Other species of Gram-positive cellulolytic rumen bacteria are known to include Eubac — terium cellulosolvens (32,33). Some strains of Butyrivibrio fibrisolvens have also been reported to be cellulolytic, although most strains of this species are actively hemicellulolytic, but not cellulolytic (34-36). Interestingly, there is evidence for cell-associated enzyme complexes in B. fibrisolvens (37). Sortase-mediated anchoring of individual enzymes to the bacterial cell wall has also been reported for amylases from a human colonic B. fibrisolvens strain and from the related Roseburia inulinivorans (38). It is likely, however, that many other species whose primary niche is not in plant cell wall breakdown possess enzymes for the transport and utilization of oligosaccharides released from plant cell walls. Selenomonas ruminantium, for example, has a xylan utilization operon (39) and is involved in interactions with cellu­lolytic species (40). Even Streptococcus bovis, assumed to be primarily a starch-degrading species, was found to possess a mixed link p-glucanase (41). The role of this enzyme was postulated to be in gaining access to starch through the removal of p-glucan-rich walls of cereal endosperm cells.

Bacteria are by far the most numerically abundant and taxonomically diverse microorgan­isms in bulk soil and are found both free-living and attached to the surface of soil particles. Many soil bacteria also interact with the roots of plants within the rhizosphere. Many of these bacteria have also been shown to possess hydrolytic enzymes that enable them to colonize and degrade plant biomass. They are widely distributed across phylogenetic groups that include diverse functional groupings. Many are native to soil, but have also been isolated from such diverse environments as hot springs, rumen contents, compost piles, termite guts as well as various other sites. The biochemical characteristics and physiology of biomass de­grading prokaryotes vary according to the environmental niche where they are found. Due, in part, to the diversity of prokaryotic microorganisms that produce cellulolytic enzymes and the promise of newer and better high-specific activity enzymes for industry, (Diversa is now called Verenium Corporation) these systems remain the focus of considerable study. Companies like Diversa Corporation in San Diego, California, collect DNA from hydrother­mal and other habitats worldwide, and then screen-extracted genomic DNA for the ability to produce useful enzymes for biomass conversion applications. Diversa’s business platform since 1995 has been to discover novel enzyme by creating gene libraries from bioprospecting globally diverse environments. Their ultimate goal is to develop enzymes capable of sur­viving at extreme process conditions for industry. Another company, Dyadic International, Inc, uses an integrated high-throughput technology platform targeted specifically at the discovery, expression, and modification of both prokaryotic and eukaryotic genes.

For microorganisms to degrade and metabolize the insoluble polysaccharides found in plant cell walls they must produce several extracellular hydrolytic enzymes. These enzymes can be secreted and act free in solution, or are cell associated. Adherence or colonization to insoluble substrates by microorganisms is common. Attachment can also be a factor in the control of enzyme expression and can be a precondition to the production of hydrolytic enzymes. Commonly, microorganisms act synergistically to enable efficient conversion of the substrate by the concerted action of several species. Synergism may involve the production of bacterial communication signals (quorum sensing) such as acylated homoserine lactones (AHLs) to establish these communities (20, 21). Quorum sensing has also been reported as a regulatory system for the control of extracellular enzyme synthesis in phytopathogenic bacteria, and in nitrogen-fixing rhizobia (22, 23). Bacterial communication signals may also allow the cells to regulate secretion of hydrolytic enzymes to reduce losses because of diffusion.

Because cellulose is a large insoluble polymer its use first requires binding of the enzymes either as a binary enzyme-substrate complex or as a ternary enzyme-substrate-microbe complex. Adhesion is most pronounced in the Gram-positive, thermophilic anaerobes such as Clostridium thermocellum or Clostridium cellulolyticum, which secretes an active and thermostable high molecular weight cellulase complexes (cellulosome) responsible for de­grading crystalline cellulose (24,25). Cellulosomes contain at least 30 polypeptides, most of the enzymes are endoglucanases (EC:3.2.1.4), but there are also some xylanases (EC:3.2.1.8), p-glucosidases (EC:3.2.1.21), and endo-p-1,3-1,4-glucanases (EC:3.2.1.73). Hydrolysis of cellulose by cellulosome producing organisms is dependent on adherence of the organism to the substrate through specific cellulose-binding proteins.

At least 46 unique bacterial producers of cellulases have been identified from many aerobic bacterial systems, including species within the genera Acidothermus, Bacillus, Cellulomonas, Cellvibrio, Cytophaga, Microbispora, Pseudomonas, and Thermobifida (26). Anaerobic bac­teria identified as biomass degraders include members of the genera Acetivibrio, Bacteroides, Clostridium, Micromonospora, and Ruminococcus. Bacteria that decay biomass have been isolated from temperature extremes that include psychrophilic, mesophilic, thermophilic, and hyperthermophilic conditions. The actinobacteria, for example, are widespread mi­crobial components of terrestrial and aquatic communities that have been demonstrated to play key roles in biomass turnover and nitrogen dynamics. Like fungi the actinobac­teria have the ability to penetrate lignocellulosic biomass which gives them the ability of secreting hydrolytic enzymes in confined cavities allowing for higher concentrations of free enzymes.

