Phytochemistry 70 (2009) 147–155

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Phytochemistry

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On the role of the monolignol c-carbon functionality in lignin biopolymerization

Anders Holmgren a, Magnus Norgren b,c, Liming Zhang a, Gunnar Henriksson a,*

a Division of Wood Chemistry, Department of Fibre and Polymer Technology, Royal Institute of Technology, KTH, Teknikringen 56, Stockholm, Swedenb Division of Fibre Technology, Department of Fibre and Polymer Technology, Royal Institute of Technology, KTH, Stockholm, Swedenc Centre for Fibre Science and Communication Network, Mid Sweden University, Sundsvall, Sweden

a r t i c l e i n f o

Article history:Received 2 November 2007Received in revised form 6 October 2008Available online 4 December 2008

Keywords:LigninMonolignol biosynthesisLignin-polysaccharide networksPlant cell wallBiopolymer

0031-9422/$ - see front matter � 2008 Elsevier Ltd. Adoi:10.1016/j.phytochem.2008.10.014

* Corresponding author. Tel.: +46 87906163; fax: +E-mail address: ghenrik@polymer.kth.se (G. Henri

a b s t r a c t

In order to investigate the importance of the monomeric c-carbon chemistry in lignin biopolymerizationand structure, synthetic lignins (dehydrogenation polymers; DHP) were made from monomers with dif-ferent degrees of oxidation at the c-carbon, i.e., carboxylic acid, aldehyde and alcohol. All monomersformed a polymeric material through enzymatic oxidation. The polymers displayed similar sizes by sizeexclusion chromatography analyses, but also exhibited some physical and chemical differences. The DHPmade of coniferaldehyde had poorer solubility properties than the other DHPs, and through contact angleof water measurement on spin-coated surfaces of the polymeric materials, the DHPs made of coniferal-dehyde and carboxylic ferulic acid exhibited higher hydrophobicity than the coniferyl alcohol DHP. Astructural characterization with 13C NMR revealed major differences between the coniferyl alcohol-basedpolymer and the coniferaldehyde/ferulic acid polymers, such as the predominance of aliphatic doublebonds and the lack of certain benzylic structures in the latter cases. The biological role of the reductionat the c-carbon during monolignol biosynthesis with regard to lignin polymerization is discussed.

� 2008 Elsevier Ltd. All rights reserved.

1. Introduction

Lignin is a main constituent of the cell walls of vascular plants,especially in woody tissues where it is present in high amounts. Itacts as a binding agent, provides compressive strength andbending stiffness (Sederoff and Chang, 1991), functions as aphysico-chemical barrier against pathogens (Baucher et al.,1998), and conveys hydrophobicity to the cell wall, inhibitingswelling and boosting water transport through the cell lumen(Sederoff and Chang, 1991; Brett and Waldron, 1996). From achemical point of view, lignin is a branched polymer built mainlywith p-hydroxycinnamyl alcohols with different degrees of meth-oxylation (also called monolignols: p-coumaryl alcohol, coniferylalcohol and sinapyl alcohol; Fig. 1a). Moreover, lignin is believedto be shaped like a network that connects different cell wall poly-mers (Lundquist et al., 1983; Xie et al., 2000; Lawoko et al., 2006).

The polymerization of monolignols into lignin involves an oxi-dation step that is catalyzed by peroxidases, creating resonance-stabilized radicals, and an uncatalyzed radical–radical couplingstep. The polymer grows as its free phenolic groups are oxidizedand couple new radicals (Fig. 1b). In plants, the monolignols arebiosynthesized from the amino acid phenylalanine through thephenylpropranoid pathway (Boerjan et al., 2003) (Fig. 2); as one ofthe building blocks of proteins it is ubiquitous in all life forms.

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46 87906166.ksson).

Generally, the following chemical modifications are carried outon the amino acid in the pathway to becoming a monolignol:

1. The amino group is removed; simultaneously, a double bond isintroduced between the a- and b-carbons.

2. Carbons 3 and 5 may be methoxylated via a hydroxylation andcarbon 4 is hydroxylated only.

3. The carboxylic acid at the c-carbon is reduced first to an alde-hyde and then to an alcohol.

The reasons for the first two modifications are rather obvious;by removing the amino group and introducing the double bond,nitrogen, which is often a limiting nutrient for plants, is recoveredfor other uses. In addition, the double bond gives possibilities forradical stabilization and radical couplings, while the methoxyla-tions are a way for the plant to regulate the coupling pattern bysterically blocking formation of bonds to the 50 and 30 carbons ofthe aromatic ring. However, the reason for reducing the monomersat the c-carbon is less obvious, since this carbon is not directly in-volved in the polymerization, except in stabilizing ether bonds ine.g., the resinol structure. Nevertheless, this reduction probablyconveys an important function. For each biosynthesized monolig-nol, two nicotinamide adenine dinucleotide phosphates (NADPHs)and one adenosine trisphosphate (ATP) are consumed to reducethe carboxylic acid to the aldehyde and the aldehyde to the alcohol(Fig. 2) (Sjöström, 1981). Taking into account the fact that 15–36%or more of the cell wall in woody plants consists of lignin, a

