Non-enzymatic reduction of quinone methides during oxidative coupling of monolignols: Implications for the origin of benzyl structures in lignins

Article abstract of DOI:10.1039/b606369a

Lignin is believed to be synthesized by oxidative coupling of 4-hydroxyphenylpropanoids. In native lignin there are some types of reduced structures that cannot be explained solely by oxidative coupling. In the present work we showed via biomimetic model experiments that nicotinamide adenine dinucleotide (NADH), in an uncatalyzed process, reduced a β-aryl ether quinone methide to its benzyl derivative. A number of other biologically significant reductants, including the enzyme cellobiose dehydrogenase, failed to produce the reduced structures. Synthetic dehydrogenation polymers of coniferyl alcohol synthesized (under oxidative conditions) in the presence of the reductant NADH produced the same kind of reduced structures as in the model experiment, demonstrating that oxidative and reductive processes can occur in the same environment, and that reduction of the in situ-generated quinone methides was sufficiently competitive with water addition. In situ reduction of β-β-quinone methides was not achieved in this study. The origin of racemic benzyl structures in lignins therefore remains unknown, but the potential for simple chemical reduction is demonstrated here. The Royal Society of Chemistry 2006.

Full text of DOI:10.1039/b606369a

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www.rsc.org/obc | Organic & Biomolecular Chemistry
Non-enzymatic reduction of quinone methides during oxidative coupling of
monolignols: implications for the origin of benzyl structures in lignins†
Anders Holmgren,a Go¨sta Brunow,b Gunnar Henriksson,a Liming Zhang*a and John Ralphc
Received 5th May 2006, Accepted 30th June 2006
First published as an Advance Article on the web 7th August 2006
DOI: 10.1039/b606369a
Lignin is believed to be synthesized by oxidative coupling of 4-hydroxyphenylpropanoids. In native
lignin there are some types of reduced structures that cannot be explained solely by oxidative coupling.
In the present work we showed via biomimetic model experiments that nicotinamide adenine
dinucleotide (NADH), in an uncatalyzed process, reduced a b-aryl ether quinone methide to its benzyl
derivative. A number of other biologically significant reductants, including the enzyme cellobiose
dehydrogenase, failed to produce the reduced structures. Synthetic dehydrogenation polymers of
coniferyl alcohol synthesized (under oxidative conditions) in the presence of the reductant NADH
produced the same kind of reduced structures as in the model experiment, demonstrating that oxidative
and reductive processes can occur in the same environment, and that reduction of the in situ-generated
quinone methides was sufficiently competitive with water addition. In situ reduction of b–b-quinone
methides was not achieved in this study. The origin of racemic benzyl structures in lignins therefore
remains unknown, but the potential for simple chemical reduction is demonstrated here.
Introduction
The cell walls of mature secondary xylem cells in woody plants are
generally strong, stiff, relatively resistant to microbial degradation,
and have low water absorbing capacity. These properties are
necessary for fulfilling the functions of water conductivity and
mechanical support of various types of wood cells, as tracheids,
vessels and libriform fibers. Plants have developed a composite
type material suitable for these functions consisting mainly of
cellulose, a b-1,4-homoglucan, hemicelluloses, a group of often
branched heteropolysaccharides, and lignin, a racemic, branched
phenylpropanoid polymer. The woody cell walls also contain
smaller amounts of proteins, extractives (various types of low
molecular mass organic compounds), and inorganic materials.2,3
The cellulose chains form crystalline bundles, fibrils, that function
as enforcing fibers, while hemicelluloses and lignin play the
role of a flexible connecting plastic.3,4 The biosynthesis of this
composite is complex: cellulose is synthesized directly outside the
cell-membrane by complexes of membrane proteins that create
a cellulose fibril consisting of about 64 or more chains,3 while
hemicelluloses are synthesized in the Golgi apparatus, excreted
to the cell wall through budding of Golgi vesicles, and thereafter
precipitated.5 As for cellulose biosynthesis, hemicellulose
polymerization is thermodynamically-driven by
a coupled
hydrolysis of nucleotide triphosphates.6,7 Lignin, on the other
hand, is believed to be synthesized from its monomers, primarily
the three monolignols (Fig. 1a), in situ into a gel consisting
of mainly cellulose and hemicellulose. According to a widely
aDepartment of Fiber and Polymer Technology, School of Chemical Sci-
ences, Royal Institute of Technology, KTH, 100 44 Stockholm, Sweden.
