension. The possibility of shear stresses developing within a fibril increases with lateral dimension also. This may well be why load-bearing structures of higher plants have fibrils with such small lateral dimensions. Because of small diameters, they are not likely to de­velop significant internal shear stresses. Because their association with neighboring fibrils is mediated by water, they can move parallel to each other when under load. Panel B in Figure 6.3 clearly suggests that as the lateral dimensions are reduced, the long-period helical twist can be more easily accommodated. The implications of the long-period twist for the subject of cell wall deconstruction will be discussed further in a following section.

CCR: tyramine derivatives are not chemical signatures of CCR downregulation/mutation, and abnormal lignins are not formed

CCR has recently been both downregulated and mutated in tobacco (Nicotiana tabacum) (232) and Arabidopsis (233), respectively. The phenotypes obtained for each line were, at first glance, quite similar: both were considerably dwarfed (131, 232) (Figures 7.13C and 7.13D). Such effects, as already noted earlier, are not always a typical consequence of downregulating lignin amounts and/or compositions. Nevertheless, by comprehensively examining the lignin contents and compositions, stem diameters/lengths and anatomy, it was considered that the Arabidopsis irx4 mutant was delayed in overall development (Figure 7.12F), particularly as regards lignification (131). Specifically, the deposition of S-lignin components in the irx4 mutant line initially lagged behind that of wild type. At maturation, the lignin S/G compositions (ratios) were similar though for both lines, with the overall lignin amounts only being 10-15% lower in the mutant (131). On the other hand, the lignin levels reported for the CCR downregulated tobacco stems were circa 50% of wild-type levels, with the xylem cells collapsed (Figure 7.12E), indicative of a much compromised vascular apparatus (77, 232). At the anatomical level, it was of interest that both the Arabidopsis and tobacco stem cross-sections apparently differed substantially in their overall effects on vascular integrity; in neither case were “perfectly viable” plants obtained, given the defects/pleiotropic effects noted.

Detailed analysis of lignin deposition in the Arabidopsis irx4 line had thus established that initially the mutant had a delayed but coherent (normal) program of lignification (131, 132). By contrast, a previous study by other researchers (233) had reported that this plant line contained an “abnormal lignin,” derived from “alternative” phenolics and whose lignin levels were reduced by circa 50%. The “abnormal” nature of the lignin was apparently determined by NMR spectroscopic analyses, although no data were actually provided. These reports have since been revised with more in-depth analyses from our laboratory (131, 132). First, the full extent of the lignification response in the irx4 mutant line was only actually determined by examining the lignin contents and compositions up to maturation (>8 weeks), with only a small reduction (~10—15%) in deposition levels noted. Second, NMR spectroscopic analyses indicated that typical G-S lignins were being formed and not “abnormal” lignins as had been reported. Furthermore, we proposed that the dwarfing phenomenon and reduced lignin levels may be due to CoASH levels being reduced (due to build up of hydroxycinnamoyl CoA derivatives) in the irx4 line, with some sort of, for example, feedback inhibition occurring (131); however, this remains to be fully established in future studies.

Perhaps most importantly, plotting lignin contents versus (thioacidolysis) releasable monomers (representing a subset of cleavable 8- O-4′-linkages) again indicated that both irx4 and wild-type lines had a monomer invariant frequency (Figure 7.14C) of said linkages at all stages of lignin deposition, plant growth, and development. These data are thus again discussed later in terms of further indications of a non-random assembly process.

The original, albeit now incorrect, report by Jones etal. (233) was apparently based on ex­pectations that had been raised from the study of the CCR downregulation in tobacco (173, 174,226). The latter papers described bewildering findings as regards lignin macromolecular assembly and composition. Specifically, it was reported that when CCR was downregulated in tobacco, the plants compensated for reduction in monolignol (lignin precursor) supply by incorporating other “alternative” phenolics into the lignifying cell walls. Initially, the “non-traditional” phenolics reported as incorporated into lignin through CCR downregu­lation included ferulic (11) and sinapic (13) acids, as well as a variety of other phenolics, including acetosyringone (61) (173, 174, 226). It was also reported that feruloyl tyramine (60) was “heavily incorporated” into the lignin as a consequence of CCR downregulation (173, 174, 226), and that this moiety represented a chemical “signature” for CCR downreg­ulation (174). However, no quantification of feruloyl tyramine (60) levels of any sort was carried out.