Fig. 2. The monolignol biosynthesis pathway. The starting substance is the amino acid phenylalanine. This is subjected to three major modifications in order to form themonolignols: first, the removal of the amino group with the introduction of a double bond (1); second, the introduction of methoxy groups via hydroxylations (2, 3); and third,reduction at the c-carbon carboxylic acid to an aldehyde (4) and eventually to an alcohol (5). Observe that reactions 4 and 5 consume one NADPH each. One ATP is alsoconsumed in a ‘‘preparation stage” for reaction 4, where coenzyme A forms a temporary thioester to the c-carbon, thereby activating it for reduction (6). In dashed boxes arethe monomers used in this study.

Fig. 1. (a) The most common lignin monomers, the monolignols. (b) Simplified pathway for the formation of lignin through radical coupling of a monolignol (here coniferylalcohol) to a phenolic end-group forming the most common lignin inter-unit bond, b-O-40 . Monolignols or phenolic end-groups are oxidized, probably by peroxidases, toresonance-stabilized radicals. Next, the radicals couple to each other in an uncatalyzed step. The positions that can form covalent bonds are marked with bold arrows.Observe that the c-carbon is not involved in the radical–radical coupling and that the 3 and 5 positions will be available depending on the type of monolignol (see a). Radicalcoupling can occur not only between monomer and oligomer but between oligomers.

148 A. Holmgren et al. / Phytochemistry 70 (2009) 147–155

considerable part of the reducing power and free energy producedduring photosynthesis is consumed in these reductions. Given thatsome of the aldehyde and carboxylic acid products of the phenyl-

propanoid pathway can polymerize through the same basic stepsas the monolignols (dehydrogenative polymerization), are thereany advantages to an alcohol-based lignin?

Table 1Molecular weight averages relative to polystyrene standards from SEC analysis ofacetylated DHP products.

Polymer Mw Mn P MP

Ferulic acid DHP 4489 3395 1.3 5017Coniferaldehyde DHP 2709 1898 1.4 2222Coniferyl alcohol DHP 4464 3206 1.4 2725

Mw, molecular weight average (g/mol); Mn, Number average molecular weight(g/mol); P = Mw/Mn, polydispersity; MP, peak molecular weight (g/mol).

A. Holmgren et al. / Phytochemistry 70 (2009) 147–155 149

In this study, we generate dehydrogenation polymers (DHPs)from coniferyl alcohol, coniferaldehyde and ferulic acid (Fig. 2),and characterize the products in order to compare them and inves-tigate the biological reasons for the reduction at the c-carbon.

2. Results

2.1. DHP synthesis

Dehydrogenated polymers (‘‘synthetic lignin”; DHP) from feru-lic acid, coniferaldehyde and coniferyl alcohol (Fig. 2) were synthe-sized by peroxidase oxidation. After extraction of low molecularweight compounds, the yields were 58%, 47% and 35%, respectively.The relatively low coniferyl alcohol DHP yield could be due to ten-dency of coniferyl alcohol to precipitate at the beginning of thereaction in the conditions used here. Interestingly, the ferulicacid-based DHPs did not precipitate during the reaction, in con-trast to the other two DHPs.

2.2. Size exclusion chromatography (SEC)

Size exclusion chromatography analysis (Fig. 3) revealed thatthe three DHPs contained dispersed polymeric material. Theconiferyl alcohol and coniferaldehyde DHPs displayed a similarpolydispersity (Table 1, Fig. 3), while the ferulic acid DHP showeda somewhat lower polydispersity. The molecular weight average ofthe coniferyl alcohol and the ferulic acid DHPs were similar, butthe molecular weight of the peak (MP) of the latter was almosttwice the MP of the other DHPs, indicating that the ferulic acidpolymerization generally produced large populations of largerpolymers. This could be partly explained by the polymers stayingin solution throughout the reaction, making the larger speciesavailable for further polymerization, as opposed to the other DHPs,which commonly precipitated. Moreover, the SEC analysis of theconiferaldehyde DHP revealed that there was a significant amountof low molecular weight species in the coniferaldehyde DHP,meaning they might not have been washed out as effectively asin the case of the two other DHPs.

