Lignins, complex and irregular polymers present in the cell walls of vascular plants, are built from three basic monolignols. An understanding of their nature is evolving as a result of detailed structural investigations aided by improvements in analytical methodologies and the availability of mutant and transgenic plants. Oxidative phenolic coupling reactions, where monomers primarily couple endwise with the growing chain, generate the polymer. The combinatorial linkage synthesis, the random generation of new optical centres each time a monolignol couples via its sidechain, and the inclusion of monomers other than the monolignols, cascade to create polymers with enormous variation in primary structure. Lignification is a strategic process that has evolved to allow plants considerable flexibility in dealing with various environmental stresses. The malleability offers significant opportunities to engineer the structures of lignins beyond the limits explored to date.

Keywords: biosynthesis; lignification; monolignol; oxidative coupling; polymerization

Figure 1.

Phenylpropanoid and monolignol biosynthetic pathways. The blue‐highlighted route towards the production of monolignols is considered to be most favoured in angiosperms. The lighter blue routes also occur, depending on the species and conditions. Nonhighlighted pathways do not play a significant role. CAD, cinnamyl alcohol dehydrogenase; 4CL, 4‐coumarate:CoA ligase; C3H, p‐coumarate 3‐hydroxylase; C4H, cinnamate 4‐hydroxylase; CCoAOMT, caffeoyl‐CoA O‐methyltransferase; CCR, cinnamoyl‐CoA reductase; COMT, caffeic acid O‐methyltransferase; HCT, p‐hydroxycinnamoyl‐CoA:d‐quinate or shikimate p‐hydroxycinnamoyltransferase; F5H, ferulate 5‐hydroxylase; PAL, phenylalanine ammonia‐lyase; SAD, sinapyl alcohol dehydrogenase. ?, conversion demonstrated; ??, direct conversion not convincingly demonstrated; F5H?, substrate not tested; others, enzymatic activity shown in vitro. Other pathways continue to be revealed. Modified from Boerjan ().

Figure 2.

Lignification differs substantially from dimerization of monolignols. (a) Dehydrodimerization of coniferyl alcohol produces three dehydrodimers in comparable amounts. The new bond formed by the radical coupling reaction is drawn bolder. (b) Dehydrodimerization of sinapyl alcohol produces only two products. (c) Dehydrodimerization does not produce these structures. (d) Cross‐coupling of a hydroxycinnamyl alcohol with a G unit gives only two main products. (e) Cross‐coupling of a hydroxycinnamyl alcohol with an S unit leads almost exclusively to β‐ether units A. When the polymer phenolic end unit is a β‐ether, β–1‐coupling may also occur to a relatively minor extent. (f) Coupling of preformed oligomers is the source of the 5–5‐ and 4–O–5 units. Red arrows indicate sites at which further radical coupling can occur during lignification; the lighter arrow to the 5‐positions in (d) and (e) structures shows where coupling can occur in G units but not in S units (where the 5‐position is occupied by a methoxyl group).

Figure 3.