Recently sequenced genomes representative of the actinobacteria include organisms that are widespread in plant rhizospheres, plant tissues, and compost, including Streptomyces

spp., Thermobifida fusca, and Leifsonia xyli. A close relative to Acidothermus cellulolyticus and Frankia, Kineococcus radiotolerans is a drought-tolerant soil microbe that is also radiation tolerant, while Rubrobacter is a thermophilic genus containing radiotolerant species. The genomes of all these taxa contain cellulose — and other plant biomass-degrading metabolic pathways. The high-GC actinobacteria represent a large phylogenetic group and serve as a reservoir of organic carbon and nitrogen cycling capabilities. Many species of actinobacteria have adapted to diverse and often extreme environmental conditions. For example, Acti­nobacteria have been identified a the major group of cellulolytic organisms in ecosystems such as acid Sphagnum peat bogs (27).

The genomic sequence of A. cellulolyticus, a major cellulose degrader (see also Figure 15.2) originally isolated from the hot springs samples has recently been compiled and published by the Department of Energy Joint Genome Institute, accession NC-008578. A. cellulolyticus was originally isolated from submerged woody debris in a Yellowstone National Park hot spring, because of its efficient cellulose degradation (28). The organism was isolated using selective culturing methods of mud and decaying wood samples from thermal features in and around the Norris Geyser Basin region of the park. The microorganism was found to grow at 37-65°C with an optimum of 55°C. A type strain was selected and deposited in the American Type Culture Collection and is part of their National Park Service special collection (http://www. atcc. org/SpecialCollection/NPS. cfm). The genome contains a suite of glycosyl hydrolases responsible for cellulose degradation that have already been charac­terized as thermostable (29) as well as a xylanase and additional multifunctional hydrolytic activities.

The genome of the closely related thermophile, Thermobifida fusca, has also been com­pleted. The T. fusca genome shares a high number of gene homologs with A. cellulolyticus. Also of interest are members of the genus Frankia, which are filamentous actinobacteria that form nitrogen-fixing root nodules on woody trees and shrubs in a symbiosis termed “actinorhizal.” Genome sequence data are now available for three strains of Frankia, an eco­logically important nitrogen-fixing root nodule symbiont which is the closest phylogenetic relative to A. cellulolyticus. The sequenced Frankia genomes contain homologs for cellulases, xylanases, peroxidases, and aromatic degradative enzymes, as well as genes for secondary metabolite biosynthesis. Similar metabolic capabilities have been identified in the genomes of Streptomyces spp., suggesting that actinobacteria maybe a rich source of enzymes for bio­conversion. Members of the Frankia have been found to inhabit root nodules, rhizosphere (termed actinorhizal), and the soil as a saprophyte. Frankia — actinorhizal plant symbio­sis has been reported within a range of actinorhizal plants scattered among eight families consisting of over 200 species of angiosperms (30, 31).

Thermobifida fusca is a moderate thermophilic soil actinomycete (growth temperature ranging from minimal 30°C to maximal 55°C) that is a major degrader of plant cell walls in heated organic materials such as compost heaps, rotting hay, manure piles, or mushroom growth medium. It produces spores that can be allergenic and causes a condition called farmer’s lung. Its extracellular enzymes, including cellulases, have been studied extensively because of their thermostability, broad pH range (32-37), and high-specific activity. It degrades all major plant cell wall polymers except lignin and pectin and can grow on most simple sugars and carboxylic acids. Its genome has been sequenced and closed by the JGI; it has a size of 3.4 Mb and encodes seven known cellulase genes, all which have been expressed and characterized. Four of the cellulases are endocellulases (Cel5A, Cel5B, Cel6A, Cel9B), one is an exocellulase attacking the nonreducing ends of cellulose molecules (Cel6B), one is an exocellulase attacking the reducing ends of cellulose molecules (Cel48A), and one is a new class of cellulase, a processive endocellulase, which have now been found in several cellulolytic bacteria. All the cellulases contain a family 2 cellulose-binding module (CBM) attached by a linker peptide. There are many hemicellulose genes in T. fusca and five of them have been expressed and characterized: Xyl11A, Xyl10A, Xyl10B, XG74A, and Gn81A.

There have also been extensive studies of the regulation of T. fusca cellulases that have shown that their synthesis is induced by cellobiose and repressed by any good carbon source. All the six-cellulase genes encoding cellulases that are regulated by cellobiose contain at least one copy of a 14-base inverted repeat sequence upstream of their start codon. This sequence is the binding site for a lacI family regulatory protein, CelR. CelR has been expressed and characterized in E. coli. Its binding to DNA containing the 14 base sequence is inhibited by cellobiose. There are a number of copies of this 14 base sequence in T. fusca DNA, including one copy upstream of an operon that contains a p-glucosidase gene and genes for a binding protein transport system. The two exocellulases make up about 75% of the total cellulase produced by induced T. fusca and both exocellulase genes contain a second copy of the CelR-binding site upstream of their start codon. It is not clear if this regulatory site is responsible for their higher transcription rate. It is also not known if CelR plays a role in carbon source repression.