2.3. Solubility panel

The DHPs were characterized with a solvent panel (Table 2).Generally, the coniferaldehyde DHP had the lowest solubility,while the ferulic acid and coniferyl alcohol DHPs showed rathersimilar patterns, although there was a tendency for the DHP madeof ferulic acid to be more soluble in polar solvents, and the DHP

Fig. 3. SEC chromatogram of acetylated DHP products. See Table 1 for molecularweight approximations.

made of coniferyl alcohol to be more soluble in apolar solvents (Ta-ble 2). Thus the DHP products displayed different solubilities; theconiferaldehyde DHP in particular was resistant to complete disso-lution in any of the solvents.

2.4. NMR analyses

All three DHPs were analyzed with 13C NMR, and the spectra areshown in Fig. 4. A more complete and unambiguous structuralcharacterization, although relevant, is outside the scope of thisstudy. Instead, signal assignments are suggested for several sub-structures (Table 3 and Fig. 5) based on both reported values forsynthesized model components and NMR studies on isolated lig-nins (Ralph et al., 1994, 2004; Landucci et al., 1998; Kim et al.,2003). Several observations were made from the comparison ofthe three spectra:

� The different DHPs could generally form the same type of bond-ing structures: b-O-40, b-50, b–b0 and 5–50.

� Although equivalent bonding structures were found in all DHPs,chemical differences were obvious when comparing the dc 60–90 ppm region of the three spectra. The coniferyl alcohol DHPdisplayed a number of signals in this region corresponding tosaturated a, b and c carbons, while the ferulic acid and conifer-aldehyde DHPs almost completely lacked any such signals. Sat-urated side-chains are formed when coniferyl alcohol radicalcoupling occurs through at least one b-carbon, i.e., b-O-40, b-50

and b–b0. Instead, although such coupling occurred in the DHPsfrom ferulic acid and coniferaldehyde, respectively, the struc-tures formed were predominantly unsaturated (Fig. 5) and theirsignals appear in the 100–140 ppm region.

� There is a general lack of cyclic structures (e.g., b-50 and b–b0) inboth the ferulic acid and coniferaldehyde DHPs compared to theconiferyl alcohol DHP. The ferulic acid DHP did display somesignals, although weak, for possibly cyclic unsaturated b-50 andb–b0.

� The ferulic acid and coniferaldehyde DHPs lacked benzyl alcoholgroups, i.e., OH-groups on the a-carbon, and non-cyclic benzylaryl ether structures, both of which are typical of coniferyl alco-hol DHPs (Freudenberg, 1959; Adler, 1977). In lignins, benzylalcohols are very common as they are formed during polymeri-zation from the addition of water to the quinone methide inter-mediate of the most abundant inter-monomeric structure: theb-O-40 bond. Non-cyclic benzyl aryl ethers are also present in lig-nins but generally in lower quantities compared to DHPs (Edeand Kilpeläinen, 1995).

2.5. Contact angle measurements on DHP model surfaces

Fig. 6 shows AFM images of very smooth spin-coated DHP mod-el surfaces. As a relative measure of the hydrophobicity of the dif-ferent polymeric materials, the contact angle of water wasdetermined on the DHP model surfaces. The results (Table 4) indi-cated that the coniferyl alcohol DHP was less hydrophobic than the

Table 2Ocular inspection of the solubility of the different unmodified DHP materials in organic and aqueous solvents with increasing dielectric constants (excluding dioxane:water andacetone:water, which are mixes of solvents at 1:1 ratio). +/� means partly soluble. EtoAc is ethyl acetate.

Polymer Dioxane EtoAc THF CH2CL2 Pyridine Acetone DMF DMSO Water Dioxane:water Acetone:water

Ferulic acid DHP No No Yes No Yes +/� Yes Yes No No YesConiferaldehyde DHP No No No No +/� No +/� Yes No No NoConiferyl alcohol DHP +/� No No No Yes No Yes Yes No Yes Yes

Fig. 4. 13C NMR spectra of (a) ferulic acid DHP; (b) coniferaldehyde DHP and (c) coniferyl alcohol DHP in DMSO-d6. Suggestions for the signal assignments are found in Table 3and those for the types of inter-unit structures in Fig. 5.

