Opportunities for Biotechnological Modification of

Opportunities for Biotechnological Modification of Lignin Structure and Composition. Extracellular Enzymes as Targets for Genetic Engineering of Forest Trees.

Jeffrey F.D. Dean and Karl-Erik L. Eriksson Department of Biochemistry University of Georgia Athens, Georgia

Introduction Lignin, a highly-stable polymer of mostly methoxylated phenolic residues synthesized as part of the cell wall of vascular plants, constitutes the second most abundant organic polymer on Earth after cellulose. The abundance and stability of lignin impacts upon mankind and the environment in several ways. In the course of pulp bleaching, residual lignin is most often separated from the polysaccharide components of wood using chlorine and chlorine derivatives, resulting in the generation of numerous toxic compounds which constitute an environmental hazard (Eriksson, 1990). Lignin is also a barrier to efficient bioconversion of cell wall polysaccharides in waste biomass to useful sugars and alcohols (Brown, 1985), and it limits the digestibility of forage crops by cattle and other ruminants (Hartley and Ford, 1989). On the other hand, degraded lignocellulosic materials are significant and beneficial constituents of soil humus (Brown, 1985), and the phenolic residues contained in lignin offer an attractive chemical feedstock (Bergeron and Hinman, 1990; Harvey et al. 1985). In light of these features, a better understanding of the structure, localization and biosynthesis of lignin would not only allow us to manipulate plants to be more resistant to disease and environmental stresses, but would also provide opportunities for more economical paper and chemical manufacture, more efficient recycling of agricultural wastes, and improvement of our environment. Lignin appears to serve several functions in the extracellular matrix of plants: it lends the cell wall mechanical support (Monties, 1989; Nelmes et al. 1973; Northcote, 1972); it serves as a barrier against microbial attack (Moerschbacher et al. 1990; Vance et al. 1980); and it acts as a water-impermeant seal for the xylem vessels of the plant vasculature (Northcote, 1989). The lignin polymer permeates xylem cell walls, surrounding and becoming covalently coupled to polysaccharides and other cell wall components during polymerization, serving much the same purpose as the resin in fiberglass (Northcote, 1972). It is this continuous, interlocking structure of polysaccharides and protein enmeshed in and covalently bound to lignin, which imparts strength and stability to the plant cell wall. Unfortunately, this same strength and stability has been a substantial barrier to the study of lignin structure and composition because it

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necessitates mechanical disruption and/or oxidation of the cell wall before lignin fragments can be extracted for analysis. In a recent review on the feasibility of using genetic engineering techniques to produce trees with altered lignin, Timmis and Trotter (1989) suggest that our ignorance of the molecular basis for particular tree characteristics will probably prove to be the limiting factor in our ability to apply biotechnology to modify trees. In this paper, we will review how new information about lignin structure in vivo is changing current thinking about lignin biosynthesis, particularly in respect to the enzymes involved in the extracellular assembly of lignin. We will also examine some of the current and future opportunities for using biotechnology to alter lignin biosynthesis in trees and other plants.

Lignin Chemistry The historic basis for our knowledge of lignin structure and composition comes primarily from chemical analyses performed to improve paper production, and are consequently slanted toward a description of tree lignins. Chemical and spectrometric analyses of extracted lignin have revealed that lignin content varies from 15 to 36% of total wood weight, with gymnosperms generally containing higher, and angiosperms lower, amounts (Fengel and Wegener, 1983; Sarkanen and Ludwig, 1971). The concentration of lignin in plant tissues also varies at the organismal level (secondary walls of vascular tissues contain the majority of lignin in plants), as well as the subcellular level (the middle lamella of the cell wall contains a higher concentration of lignin than the primary or secondary cell walls) (Saka and Goring, 1985). Studies have also demonstrated wide variation in the p-coumaryl, guaiacyl, and syringyl residue composition of lignins from gymnosperms, angiosperms, and grasses (Grand et al. 1982; Higuchi, 1990; Monties, 1985). Lignin composition is also affected by cell type (guaiacyl > syringyl in parenchyma versus vascular tissue) (Monties, 1985; Musha and Goring, 1975), subcellular localization (guaiacyl > syringyl in the middle lamella versus secondary cell wall) (Terashima and Fukushima, 1989), and various environmental stresses (guaiacyl > syringyl in diseased versus healthy tissue) (Vance et al.