The genome of Cytophaga hutchinsonii, another eubacterial cellulose degrader, has also recently been sequenced. C. hutchinsonii, an aerobic Gram-negative bacterium commonly found in soil that has been shown to rapidly digests crystalline cellulose (38). Molecular analysis of cellulose degradation by C. hutchinsonii is now feasible, since techniques for the genetic manipulation of this organism have recently been developed (39). C. hutchin­sonii belongs to the Cytophaga-Flavobacterium branch of the eubacterial phylogenetic tree. Members of this group are widely distributed in many environments and have the ability to move rapidly over surfaces by gliding motility (40, 41). Gliding motility is thought to be important in allowing C. hutchinsonii to colonize its insoluble growth substrate. The genome of C. hutchinsonii was sequenced by the JGI and has been shown to differ from most known cellulolytic microorganisms in that none of its cellulase genes encode processive cellulases and few of them encode a CBM. These results provide strong evidence that C. hutchinsonii does not use either of the two known mechanisms for cellulose degradation: secretion of a set of individual synergistic cellulases containing CBMs or production of cellulosomes, since processive cellulases and CBMs are key for both mechanisms. Determining the detailed mechanism of cellulose degradation by C. hutchinsonii is a major unsolved problem in plant cell wall degradation and this pathway might provide new proteins that improve the rate of cellulose degradation.

Recently, an interesting group of marine microorganisms, the marine complex polysac­charide (CP)-degrading bacteria from the Microbulbifer, Teredinibacter, and Saccharopha — gus group have been described. This group of organisms produces an array of enzymes to degrade complex polysaccharides including cellulose (42). The genome of the marine bac — terium, Saccharophagus degradans, has been completed and reportedly contains more than 180 open reading frames that encode carbohydrate-degrading proteins (43). S. degradans is a pleomorphic, Gram-negative, aerobic member of the у — proteobacterium isolated from decaying salt marsh cord grass.

C. phytofermentans is a recently discovered member of the order Clostridiales. Its genome is presently being sequenced by the JGI. It was found in forest soil and has a broad growth substrate range and is able to rapidly degrade and ferment several plant polymers, including cellulose, pectin, starch, and xylan (44). A remarkable property of cellulose-fermenting C. phytofermentans cultures is the production of high concentrations of ethanol, typically more than twice the concentration produced by other cellulolytic clostridia, and hydrogen.

4.4.1 Primary cell walls

4.4.1.1 Pectin-rich walls

Primary walls that are rich in pectic polysaccharides occur in the gymnosperms, eudicotyle — dons, non-commelinid monocotyledons, and palms (Arecaceae) (3) (see Table 4.1). The first model for this type of wall was proposed by Keegstra and coworkers (142) based on detailed analyses of the walls of cell suspension cultures of sycamore (Acer pseudoplatanus). This model depicted the pectic polysaccharides as being covalently linked to the xyloglucans and to the glycoprotein extensin. Hydrogen bonding between the xyloglucans and the cellulose microfibrils provided the link between the matrix complex and the cellulose. However, evi­dence for possible covalent linkages between pectic polysaccharides and xyloglucans was not obtained until recently (86-88). Instead, models, often referred to as “tethered or sticky net­work models,” were developed in which xyloglucans were postulated to form non-covalent bridges between cellulose microfibrils in walls through hydrogen bonding (143, 144).

Evidence for these non-covalent bridges was obtained using transmission electron mi­croscopy after preparation of walls of the non-commelinid monocotyledon onion (Allium cepa) by fast freezing, deep etching, and rotary shadowing (145). From these studies, they developed a model in which there are two co-extensive, but independent, polymer networks: a cellulose-xyloglucan network and a pectic polysaccharide network. The first network is thought to be the main load-bearing structure of the wall, and the second is thought to deter­mine wall porosity (Figure 4.9). A third network composed of extensin may also be present (146). Using the same technique, similar bridges between cellulose have since been observed by other researchers in primary walls of this type from a variety of species (147-149).

In these wall models, it is usually assumed that in addition to cross-linking the cellulose microfibrils, xyloglucans completely coat the surfaces of these microfibrils. However, solid — state 13C NMR spectroscopy on isolated cell walls of mung bean (Vigna radiata) indicated a maximum of only 8% of the surface of the cellulose microfibrils had adsorbed xyloglucan (150).

