Engineering traditional monolignols out of lignin by concomitant up-regulation of F5H1 and down-regulation of COMT in Arabidopsis


  • Ruben Vanholme,

    1. Department of Plant Systems Biology, VIB, 9052 Gent, Belgium
    2. Department of Plant Biotechnology and Genetics, Ghent University, 9052 Gent, Belgium
    Search for more papers by this author
  • John Ralph,

    1. Department of Biochemistry and Great Lakes Bioenergy Research Center, University of Wisconsin, Madison, WI 53706, USA
    Search for more papers by this author
  • Takuya Akiyama,

    1. Department of Biochemistry and Great Lakes Bioenergy Research Center, University of Wisconsin, Madison, WI 53706, USA
    Search for more papers by this author
  • Fachuang Lu,

    1. Department of Biochemistry and Great Lakes Bioenergy Research Center, University of Wisconsin, Madison, WI 53706, USA
    Search for more papers by this author
  • Jorge Rencoret Pazo,

    1. Department of Biochemistry and Great Lakes Bioenergy Research Center, University of Wisconsin, Madison, WI 53706, USA
    Search for more papers by this author
  • Hoon Kim,

    1. Department of Biochemistry and Great Lakes Bioenergy Research Center, University of Wisconsin, Madison, WI 53706, USA
    Search for more papers by this author
  • Jørgen Holst Christensen,

    1. Department of Plant Systems Biology, VIB, 9052 Gent, Belgium
    2. Department of Plant Biotechnology and Genetics, Ghent University, 9052 Gent, Belgium
    Search for more papers by this author
  • Brecht Van Reusel,

    1. Department of Plant Systems Biology, VIB, 9052 Gent, Belgium
    2. Department of Plant Biotechnology and Genetics, Ghent University, 9052 Gent, Belgium
    Search for more papers by this author
  • Véronique Storme,

    1. Department of Plant Systems Biology, VIB, 9052 Gent, Belgium
    2. Department of Plant Biotechnology and Genetics, Ghent University, 9052 Gent, Belgium
    Search for more papers by this author
  • Riet De Rycke,

    1. Department of Plant Systems Biology, VIB, 9052 Gent, Belgium
    2. Department of Plant Biotechnology and Genetics, Ghent University, 9052 Gent, Belgium
    Search for more papers by this author
  • Antje Rohde,

    1. Department of Plant Systems Biology, VIB, 9052 Gent, Belgium
    2. Department of Plant Biotechnology and Genetics, Ghent University, 9052 Gent, Belgium
    Search for more papers by this author
  • Kris Morreel,

    1. Department of Plant Systems Biology, VIB, 9052 Gent, Belgium
    2. Department of Plant Biotechnology and Genetics, Ghent University, 9052 Gent, Belgium
    Search for more papers by this author
  • Wout Boerjan

    Corresponding author
    1. Department of Plant Systems Biology, VIB, 9052 Gent, Belgium
    2. Department of Plant Biotechnology and Genetics, Ghent University, 9052 Gent, Belgium
      (fax +32 9 3313809; e-mail
    Search for more papers by this author

(fax +32 9 3313809; e-mail


Lignin engineering is a promising strategy to optimize lignocellulosic plant biomass for use as a renewable feedstock for agro-industrial applications. Current efforts focus on engineering lignin with monomers that are not normally incorporated into wild-type lignins. Here we describe an Arabidopsis line in which the lignin is derived to a major extent from a non-traditional monomer. The combination of mutation in the gene encoding caffeic acid O-methyltransferase (comt) with over-expression of ferulate 5-hydroxylase under the control of the cinnamate 4-hydroxylase promoter (C4H:F5H1) resulted in plants with a unique lignin comprising almost 92% benzodioxane units. In addition to biosynthesis of this particular lignin, the comt C4H:F5H1 plants revealed massive shifts in phenolic metabolism compared to the wild type. The structures of 38 metabolites that accumulated in comt C4H:F51 plants were resolved by mass spectral analyses, and were shown to derive from 5-hydroxy-substituted phenylpropanoids. These metabolites probably originate from passive metabolism via existing biochemical routes normally used for 5-methoxylated and 5-unsubstituted phenylpropanoids and from active detoxification by hexosylation. Transcripts of the phenylpropanoid biosynthesis pathway were highly up-regulated in comt C4H:F5H1 plants, indicating feedback regulation within the pathway. To investigate the role of flavonoids in the abnormal growth of comt C4H:F5H1 plants, a mutation in a gene encoding chalcone synthase (chs) was crossed in. The resulting comt C4H:F5H1 chs plants showed partial restoration of growth. However, a causal connection between flavonoid deficiency and this restoration of growth was not demonstrated; instead, genetic interactions between phenylpropanoid and flavonoid biosynthesis could explain the partial restoration. These genetic interactions must be taken into account in future cell-wall engineering strategies.