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1980). Thus, while it appears impossible to grow plants devoid of lignin in the terrestrial environment, reductions of 10-15% in total lignin content, as well as significant changes in lignin composition, should be attainable without drastic reductions in biomass yield. Employing a synthetic chemistry approach, Freudenberg (1968), Higuchi (1985; 1990) and others have determined that lignin is probably synthesized in an enzyme-initiated, free radical-mediated, dehydrogenative polymerization of three aromatic precursors or monolignols -- p-coumaryl, coniferyl, and sinapyl alcohol. The chemical bonding patterns of the dehydrogenation polymers, or DHPs, synthesized in vitro appear to be random and display no stereospecificity, traits they share with lignins extracted from plant cell walls using a variety of techniques. Nearly all of the linkages found between the aromatic rings in lignin fragments extracted from plant tissues have been reproduced in vitro by manipulating polymerization conditions (Freudenberg, 1968). However, as every technique available for the extraction of lignin from plant tissues appears to alter the structure of lignin to varying degrees, it is unknown how many of these linkages actually occur in the plant (Lai and Sarkanen, 1971; Lewis and Yamamoto, 1990). A further troubling aspect to structural comparisons between lignin in vivo and synthetic DHPs is that DHPs are usually synthesized using plant and fungal oxidative enzymes (peroxidase or laccase) which are not responsible for lignin biosynthesis in vivo. It is perhaps not surprisingly, then, that DHPs display numerous structural differences from "native" lignins (Faix and Schweers, 1974; Lewis et al. 1989; Nimz et al. 1974; Schweers and Faix, 1973). Much of our current thinking about lignin structure and function is based on chemical analyses of aromatic fragments extracted from pulverized wood, as well as the bonding patterns of polymers synthesized under non-physiological conditions with enzymes of unknown relationship to those acting in vivo. This has led to a common description of lignin as a randomly-linked polymer of aromatic rings; however, as pointed out by Lewis and Yamamoto (1990), such an ill-defined structure would be unique among biomolecules. The chemical paradigm for lignin is thus insufficient to represent the reality of lignin as it exists in the plant. Particular care should be taken when illustrating the molecular structure of lignin

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using one of the figures most commonly found in texts (Fig. 1, for example) as it is probably more than a little misleading and, at best, should only be used to illustrate possible chemical bonds within the lignin molecule.

Lignin Biology In contrast to a chemist's view of monolignol free radicals bonding randomly to form lignin, from a biological point of view, tight control of lignin biosynthesis appears imperative. Not only are the monolignols derived from the most bioenergetically-expensive anabolic pathway the cell has to offer (Herrmann, 1983; Kremers, 1957), but free radicals are highly toxic to cells (Elstner, 1982). It seems unlikely that plant cells can afford to waste energy and generate toxic substances in a manner similar to that generally employed to synthesize DHPs in vitro. However, integration of biological observations with the chemical analyses is pointing the way to new thinking about lignin structure and biosynthesis. Through the use of techniques such as ultraviolet microscopy, as well as scanning electron microscopy/energy-dispersive x-ray analysis (SEM/EDXA), lignin biosynthesis has been shown to begin in the corners of the primary cell wall and proceed along the radial walls of the middle lamella to the intercellular region of the corners (Saka and Thomas, 1982; Saleh et al. 1967; Wardrop and Bland, 1959). Lignification of the secondary cell wall then begins in the corners of the S1 layer and proceeds toward the lumen, lagging behind in sites where the active deposition of polysaccharides is occurring (Takabe et al. 1989). This pattern holds for secondary xylem elements, but primary xylem tracheids only deposit lignin in the gyres of thickening, signifying that cells are able to control which parts of the wall become lignified in a very precise manner (O'Brien, 1974). Raman microprobe spectroscopy has shown that rather than being a randomly-ordered structure in vivo, the aromatic rings in lignin are oriented parallel to the cell wall surface (Atalla and Agarwal, 1985). Similarly, nuclear magnetic resonance [NMR] spectroscopy has demonstrated that only specific linkages develop in lignin in vivo, and that the pattern of linkage development appears to vary between plant species (Lewis