E-mail: liming@polymer.kth.se; Fax: +46 8 7096166
bDepartment of Chemistry, University of Helsinki, Helsinki, Finland
cUS Dairy Forage Research Center, USDA-Agricultural Research Service,
Madison, WI 53706, USA
Fig. 1 (a) Monolignols (the lignin monomers) and (b) the major
† A new model in lignin biosynthesis II (for part I, see ref. 16).
structures in lignins.
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Scheme 1 Mechanism for formation of a covalent bond, the b-O-4ꢀ-bond, in lignins. In the last structure, the nomenclature for inter-monomer bonds
in lignins is explained; the aliphatic side-chain carbons are denoted a, b and c with a being the benzylic position, i.e. closest to the aromatic ring. The
carbons in the aromatic ring are numbered with ‘1’ for the carbon connected to the aliphatic carbon chain. MG is coniferyl alcohol, PG is a guaiacyl
end-group on the lignin polymer and (P + 1)G is the coupling product of coniferyl alcohol to the polymer.
accepted model originally presented by Freudenberg and
Neish,8 this takes place by an uncatalyzed radical coupling of
resonance-stabilized monolignol radicals and phenolic radicals
on the lignin polymer, followed by nucleophilic additions to
the quinone methide intermediates, as explained in Scheme 1.
Aromatization of b-ether quinone methides, usually via water
addition, yields b-aryl ethers A (Fig. 1b). Covalent bonds (ethers
and esters) between lignin and polysaccharide can form if
quinone intermediates are trapped by polysaccharide hydroxy
groups.9–11 Phenolic or aliphatic hydroxy groups on the lignin
itself can also perform the nucleophilic attack creating intra-lignin
ethers in phenylcoumaran (b-5ꢀ-coupled) units B and in resinol
(b–bꢀcoupled) units C.8 The phenolic radicals are probably
generated either directly by an enzyme such as peroxidase,12–14 or
possibly indirectly via a redox shuttle of some kind.15,16
Although the Freudenberg model explains the large majority of
lignin structures, a number of structures that cannot be explained
by this chemistry alone have long been noted in lignins, including
the reduced lignin structures1,17 described in Fig. 2. Reduced b–
bꢀ-structures are especially interesting, since they are routinely
observed in softwood lignins by NMR,1,18 and by degradative
methods.19 There is some evidence that the common type of b–
bꢀ-structure, i.e. pinoresinol units C (Fig. 1 and Scheme 1 for a
nomenclature explanation of lignin bonds), is frequently 5-linked
in the structure of lignin, whereas the reduced b–bꢀ-structures
normally seem to be simply etherified (i.e. by b-O-4-bonds ).1
A possible explanation for the presence of reduced structures
in native lignins is that the quinone methide intermediate formed
during lignin biopolymerization is, in some way, two-electron re-
duced (Scheme 2), instead of being subject to a nucleophilic attack
by a hydroxy group. Although enzyme-driven reductions could
possibly explain reduced structures in lignin, secoisolariciresinol
products released from lignins (by methods shown to not scramble
the b–bꢀ-stereochemistry) have been shown to be racemic.18
‡
There is a number of potential reducing agents synthesized by
living organisms that could potentially reduce quinone methide
structures, generated during lignification, in uncatalyzed reactions.