Lignins are aromatic heteropolymers that are predominantly deposited in secondarily thickened plant cell walls, where they provide strength and imperviousness to the wall, and protect the cellulose microfibrils from microbial attack. At the same time, lignins limit the efficiency of several industrial processes that use plant cell walls as a feedstock. Engineering of lignin amount and structure through biotechnology is a promising strategy to improve the industrial processing of lignocellulosic plant biomass. During the last two decades, many details of the biochemical pathway producing coniferyl and sinapyl alcohol, the primary monomers from which polymeric lignins derive, have been established. The genes for each of the steps have been identified, and the activities of the derived enzymes have been reasonably well characterized (Boerjan et al., 2003; Ralph et al., 2004; Vanholme et al., 2010).

Many of the recent advances in lignin research have come from examining the consequences on overall lignin structure of up- and down-regulating the various genes of the pathway. Such studies have also revealed the rather striking metabolic malleability of the lignification process. Massive compositional changes (i.e. changes in the ratio of guaiacyl (G) and syringyl (S) moieties resulting from incorporation of coniferyl and sinapyl alcohol monomers, respectively) in angiosperm lignins can result from mis-regulation of single genes in the pathway. For example, lignins consist largely (or almost entirely) of G units when F5H, the gene encoding ferulate 5-hydroxylase, is down-regulated (Figure 1) (Meyer et al., 1998; Marita et al., 1999). Conversely, up-regulation of F5H using a xylem-specific promoter in Arabidopsis, poplar (Populus tremula×alba) or tobacco (Nicotiana tabacum) leads to lignins with extraordinarily high S:G ratios (Meyer et al., 1998; Marita et al., 1999; Franke et al., 2000; Stewart et al., 2009). Despite containing essentially linear and apparently low-molecular-weight lignins, poplar plants over-expressing F5H displayed no visible growth phenotypes (Stewart et al., 2009). However, the wood was markedly easier to pulp than that of the wild type (Huntley et al., 2003), demonstrating that altering lignin composition can be a viable approach towards improved processing.

Figure 1.

 Biosynthetic pathway for coniferyl and sinapyl alcohol (green box) and the metabolites that accumulate during concomitant up-regulation of F5H1 and down-regulation of COMT (red boxes).
The black route is expected to be the favored route; the gray routes may also be followed depending on the genetic background and growth conditions. Dashed arrow: route suggested by Mir Derikvand et al. (2008), Lepléet al. (2007) and Nair et al. (2004); ?, conversion not demonstrated. Possible routes towards the four classes of compounds are indicated by thin red arrows. CCR, cinnamoyl CoA reductase; COMT, caffeic acid O-methyltransferase; HCALDH, hydroxycinnamaldehyde dehydrogenase; F5H, ferulate 5-hydroxylase. For convenience, metabolites are classified in four classes based on their chemical identity: class I, oligolignols; class II, hexosylated oligolignols; class III, 5-hydroxyferulic acid-containing dimers; class IV, derivatives of 5-hydroxylated phenylpropanoid monomers (see Table 3).