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et al 1989). Elegant studies by Terashima and co-workers (Fukushima and Terashima, 1991; Takabe et al. 1989; Terashima, 1989; Terashima et al. 1988) suggest that the specificity for monolignols of the extracellular enzyme complex which assembles the lignin polymer changes during the course of cell wall development; however, the exact mechanism for discrimination between monolignols is still unknown. These particular observations emphasize the role of extracellular processes in regulating the deposition and structure of lignin; however, there is little doubt that the intracellular biosynthetic pathways which produce the monolignols play a major role in determining the composition, and thereby the structure, of lignin. Numerous laboratories are studying regulation of the intracellular, biosynthetic enzymes of the aromatic amino acid (Jensen et al. 1989; Singh et al. 1990), phenylpropanoid (Hahlbrock and Griesebach, 1979), and monolignol pathways (Griesebach, 1981), and initial efforts to alter lignin deposition through genetic engineering will probably attempt to modify enzymes within one of these portions of the biosynthetic pathway (Timmis and Trotter, 1989). Only very limited study has been made of the mechanisms regulating transport and secretion of the monolignols or their glucosides to the apoplasm, even though these processes are likely to play significant roles in tissue lignification (Lewis and Yamamoto, 1990; Takabe et al. 1989; Yamamoto et al. 1989). We will now consider some of the possible roles of extracellular enzymes in controlling lignin quantity and structure, paying particular attention to areas which are in need of clarification. ß-Aryl Glucosidase. Glycosidic derivatives of the monolignols are easily isolated from the sap of many sofftwood species, but are found in trace amounts, if at all, in hardwoods. Because the monolignol glucosides cannot be polymerized directly to form lignin, they have been proposed to function as the primary transport form of the monolignols, both intracellularly and intercellularly (Freudenberg, 1968; Wardrop, 1971). However, recent experiments which determined the turnover rate for coniferin (coniferyl alcohol glucoside) in Norway spruce seedlings suggest that only a portion of the coniferyl alcohol for lignin biosynthesis proceeds through the glucoside pool in these tissues (Marcinowski and Grisebach, 1978). Thus, the monolignol glucosides may not be obligatory for intercellular transport, but they may still

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function in intracellular transport, as well as acting as a storage pool for lignin precursors (Grisebach, 1981; Lewis et al. 1988). ß-Aryl glucosidases (EC 3.2.1.21) need to be present in lignifying tissues if these storage forms of the monolignols are to be mobilized for lignin biosynthesis. Such glucosidases were identified by Freudenberg (1968), and have since been immunolocalized to lignifying cell walls (Burmeister and Hösel, 1981; Marcinowski et al. 1979). Investigations into the substrate specificities of these glucosidases have not demonstrated this to be a significant regulatory mechanism for lignin biosynthesis (Grisebach, 1981; Hösel et al. 1982). However, as coniferin has been shown to accumulate in gymnosperm cambium prior to the start of lignification (Terazawa and Miyake, 1984; Savidge 1988; 1989), it seems likely that the level of aryl ß-glucosidase activity in these tissues must fluctuate in a manner reflective of tissue lignification. Peroxidase. Peroxidases (EC 1.11.1.7) are heme-containing enzymes which mediate the oxidation of a variety of molecules using hydrogen peroxide as an the electron acceptor. Peroxidases are widely distributed in higher plants (Greppin et al. 1986), and have been the enzymes most commonly associated with lignin biosynthesis since Higuchi and Ito demonstrated that plant peroxidases could form DHPs from coniferyl alcohol and H2O2 (Higuchi, 1959; Higuchi and Ito, 1958). Several studies have demonstrated a strong link between peroxidase activity and tissue lignification in vivo (Church and Galston, 1988b; Greppin et al. 1986; Harkin and Obst, 1973; Lewis and Yamamoto, 1990). Peroxidase often occurs as a variety of isozymes which may vary with tissue type, stage of development, or in response to environmental factors such as disease (Bruce and West, 1989; Greppin et al. 1986; Kay and Basile, 1987). Although these isozymes often demonstrate relatively broad substrate specificities, there is strong evidence suggesting that certain isozymes with increased specificity for particular monolignols may influence the composition of native lignin (Church and Galston, 1988b; Imberty et al. 1985; Masuda et al. 1983; Stich and Ebermann, 1988; Tsutsumi and Sakai, 1990). Evidence suggests that some peroxidase isozymes represent different gene products (Mäder,