Lignin precursors, and structures in the polymer (Boerjan et al., ). (a) Monomers’ (Lignin Precursors): Lignins derive primarily from the three monolignols M1, namely M1h, M1g and M1s (where the subscripts indicate the type of aromatic nucleus, p‐hydroxyphenyl, guaiacyl or syringyl, resulting from incorporation of the monomer). M15H is a monomer in COMT‐deficient plants resulting in 5‐hydroxyguaiacyl units in the form of benzodioxanes J in the polymer. Other precursors M2M12 incorporate into lignins in varying degrees. Bracketed compounds have not been firmly established as authentic monomers or, in the case of M12, are of unknown derivation. (b) Lignin Polymer Units: Units are generally denoted based on the methoxyl substitution on the aromatic ring as H, G, S (and 5H); dashed bonds represent other potential attachments (via coupling reactions). The most common structures in lignins from normal and transgenic plants are shown as structures AL with the bond formed during the radical coupling step; p‐hydroxyphenyl units are not shown. The dashed bonds indicate substitutions by methoxyl (in syringyl components) or other attachments from coupling reactions; generic side‐chains are shown truncated (zigzag lines). Most units arise from cross‐coupling reactions of a monomer with the growing polymer or by polymer–polymer coupling reactions. Resinol units C are from monolignol–monolignol coupling (followed by further cross‐coupling reactions). Most 5–5‐linked units D are in the form of dibenzodioxocins D1. bis‐Aryl ether units A2 are rare in most lignins, but relatively prevalent in tobacco. Units F, β–1‐structures occur mainly as spirodienones, but may partially cleave to give units F2. Benzodioxanes J result from the incorporation of 5‐hydroxyconiferyl alcohol M15H monomers (see Figure ). Units K (from coupling of hydroxycinnamaldehydes M2) are prevalent in CAD‐deficient angiosperms. Units L are from ferulate incorporation in grass lignins, for example. Note that hydroxycinnamates and ferulates typically have their side‐chain carbons labelled 7–9, whereas the hydroxycinnamyl alcohols are labelled α, β and γ. Endgroups arise from coupling reactions that are not at the side‐chain β‐position. Hydroxycinnamyl endgroups X1 arise from dimerization reactions. Endgroups X2X6 derive from the corresponding monomers M2M6; X6b may result from oxidation of X6 units. Glycerols X7 may be from monomers M7 or may be produced during ball milling from β‐ether units A. Any of the units AL bearing a γ‐OH may also bear an acyl group, partial structures Y9Y11, and arise from the corresponding monolignols M9M11. Finally, some other groups resulting from incorporation reactions are not accommodated by the other structures. Partial structures Z8 are from incorporation of monomer M8; general aldehydes Z2 are from hydroxycinnamaldehyde M2 or hydroxybenzaldehyde monomers M3 and include structures K; general esters Z4 result from the incorporation of hydroxycinnamates M4 and their dehydrodimers and include structures L. Modified from Boerjan ().

Figure 4.

Lignin Polymer Models for (a) a softwood (spruce) lignin with 25 units, redrawn from (Brunow, ), and (b) a hardwood (poplar) lignin with 20 units, redrawn from (Boerjan et al., ). Colour coding is uniform across the two models. Bold, black bonds indicate the bonds formed by radical coupling during lignification; lighter (grey) bonds result from postcoupling internal rearomatization reactions; α‐OH groups from nucleophilically added water assume the colours of their parent structure. The softwood lignin is more branched and contains a lower proportion of β‐ether units A. The branch points (4–O–5‐units E, orange; dibenzodioxocin units D, dark blue) are differentiated by unique colouring, even though such units may also be β‐ethers, for example. Note that each of these structures represents only one of billions of isomers (Ralph et al., ). Caution: these are ONLY MODELS! They do not imply any primary structure or sequencing in the lignins themselves but attempt to accommodate the main linkage types and their approximate relative frequencies. Modified from Brunow G and Ralph et al. .

Figure 5.

Partial poplar lignin NMR HMQC spectra (at 360 MHz) showing major lignin peaks and highlighting new peaks for benzodioxane units J in COMT‐downregulated plants. Acetylated lignins were from (a) a control poplar, (b) a COMT‐downregulated poplar transgenic. (c) Scheme for production of benzodioxanes J in lignins via incorporation of 5‐hydroxyconiferyl alcohol M15H into a guaiacyl lignin. Lignin unit designations are the same as in Figures . Modified from Ralph J et al. (2001) Elucidation of new structures in lignins of CAD‐ and COMT‐deficient plants by NMR. Phytochemistry 57: 993–1003.



Atalla RH and Agarwal UP (1985) Raman microprobe evidence for lignin orientation in the cell walls of native woody tissue. Science 227: 636–638.

Boerjan W, Ralph J and Baucher M (2003) Lignin biosynthesis. Annual Reviews in Plant Biology 54: 519–549.