150 A. Holmgren et al. / Phytochemistry 70 (2009) 147–155

coniferaldehyde and the ferulic acid DHPs. All three DHPs had amuch higher hydrophobicity than was previously determined forkraft lignin (Norgren et al., 2006), but this was expected sincethe latter is a lignin derivative where both free phenolic groupsand carboxyl groups are present. There are reasons to believe thatthe hydrophobicity of the coniferaldehyde DHP is somewhatunderestimated, since this material was incompletely dissolvedin pyridine, the solvent used for surface preparation. Therefore,the fraction of the DHP that was dissolved by the pyridine and usedfor the model surface preparation most likely represented a some-what lower molecular weight DHP that was less hydrophobic.

3. Discussion

In summary, our investigation has demonstrated that mono-mers with carbonyl c-carbons can form dehydrogenated polymerswith generally the same types of inter-unit bonds as natural lig-nins. Although synthetic polymers from coniferaldehyde and feru-lic acid have been produced through dehydrogenation in variousworks (Karmanov et al., 1991; Higuchi et al., 1994; Ward et al.,2001), no comparison of physical and chemical properties of theDHP products with the more common coniferyl alcohol DHP hasbeen presented to our knowledge. In this work, we raise the ques-tion of why the alcohol-type monomers are seemingly preferred inlignin biosynthesis and suggest some answers based on the com-parison of the different DHPs obtained.

The DHP made of ferulic acid seemed even to form somewhatlarger polymers than the DHP from coniferyl alcohol, a commonmonolignol (Fig. 1), most likely due to the charges on the carbox-ylic acid groups, which increase the solubility (and thereby thereactivity) of lignin oligomers. Thus, there is no reason to believethat reduction of the monolignol precursors at the c-carbon is nec-essary to obtain a polymer. We investigated the hydrophobicity/hydrophilicity of the polymers by a solvent panel and by contactangle measurement of DHP surfaces. The results indicated thatconiferyl alcohol DHP was the least hydrophobic. Therefore, thereduction at the c-carbon could be a way to make the ligninslightly more hydrophilic. This is surprising for two reasons. First,one of the biological roles of lignin is to make the cell wall hydro-phobic and waterproof, which is probably a prerequisite for thedevelopment of water conducting tissues in plants. However, a lig-nin structure that is too hydrophobic may interact poorly with cel-lulose and hemicelluloses, and therefore not fulfill its biologicalfunctions efficiently. Second, in contrast to the primary alcohol ofconiferyl alcohol, the carboxylic acid in ferulic acid is a chargedstructure at physiological pH. Thus, it was expected that the DHPmade of ferulic acid would be considerably more hydrophilic thanthe DHP made from coniferyl alcohol, not the opposite as was seenin our results.

The NMR characterization demonstrated that, in addition to theexpected differences from the c-carbon signals (carboxylic acidsfor the ferulic acid DHP, aldehydes for the coniferaldehyde DHP

Table 3Signal assignments from 13C NMR spectra of ferulic acid (FA), coniferaldehyde (CAld) and coniferyl alcohol (CA) DHPs in DMSO-d6.

Signal d Assignments

Peaks in 13C NMR, ferulic acid DHP1 171.7 Cc in b-50 c2 170.4 Carbonyl carbon from ethyl acetate (solvent)3 167.8 Cc0 (FA) in b-O-40 , Cc in b–b0 nc; Cc and Cc0 (FA) in b-50 c4 150.0 C3 in 4-O-50

5 149.1 C30 and C3 in 4-O-50 and C40 in b-50 c; C30 and C4 in b-O-40

6 147.7 C3, C40 in b-O-40; C4 and C30 in b-50 nc; C3 in b-50; C3 in 5–50; C4 in 4-O-50

7 147.1 C3 and C40 in b-O-40; C4 in b–b0 c; C3 in b–b0 nc; C40 in b-50 nc; C4 in b-50 c8 145.4 C3 in b-50 nc9 144.2 Ca0 (FA) in b-50 nc; Ca in 5–50; Ca0 (FA) and C30 in b-50 c10 143.8 Ca0 (FA) in b-O-40; Ca0 (FA) and C30 in b-50 c, Ca and Ca0 (FA) 4-O-50

11 138.7 Ca in b-50 nc; Cb in b-O-40; C1 in b-5 c12 129.5 C1 in b�b0

13 128.4 C10 in b-O-40; C1 in b–b0 c14 126.7 Ca in b-O-40; Cb in b-50 nc15 124.2 C6 in b-O-40; C6 in b–b0 nc; C50 and C60 in b-50 nc16 121.8 C60 in b-O-40; C6 in 4-O-50

17 118.1 C60 in b-50 c18 116.9 Cb0 (FA) in b-50 c; Cb0 (FA) in 4-O-50; Cb0 (FA) in b-O-40