The organization of the pectic polysaccharide network in these wall models is not clear but is probably in the form of a gel. Thus, the anionic HGA and RG-I have strong gel forming capabilities (53). HGA in the solid/gel state has an extended flexible conformation and may adopt a double or triple helical structure depending on the degree of hydration and the nature of the counter ion. Gel formation is due to the coordination of Ca2+ ions by carboxylic acid groups on adjacent chains enabling the formation of junction zones between HGA or RG-I chains. The strength of the HGA gel depends on the extent of methyl esterification of the GalAp residues.

50 nm

Figure 4.9 A simplified schematic representation of the spatial arrangement of polymers in a pectin-rich primary cell wall, e. g. of a parenchyma cell, as occurs in gymnosperms, eudicotyledons, non-commelinid monocotyledons, and palms (Arecaceae). The cellulosic microfibrils are embedded in a network of non — cellulosic polysaccharides [xyloglucans, pectic polysaccharides, and proteins (not shown)]. The xyloglu — cans are associated with the microfibril surfaces and form bridges between them. The immature primary wall may contain ~60% water but during development of the wall in some cell types (e. g., xylem fiber or tracheid), the water is replaced by lignins, which encrust the cellulosic microfibrils and non-cellulosic polysaccharides and may be covalently linked to them. (Reprinted with permission, from McCann, M. C. & Roberts, K. (1991) Architecture of the primary cell wall. In: The Cytoskeletal Basis of Plant Growth and Form (ed. C. W. Lloyd), pp. 109-129, Fig. 9.19, p. 126. Academic Press, London.)

The conformation of RG-I depends on the length of the alternating ь-Rhap insertions into the galacturonan chain (Figure 4.2). A single ь-Rhap causes a kink in the chain but two or three alternating ь-Rhap residues results in a chain having a fully extended conformation comparable with HGA. RG-I side chain galactans, arabino-4-galactans etc. (see Section 4.2.1.2.4) may not seriously restrict the stereochemistry of the backbone permitting junction zone formation (53) HGA and RG-I gels are important structurally in primary cell walls and in interfaces between walls in the middle lamella, the interfacial layer between adjacent cells.

In addition to calcium bridges between HG domains, cross-linking of the pectic polysac­charide network can occur by formation of borate esters between RG-II substituents, and in the walls of “core” families of the Caryophyllales (eudicotyledons) by DDFA cross-linking of RG-I (3, 52,62). Covalent linking of pectic polysaccharides to xyloglucans may also possibly occur (86-88).

Detergent-solubilized microsomal proteins from flax can use an RG-I-enriched fraction as an exogenous acceptor for methylation in the presence of S-adenosylmethione (334). The pectin methyltransferase activity was stimulated by the addition of the enriched RG-I fraction 1.5- to 1.7-fold above levels recovered using endogenous acceptor, and the radio­labeled product had a size similar to RG-I. However, since it was not shown where in RG-I the methylation occurred, it is not clear whether the methylation occurred on GalA in the RG-I backbone or, rather, on possible HG tails that may have been covalently linked to RG-I. Also, it was not established whether some of the methylation may have occurred on a non-galacturonic substituent such as methylation at the 4-position of glucuronic acid in the side branched of RG-I (278).

5.4.10.2 RG-I:acetyltransferase (RG-I:AT)

GalA in the alternating [^4)-a-D-GalpA-(1^2)- a-L-Rhap-(1^] backbone of RG-I may be acetylated on C-2 and/or C-3 (289). Microsomal membranes from suspension — cultured potato cells (339) contain an RG-I acetyltransferase that transfers [14C]acetate from [14C]acetyl-CoA onto endogenous RG-I acceptor to yield a >500-kDa radiolabeled product (339), based on the release of [14C]acetate following incubation of the product with a purified rhamnogalacturonan O-acetyl esterase, and fragmentation of the product by rhamnogalacturonan lyase (RGase B) (339). The RG-I acetyltransferase has an apparent Km for acetyl-CoA of 35 ^M and a pH optimum of 7.0.

Traditional protein biochemistry and classical purification was instrumental in identify­ing key wall biosynthetic enzymes, for example, XG:1-2,fucosyltransferse (116, 117) and GM:1-6,galactosyltransferase (88). As most of the glycosyltransferases involved in polysac­charide synthesis are membrane bound and are of low abundance, i. e., estimated 2-6 molecules per cell, it is clear that other methods should be sought to identify wall biosyn­thetic proteins. Classical genetics has already contributed immensely to the identification of large numbers of genes involved in glycan synthesis such as cellulose synthase and their large gene family (507,508) andxyloglucan synthesis (120). The genome sequence facilitated the identification of mannan synthase (85) and cellulose-like proteins (154). A combined approach of partial protein purification and proteomics was useful in the identification of a large family associated with the pectin biosynthetic HG:galacturonosyltransferases (137). The classification of GTs in the CAZY database was instrumental in identifying GT gene candidates targeted for a reverse genetic approach using the SALK T-DNA or other mutant library collections. Lastly, microarray analyses of tissues or cell types at different devel­opmental stages were useful in identifying secondary wall synthesis candidates (139, 509). Similar biochemical, genetic, and genomic approaches were successful in identifying biosyn­thetic genes involved in nucleotide-sugar synthesis and NDP-sugar transporters. The last decade has been a very fruitful and exciting time for the community of wall researchers (finally a crack in the wall).