As regards the reported increases in amounts of hydroxycinnamic acids/benzaldehydes, etc. in the lignins of the highly dwarfed, lignin reduced (~50%), CCR downregulated tobacco lines, relative to the wild type, there was apparently no significant difference in total amounts measured quantitatively (77). For example, the reported levels of such moieties ranged from 0.04 to 0.07% of stem cell wall residues in both wild type and downregulated lines (77). Such minute levels would not constitute compelling evidence for “abnormal” lignin and “aberrant” lignins being formed from “non-traditional monomers.” Indeed, since the lignin contents of CCR downregulated and wild type were ~11 and ~22% of the plant stem dry weight, respectively (77), the amounts of the aldehydes/acids (0.04-0.07%), etc. were minuscule relative to actual lignin levels.

Other studies by Ralph and colleagues (173, 174), which reported that there were ele­vated levels of feruloyl tyramine (60) covalently attached to tobacco lignins, and that these were chemical signatures of CCR downregulation/mutation, could not be independently verified either. Furthermore, the lack of any feruloyl (p-hydroxycinnamoyl) tyramine (60) resonances in the Arabidopsis CCR-irx4 lignin-derived isolates (77, 132) eliminated these as being generic chemical “signatures” for CCR mutation/downregulation as had been pro­posed (174). By contrast, careful isolation (by dissection) ofthe vascular (lignified) apparatus and subsequent detailed NMR spectroscopic analyses of the resulting lignin isolate(s) gave no evidence for the presence of feruloyl tyramine (60) moieties in the lignins from the wild — type lines of tobacco (177). Nor were feruloyl tyramine moieties (60) observed in the lignin preparations by Bernard-Vailhe et al. (234). Additionally, in contrast to reports (173, 174) of feruloyl tyramine (60) moieties being incorporated into lignin, these researchers had never demonstrated that the feruloyl tyramine-like resonances in the lignin-enriched isolate were covalently linked to lignin, or came from the same cell wall types harboring lignins. Nor was evidence provided that the overall amounts of feruloyl tyramine (60) moieties had increased in the isolates from the CCR downregulated lines, relative to the original levels in the wild-type line.

Force fields

From the discussion above it should be obvious that the accuracy of any molecular mechanics method is entirely dependent on the quality of the parameterization used. In theory one could parameterize specifically for each simulation that is to be run but this would be extremely time-consuming and would also make the parameters non-transferable. Instead, in order to preserve the transferability of the force field one attempts to minimize the number of parameters by reducing the system to a set of building blocks. For example, in proteins this is typically the set of individual amino acid residues.

There are a large number of different force fields that have been developed over the years and all have their advantages and disadvantages. For example, the CHARMM force fields include charmm19 (15), charmm22 (27), charmm27 (28), and AMBER force fields include FF94 (6), FF96 (29), FF99 (30), FF99SB (31), FF03 (32, 33). A comprehensive review of the various force fields is beyond the scope of this chapter. For further information the reader is encouraged to read the various papers referring to each force field. In line with the topic of this book, which is the study of cellulose, we will concentrate our discussion on the force fields available for carbohydrates.

A — D-galactosidase

a-D-galactosidase activity is required for hydrolysis of softwood mannans, specifically galac — tomannans and galactoglucomannans (71). The enzyme acts on the a — galactosyl side groups attached to the O-6 position of the backbone mannose units (65). Little work has been collected regarding this enzyme, though its importance in softwood pulping has been considered.

10.4.3 Acetyl xylan esterase

Acetyl groups occur on several hemicelluloses although the primary examples are xylan and galactoglucomannans. Cereal and hardwood xylans have much higher levels of acetylation than softwood xylans. Softwood acetylation occurs principally in the galactoglucomannans. The most likely reason for acetylation is to keep the hemicelluloses soluble and hydrated. Deacetylation ofxylan and glucogalactomannan results in significantly decreased solubility of the polymer. Acetyl groups also cause problems for microorganisms when they are released from the main chain, resulting in decreased pH. The release of acetate is inhibitory to many microbes and is a considerable problem in the conversion of biomass to fermentation products (72-75).