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1980; Morgens et al. 1990; van Huystee and Cairns, 1982), and four different plant peroxidase genes have been cloned so far (Lagrimini et al. 1987; Morgens et al. 1990; Roberts et al. 1988). However, there is still doubt as to whether any of these are directly involved in lignin biosynthesis. Studies have documented peroxidase localization in isolated and washed cell walls (Goldberg et al. 1983, 1985; Mäder et al. 1975), and a probe based on the protein sequence of one of these "cell-wall peroxidases" was used to isolate a gene for lignin-specific peroxidase (Lagrimini et al. 1987). Susequent work suggests, however, that this peroxidase isozyme may be localized in the vacuole, and becomes tenaciously bound to the cell wall only during tissue maceration (Schlob et al. 1987). Thus, this enzyme is unlikely to be involved in lignin biosynthesis (Church and Galston, 1988b). Efforts to establish direct roles for particular peroxidase isozymes in lignin biosynthesis have suffered from a lack of indisputable localization data, e.g. immunolocalization using monoclonal antibodies (van Huystee, 1987), as well as problems of substrate specificity. Substrate specificity is particularly important given that specific peroxidase isozymes probably exist within cell walls to couple not only the phenolic residues esterified to hemicelluloses, but also the tyrosine residues in extensin precursors (Fry and Miller, 1989; Hartley and Ford, 1989). Lewis and Yamamoto (1990) have correctly pointed out that in order to assign specific peroxidase isozymes to play particular roles in lignin biosynthesis, we must determine at least four parameters: substrate specificity using monolignols, primary structure (sequence), subcellular localization, and temporal correlation with lignification. Regrettably, no study of peroxidase isozymes has yet fulfilled all four of these criteria. Definitive proof will probably result only from genetic engineering experiments which "knock out" synthesis of the specific isozyme, and subsequently change the structure or quantity of lignin in the transformed plant. Hydrogen Peroxide Generation. As mentioned above, peroxidases appear to catalyze lignin polymerization by using H2O2 to oxidize the monolignols which have been secreted from the cell. Hydrogen peroxide has been demonstrated in xylem tissues undergoing lignification (Angelini and Federico, 1989; Sagisaka, 1976), but whereas glycosylated or free monolignols are probably transported

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intracellularly in membrane-delimited vesicles, H2O2 is too reactive to be similarly transported. Reactivity also limits the feasibility of generating H2O2 for lignin biosynthesis at the plasmalemma as it would be difficult to regulate the final concentration of H2O2 arriving to the peroxidase after diffusion through the cell wall. Thus, it seems most likely that H2O2 for lignin biosynthesis is being generated within the cell wall by specific wall-bound enzymes. The current prevailing hypothesis (Gross, 1979; Higuchi, 1990), developed using cell walls isolated from macerated tissues, suggests that malate released by the cell is oxidized by a wall-bound malate dehydrogenase to oxaloacetate with the transfer of electrons to NAD(P)+. Then, in a reversal of the commonly described lignin biosynthetic process, the reduced NAD(P)H combines with an unidentified peroxidase, Mn2+, and a monolignol to produce H2O2 (Elstner and Heupel, 1976; Gross and Janse, 1977; Gross et al. 1977). Several problems appear to exist with this scheme. Although low levels of malate dehydrogenase can be detected in cell walls isolated from some species, such as horseradish, the enzyme cannot be found in cell walls from many other plant species (Goldberg et al. 1985). It is possible, in fact, that the wall-associated malate dehydrogenase activity actually represents a mitochondrial enzyme contaminant released during tissue maceration, similar to the situation previously suggested above for certain peroxidases. Another problem with this model is that use of malate/oxaloacetate as an electron shuttle, such as exists in mitochondria, requires directed transport mechanisms for both metabolites. Although a system for malate secretion exists, there is no evidence for an oxaloacetate uptake system in the apoplast (Goldberg et al. 1985). And finally, there is little or no evidence supporting the existence of NAD(P)+ in the cell wall in vivo (McNeil et al. 1984; Crane and Barr, 1989). An alternative hypothesis for cell wall H2O2 production involves the oxidation of putrescine or polyamines by diamine oxidase (DAO; EC 1.4.3.6) or polyamine oxidase (PAO; 1.5.3.3), respectively (Angelini et al. 1990). The reaction catalyzed by these enzymes requires the respective di- or polyamine and O2 as substrates, and releases H2O2, NH3, and pyrroline as products. Both PAO (Kaur-Sawhney et al. 1981) and DAO (Federico and Angelini, 1986) have been localized in the cell wall, and a recent study