In this work we study and discuss the formation of the reduced
lignin structures based on results of model experiments and
synthetic dehydrogenation polymers.
Scheme 2 Reduced structures can be created by two-electron reduction
of quinone methide intermediates.
Results and discussion
We tested four low molecular mass reducing agents produced by
living cells for their abilities to reduce a model compound for the
b-ether quinone methide intermediate in lignin biopolymerization:
nicotinamide adenine dinucleotide (NADH), reduced glutathione,
‡ Note that in this reference, as detailed recently,16 the DFRC prod-
uct of secoisolariciresinol units was originally incorrectly attributed to
pinoresinol units.
Fig. 2 Reduced lignin sub-structures that cannot be directly explained by
radical coupling reactions and nucleophilic attack, according to Scheme 1.
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both being common reducing agents produced in virtually all
living cells,20 ascorbic acid, known as a redox buffer agent in
plant cell walls,21 and a-ketoglutaric acid, intermediate in the citric
acid cycle. Furthermore, the extracellular enzyme cellobiose dehy-
drogenase (CDH), that carries both flavin and heme prosthetic
groups,22 known to have a broad specificity for reducing various
compounds including quinones, was included in the screen as an
example of a reducing enzyme. Sodium borohydride was used as
a positive control reducing agent.
Reduction of the b-ether quinone methide model was achieved
using NADH or NaBH4, but with none of the other reducing
agents tested, including the enzyme CDH. The treated model
compound was analyzed by GC–MS. The gas chromatogram of
the sodium borohydride-treated sample displayed a single peak.
An equivalent peak was found in the chromatogram of the NADH-
treated sample (peak labeled with 2, Fig. 3). In each case the
peak exhibited a mass spectrum corresponding to the silylated
reduced model compound 2 (Scheme 3). To confirm the identity
of the product, three NMR experiments (1H, 13C-DEPT-135 and
2-D HSQC) were run on the sodium borohydride-treated sample.
DEPT-135 spectra confirmed that the a-carbon was a methylene
group by its inversion, like the c-carbon, relative to the methoxy,
b- and aromatic C–Hs.
Scheme 3 Model compound guaiacylglycerol-b-guaiacyl ether 1 is con-
verted to a quinone methide by a two-step reaction and then exposed to a
reductant in the presence of ethanol to give the benzyl compound 2 along
with ethanol addition product 3 (as a mixture of isomers).
attacked by some intra- or extra-molecular nucleophile. In this
work, HSQC spectra of the DHPs showed the presence of b-aryl
ether A, phenylcoumaran B, resinol C, and a,b-diaryl ether A2
structures (Fig. 4) which are common in conventional DHPs.10,23
Furthermore, reduced b-ether structures AR (Fig. 4) of the same
type as compound 2 (Scheme 3) were observed from DHPs made
in the presence of NADH. The diagnostic coupling network
is elegantly revealed by HSQC-TOCSY experiments (Fig. 5).
Disappointingly, secoisolariciresinol units CR (Figs. 2 and 4)
were not observed, nor were dihydroconiferyl alcohol (X5, Fig. 2)
moieties found.
Biological significance
The results in this work demonstrate that it might be possi-
ble to obtain lignin-reduced structures in uncatalyzed protein-
independent reactions using a biologically significant reductant
such as NADH. Whether it is this compound that in vivo is
responsible for the reduction, a similar compound, or an enzyme
system, is not known. The inability of CDH to reduce the quinone
methide here does not imply that there are no other enzymes
that can catalyze the reaction. The present results show that
direct involvement of enzymes may not be necessary to clarify
the occurrence of these lignin structures. The detection of reduced
b-aryl ether units AR, at substantial levels (ca. 48% of the b-
ethers) in the synthesized lignins, indicates that NADH was
able to trap and reduce the b-aryl ether quinone methide in an
oxidative radical–radical polymerization system. NADH was not,
however, able to produce reduced b–bꢀ-structures in detectable
amounts, so conditions and/or reductants for such a process
remain elusive. Presumably the reduction of the a-carbon of
a b-aryl ether quinone methide competes with a nucleophilic
attack by an external nucleophile (water or a phenol), whereas
Fig. 3 GC chromatogram of the silylated products of the quinone methide
reduction with NADH. Peaks are labeled with their structures from
Scheme 3; 1 is guaiacylglycerol-b-guaiacyl ether, 3a and 3b are the two
isomers of the ethanol addition product, and 2 is the reduced quinone
methide.