Can bioinformatics assist in the prediction of GTs? Would the knowledge of UDP-binding sites in several plant GTs be useful to distinguish between GDP-, or ADP-transferases? Would the knowledge of the sugar moiety-binding site be similarly helpful to identify, for example, putative GalTs? Is the binding pocket for UDP-Rha in a flavonoid: RhaT the same as for pectin:RhaT? Clearly, a critical mass of biochemical knowledge is required to start to predict gene function by computer. We are not there yet__________________________

Acknowledgments

A review of this magnitude is not possible without the input, efforts, and patience of countless individuals. The authors express sincere gratitude to the multiple students, researchers, colleagues, and staff who contributed to this chapter through engaging discussions and editorial effort. Special thanks go out to our families who graciously endured our too frequent absences. We also thank the Department of Energy, the National Science Foundation and

the NRI, CSREES, USDA who provide the invaluable funding that supports plant cell wall research.

A similar situation to that for PAL downregulation/mutation holds for C4H. Although this enzyme is considered to be one of the important rate-limiting processes in phenylpropanoid metabolism (such as for differential carbon allocation) (34, 35), it is also a common step leading to various metabolites, including monolignols, lignins, flavonoids, and so forth. Downregulation of C4H has though only been studied in a very preliminary way thus far

Wild type B3 Tobacco Cinnamoyl CoA reductase
Wild type AOPP — treated Mungbean Phenylalanine ammonia lyase
Normal > development	Delayed development
CCRt-irx4
Wild type cad-4 cad-5 Arabidopsis Cinnamyl alcohol dehydrogenase
Wild type pC3H-l Alfalfa p-Coumarate-3-hydroxylase
Larger vessels
Normal vessels
Wild type CS-1 Tobacco l	Wild type 1074 Tobacco s p

(77, 207, 210-212) as regards overall effects on lignification and the vasculature. As noted above for PAL, the methodologies employed in two of these studies (207,212) were again very questionable (i. e., estimations of Klason lignin of neutral detergent fiber), and ultimately found to be unreliable as analytical protocols. Nevertheless, in all three studies the lignin levels provisionally appeared to be reduced — at least by >25% (207, 210, 211). Various alterations were also noted in S/G ratios of the lignified tissues, the reasons for which are yet unknown. As for PAL modulation, no evidence was obtained indicating deployment of non-monolignol moieties to “compensate” for reductions in lignin levels. There were also no experiments conducted to quantitatively ascertain the effects on the vascular integrity through biomechanical analyses. In hindsight, such studies need to be expanded significantly in order to ascertain, at the very least, the true effects of reductions in lignin levels on plant growth/development, on overall vascular structure/anatomy, and on lignin macromolecular configuration, etc.

<———————————————————————————————————————

Figure 7.12 Examples of effects on vascular anatomy of inhibiting/mutating/downregulating various en­zymes and/or genes in the phenylpropanoid pathway. (A) AOPP-treatment of mungbean (204), results in collapsed xylem. (B) Similar effects occurred in a pC3H mutated Arabidopsis line (ref8). (C) On the other hand, formation of gelatinous fibers such as in wild-type alfalfa (Medicago sativa) apparently help mechanically offset deleterious effects of pC3H downregulation without forming "abnormal" lignins (72).

(D) Collapsed xylem observed in tobacco (Nicotiana tabacum) lines 4CL downregulated (223), as was