19 115.6 C5 in b-50 nc20 115.2 C5 in b-O-40; C5 in b–b0 c; C5 in b–b0 nc; C5 in b-50 c21 112.4 C2 in b-O-40; C2 in b-50 nc; C20 in b-50 c; C2 in 4-O-50

22 111.5 C20 in b-O-40; C2 in 4-O-50

23 110.3 C2 in b–b0 c; C20 in b-50 nc; C2 in b-50 c24 86.7 Ca in b-50 c25 80.3 Ca in b–b0 c26 73.5 Cb in b-O-40 trimer (see b1 in Fig. 6)27 59.8 CH2–C@O ethyl acetate (solvent)28 55.9 Methoxy carbons29 52.7 Cb in b-50 c; b in b–b0 c30 39.4 Methyl carbon, DMSO

Peaks in 13C NMR, coniferaldehyde DHP1 194.3 Cc0 (CAld) in b-O-40; Cc and Cc0 (CAld) in b-50 nc; Cc0 (CAld) in b-50 c2 193.1 Cc0 (CAld) in b-50 c; Cc in b–b0

3 163.3 Unknown4 153.2 Ca0 (CAld) in b-O-40; Ca0 (CAld) in b-50 c; Ca in b–b0

5 152.9 Ca0 (CAld) in b-O-40; Ca in b–b0

6 150.0 C3 in b-O-40; C40 in b-50 nc; C4 in b–b0; C5 in 5–50

7 147.6 Cb in b-O-40; C3 in b-50 nc; C3 and C30 in b-50 c8 134.2 Cb in b�b0

9 132.6 C1 in b-50 c10 128.8 Cb0 (CAld) in b-O-40

11 126.5 C6 in b-O-40; C6 and C60 in b-50 nc; C6 in b–b0 , b in 5–50

12 125.8 C1 in b-O-40; C60 in b-50 nc; C50 in b-50 c; C6 in b–b0 , C4 in 5–50

13 125.1 C1 in b-O-40

14 123.2 Cb in b-50 nc; C60 in b-O40

15 115.9 C5 in b-O-40; C5 in b-50 nc; C5 in b-50 c; C5 in b–b0

16 113.6 C2 in b-O-40; C2 in b-50 nc; C2 in b–b0

17 111.5 C20 in b-O-40; C20 in b-50 nc; C2 in b-50 c18 109.5 C2 in 5–50

19 55.6 Methoxy carbons20 39.2 Methyl carbon, DMSO

Peaks in 13C NMR, coniferyl alcohol DHP1 194.2 Cc, coniferaldehyde2 152.9 C30 in b-O-40; phenolic C3 in 5–50

3 151.8 C40 in b-O-40 , C30 in a-O-40

4 150.7 Ca in coniferaldehyde5 149.8 C40 in b-50; CA C30 in b-O-40; C30 in 4-O-50

6 147.5 C40 in b-O-40; C40 in a-O-40; C3 in b–b0

7 147.1 C3 in b-O-40 a-O-40 b-O-50

8 146.1 C4 in a-O-40; C4 in a-O-40 in b-50; C4 in b–b0

9 143.7 C30 in b-50; C4 in 4-O-50

10 143.1 Unknown11 138.1 C4 in 5–50

12 136.5 C1 and C10 in b-O-40 , b-50; C1 in 5–50

13 134.6 Ca0 (CA) in b-50

14 132.2 C1 in b-O-40 , b-50 and b–b0; C5 in 5–50

15 130.6 C10 in b-O-40 and b-50; C1 in a-O-40

16 129.0 Ca0 (CA) in b-50; C50 (CA) in b-50

17 128.6 Ca0 (CA) in b-O-40 and a-O-40

18 128.1 Cb0 (CA) in b-O-40 and a-O-40

19 125.4 C1 in 5–50

20 120.4 C6 in b-O-40

21 119.0 C6 in b–b0

(continued on next page)

A. Holmgren et al. / Phytochemistry 70 (2009) 147–155 151

Table 3 (continued)

Signal d Assignments

22 118.6 C6 in b-50

23 116.1 C50 in b-O-40 and a-O-40

24 115.0 C50 and C5 in b–b0; C60 in b-50

25 111.9 C2 in b-O-40

26 110.4 C2 and C20 in b-50 and b–b0

27 109.9 C20 in b-O-40 and a-O-40

28 105.1 Unknown29 103.6 Unknown30 86.9 Ca0 (CA) in b-50

31 85.2 Ca0 (CA) in b–b0

32 82.6 Cb0 (CA) in b-O-40; Cb in a-O-40

33 79.0 Ca in a-O-40

34 71.0 Ca in b-O-40; c in b–b0

35 67.0 Unknown36 63.0 Cc in b-50

37 61.7 Cb0 (b carbon in CA endgroup)38 59.8 Cc in b-O-40 and a-O-40

39 55.7 Methoxyl C40 53.6 Cb in b–b0

41 53.1 Cb in b-50

42 39.8 Methyl carbon, DMSO

c, cyclic; nc, non-cyclic. (FA) (CAld) and (CA) are used when the signals come ferulic acid, coniferaldehyde and coniferyl alcohol, respectively, as end-groups, with unsaturateda and b carbons.