Brunow G (2001) Methods to Reveal the Structure of Lignin. In: Hofrichter M and Steinbüchel A (eds) Lignin, Humic Substances and Coal, vol. 1, pp. 89–116. Weinheim: Wiley‐VHC.

Donaldson LA (1994) Mechanical constraints on lignin deposition during lignification. Wood Science Technology 28: 111–118.

Freudenberg K and Neish AC (1968) Constitution and Biosynthesis of Lignin. Berlin‐Heidelberg‐New York: Springer‐Verlag.

Huntley SK, Ellis D, Gilbert M, Chapple C and Mansfield SD (2003) Significant increases in pulping efficiency in C4H‐F5H‐transformed poplars: Improved chemical savings and reduced environmental toxins. Journal of Agricultural Food Chemistry 51: 6178–6183.

Lapierre C, Pilate G, Pollet B et al. (2004) Signatures of cinnamyl alcohol dehydrogenase deficiency in poplar lignins. Phytochemistry 65: 313–321.

Ralph J, Akiyama T, Kim H et al. (2006) Effects of coumarate‐3‐hydroxylase downregulation on lignin structure. Journal of Biological Chemistry 281: 8843–8853.

Ralph J, Bunzel M, Marita JM et al. (2004) Peroxidase‐dependent cross‐linking reactions of p‐hydroxycinnamates in plant cell walls. Phytochemistry Reviews 3: 79–96.

Reddy MSS, Chen F, Shadle GL et al. (2005) Targeted down‐regulation of cytochrome P450 enzymes for forage quality improvement in alfalfa (Medicago sativa L.). Proceedings of the National Academy of Sciences of the USA 102: 16573–16578.

Terashima N, Fukushima K, He L‐F and Takabe K (1993) Comprehensive model of the lignified plant cell wall. In: Jung HG, Buxton DR, Hatfield RD and Ralph J (eds) Forage Cell Wall Structure and Digestibility, pp. 247–270. Madison, WI: ASA‐CSSA‐SSSA.

Further Reading

Baucher M, Halpin C, Petit‐Conil M and Boerjan W (2003) Lignin: Genetic engineering and impact on pulping. Critical Reviews in Biochemistry and Molecular Biology 38: 305–350.

Brunow G, Lundquist K and Gellerstedt G (1999) Lignin. In: Sjöström E and Alén R (eds) Analytical Methods in Wood Chemistry, Pulping, and Papermaking, pp. 77–124. Germany: Springer‐Verlag.

Halpin C and Boerjan W (2003) Stacking transgenes in forest trees. Trends in Plant Science 8: 363–365.

Morreel K, Ralph J, Kim H et al. (2004) Profiling of oligolignols reveals monolignol coupling conditions in lignifying poplar xylem. Plant Physiology 136: 3537–3549.

Pilate G, Guiney E, Holt K et al. (2002) Field and pulping performances of transgenic trees with altered lignification. Nature Biotechnology 20: 607–612.

Ralph J, Lundquist K, Brunow G et al. (2004) Lignins: natural polymers from oxidative coupling of 4‐hydroxyphenylpropanoids. Phytochemistry Reviews 3: 29–60.

Sarkanen KV and Ludwig CH (1971) Lignins, Occurrence, Formation, Structure and Reactions. New York: Wiley‐Interscience.

Whetten R and Sederoff R (1995) Lignin Biosynthesis. Plant Cell 7: 1001–1013.

Zhang L, Henriksson G and Gellerstedt G (2003) The formation of β–β structures in lignin biosynthesis – are there two different pathways? Organic and Biomolecular Chemistry 1: 3621–3624.

Contact Editor close
Submit a note to the editor about this article by filling in the form below.

* Required Field

How to Cite close
Ralph, John, Brunow, Gösta, and Boerjan, Wout(Sep 2007) Lignins. In: eLS. John Wiley & Sons Ltd, Chichester. [doi: 10.1002/9780470015902.a0020104]