Acetyl groups are released from hemicelluloses both from high degree of polymerization (i. e., native) substrates and from acetylated oligomers resulting from depolymerization. Acetyl xylan esterases (AXEs) may exhibit a preference for one or the other form, or may act on both types (76). Synergy studies between AXE, xylanases, and other hemicellulase enzymes have repeatedly demonstrated that the most effect digestion occurs with the appro­priate ratios of all enzymes acting simultaneously (70, 76-78). Debranching in the absence of depolymerization results in insoluble long-chain hemicelluloses that can be more diffi­cult for the depolymerases to access (78). Depolymerization without debranching maybe limited by depolymerase access to the polymer main chain. Most AXEs are low molecular weight and may or may not have a carbohydrate-binding module.

Pretreatments for Enhanced Digestibility of Feedstocks

David K. Johnson, and Richard T. Elander

This chapter reviews how pretreatment is used to decrease the recalcitrance of biomass, and make the cellulose in the feedstock more susceptible to digestion by cellulase enzymes. Pretreatments have been designed to operate under very different chemistries, at widely varying temperatures, and for very different reaction times. With some pretreatments there is almost no change in the composition of the feedstock whereas in others the composition is changed considerably by dissolving the hemicelluloses, the lignin, or both. Pretreatment can also cause changes in the physicochemical structure of the biomass with changes in cellulose crystallinity, molecular weight, and accessibility, plus changes in biomass porosity and particle size. This chapter will focus on more recent developments in pretreatment and describe how these pretreatment processes attempt to overcome the natural recalcitrance of biomass to digestion by enzymes.

14.1 Introduction

Prior to the discovery of cellulolytic enzyme systems, processes that could thermochem­ically hydrolyze lignocellulose to produce soluble sugars were investigated and developed with some commercial use during wartime periods. Most of these processes were operated under fairly severe conditions and typically utilized concentrated acids (primarily sulfuric or hydrochloric) at relatively low temperatures (under 100°C) or dilute acids (again, typically sulfuric of hydrochloric) at much higher temperatures (often above 200°C). Such harsh conditions generally resulted in relatively low recoverable sugar yields due to sugar degra­dation reactions that produced primarily aldehydes and organic acids, along with other undesirable compounds. Also, the harsh conditions and, in the case of concentrated acid processes, the large amounts of catalyst used, caused such processes to be highly capital intensive, either from the requirements for pressurized corrosion resistant reactors capable of processing solid materials or the economic requirement to recover and recycle the acid catalysts. Advances have been made in thermochemical acid hydrolysis processes to improve their commercialization potential in some niche opportunities, especially for concentrated acid processes. These improvements are primarily related to acid catalyst recovery processes that allow for the cost-effective recycling of the large quantities of required acid catalysts (1).

Biomass Recalcitrance: Deconstructing the Plant Cell Wall for Bioenergy. Edited by Michael. E. Himmel © 2008 Blackwell Publishing Ltd. ISBN: 978-1-405-16360-6

The discovery of cellulase enzymes and the commercial availability of such enzyme sys­tems have dramatically changed the context of the thermochemical step in the conversion of lignocellulosic biomass to sugars. Rather than requiring this thermochemical step to directly produce all resulting sugars from biomass, this step can now be viewed as a true pre­treatment step, whose purpose is to prepare the biomass for rapid and complete enzymatic hydrolysis to produce a monomeric sugar stream. Conceptually, the enzymatic hydrolysis approach has several inherent advantages over thermochemical hydrolysis, including low — temperature, mild-pH conditions leading to less expensive and less complex reactor systems and significantly lower or no losses of resulting sugars to degradation products.


The requirement for high level cellulase synthesis in order for microbes to grow on cellulose raises the question: how does an anaerobic cellulolytic bacterium generate enough ATP for cellulase synthesis? We determined that C. thermocellum assimilated cellodextrins with a mean degree of polymerization (DP) of about four during growth on cellulose and that these cellodextrins were subsequently cleaved by intracellular phosphorolytic enzymes (44). The process is presented in Figure 16.4.