(E) xylem in a tobacco line CCR downregulated (232). (F) Detailed analysis of the Arabidopsis irregular xylem4 mutant (CCR1-irx4) identified pleiotropic effects, including delayed xylem formation (131). (G) An Arabidopsis CAD double mutant, AtCAD4/5, (cad-4, cad-5) with unusual red coloration in the xylem attributed to presence of p-hydroxycinnamyl aldehydes (71). (H) Qualitatively larger vessels observed in a tobacco line downregulated for a lignin-specific peroxidase (257). All images are of transverse sections and taken using brightfield microscopy, unless otherwise specified. Images shown in (A) are SEM analyses, while images (B and F) are images of semi-thin resin embedded sections stained with either toluidine blue O or Stevenel’s Blue. Image (C) was taken of cryosections stained with zinc chloro-iodide. Image (D) was taken of hand-cut sections stained with phloroglucinol-HCl. Image (E) was taken using UV light microscopy. Image (G) was recorded using unstained hand-cut sections, while image (H) was taken of phloroglucinol-HCl stained sections. [Reprinted from (A) The European Journal of Cell Biology, vol. 29, Amrhein, N., Frank, G., Lemm, G. & Luhmann, H.-B., Inhibition of lignin formation by L-a-aminooxy-P — phenylpropionic acid, an inhibitor of phenylalanine ammonia-lyase, pp. 139-144, Copyright 1983, with permission from Elsevier. (D) Plant Science, vol. 128, Kajita, S., Mashino, Y., Nishikubo, N., Katayama, Y. & Omori, S., Immunological characterization of transgenic tobacco plants with a chimeric gene for 4-coumarate:CoA ligase that have altered lignin in their xylem tissue, pp. 109-118, Copyright 1997, with permission from Elsevier. (E) Plant Journal, vol. 13, Piquemal, J., Lapierre, C., Myton, K., O’Connell, A., Schuch, W., Grima-Pettenati, J. & Boudet, A.-M., Downregulation of cinnamoyl-CoA reductase induces significant changes of lignin profiles in transgenic tobacco plants, pp. 71-83, Copyright 1998, with per­mission from Blackwell. (F) Phytochemistry, vol. 66, Patten, A. M., Cardenas, C. L., Cochrane, F. C., Laskar, D. D., Bedgar, D. L., Davin, L. B. & Lewis, N. G., Reassessment of effects on lignification and vascular devel­opment in the irx4 Arabidopsis mutant, pp. 2092-2107, Copyright 2005, with permission from Elsevier. (G) Phytochemistry, vol. 68, Jourdes, M., Cardenas, C. L., Laskar, D. D., Moinuddin, S. G.A., Davin, L. B. & Lewis, N. G., Plant cell walls are enfeebled when attempting to preserve native lignin configuration with poly-p-hydroxycinnamaldehydes: Evolutionary implications, pp. 1932-1956, Copyright 2007, with per­mission from Elsevier;and (H) Phytochemistry, vol. 64, Blee, K. A., Choi, J. W., O’Connell, A. P., Schuch, W., Lewis, N. G. & Bolwell, G. P., A lignin-specific peroxidase in tobacco whose antisense suppression leads to vascular tissue modification, pp. 163-176, Copyright 2003, with permission from Elsevier.] (Reproduced in color as Plate 21.)

Figure 7.13 Gross phenotypical changes/effects of either mutating or downregulating various phenyl — propanoid genes, as well as vascular related transcription factors. (A) p C3H mutant with various pleiotropic effects, resulted in a severely dwarfed line (ref8) (213, 215) (shown 23 days post seed-sown). (B) pC3H downregulation in alfalfa (Medicago sativa) resulted in a phenotype (pC3H-I) very similar to wild type without visible pleiotropic effects (shown 4 weeks post-cut-back). (C) Dwarfed Arabidopsis CCR irx4 mutant with pleiotropic effects (shown 6 weeks post seed-sown) (131). (D) CCR downregulation in to­bacco, with stunted morphology relative to wild type (232). (E) CAD double mutation in Arabidopsis (AtCAD4/5, cad-4, cad-5) resulted in a limp to prostrate stem phenotype (shown after 4 weeks growth) (71). (F) Double T-DNA tagged Arabidopsis nst1 nst3 line (199). [Reprinted from (B) The American Journal

The accurate treatment of non-bonded interactions is paramount in a force field descrip­tion. The philosophy behind the AMBER force field is to deal with these interactions in the most accurate way possible within the limitations of the available hardware. Non-bonded interactions do not depend upon a specific bonding relationship between atoms. They are “through-space” interactions, the number of which scales roughly as the square of the num­ber of atoms. Unsurprisingly, the non-bonded interactions form the most time-consuming component of molecular mechanics simulations. Molecular mechanics force fields typically consider the non-bonded interactions as two groups, one comprising electrostatic interac­tions and the other van der Waals interactions.

8.2.2.4.1 ELECTROSTATIC INTERACTIONS

The last term in Equation (8.3) describes the electrostatic interactions within the system. There are a number of ways to represent the charge distribution within a molecule, the simplest being the use of point charges. This is the method utilized by most biologically oriented force fields. In the point charge model a series of fractional charges are distributed throughout the molecule. If the charges are centered on atoms then they are referred to as partial atomic charges. The interaction energy is calculated using Coulomb’s law.

Since the charge on an atom is not experimentally observable the partial atomic charges have to be assigned in an analogous fashion to the parameters used in the bonding interaction terms. The partial charges are generally obtained by fitting to a charge potential obtained from an ab initio electronic structure calculation.

Separation

10.3.1 Xylanases

Xylans from different plant sources are categorized by the substituents of their side chains (i. e., arabinoxylan, glucuronxylan, etc.); however, all xylans are built upon apoly-p-(1^4)- xylopyranose backbone. Xylanases depolymerize the (3-(1^4)-xylopyranose backbone of
a variety of xylans, with the endoxylanases classified in 3.2.1.8 [P-(1—>4) hydrolysis] or 3.2.1.32 [p-(1—4) or (3-(1—— 3) hydrolysis]. In both groups, the anomeric configuration of the reducing-end xylan is retained. Most xylanases belong to the two structurally different glycosyl hydrolase groups (families 10 and 11) and differ from each other with respect to their catalytic properties (17). Xylanases with high Mr/low pI (family 10) seem to exhibit greater catalytic versatility than the low Mr/high pI-xylanases (family 11) and thus they are, for example, able to more efficiently hydrolyze highly substituted xylans. Few xylanases have been classified in family 5, which is a very versatile family containing various types of glycanases. Some xylanases have been reported to contain either a xylan-binding domain (18) or a cellulose-binding domain (19). Some of the binding domains have been found to increase the degree of hydrolysis of fiber-bound xylan, whereas others have no effect thereon.