In addition to the compositional shifts in S:G ratios, it has become apparent that lignification may tolerate the (partial) substitution of monolignols by various other phenolic precursors. Such substitution is compatible with the current theory of lignification, in which the polymer derives via chemical radical coupling reactions, primarily coupling of a monomer with the growing polymer; any phenolic precursor within the lignification zone may co-polymerize with other available phenolic precursors to the extent allowed by chemical compatibility (Ralph et al., 2004). Less obvious, but apparently possible, is that phenolic precursors, which are often formed in cells with perturbed phenylpropanoid biosynthesis, can be transported to the wall for lignification. For example, angiosperms deficient in caffeic acid O-methyltransferase (COMT) incorporate 5-hydroxyconiferyl alcohol into their lignins, forming benzodioxane units that are readily apparent by NMR, thioacidolysis and phenolic profiling of lignin oligomers (Atanassova et al., 1995; Van Doorsselaere et al., 1995; Jouanin et al., 2000; Guo et al., 2001; Ralph et al., 2001a,b; Marita et al., 2003; Morreel et al., 2004b; Do et al., 2007; Lu et al., 2010).

The goal of the current study was to develop plants in which the lignin composition approaches the state whereby none of its constituents are derived from traditional monolignols. Despite the already overwhelming evidence that various non-traditional monolignols are readily incorporated into the lignin polymer in a variety of natural and transgenic plants, a plant with a minor complement of traditional monolignols would provide insight into how far and in which directions lignin might be redesigned. One simple strategy to produce non-traditional lignins consists of up-regulating F5H while blocking COMT expression (Figure 1). If there is no feedback inhibition on F5H specifically, or within the phenylpropanoid pathway in general, it is anticipated that the cells will primarily produce 5-hydroxyconiferyl alcohol for lignification. Substantial levels of 5-hydroxyconiferyl alcohol can apparently be tolerated in lignification without obvious adverse effects on plant growth and development (Figure S1). Here we describe comt C4H:F5H1 plants in which the lignin comprises approximately 92% benzodioxane units, far more than ever reported previously and how plants respond to such a massive replacement of traditional monolignols with 5-hydroxyconiferyl alcohol.


comt C4H:F5H1 plants have an altered development

In order to design lignins that are derived predominantly from non-traditional 5-hydroxyconiferyl alcohol monomers, Arabidopsis plants over-expressing F5H1 under the control of the cinnamate 4-hydroxylase (C4H) promoter (C4H:F5H1 plants) were crossed with Arabidopsis comt mutants. Plants homozygous for the comt mutation and the C4H:F5H1 construct (comt C4H:F5H1 plants) were identified in the F2 progeny. Although the comt mutant grew normally and the C4H:F5H1 over-expressing line showed only moderate growth effects, plant development in comt C4H:F5H1 plants was severely affected. When grown on agar medium, leaves of the comt C4H:F5H1 seedlings were smaller compared to those of the wild type (Figure 2a). The inflorescence stems of agar-grown comt C4H:F5H1 plants remained small, with a maximal observed final length of 8 cm, whereas wild-type stems were over 30 cm at maturity. Extensive phenotypic variation was observed for the length, number and color of the inflorescence stems (Figure 2b–d), with the color ranging from green to purple red. In many cases, additional inflorescences developed, resulting in a bushy phenotype (Figure 2d). The total stem biomass of senesced soil-grown comt C4H:F5H1 plants was only a fraction of that of the wild type (Table 1). Although comt C4H:F5H1 plants developed flowers, they did not set seeds.

Figure 2.

 Phenotype and inflorescence stem morphology of comt C4H:F5H1 and comt C4H:F5H1 chs plants.
(a) Fourteen-day-old seedlings, segregating from a comt/COMT C4H:F5H1/C4H:F5H1 parental line. The double homozygous comt C4H:F5H1 plant is indicated with an arrow. (b–d) comt C4H:F5H1 plants with a wide phenotypic diversity: (b,c) 13-week-old and (d) 24-week-old plants.
(e) Mäule staining of comt C4H:F5H1 plant grown for 28 weeks in vitro. A section of a greenhouse-grown wild-type plant is visible in the upper left corner.
(f, g) Light microscopy of toluidine blue-stained sections of wild-type and comt C4H:F5H1 tissue culture-grown plants, respectively. Arrows indicate xylem cells. Note the collapsed xylem vessels in the comt C4H:F5H1 plants.
(h–k) TEMs of cell walls stained with uranyl acetate, showing interfascicular fibers of the wild type (h), interfascicular fibers of comt C4H:F5H1 (i), the xylem region of the wild type (j), and the xylem region of comt C4H:F5H1 (k).
(l) Partially recovered phenotype of a comt C4H:F5H1 chs plant (see Table 1 for biomass data).