Fig. 5. Types of inter-monomeric structures found in the three different DHPs as deduced from the 13C NMR spectra. The structures displaying unsaturated a–b bonds wereprevalent in the coniferaldehyde and ferulic acid DHPs.

Fig. 6. AFM tapping mode height images (1 � 1 lm2) and rms roughness determinations of the different spin-coated DHP model surfaces that were used in themeasurements of the contact angle of water. (a) DHP from coniferyl alcohol; rms roughness 0.349 nm. (b) DHP from coniferaldehyde; rms roughness 0.411 nm. (c) DHP fromferulic acid; rms roughness 0.404 nm.

152 A. Holmgren et al. / Phytochemistry 70 (2009) 147–155

and primary alcohol for the coniferyl alcohol lignin), there was onemain difference between the coniferyl alcohol DHP and ferulic

acid/coniferaldehyde DHPs: the latter DHPs displayed nearly onlya–b unsaturated side-chains, whereas the coniferyl alcohol DHP

Table 4Initial equilibrium contact angles of water on DHP model surfaces.

Sample Contact angle (�)

DHP from ferulic acid 63 ± 0.3DHP from coniferaldehyde 60 ± 0.2DHP from coniferyl alcohol 58 ± 0.5Kraft lignin (spruce) 46*

* From Norgren et al. (2006).

Fig. 7. a1 and a2 are products of decarboxylation in ferulic acid dehydrogenativepolymerization; b1 and b2 are products of further oxidative coupling of a1 and a2,respectively.

A. Holmgren et al. / Phytochemistry 70 (2009) 147–155 153

displayed both unsaturated and saturated a–b bonds. In coniferylalcohol DHPs, unsaturated a–b side-chains commonly exist inconiferyl alcohol end-groups, whereas the saturated structuresare formed through b-coupling. Additionally, benzyl alcohols, a fre-quent product of b-coupling of coniferyl alcohol DHP (b-O-40;Fig. 5) may partially explain the lower hydrophobicity. Neverthe-less, it is puzzling that the DHP made of ferulic acid is as hydropho-bic as it is. One possible explanation is that it has undergonesubstantial decarboxylation, and thus loss of charge. Apparently,there is a tendency for decarboxylation of certain structures inthe radical polymerization of ferulic acid (Ralph et al., 1994,2007; Ward et al., 2001), which can lead to new structures throughfurther oxidation and coupling (Fig. 7). If and to what extent decar-boxylation has occurred in our polymerisate is difficult to ascertainfrom the NMR measurements, since signals from such structureswere difficult to distinguish from the non-decarboxylated ones.

Why do the coniferaldehyde and ferulic acid DHPs display thesedifferences compared with the coniferyl alcohol DHP, when the

Fig. 8. Hypothetical mechanisms for reactions of quinone methides formed by radical coconiferyl alcohol is relatively stable, and water can perform a nucleophilic attack on thedeprotonation of the b-proton, which is relatively acidic due to the neighbouring carbdecarboxylation caused by the relatively electron abstracting b-carbon, leading to a rearofrom ferulic acid system might also react similarly to that of case (b).

coupling reactions do not directly involve the-carbon? An explana-tion might be the acidity of the b proton that is released upon therearomatization of the quinone methide intermediates in the caseof the coniferaldehyde and ferulic acid DHPs (Fig. 8). In the latter

uplings of the monolignols used in this work. (a) The quinomethide generated froma-carbon. (b) The quinone methide generated from coniferaldehyde is stabilized byons with d+. (c) The quinone methide generated from ferulic acid may undergomatized product (Ralph et al., 2007). Alternatively, the quinone methide generated