Synthetic dehydrogenation polymers (DHPs) were prepared in
the presence NADH to test the reducing capabilities of NADH
in an oxidizing environment such as that involved in lignin
polymerization. The interest here was to determine if oxidation
(radical generation from the phenol) and reduction could proceed
in the same reaction medium. Even though conventional dehydro-
genation polymers differ in unit proportions from isolated lignins,
as reviewed,19 they reproduce essentially all of the known inter-unit
linkages, and derive from the same chemical steps to achieve poly-
merization: oxidation of monolignols/oligomer, radical–radical
coupling forming a quinone methide intermediate which is finally
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Fig. 5 HSQC-TOCSY of a DHP synthesized in the presence of
NADH. The dashed lines represent the proton–carbon correlations on
the propanoid chain of the reduced b-aryl ether structure AR (cf. 2 in
Scheme 3).
reduction, remain to be eliminated. One possibility is that the
conditions during part of the natural biosynthesis of lignin in
some way give more long-lived b–bꢀ-quinone methides than in our
in vitro experiments.
A reduction of a quinone methide with NADH costs the
plant three ATP equivalents of energy.27 It is therefore logical
to question the reason for reduced structures in lignins unless
they are simply to be attributed to the results of coincidental
side-reactions. In an earlier work1 it was shown that the reduced
b–bꢀ-units seemed to have been created as dimers, i.e. according
to bulk polymerization. Thus, it might be possible that under
some period in the cell wall synthesis, high biochemical activity
includes the presence of reducing agents in concert with fast lignin
polymerization leading to bulk polymerization and reduction of
quinone methides, whereas under other periods, reducing agents
are absent and the lignification rate is slower, giving an end-wise
polymerization. A lignin polymer with reduced structures would
have different properties than ‘normal’ lignins, such as higher
hydrophobicity and a lower propensity for hydrogen bonding or
formation of covalent bonds to polysaccharides through a-ethers
and a-esters. Additionally, a higher flexibility of the b–bꢀ-bonds
would be expected due to the absence of the tetrahydrofuran ring
structures. Therefore, reduction might be the way for the plant to
manipulate the properties of some of its lignin.
Fig. 4 HSQC spectra of a DHP synthesized in the presence of NADH.
A corresponds to b-aryl ether, A2 to a,b-diaryl ether, AR to reduced
b-aryl ether, B to phenylcoumaran, C to resinol, X1 to coniferyl alcohol
end-groups. CR and X5 correspond to secoisolariciresinol (reduced
b–bꢀ) units and dihydroconiferyl alcohol end-groups, with their expected
correlations according to data from Zhang et al.1
the b–bꢀ- or b-5ꢀ-quinone methides have intramolecular hydroxy
groups out-competing the attack on the a-carbon. Since we
observed resinol and phenylcoumaran structures in the DHPs,
NADH was apparently not effective in competing against the fast
intramolecular reactions in the polymerization process under these
conditions.
So far, reduced structures have only been discovered in end-
groups X5 and in b–bꢀ CR units in natural lignins.1,17,24,25 Un-
fortunately, in this work we have not detected any reduced b–
bꢀ-structures CR, but succeeded in producing reduced b-O-4ꢀ-
structures AR. Whether this is due to limitations in the detection
methods or, more probably, to a difference in the structure and the
polymerization environment between lignin and DHPs cannot be
definitively confirmed. DHP systems are known to be sensitive
to polymerization conditions.26 If the latter explanation is the
case, then there must be some kind of mechanism directing
the reducing effect towards b–bꢀ-units in lignin polymerization.