Most of the xylanases characterized are able to hydrolyze different types of xylans, show­ing only differences in the spectrum of end products. However, no systematic studies on the substrate specificity of xylanases belonging to different families on fiber-bound sub­strates have been carried out. The majority of the endoxylanases produce mainly xylobiose, xylotriose, and substituted oligomers of two to four xylosyl residues as products upon ex­tended incubation. Xylose, xylopentaose, and higher oligomers may also be produced, with specific product patterns being dependent upon the individual enzyme and the substrate. Most of the endoxylanases hydrolyze unsubstituted xylose polymers, with the tolerance of side chains again being dependent on the specific xylanase.

The exoxylanases are found in 3.2.1.37,3.2.1.72, and 3.2.1.156, though enzymes in 3.2.1.72 are active on (3-(1—— 3)-linkedxylosides and have limited activity on the majority of xylans. The other two categories, though similar in activity, are distinguished by their difference in preferred substrate degree of polymerization, with the p-xylosidase (1,4-p-D-xyloside xylohydrolase, EC 3.2.1.37) acting more effectively on polymeric xylan, while enzymes in 3.2.1.156 have a greater affinity for xylo-oligomers. Both groups act at the reducing end of the chain, yielding xylose as their product. Exoxylanases are less abundant and consequently less well known and understood than the endoxylanases, as only a few examples of the former have been characterized. Taking into account the high degree of substitution of native xylans, the exo-mode of action maybe less dominant among hemicellulases than cellulases. Exoglycanases are generally larger proteins than endoglycanases, with molecular weights above 100 kDa and they are often built up by two or more subunits. The p-xylosidases, which have a few well-studied examples and though active primarily on xylobiose, may have a processive activity on xylo-oligomers.

Early on, it was recognized that cellulosomes were intimately associated with the bacterial cell surface of C. thermocellum (88, 108, 120). As mentioned above, the molecular mechanism for this phenomenon was later determined. The type-II cohesin-dockerin complex fixes the primary scaffoldin and its complement of enzymes to one of the anchoring proteins, which also contains an S-layer homology (SLH) module that mediates attachment to the peptidoglycan of the cell surface (Figure 13.1) (36, 40-43, 121). Other species of bacteria, e. g., Acetivibrio cellulolyticus and Bacteroides cellulosolvens, possess anchoring protein(s) that exhibit similar type-II cohesins and SLH modules for attachment of the enzyme-bearing primary scaffoldin to the cell surface (88, 108, 120, 122-124). However, this is not the only strategy. As described above, ScaE of R. flavefaciens contains a sortase signal motif through which the scaffoldin is covalently attached to the peptidoglycan (Figure 13.2) (68). Other bacteria, notably C. cellulolyticum and C. cellulovorans, are also known to bear cellulosomes on their cell surface (125, 126), but there is as yet no evidence for involvement of an SLH module or sortase-mediated covalent attachment. The known scaffoldin of both of the latter species includes two and four copies of a hydrophilic domain (37, 46), which, in the case of C. cellulovorans, has been implicated in a cell-attachment function (127), but the true function of this particular domain awaits further experimental verification.

The attachment of the cellulosome to the cell surface, coupled with the CBM of the pri­mary scaffoldin, inferred that the cellulosome is intimately involved in cell adhesion to the insoluble substrate. Indeed, the initial demonstration of the cellulosome in C. thermocel — lum was assisted by the isolation of an adherence defective mutant that was also impaired in its arrangement of cellulosomes on the cell surface (108). Cellulosomes are packaged into polycellulosome-containing protuberances (Figure 13.4), which undergo a dramatic conformational change upon interaction with the cellulose surface (27-29). The protuber­ances thus protract to form “contact corridors” laden with fibrous material that connect between cellulose-bound cellulosomes and the cell surface thus promoting cell uptake of the cellulose degradation products. The cellulosomal enzymes are subject to potent product inhibition (89,111), and cell uptake and assimilation of the cellulose-degradation products (i. e., cellobiose and higher order soluble cellodextrins) relieves the inhibitory effect, thereby allowing facile hydrolysis of the insoluble substrate to proceed unhindered. Moreover, addi­tional saccharolytic bacteria, which share the same ecosystem with the polymer-degrading strains (e. g., C. thermocellum), further purge the immediate environment of the inhibitory sugars, thus propagating substrate degradation even more (19, 128).