154 A. Holmgren et al. / Phytochemistry 70 (2009) 147–155

cases, a decarboxylation is an additional way of rearomatizing aquinone methide. These additional possibilities for the rearomati-zation of the quinone methide intermediate during ferulic acidand coniferaldehyde dehydrogenative polymerization apparentlylimit the nucleophilic attacks by water or a phenol, as comparedto coniferyl alcohol polymerization. Due to the carbonyl groupsat the c position, the formation of a Ca–Cb double bond is predom-inant during coupling in both cases. In Fig. 8, the hypotheticalmechanisms are summarized for the creation of the most commonintermonolignol bond in lignin, the b-O-40 bond. Another way ofrearomatizing the quinone methide intermediate is by an intramo-lecular nucleophilic attack leading to a cyclic structure. Althoughthe possibility of forming cyclic structures (i.e., b-50 and b–b0 forthe ferulic acid and b-50 for the coniferaldehyde DHP), producingsaturated a–b side-chains, has been shown elsewhere (Wardet al., 2001; Kim et al., 2003), in this work, only weak signals frompossible cyclic b-50 and b–b0 structures were produced in the ferulicacid DHPs. Other reports of coniferaldehyde dehydrogenative poly-merization support the lack of cyclic b-50 structure (Connors et al.,1970; Higuchi et al., 1994). However, it has been proposed that thiscyclic b-50 is formed first, but that it is not thermodynamically sta-ble and is subsequently transformed into the non-cyclic form (Kimet al., 2003). In the case of the ferulic acid DHP, a suggestion hasbeen made that cyclic and non-cyclic dimers of b-50 and b–b0 struc-tures could both be present and in some sort of equilibrium (Wardet al., 2001; Arrieta-Baez and Stark, 2006).

Can these differences in the structure provide an explanationfor the large investment by plants in a reduction of the mono-mers at the c-carbon? Apart from an increased hydrophobicity,the polymerization of coniferaldehyde and ferulic acid (Fig. 8)could be expected to lead to less covalent bonding between ligninand cell wall polysaccharides. Such covalent bonds, which yieldlignin-carbohydrate complexes (LCC), may be created by nucleo-philic attacks on the a-carbon of the quinone methide intermedi-ate by alcohols or carboxylic acids on the polysaccharide, i.e., byreplacing water in the mechanism of Fig. 8a (Sarkanen and Lud-wig, 1971; Eriksson et al., 1980). As the mechanisms in Figs. 8band c describe, and the results of this studies show, the possibil-ities for extramolecular nucleophilic attacks might be limited andconsequently, limit the formation of this type of LCC. Recently, itwas demonstrated that LCC form networks in wood, where lignincross-links different kinds of polysaccharides (Lawoko et al.,2006). These networks have been suggested to play an importantrole in the mechanical properties of the wood. Thus, it might bepossible that one reason for the reduction at the c-carbon is toimprove the formation of lignin-polysaccharide covalent net-works. Interestingly, a mutant loblolly pine was discovered thathad deficient expression of the cinnamyl alcohol dehydrogenaseenzyme of the phenylpropanoid pathway (step 5, Fig. 2). The via-ble plant exhibited a decreased lignin content, but also hadabnormally high amounts of aldehyde groups from coniferalde-hyde end-groups, among others (Ralph et al., 1997). Coniferalde-hyde was able to copolymerize with normal lignin monomers tosome degree and keep the lignin polymer somewhat functional.However, this does not indicate that the reduction is unnecessaryfor obtaining the fully functional lignin, since the lignin of thismutant still contained mostly alcohol monomers, which can formthe a-carbon LCC.

Lignin has a unique ability among biopolymers of ‘‘metabolicplasticity”, i.e., that non-conventional monomers can be incorpo-rated into the polymer (Ralph et al., 1997; Pilate et al., 2002). How-ever, this should not be understood as any potential monomerbeing able to form a highly functional lignin. The standard mono-mers of lignin have been very well chosen by nature to fulfill itsbiological functions, as discussed here for the reduction at thec-carbon.

4. Conclusions

� Coniferaldehyde and ferulic acid form polymers equivalent toconiferyl alcohol polymers by dehydrogenative polymerization.

� The DHPs of coniferaldehyde and ferulic acid were more hydro-phobic than the DHP made of coniferyl alcohol.

� The DHPs of coniferaldehyde and ferulic acid displayed mostlyunsaturated inter-unit structures and only small amounts ofcyclic inter-unit structures for the ferulic acid DHP, whereasconiferyl alcohol DHP displayed a variety of saturated and cyclicinter-unit structures.

� Lignin made of coniferyl alcohol may be able to form LCC easierthan lignin made of coniferaldehyde or ferulic acid.

5. Experimental

5.1. Materials

Coniferaldehyde (4-hydroxy,3-methoxy-cinnamaldehyde),ferulic acid (4-hydroxy,3-methoxy-cinnamic acid), and horseradishperoxidase (HRP) type VI were purchased from Sigma–AldrichSweden AB, Stockholm, Sweden. Coniferyl alcohol was obtainedfrom the reduction of coniferaldehyde with NaBH4 (Ludley andRalph, 1996). All other chemicals were of analytical grade.