Enzymatic catalysis remains possible, but other options such as
low molecular weight catalysts, or unusual conditions during the
Conclusion
The reduction of a b-aryl ether quinone methide model compound
was achieved by non-enzymatic catalysis using the biologically
significant reducing agent, NADH. Moreover, synthetic lignins
(DHPs) synthesized in the presence of NADH produced anal-
ogous reduced structures, indicating that reduction of quinone
methides can be achieved by biological reductants present in
the oxidative environment of lignin polymerization. We have so
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far failed to demonstrate enzyme-free reduction of b–bꢀ-quinone
methides. The origin of racemic benzyl structures occurring in
lignins therefore remains unknown, but we have been able to
demonstrate the potential for simple chemical reduction as a part
of the Freudenberg oxidative model for lignin biosynthesis.
acetone (10 ml) and mixed together with potassium phosphate
buffer (90 ml, 25 mM, pH 6.0) containing NADH (310 mg)
and HRP (580 units). The pH was chosen according to the
activity pH range of the peroxidase, and following similar DHP
synthesis systems in the literature.26 A hydrogen peroxide solution
(29 mM, 100 ml) was added at a rate of 6 ml h−1 into the
coniferyl alcohol/HRP/NADH solution. The reaction was stirred
for a total of 22 h at room temperature. Then, the suspended
polymerisate was centrifuged at 5700 rpm for 10 min and the
pellet washed twice, resuspended in ultrapure water and freeze-
dried and washed with dichloromethane for one hour to extract
low molecular weight DHP compounds. The product obtained was
a light-brown powder that was subsequently prepared for NMR
analysis.
Experimental
Materials
Guaiacylglycerol-b-guaiacyl ether was purchased from TCI Eu-
rope NV, Zwijndrecht, Belgium, L-glutathione (reduced form),
a-ketoglutaric acid, b-nicotinamide adenine dinucleotide re-
duced disodium salt (NADH), N,O-bis(trimethylsilyl) trifluo-
roacetamide (BSTFA), D-(+)-cellobiose, horseradish peroxidase
(HRP) type VI and coniferaldehyde were purchased from Sigma-
Aldrich Sweden AB, Stockholm, Sweden. Bromotrimethylsilane
(TMSBr) and L-(+)-ascorbic acid were from E. Merck AB,
Sweden. Cellobiose dehydrogenase (CDH) was a kind gift from
Gunnar Johansson at the Department of Biochemistry, Univer-
sity of Uppsala, Sweden, and was purified from Phanerochaete
chrysosporium according to Henriksson et al.28 All other chemicals
were of analytical grade. Ultrafiltration units (for centrifugation)
Nanosep 10 K were from Pall Corporation, New York.
Gas chromatography–mass spectrometry (GC–MS) and nuclear
magnetic resonance (NMR)
The GC/MS instrument was from Thermo Finnigan Trace GC–
MS, 2000 series. The column used was RTX 5MS (30 m, 0.32 mm,
0.25 lm). The silylated products of the quinone methide reduction
were analyzed with◦a carrier gas flow of 0.5 ml min−1, an initial
temperature of 120 C, a ramp rate of 5.0 ◦C min−1 until 200 ◦C,
◦
and then a rate of 4 C min−1 until 280 ◦C and held there for
30 min. The MS ionization method used was electron impact
operating at 70 eV. For the NMR analyses from the quinone
methide model experiments, the unmodified reference sample was
dissolved in CDCl3. From the DHP experiment the unmodified
product was dissolved in acetone-d6–D2O (3 : 1). All NMR
experiments were run on a Bruker Avance 400 MHz instrument.