Table 1.   Inflorescence height and the total inflorescence stem dry weight, as a measure for biomass, of the various lines
LineHeight (cm)Dry weight (mg)
Mean ± SDnMean ± SDn
  1. n, number of plants analyzed.

  2. *Significantly different from the wild type (< 0.001).

Wild type47.7 ± 3.529104.9 ± 32.010
comt44.2 ± 4.930125.4 ± 27.110
C4H:F5H129.4 ± 4.7*2977.6 ± 21.88
chs48.1 ± 4.827130.4 ± 58.010
comt C4H:F5H11.1 ± 1.4*141.4 ± 3.4*10
comt C4H:F5H1 chs11.0 ± 2.1*5010.9 ± 3.0*12

comt C4H:F5H1 plants have an irregular xylem phenotype

To reveal the morphological consequences of F5H1 over-expression in the comt mutant background at the tissue level, sections of the inflorescence stem were analyzed by light microscopy. Toluidine blue-stained sections of comt C4H:F5H1 plants showed collapsed xylem vessels, a phenotype typically known as the irregular xylem (irx) phenotype and observed in a range of mutants with perturbed secondary cell-wall development (Figure 2f,g) (Turner and Somerville, 1997; Jones et al., 2001; Franke et al., 2002; Goujon et al., 2003; Hoffmann et al., 2004; Brown et al., 2005; Besseau et al., 2007; Mir Derikvand et al., 2008; Schilmiller et al., 2009; Li et al., 2010). No obvious differences were noted in the other stem tissues. In contrast to the wild type, Mäule staining, which is indicative of S units in lignin, generally did not result in red coloration of the secondary cell wall, indicating a strong reduction in, or absence of, S units in the comt C4H:F5H1 plants (Figure 2e). However, a slight red coloration was observed in a few of these plants, suggesting the presence of residual COMT.

Transmission electron microscopy (TEM) of the cell walls revealed multiple concentric cell-wall sub-layers in the comt C4H:F5H1 plants (Figure 2i,k). This stratification appeared to be restricted to the S2 region of the cell wall, and was more obvious in interfascicular fibers than in xylem vessels. However, a wide variability in the extent of cell-wall stratification, ranging from absent to extensive, was observed in the fibers. This remarkable phenotype, already described for ccr1 Arabidopsis mutants and CCR down-regulated poplars, has been suggested to be caused by an altered assembly of cellulose microfibrils (Goujon et al., 2003; Lepléet al., 2007).

NMR analyses reveal massive incorporation of 5-hydroxyconiferyl alcohol into lignin

Although acetylbromide lignin levels in the senesced inflorescence stems of either comt or C4H:F5H1 plants were not significantly different from those of the wild-type, lignin amounts in comt C4H:F5H1 plants were reduced to approximately 56% of wild-type levels (Table 2). Cellulolytic enzyme lignins (CEL) were isolated from senesced inflorescence stems and analyzed by NMR. The aromatic regions of the 2D 13C–1H correlation (heteronuclear single quantum coherence, HSQC) spectra highlight differences in the distribution of guaiacyl, 5-hydroxyguaiacyl and syringyl (G, 5H and S) units in the lignins. The resulting spectra (Figures 3 and S2) clearly show that G signals from comt C4H:F5H1 plants are low, representing approximately 13.8% of the total aromatic content (based on volume integration), compared to 82.7% in the wild type. The dominant signals are from 5H units. It is not immediately clear that S units are depleted, because the S2/6 contours overlap with those from 5H6. However, the level of the 5H2 contours suggests that most of the 5H6/S2/6 contour region must belong to 5H6. The S2/6 volume integral can be estimated by subtracting the volume integral of the 5H2 region from that of the 5H6+S2/6 region; the volume integrals for 5H2 and 5H6 regions should be equal. This assumption results in a G:5H:S ratio of approximately 13.8:71.0:15.2; i.e. we estimate that the lignin is derived from approximately 71% of 5-hydroxylated monomers.