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suggests this localization to be exclusive in corn and pea, respectively (Slocum and Furey, 1990). However, other reports suggest that PAO may be symplastic in some plants (Li and McClure, 1989). The work of Slocum and Furey (1990) also documented a strong association between PAO or DAO and tissues undergoing lignification, as well as localization in the middle lamella of the cell wall, where lignin concentration is the highest. The involvement of polyamines in lignification might explain the inhibition of xylogenesis, but not cell division, after blockage of polyamine biosynthesis in differentiating tissue explants (Phillips et al. 1988). Experimental evidence supports the existence of appropriate transport mechanisms for the substrates and products of DAO and PAO -- diamine and polyamines, NH3, and pyrroline -- in the plasmalemma (Flores and Filner, 1985). The concentration of putrescine and polyamines can approach millimolar levels in the xylem saps of some plants (Friedmann et al. 1986), and these concentrations can vary developmentally and seasonally (Königshofer 1990), just as does lignification. Levels of polyamines, like peroxidase levels and lignin deposition, also vary in response to environmental stresses (Dohmen et al. 1990). Extracellular polyamines become covalently bound to the cell wall in a developmentallyregulated manner (Goldberg and Perdrizet, 1984), and this binding appears to involve pectin and lignin (Mariani et al. 1989; Vallée et al. 1983). Interestingly, polyamines conjugated to phenolic residues resembling the monolignols are widespread in plants (Martin-Tanguy et al. 1978), and these conjugates evidently play a role in the differentiation of certain tissues (Evans and Malmberg, 1989; Wyss-Benz et al. 1990). It is interesting to speculate on whether these conjugates might provide for the co-transport of stoichimetric amounts of polyamines and phenolic residues to the apoplasm for incorporation into the cell wall during lignification. An association between lignin biosynthesis and polyamine metabolism also suggests some possible regulatory interactions with the phytohormone, ethylene (Dean and Mattoo, 1990; Mattoo and Suttle, 1991; Yang and Hoffman, 1984). Ethylene stimulates the lignification of many tissues, and one of its precursors, S-adenosylmethionine (SAM), is important for methylation of the monolignols. Ethylene

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also regulates the activity of key polyamine biosynthetic enzymes, including SAM synthetase, and SAM is a substrate for polyamine biosynthesis, acting as a propylamine donor (Slocum et al. 1984). Ethylene also affects peroxidase activity in a variety of plant tissues (Greppin et al. 1986). Given these relationships, coordinated increases in lignin biosynthesis, putrescine levels, peroxidase activity, and ethylene production in response to various biotic and abiotic stresses may not be surprising. Laccase (Polyphenoloxidase). The first in vitro polymerization of coniferyl alcohol to form a lignin-like DHP used a fungal laccase (p-diphenol: O2 oxidoreductase; EC 1.10.3.1) to catalyze the reaction (Freudenberg, 1965; Higuchi, 1958). Freudenberg and co-workers subsequently demonstrated slow polymerization of monolignols in oxygenated extracts from a gymnosperm (Araucaria excelsa), but a much more rapid polymerization occurred upon the addition of hydrogen peroxide (Freudenberg et al. 1958). In contrast to the studies with Araucaria extracts, partially purified laccase from Norway spruce (Picea abies) cambium extracts was much more effective at polymerizing coniferyl alcohol than was spruce peroxidases (Freudenberg, 1959). From this Freudenberg concluded that the combined actions of laccases and peroxidases are required for lignin biosynthesis in plants. An observation which might be interpreted to support Freudenberg's contention that laccase is important for lignification was made by Siegel et al. (1962), who showed that bean plants grown under conditions of reduced oxygen tension contained less lignin, less fiberous vascular tissue, and were more readily dessicated than were plants grown under ambient conditions. This was the case even though these same plants contained substantially more peroxidase activity. On the other hand, Nakamura (1967) demonstrated that although a laccase purified from exudates of the Japanese laquer tree could oxidize pyrocatechol, hydroquinone, and ascorbic acid, it could not oxidize p-cresol or guaiacol, nor could it form DHP from coniferyl alcohol. These results provided the fundamental basis for dismissal of laccase invovlement in lignin biosynthesis. However, we have recently purified a laccase from sycamore cell suspension cultures (Bligney and Douce, 1983), and in contrast to the results of Nakamura (1967), we have found this enzyme quite capable of oxidizing