2-D NMR experiments were performed with a proton–carbon-
selective inverse probe equipped with a Z-gradient coil. Standard
Bruker pulse programs were used in all experiments. Pulse lengths
of the NMR probes were carefully calibrated and optimum
tuning and matching as well as shimming of the NMR probe
was performed with each run. The 2-D HSQC spectrum (pulse
program ‘hsqcetgpsi’) was acquired with a spectral window of
12.8 ppm in F2 and 150 ppm in F1 with 512 increments of 2 K
points each, giving an acquisition time of 0.2 s in F2. Other
parameters for data acquisition included: a delay time of 2 s,
an average assumed coupling constant of 150 Hz, 32 scans per
increment. The 2-D HSQC-TOCSY spectrum (pulse program
‘hsqcetgpml’) was acquired with a delay time of 1.3 s, a mixing
time of 60 ms and 128 scans per increment. Other acquisition
parameters were similar to those of the HSQC experiment. The
Quinone methide reduction
Guaiacylglycerol-b-guaiacyl ether [1-(4-hydroxy-3-methoxyphe-
nyl)-2-(2-methoxyphenoxy)-propane-1,3-diol] 1 (Scheme 3) was
used to prepare the quinone methide via the benzylic bromide
generated via TMSBr.29,30 Dichloromethane was evaporated at
high vacuum on ice and the quinone methide dissolved in 99.5%
ethanol and kept on ice. The reducing agent was dissolved in
potassium phthalate buffer (50 mM, pH 5) and the quinone
methide was slowly added to the buffer. The final reaction mixture
contained 10% ethanol and the ratio of quinone methide to
reducing agent was 1 : 10. The reaction was stirred overnight
at room temperature under nitrogen. The mixture was extracted
twice with dichloromethane, and dried with Na2SO4. The solvent
was evaporated and the crude product was silylated with BSTFA
for 1 h, as described elsewhere.31 When testing the enzyme
CDH, 0.13 ml of the enzyme solution (10.9 lM) and 6 mg
D-cellobiose (co-factor) were dissolved in potassium phthalate
buffer prior to quinone methide addition. After the reaction,
the enzyme was separated from the mixture by centrifugation
in ultrafiltration units and the filtrate extracted and dried as
previously described.
1
spectral centre was set at 5.3 ppm in the H dimension and at
91 ppm in the 13C dimension. The acquired 2-D NMR data sets
were processed with 2 K × 1 K data points for the HSQC and
HSQC-TOCSY spectra, using the p/2-shifted sine-bell window
functions in both dimensions.
A reference sample of compound 2 was prepared by reduc-
ing the quinone methide model with NaBH4.32,33 The quinone
methide was prepared as described previously, dissolved in
dichloromethane and stirred with aqueous NaBH4 until the pale
yellow color disappeared. The organic phase was separated, dried
with Na2SO4 and evaporated. For GC–MS, the crude product was
silylated as described previously.
Compound 2, 4-[3-hydroxy-2-(2-methoxyphenoxy)-propyl]-2-
methoxyphenol (Scheme 3). 1H NMR (400 MHz; CDCl3): d 2.87–
3.09 (a-CH2), 2.97 (c-OH), 3.60–3.68 (c-CH2), 3.85–3.89 (OCH3),
4.20–4.26 (b-CH), 5.52 (phenolic H), 6.75–7.03 (aromatic H). 13
C
Synthesis of dehydrogenation polymers (DHPs) under reductive
conditions
NMR (in CDCl3): d 37.52 (a-CH2), 55.86 and 55.96 (2 OCH3),
63.61 (c-CH2), 85.32 (b-CH), 112.14–123.51 (aromatic-C). MS
(m/z) silylated: 448 (M+, 1%), 324 (2%), 293 (2%), 209 (100%),
179 (12%), 166 (12%), 103 (5%), 73 (72%).