Table 2.   Lignin composition as measured by NMR and total acetyl bromide lignin content
LineMonomeric composition (%)Unit types (%)Acetyl bromide lignin (%)
  1. A, β-aryl ether; B, phenylcoumaran; C, resinol; D, dibenzodioxocin; J, benzodioxane (see Figure S3). n, number of plants analyzed.

  2. Statistical tests relative to the wild type were performed for total lignin only. Asterisks indicate values that are significantly different from the wild type at **0.001 < < 0.01 and ***< 0.001, respectively.

Wild type82.480.0017.5275.8817.216.180.730.0012.0 ± 1.77
comt94.791.283.9263.5917.673.942.9211.8710.5 ± 1.04
C4H:F5H16.551.9191.5583.840.003.500.0012.669.4 ± 1.74
chs84.200.0015.8073.7018.406.701.300.0011.7 ± 0.95
comt C4H:F5H113.7870.9915.236.440.850.790.0091.936.8 ± 0.8***5
comt C4H:F5H1 chs33.5420.0246.4459.071.421.500.0038.028.8 ± 1.1**8
Figure 3.

 Partial short-range 13C–1H (HSQC) spectra (aromatic regions) of acetylated enzyme lignins.
See Figure S2 for spectra of the complete set of transgenic lines and controls.

The inter-unit linkage distribution in the lignin fraction can be determined from the aliphatic side-chain region of the HSQC spectra (Table 2 and Figure S3). The correlation contours in the spectra (Figure S3), such as those for β-aryl ether (β–O–4) units (A), phenylcoumaran (β–5) units (B), resinol (β–β) units (C), dibenzodioxocin (5–5/β–O–4) units (D) and benzodioxane units (J), can be readily assigned by comparison with model data and from lignin and cell-wall spectra obtained previously (Ralph et al., 1999; Ralph and Landucci, 2010). As for the aromatic regions in Figures 3 and S2, the side-chain region of the comt C4H:F5H1 plants (Figure S3e) is strikingly different from that of the wild type (Figure S3a). Even though they are plotted at a lower contour level (closer to noise level, to show the weak residual signals from the traditional lignin structures), it is evident that proportions of the various ‘normal’ lignin structures (AC) are minor. The obvious major difference is the overwhelming presence of benzodioxane units (J) in the comt C4H:F5H1 plants, for which the α- and β-correlations are well resolved. No such correlations exist in the wild-type spectrum. The volume integrals of the correlation contours provide a measure of the relative contribution of each unit type in the lignin polymer (Table 2). In the comt C4H:F5H1 plants, benzodioxanes account for approximately 92% of the inter-unit linkage integrals measured. As noted, the 5H level is approximately 71%; benzodioxane levels are higher because they may result from addition of coniferyl or sinapyl alcohol, as well as 5-hydroxyconiferyl alcohol, to the 5-hydroxyguaiacyl end of the growing polymer, i.e. they can be S, 5H or G benzodioxanes (see also Figure S1).

Phenolic profiling reveals benzodioxane-rich soluble phenolics

To investigate whether perturbation of the monolignol biosynthetic pathway resulted in metabolites derived from 5-hydroxyconiferyl alcohol, a semi-quantitative comparison of the profiles of methanol-soluble phenolics from comt C4H:F5H1 and wild-type inflorescence stems was performed. The profiles of wild-type stems were dominated by sinapate esters and three flavonol glycosides (Figure S4). In contrast, the comt C4H:F5H1 plants had lower levels of sinapate esters and higher levels of flavonol glycosides, and accumulated a large number of phenolic molecules that were below the detection limit in wild-type plants (Figure S4). Elucidating their identities by MS2 spectral analyses (Figure S5) (Morreel et al., 2004a,b; 2010a,b) revealed that these metabolites were 5-hydroxylated phenylpropanoids. Based on their chemical identity, they were grouped into four main classes (Figure 1 and Table 3). Class I consists of oligolignols with at least one and up to four 5H units (111). Class II compounds (1218) consist of hexosylated di- and trilignols with at least one 5H unit. Class III contains heterodimers of 5-hydroxyferulic acid with coniferyl, 5-hydroxyconiferyl or sinapyl alcohol, and their hexose and malate derivatives (1928), and class IV compounds (2938) are a series of derivatized 5-hydroxylated phenylpropanoids.