5.2. Making of synthetic lignin (DHP)

Coniferaldehyde-based DHP and coniferyl alcohol-based DHPwere synthesized using the following method: 1 g of monomerwas dissolved in 10 ml acetone and then mixed together withHRP (5 mg, 1450 U) and 190 ml 50 mM K2HPO4 buffer (pH 6.5). Asolution of hydrogen peroxide (74 mM) in 200 ml phosphate bufferwas dropped at 10 ml � h�1 into the monolignol solution. After24 h, the reaction was stopped and the suspended polymerisatewas centrifuged at 10,000 rpm and the pellet washed with deion-ized water 2–3 times. Finally, the pellet was suspended in waterto be freeze-dried. Low molecular weight compounds were ex-tracted for 1 h with 50 ml CH2Cl2. The ferulic acid-based DHPwas similarly prepared except for the following modifications:ferulic acid was dissolved in 10 ml of acetone:K2HPO4 buffer(1:1) before adding to the rest of the buffer. The pH of the reactionmixture was adjusted back to 6.5 with diluted NaOH after theaddition of ferulic acid. After the reaction, diluted H2SO4 was addeduntil the pH was 2–3 to precipitate the polymerisate. Lowmolecular weight compounds were extracted with ethyl acetatefor 1 h.

5.3. Solubility panel

The solubility of the three DHP materials was analyzed by dis-solving 1–2 mg into 0.5 ml solvent and determining by simple ocu-lar inspection if the material was completely, partly or poorlydissolved.

5.4. Molecular mass determinations

Samples of each DHP product were acetylated in acetic anhy-dride:pyridine (1:1) overnight and analyzed by size exclusionchromatography (SEC), performed on a system of three cross-linked (PS) columns in series (Styragel, Waters HR4 + HR2 + HR0.5)using THF as the effluent at 0.8 ml �min�1. Detection was per-formed with a UV-light detector set at 280 nm. External calibrationwas made with a kit of monodisperse polystyrene standards in themolar mass range of 580–3,053,000 g �mol�1.

A. Holmgren et al. / Phytochemistry 70 (2009) 147–155 155

5.5. NMR

NMR experiments were performed on a Bruker Avance 400 MHzinstrument. 13C NMR was performed on the DHP polymeric frac-tion by dissolving the dried material in DMSO-d6. A pulse programwith inverse-gated proton decoupling was applied to acquire 13CNMR data. The measurements were acquired at 24 �C, with a spec-tral window of 240 ppm centered at 110 ppm. The data points ac-quired in the time domain were set to 32 K, which gave anacquisition time of 0.68 s. A relaxation delay of 14 s was used toavoid signal saturation. Five thousand and eight hundred scanswere collected for each measurement.

5.6. Preparation of DHP model surfaces

In the preparation of DHP model surfaces, a Chemat TechnologyKW-4A spin-coater was used. Silicon wafers cut in pieces ofapproximately 1 cm2 were used as substrates. The DHP sampleswere dissolved in pyridine overnight to yield concentrations of2.0 wt%. Thereafter, the DHP solutions were gently passed througha 0.45 lm Millex�-HV Durapore PVDF syringe-driven filter unit toremove possible dust and undissolved DHP particles. Immediatelybefore the spin-coating, the silicon wafers were exposed to 0.1 MNaOH for twenty seconds to hydroxylate the SiO2 layer, rinsed withplenty of Milli-Q water, and finally dried by a stream of nitrogengas. In the spin-coating, a rotation speed of 3500 rpm was appliedfor 1 min.

5.7. Contact angle measurements

The contact angle measurements were performed on a FibroDAT dynamic angle tester. This instrument uses image analysiswhen monitoring the spreading of a liquid on a substrate as a func-tion of time. In this work, water was used as the fluid. The initialequilibrium contact angle between the water drop and the surfacewas evaluated and reported as an average of 4 measurements.

5.8. AFM surface imaging and roughness determinations

A Veeco Picoforce scanning probe microscopy equipped with aVeeco silica cantilever was used in tapping mode to image theDHP model surfaces. The root-mean-square (rms) roughness val-ues of the films were determined from the height images. All ofthe images were recorded in ambient air conditions (25 �C and50% relative humidity).

Acknowledgements

This work was financially supported by the Swedish ResearchCouncil for Environmental, Agricultural Science and Spatial Plan-ning (FORMAS), contract #229-2004-1240. Mr. Erik Johansson isthanked for helpful assistance in the AFM characterisation.

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