Coniferyl alcohol was obtained from the reduction of coniferalde-
hyde with NaBH4.34 Coniferyl alcohol (200 mg) were dissolved in
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Compounds 3 (a and b), syn- and anti-4-[1-ethoxy-3-hydroxy-
2-(2-methoxyphenoxy)-propyl]-2-methoxyphenol (Scheme 3). MS
(m/z) silylated: 492 (M+, <1%), 323 (1%), 293 (1%), 253 (100%),
225 (6%), 209 (5%), 181 (7%), 166 (22%), 103 (6%), 73 (62%).
15 D. Fournand, B. Cathala and C. Lapierre, Phytochemistry, 2003, 62,
139–146.
¨
16 H. Onnerud, L. Zhang, G. Gellerstedt and G. Henriksson, Plant Cell,
2002, 14, 1953–1962.
17 G. Brunow, K. Lundkvist and G. Gellerstedt, in Analytical Methods
in Wood Chemistry, Pulping and Papermaking, ed. E. Sjo¨stro¨m and
R. Alen, Springer-Verlag, Berlin, 1999, pp. 77–124.
18 J. Ralph, J. Peng, F. Lu, R. D. Hatfield and R. F. Helm, J. Agric. Food
Chem., 1999, 47, 2991–2996.
Reduced b-O-4ꢀ-structure, AR, from DHP experiment (2-D NMR
spectra, Fig. 4). 1H NMR (acetone-d6–D2O 3 : 1; 400 MHz): d
2.91 (a-CH2), 3.62 (c-CH2), 3.75 (OCH3), 4.45 (b-CH), 6.70–6.99
(aromatic H). 13C NMR (in acetone-d6–D2O 3 : 1): d 36.22 (a-
CH2), 55.51 (OCH3), 62.22 (c-CH2), 80.91 (b-CH), 108.50–121.76
(aromatic-C).
19 J. Ralph, K. Lundquist, G. Brunow, F. Lu, H. Kim, P. F. Schatz,
J. M. Marita, R. D. Hatfield, S. A. Ralph, J. Holst Christensen and
W. Boerjan, Phytochem. Rev., 2004, 3, 29–60.
20 L. Stryer, Biochemistry, W. H. Freeman & Company, New York, 3rd
edn., 1988.
21 C. Pignocchi and C. H. Foyer, Curr. Opin. Plant Biol., 2003, 6, 379–389.
22 G. Henriksson, G. Johansson and G. Pettersson, J. Biotechnol., 2000,
78, 93–113.
Acknowledgements
23 J. Ralph, J. M. Marita, S. A. Ralph, R. D. Hatfield, F. Lu, R. M. Ede,
J. Peng, S. Quideau, R. F. Helm, J. H. Grabber, H. Kim, G. Jimenez-
Monteon, Y. Zhang, H.-J. G. Jung, L. L. Landucci, J. J. MacKay, R. R.
Sederoff, C. Chapple and A. M. Boudet, in Advances in Lignocellulosics
Characterization, ed. D. S. Argyropoulos, Tappi Press, Atlanta, GA,
1999, pp. 55–108.
24 J. Ralph, J. J. MacKay, R. D. Hatfield, D. M. O’Malley, R. W. Whetten
and R. R. Sederoff, Science (Washington, D. C.), 1997, 277, 235–239.
25 J. Ralph, C. Lapierre, J. M. Marita, H. Kim, F. Lu, R. D. Hatfield, S.
Ralph, C. Chapple, R. Franke, M. R. Hemm, J. Van Doorsselaere, R. R.
Sederoff, D. M. O’Malley, J. T. Scott, J. J. MacKay, N. Yahiaoui, A. M.
Boudet, M. Pean, G. Pilate, L. Jouanin and W. Boerjan, Phytochemistry,
2001, 57, 993–1003.
The authors would like to thank the Swedish Research Council
for Environment, Agricultural Sciences and Spatial Planning
(FORMAS), contract no 229-2004-1240, for financial support.
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