Table 3.   Metabolites in comt C4H:F5H1 plants, arbitrarily classified for convenience
Peak numberRetention time (min)m/zMolecule numberName
  1. The shorthand naming convention for oligolignols is described by Morreel et al. (2004a,b). In short, G, 5H, S and 5H’ are used for units derived from coniferyl alcohol, 5-hydroxyconiferyl alcohol, sinapyl alcohol and 5-hydroxyconiferaldehyde, respectively. Here, β-O-4 (normally used for lignins) is used instead of 8-O-4 (normally used for metabolites) to ensure consistent nomenclature throughout the text.

  2. aThe hexose is not on the γ-O position of the β-coupled unit.

  3. bThe structure of the 155 Da subunit has not yet been resolved.

Class I: oligolignols
Class II: hexosylated oligolignols
 1713.653512G(β-O-4)5H hexosidea
 1814.053512G(β-O-4)5H hexosidea
 1915.453512G(β-O-4)5H hexosidea
 2012.7551135H(β-O-4)5H hexosidea
 2114.0551135H(β-O-4)5H hexosidea
 2214.4551135H(β-O-4)5H hexosidea
 2316.6549145H(β-O-4)5H’ hexoside
 2415.453715G(β-O-4)γ-O-hexosyl dihydro5H
 2516.356716S(β-O-4)γ-O-hexosyl dihydro5H
 2617.278917G(β-O-4)5H(β-O-4)5H hexoside
 2717.578917G(β-O-4)5H(β-O-4)5H hexoside
 2817.579118G(β-O-4)5H(β-O-4)dihydro5H hexoside
 2917.679118G(β-O-4)5H(β-O-4)dihydro5H hexoside
Class III: 5-hydroxyferulic acid-containing dimers
 3019.038719G(β-O-4)5-hydroxyferulic acid
 3115.254920G(β-O-4)5-hydroxyferuloyl hexose
 3215.954920G(β-O-4)5-hydroxyferuloyl hexose
 339.754921G(β-O-4)5-hydroxyferulic acid hexosidea
 3416.1565225H(β-O-4)5-hydroxyferuloyl hexose
 359.9565235H(β-O-4)5-hydroxyferulic acid hexosidea
 3616.057924S(β-O-4)5-hydroxyferuloyl hexose
 3716.657924S(β-O-4)5-hydroxyferuloyl hexose
 3810.057925S(β-O-4)5-hydroxyferulic acid hexosidea
 3910.150326G(β-O-4)5-hydroxyferuloyl malate
 4010.750326G(β-O-4)5-hydroxyferuloyl malate
 418.0519275H(β-O-4)5-hydroxyferuloyl malate
 428.9519275H(β-O-4)5-hydroxyferuloyl malate
 439.6519275H(β-O-4)5-hydroxyferuloyl malate
 4410.853328S(β-O-4)5-hydroxyferuloyl malate
Class IV: derivatives of 5-hydroxylated phenylpropanoid monomers
 452.7371295-hydroxyferulic acid hexoside
 462.8371295-hydroxyferulic acid hexoside
 478.3371305-hydroxyferuloyl hexose
 489.2371305-hydroxyferuloyl hexose
 4910.5525315-hydroxyferuloyl hexose+155b
 5012.9525315-hydroxyferuloyl hexose+155b
 517.7533325-hydroxyferuloyl hexose hexoside
 5218.856333di-5-hydroxyferuloyl hexose
 533.437334dihydro 5-hydroxyferuloyl hexose
 547.4369355-hydroxyscopoletin hexoside
 5510.3355365-hydroxyconiferaldehyde hexoside
 568.6357375-hydroxyconiferyl alcohol hexoside
 579.2357375-hydroxyconiferyl alcohol hexoside
 585.957938hexosyl 5-hydroxyconiferyl alcohol hexoside
 597.657938hexosyl 5-hydroxyconiferyl alcohol hexoside