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coniferyl alcohol (Sterjiades et al. 1991). Experiments are currently underway to verify the cell wall localization of this enzyme in sycamore trees, as well as in cells grown in suspension culture. Besides the apparent limited distribution of laccases in plants (Higuchi, 1959), negative results from histochemical studies using syringaldezine, a chromogenic substrate for fungal laccases, provided the other primary justification for contentions that laccase does not play a role in lignification (Harkin and Obst, 1973). However, syringaldezine has since been demonstrated not to be a substrate for at least one plant laccase (Joel et al. 1978), and there is at least one report of lignifying tree stem tissue oxidizing syringaldazine in the absence of hydrogen peroxide (Goldberg et al. 1983). Compounding this confusion, recent reports have suggested that syringaldezine is not even a substrate for some peroxidase isozymes associated with lignification (Church and Galston, 1988b; Grison and Pilet, 1985). These problems highlight why the monolignols themselves must be used to determine which enzymes may or may not be involved in lignin biosynthesis. Based on the observations that syringaldazine may not be an appropriate substrate for assessing the lignin biosynthetic capability of enzymes, as well as our results with the sycamore laccase, it appears that the contribution of laccase to lignin biosynthesis may need to be reassessed. Laccases comprise a highly-conserved class of metalloenzymes, the "blue" copper oxidases (Malkin and Malmström, 1970; Mayer, 1987; Mayer and Harel, 1979), which have been subjected to a great deal of biophysical analysis. However, these enzymes have received very little in vivo study, and their function remains a mystery. Laccase could actually play several different roles in lignification. It may be that in highly-lignified portions of the cell wall, such as the middle lamella, the cell may require laccase activity as a complement to peroxidase-catalyzed lignification, as was suggested by Freudenberg (1959). As lignin concentration increases and the middle lamella becomes increasingly hydrophobic, it will exclude water and H2O2 (Northcote, 1989). Under these conditions O2 might be the only oxidant capable of diffusing through previously deposited lignin; thus, requiring laccase, or similar phenoloxidase activity (Badiani et al. 1990), to catalyze lignification of these areas. Although the >110 kD Rhus (Japanese