The cellulosome-producing cellulolytic bacterium A. cellulolyticus is characterized by an especially elaborate surface morphology; when grown on cellulosic substrates, the

Resting

Protuberances

Protracted

Protuberance

Cellulosomes

.’Ceifutose.

Figure 13.4 Proposed interaction of cellulosome-producing bacteria with cellulosic substrates. (Panel A) Schematic representation of the interaction of cell-surface cellulosome-laden protuberance-like organelles with the cellulose surface. 1) Prior to contact with the cellulosic substrate, the protuberances are compacted in a resting state;2) upon cell-substrate contact, the protuberances protract whereby the cellulosomes interact directly with the cellulose surface;3) eventually, the cell detaches from the substrate, leaving cellulosome clusters on the cellulose surface. (Panel B) Scanning electron microscopy (SEM) of Acetivibrio cellulolyticus bound to cellulose, showing the presence of large characteristic protuberance-like structures on the cell surface. The cellulose-bound cells appear to be connected to the substrate via structural extensions of the cell-surface protuberances.

cellulosome-bearing protuberances inundate the surface of the substrate, a process which appears to assist the cells in overcoming the inherent recalcitrance of the cellulose (Figure 13.4B). During stationary phase growth on cellulose, the cells detach from the substrate, leaving a legacy of attached cellulosomes that continue their hydrolytic action in the absence of the parent cell (18, 28, 129, 130).

Y.-H. Percival Zhang and Lee R. Lynd

16.1 Introduction

Accumulation of the greenhouse gas, CO2, mainly from burning of fossil fuels, and the depletion of finite fossil fuels are vital threats to the sustainable development of humans (1-3). Lignocellulose is the most abundant renewable biological resource today (ca. 2 x 1011 tons/year) and is produced by photosynthesis (i. e., plants fix atmospheric CO2) (4-8). Development of technologies for effectively converting less-costly agricultural and forestry residues for use in bio-based chemical and fuels production offers potential benefits to the national interest by improving strategic security, decreasing trade deficits, encouraging healthier rural economies, and improving environmental quality by moving closer to zero net greenhouse gas emissions and sustainable resource supplies (1-3, 9-19).

Lignocellulosic feedstock is far less costly than other energy feedstocks (i. e., crude oil, natural gas, corn kernels, and soy oils) based on energy content ($/GJ) in Figure 16.1. For example, when crude oil prices vary from $40 to $70 per barrel, equaling $7.1 to $12.1/GJ, they are much higher than those of lignocellulose ($0 to $3/GJ). Similarly, corn kernels costing from $2.25 to $4.0 per bushel equal $6.3 to $11.5/GJ. During the past 2 years (2004-2006), corn kernel prices have risen by >70% from the historically low prices (~$2.25 per bushel) to ~$4 per bushel. With high demands of corn kernels for ethanol production, higher prices of corn kernels have resulted in rising prices of animal feed and human food. For example, the Chinese government banned the establishment of new ethanol production facilities based on grains in 2006. As expected, less costly feedstock and the most abundant supplies make production of fuels and chemicals from lignocellulose appealing.

Production of commodity products (i. e., fuels, chemicals, and materials) from renewable biomass is distinct from biotechnology motivated by health care at many levels, including economic driving forces, importance of feedstock prices, processing costs and capital invest­ment, the scale of applications, and feedstock availability (10). Biocommodities have low selling prices so that raw material costs are often dominant factors in determining prices of commodity products (~30-70%), whereas raw materials usually account for a very small fraction of high selling prices of pharmaceuticals (10, 13). The production costs, including capital recovery and processing costs, are usually another dominant factor determining the

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

Energy price ($/GJ)

price of commodity products, whereas they are not nearly so important for pharmaceuticals. Market sizes for individual biocommodity products and biopharmaceuticals are of relatively similar magnitude. However, tremendous differences exist with respect to a product mass basis, e. g., the largest commodity markets exceed pharmaceutical markets by approximately 11 orders of magnitude (10). The production of high-volume/low-value biocommodity products has an absolute requirement for high-volume/low-cost feedstock and must be re­sponsive to the availability and characteristics of feedstock, whereas no such requirement exists for the production of pharmaceuticals.

Overcoming the recalcitrant structure of lignocellulose is still among the greatest chal­lenges for the emerging biofuel and bio-based chemical industries (20, 21). Currently, high conversion costs, large investment risks, and a narrow economic margin between feed­stock costs and product prices slow the realization of cellulosic ethanol production on a large scale (22-24). Effective biological conversion of recalcitrant lignocellulose to bio­commodity products involves four main sequential steps: 1) biomass size reduction, 2) pretreatment/fractionation, 3) enzymatic cellulose and hemicellulose hydrolysis, and 4) fer­mentation (11,24,25). For most types of lignocellulose biomass, the enzymatic digestibility of cellulose is very low (<20%) without some type of pretreatment that opens up the struc­ture and makes it accessible to attack by enzymes (11, 24, 25). A number of biological, chemical, and physical pretreatment techniques have been investigated (22, 23, 26, 27).