Chalcone synthase deficiency reduces C4H-driven gene expression in comt C4H:F5H1 plants

Although the parental comt and C4H:F5H1 lines had a healthy appearance, growth of the comt C4H:F5H1 plants was severely affected. This phenotype is reminiscent of that of hydroxycinnamoyl CoA:shikimate/quinate hydroxycinnamoyl transferase (HCT) down-regulated Arabidopsis plants, whose growth retardation was thought to result from inhibition of auxin transport by the high levels of flavonoids in these mutants (Besseau et al., 2007). Recent findings, however, disproved the existence of this link between flavonoids and growth retardation in HCT down-regulated Arabidopsis plants (Li et al., 2010). To test whether the increased levels of flavonol glycosides (Figure S4) were responsible for growth retardation in the comt C4H:F5H1 plants, a chalcone synthase (chs) mutation was crossed into the comt C4H:F5H1 background. Soil-grown comt C4H:F5H1 chs mutants grew significantly taller than comt C4H:F5H1 plants (Figure 2l and Table 1), and produced viable seeds. However, such restoration of the phenotype could also be caused by negative feedback of the chs mutation on the C4H promoter and thus on C4H:F5H1 expression. To investigate this, quantitative RT-PCR was performed on inflorescence stems. Expression of C4H and F5H1 was significantly reduced in comt C4H:F5H1 chs plants compared to C4H:F5H1 and comt C4H:F5H1 plants (Figure 4a). The benzodioxane level in the lignin of comt C4H:F5H1 chs plants was reduced to 38%, much less than the 92% found in comt C4H:F5H1 plants (Figure S3j and Table 2). Furthermore, comt C4H:F5H1 chs plants had higher lignin amounts compared to comt C4H:F5H1 plants, but less than the wild-type (Table 2). Thus, the partial restoration in growth can be explained by inhibition of C4H:F5H1 expression.

Figure 4.

 Expression of phenylpropanoid biosynthetic genes in the various mutants as determined by quantitative RT-PCR.
(a) Expression of C4H and F5H1 in inflorescence stems. Error bars are standard deviations. Significant differences in expression between the wild type and the mutant (< 0.05) are indicated by an asterisk.
(b) Expression in 14-day-old seedling leaves. Values are the ratio of the expression in the mutant line versus that in the wild type: a value >1 indicates up regulation. Standard deviations are given in parentheses. Red and blue backgrounds indicate higher and lower gene expression relative to the wild type, respectively. The color intensity reflects the strength of induction and reduction. Values that are significantly different from those of the wild type (< 0.05) are shown in bold.

To further explore possible feedback regulation within the phenylpropanoid pathway in comt C4H:F5H1 and comt C4H:F5H1 chs plants, we measured the expression of phenylpropanoid biosynthetic genes in 14-day-old seedling leaves. At this stage, the visual phenotype is much less pronounced (Figure 2a). Strikingly, the expression of many general phenylpropanoid biosynthetic genes was higher in C4H:F5H1, comt C4H:F5H1 and comt C4H:F5H1 chs plants compared to the wild type, suggesting a higher flux through the pathway (Figure 4b). In contrast to inflorescence stems, however, C4H gene expression was similar between comt C4H:F5H1 and comt C4H:F5H1 chs in the seedling leaves, indicating developmental differences in feedback regulation.