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laquer) laccase could not polymerize coniferyl alcohol in vitro (Nakamura, 1967), perhaps the enzyme can be activated for lignin biosynthesis in vivo by proteolytic cleavage or deglycosylation. Activation of latent phenoloxidases by similar mechanisms have been documented in some plants (Söderhäll et al. 1989; 1990). In a slightly different scenario, laccase might function under conditions in which extracellular free radical-scavenging metabolites, such as ascorbate, inhibit peroxidase activity. Free Radical Scavenging Mechanisms. The inhibition of peroxidase activity by ascorbate brings us back to the problem of toxicity associated with free radicals (Elstner, 1982). Plant cells have several mechanisms for neutralizing free radicals and activated oxygen species, some of which appear to function in the apoplasm (Castillo and Greppin, 1986; Rennenberg and Polle, 1989). Any of these free radical-scavenging systems would tend to inhibit lignification as we currently envisage it. Thus, with the onset of lignification, the cell must either reduce the overall level of reductants in the apoplasm, or provide a means for reducing them in a localized manner. Some evidence suggests that ascorbate is a major free radical scavenger in the apoplast (Castillo et al. 1987; Polle et al. 1990), and as such, its presence may limit the rate of lignin biosynthesis by neutralizing phenolic radicals in the growing polymer. Ascorbate has been shown to prevent crosslinking of tyrosine residues during the polymerization of the cell wall protein, extensin, and this inhibition was overcome in time by an unidentified oxidative enzyme, ostensibly ascorbate oxidase (EC 1.10.3.3) (Cooper and Varner, 1983; 1984). Free radical scavenging activity by ascorbate might actually be able to affect structural characteristics of lignin by preventing formation of particular types of crosslinks which might normally be formed through the action of long-lifetime phenolic radicals. Alternatively, ascorbate could slow or prevent lignin deposition by reacting directly with hydrogen peroxide destined for the lignin-specific peroxidases (Chinoy, 1984). Ascorbate oxidase, a copper enzyme related to laccase (Ohkawa et al. 1989), is reported to exist primarily in the cell walls of a wide variety of plants (Butt, 1980; Kroneck et al. 1982; Lamport, 1965). Observations that the levels of cell wall-associated ascorbate oxidase increase in elongating cells (Mertz,

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1961) suggest that this enzyme is important for some aspect of cell wall maturation. No biological function has yet been identified for ascorbate oxidase, but activity of this enzyme could possibly affect lignin biosynthesis by altering ascorbate levels. Ascorbate is also a pseudosubstrate for laccase, so it is possible that laccase, too, could function to control ascorbate levels in lignifying cell walls (Joel et al. 1978; Loewus and Loewus, 1987; Mayer, 1987). However, the regeneration of reduced ascorbate by dehydroascorbate reductase (EC 1.8.5.1) activity must also be taken into account in assessing the influence of ascorbate on lignin biosynthesis (Loewus and Loewus, 1987).

Biotechnological Approaches to Modifying Lignin Biosynthesis In vitro Cell Culture. A major impediment to determining the roles of specific enzymes in lignin biosynthesis has been the scarcity of biological systems in which the development of secondary walls and subsequent deposition of lignin (xylogenesis) can be controlled or induced in individual cells (Bengochea et al. 1983). In an elegant system identified by Kohlenbach and Schmidt (1975), developed by Komamine and co-workers (Fukuda and Komamine 1980; Ingold et al. 1988), and refined by Church and Galston (1988a) mechanically-isolated mesophyll cells from young Zinnia elegans can be induced to undergo xylogenesis in a nearly synchronous manner in vitro. This system has been exploited for studies of cytodifferentiation (Burgess and Linstead, 1984; Falconer and Seagull, 1986) and cellulose synthesis (Haigler and Brown, 1986), as well as the effect of phenylalanine ammonia-lyase inhibitors on lignin biosynthesis (Ingold et al. 1990). Perhaps the best correlations between the activity of specific peroxidase isozymes and lignin biosynthesis have been obtained with this system (Fukuda and Komamine, 1982; Church and Galston, 1988b; Masuda et al. 1983). Lignin and lignin-like materials have also been identified in in vitro cultures of Norway spruce (Brunow et al. 1990), loblolly pine (Fukuda et al. 1988), Scots pine (Ramsden and Northcote, 1987), poplar (Venverloo, 1969), sycamore (Carcellar et al. 1971), and black-locust (Fukuda and Kohmoto, 1986). These in vitro systems provide an excellent tool for investigating the enzymes involved in the early stages of lignification; however, these tools have so far