Soluble cellodextrins (G„) from cellulose hydrolysis were transported across the cell mem­brane via the ABC transportation system with one ATP expenditure per molecule (45),

Gn + ATP ^ Gn + ADP + Pi

Although the sugar ABC transportation seems bioenergetically costly, it could be very im­portant for (thermophilic) bacteria competing for low concentrations of hydrolysis prod­ucts. These ABC transportation systems are widely observed in thermophilic or hyperther­mophilic microorganisms with very high affinities of sugar

After sugar assimilation, intracellular cellodextrin and cellobiose phosphorylases, rather than p-glucosidase, cleave the (3-(1-4) bonds of cellodextrins via substrate phosphorylation (49),

Gn + Pi + H2O ^ G„-1 + G — 1 — P

This phosphorylation process is important because it conserves energy stored in p-glucosidic bonds to generate one molecule of G-1-P (convertible to ATP) per cleav­age and avoids energy dissipation resulting from hydrolysis mediated by p -glucosidase. Our bioenergetic analysis clearly indicated that by assimilating cellodextrins of average DP of 4, C. thermocellum has more ATP available for cell synthesis when growing on cellulose than on cellobiose (44). Hydrolysis products longer than cellobiose are supported by the supramolec — ular structure of the cellulosome and the distance between two adjacent catalytic subunits in the C. thermocellum cellulosome has been estimated to be eight glucosidic bonds (30). Thus, simultaneous catalytic events mediated by adjacent catalytic components would be expected to result in an insoluble G8 fragment, and any subsequent inter-cleavage of this G8 fragment would result in two soluble products with mean chain length 4. Low solubil­ity (50) and/or a tendency to bind to cellulose may well prevent yet longer cellodextrins from being assimilated, even though this would in theory offer bioenergetic benefits. In summary, assimilation of longer cellodextrins means more energy generated from substrate phosphorylation and less energy expenditure for sugar transportation.

Location of pectin synthesis

All available evidence, including autoradiographic pulse chase studies using wallbiosynthetic precursors (222, 223), immunocytochemical studies using anti-pectin-specific antibodies (224-226), and subcellullar fractionation and topology studies of pectin biosynthetic en­zymes (227-231), indicate that pectin is synthesized in the Golgi and transported to the wall in membrane vesicles. Plant cells, unlike animal cells, have multiple Golgi and thus pectin synthesis occurs simultaneously in numerous Golgi stacks in the cell (225, 232). The synthesized pectin and other macromolecules are targeted to the wall by the movement of Golgi vesicles, presumably along actin filaments that have myosin motors (233).

Immunocytochemical studies also indicate that the synthesis of different regions of the pectic polysaccharides occurs in different Golgi cisternae as pectin moves from the cis, through the medial and to the trans-Golgi. For example, the use of antibodies specific to different regions of HG and RG-I suggests that HG and RG-I synthesis begins in the cis-Golgi (225, 234, 235) and continues with more extensive decoration of the backbones as the polymers move through the medial Golgi (224, 225, 235) and into the trans-Golgi cisternae (225, 235). Additional modifications of the pectic glycan structure also appear to proceed in a more-or-less organized manner with HG (236, 237) and RG-I (106, 234, 238, 239) initially synthesized in less modified forms in the cis — and medial-Golgi and becoming more modified (e. g., methylesterified) (236) in the medial — and trans-Golgi (225, 235,240-242). HG is believed to be transported to the plasma membrane and inserted into the wall as a highly methylesterified polymer (214, 237, 243-245) and once in the wall, HG is deesterified to varying degrees by pectin methylesterases (246) in the wall or at the cell plate (245). The deesterification of HG converts it to a more negatively charged form (240, 247-250) which is then available to bind ions, enzymes, proteins, and other HG molecules through Ca++ salt bridges. It is believed that a spatial partitioning of HG esterification and deesterification occurs in the wall based on localization of esterified HG throughout the cell wall (237, 240-242, 243, 245, 250, 251), while relatively unesterified HG is more restricted to the middle lamella. This conclusion is supported by the frequently observed absence of unesterified HG epitopes in the trans-Golgi vesicles. However, since some cell types, such as melon callus cells (240), contain unesterified HG in the trans-Golgi, it is possible that HG may be inserted into the wall in a relatively unesterified form, at least in some cells. Also, since specific pectic epitopes localize to different Golgi compartments in different cell types (5, 225, 234, 237, 244), it is likely that the specific localization of the diverse pectin biosynthetic enzymes may vary in a cell type, species, and development-specific manner (226,252-255). It must be noted, however, that the interpretation ofimmunocytochemistry results can be difficult since the absence of a signal using an epitope-specific antibody may be due to masking of the epitope by additional glycosylation or some other modification (e. g., methylation, acetylation, feruloylation). Thus, to conclude that a particular pectin biosynthetic event does not occur in a cell, the lack of a particular immunocytochemical signal is not sufficient. Information on the presence of the biosynthetic enzyme activity or the actual wall carbohydrate structure itself is required.