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been underutilized. Chemical Inhibitors. Inhibitor compounds resembling enzyme substrates have been used effectively to demonstrate the function of particular enzymes in complex biological systems. Lignin biosynthesis can be interrupted in the shikimate pathway by N-phosphonomethylglycine (glyphosate) at 5-enol-pyruvylshikimic acid 3-phosphate (EPSP) synthase (Schulz et al. 1990), in the phenylpropanoid pathway by L-α-aminooxy-ß-phenylpropionic acid (AOPP) at phenylalnine ammonia-lyase (PAL) (Amrhein et al. 1983; Ingold et al. 1990), in the monolignol pathway by two organic compounds acting at cinnamyl alcohol dehydrogenase (CAD) (Grand et al. 1985), and at lignin polymerization by 2-fluoroanalogues of ferulic acid and coniferyl alcohol acting on peroxidase (Goldberg et al. 1988). The potential exists to apply specific inhibitors of polyamine biosynthesis (Phillips et al. 1988; Slocum et al. 1984), as well as inhibitors of polyamine oxidation (Bitotnti et al. 1990), as a means to determine whether these metabolites are linked to lignification. Inhibitors of extracellular lignin biosynthetic enzymes would not only be effective for analysis of these components of lignification, but could possibly provide practical agricultural agents (herbicides, lignin content reducers, etc.) which would not require cellular uptake to be effective. Molecular Biology. Genetic transformation of trees using the techniques of molecular biology holds the greatest promise for enabling us to alter lignin biosynthesis in ways that can be conferred to subsequent generations. Useful genetic constructs might include those that reduce lignin content by overexpressing gene products intended to scavenge extracellular free radicals, those that reduce lignin content by underexpressing gene products intended to polymerize monolignols, or those which introduce gene products able to modify the structure of monolignols, e.g. convert coniferyl to sinapyl alcohol in gymnosperms. Transformations leading to overexpression of lignin-associated enzymes have been successful with EPSP synthase to confer herbicide resistance (Schulz et al. 1990), as well as peroxidase (Lagrimini et al. 1990). Underexpression of lignin-associated enzymes using antisense technology (Eguchi et al.1991) has been accomplished with CAD (Grima-Pettenati et al. 1990) and a lignin-

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associated peroxidase (Morohoshi et al. 1991). Work is also proceeding to introduce an angiosperm 5'-Omethyl transferase into a gymnosperm so as to produce greater quantities of sinapyl alcohol and thereby create a softwood containing less-condensed, more easily removed lignin (Bugos et al. 1990). The greatest impediment to producing genetically-engineered trees is the dearth of effective transformation and regeneration systems, particularly for commercially-important softwood species. Transfection of angiosperm trees with Agrobacterium vectors has worked well with Populus (Filatti et al. 1987; Parsons et al. 1986) and Juglans species (McGrananhan et al. 1988). Unfortunately, this techniques has so far been unable to induce more than transient expression of genes introduced into gymnosperms (Loopstra et al. 1990). Recent successes using the microprojectile, or `biolistic', transfection process (Sanford, 1988) on both gymnosperms (McCown et al. 1991) and angiosperms (Merkle et al. 1991; Wilde et al. 1991) suggest that regeneration systems for gymnosperms will soon be the only technological factor limiting application of these techniques. Computer Modeling. Computer modeling of biopolymer structures has accelerated our understanding of their interactions in biological systems (). Although relatively few such studies have focused on lignin (Elder, 1989; Smith et al. 1988), such modeling holds great potential for illuminating the structure of lignin. For instance, it should be possible to illustrate the constraints placed on extended lignin molecules by the π-π orbital interactions of adjacent polymers. Such interactions are very likely responsible for the planar alignment of phenolic rings demonstrated in cell wall lignin by Raman microprobe spectroscopy (Atalla and Agarwal, 1985), and may well explain the disk-like structure of lignin fragments leached from various fiber pulps (Favis et al. 1984; Willis et al. 1987).

Summary In a fine review of lignin structure and DHP synthesis, Nakatsubo (1981) proposed three historic periods for our understanding of lignin structure: lignin as `encrusting material' (1838-1897), dehydrogenation theory (1897-1942), and justification of dehydrogenation theory (1942-1981). We are

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now poised on the brink of a fourth period in which we define the biological structure of lignin. Advances in non-invasive, analytic chemistry techniques, such as multi-dimensional NMR spectroscopy (), and microscopy techniques, such as confocal laser scanning microscopy (Knebel and Schnepf, 1991), as well as the ability to remove or inhibit specific lignin biosynthetic enzymes, will allow us to dissect the structure of lignin as it exists within the plant. With such knowledge in hand, we will be able to reconstruct trees and other plants to bear more fruit, yield better timber, and provide stronger fiber with less disruption to our environment.

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