Lignins primarily consist of benzodioxane structures

Plants deficient in COMT and plants over-expressing F5H were previously shown to incorporate small amounts of 5-hydroxyconiferyl alcohol into their lignins, demonstrating that 5-hydroxyconiferyl alcohol can be transported, single-electron-oxidized (‘radicalized’), and co-polymerized into lignins (Atanassova et al., 1995; Van Doorsselaere et al., 1995; Jouanin et al., 2000; Guo et al., 2001; Ralph et al., 2001a,b; Sibout et al., 2002; Marita et al., 2003; Morreel et al., 2004a,b; Do et al., 2007; Lu et al., 2010). Its incorporation gives rise to specific units, i.e. benzodioxanes. A benzodioxane results from coupling of a monolignol at its β-position with a 5-hydroxyguaiacyl unit at its 4-O position, followed by internal trapping of the resulting quinone methide by the 5-OH (Ralph et al., 2001a,b). Although these benzodioxane units are the result of typical β–O–4 coupling reactions that also take place in natural G/S-type lignins, the less flexible benzodioxane ring structures alter the macromolecular configuration of the lignin polymer. Strikingly, when Arabidopsis F5H1 is over-expressed in a comt mutant background, the lignin almost exclusively comprises benzodioxane structures, accounting for approximately 92% of the units (Figure S3 and Table 2). This study shows that lignin can be composed almost entirely from units that are undetectable in wild-type plants.

The comt C4H:F5H1 plants accumulate large amounts of 5-hydroxylated phenylpropanoids and their derivatives

Phenolic profiling of the methanol-soluble fraction revealed many 5-hydroxylated phenylpropanoids (including alcohols, aldehydes, acids, and their derivatives) that were undetectable in the extracts of wild-type plants (Figure 1 and Table 3). Class I oligolignols (111) are small lignin polymers with chains of benzodioxane structures, reflecting the high level of 5-hydroxyconiferyl alcohol monomers incorporated into the lignin polymer. Although the phenolic end unit can be a 5H unit, as seen in compound 4, such units are inherently prone to oxidation to quinones and beyond. Capping such end units by coupling with a traditional monolignol stabilizes them. Indeed, most of the oligomers identified were found to be capped in this way. Interestingly, except for the cap, all other units in the identified oligolignols were 5H units, i.e. no mixed oligolignols were found with normal G or S units within the chain. End capping was also observed in class II and III compounds. Thus, although not all differential peaks were structurally resolved, the structures of the most abundant ones suggest that 5-hydroxylated monomers do not readily couple β–O–4 to neither G or S units. Furthermore, it is important to note that these small lignin oligomers involve only β–O–4 binding, because β–5 coupling cannot occur due to the hydroxylation at C5 in the 5H units. Because of the instability of 5H end groups, the only β–β-coupled compounds likely to be found would be tetramers at least, in which both phenolic ends of a β–β-coupled dimer are capped.

Classes II, III and IV comprise hexose and malate derivatives of 5-hydroxyconiferyl alcohol, 5-hydroxyconiferaldehyde or 5-hydroxyferulate, with the latter compound potentially derived from 5-hydroxyconiferaldehyde via hydroxycinnamaldehyde dehydrogenase/reduced epidermal fluorescence 1 (HCALDH/REF1) as described for the conversion from coniferaldehyde and sinapaldehyde to ferulic acid and sinapic acid, respectively (Nair et al., 2004). The class III molecules (1928) are dimers of 5-hydroxyferulate with coniferyl or sinapyl alcohol and their hexose and malate derivatives. These molecules are 5-hydroxy analogs of the G(8-O-4)feruloyl malate that is found in wild-type Arabidopsis (Rohde et al., 2004). Although the biological role of this compound is unknown, G(8-O-4)feruloyl malate has been suggested to be involved in the coupling of hemicellulose to lignin (Rohde et al., 2004). The high abundance of 5-hydroxyferulate esters (3033) in class IV is also noteworthy. The 5-hydroxyferulate esters might functionally replace the biological role of sinapate esters in vivo, because epidermal fluorescence upon illumination with UV (366 nm) is observed in leaves of comt C4H:F5H1 plants, indicative of their UV protective role (Ruegger and Chapple, 2001). This fluorescence is absent from f5h1 (fah1) mutants that produce neither 5-hydroxyferulate nor sinapate esters (Ruegger and Chapple, 2001). We deduce that 5-hydroxylated phenylpropanoids and their derivatives originate partly from passive metabolism via existing biochemical routes, and partly from active detoxification. In passive metabolism, 5-hydroxylated phenylpropanoids are surrogates for their 5-methoxylated and 5-unsubstituted analogs. For example, the presence of di-5-hydroxyferuloyl hexose (33) suggests that the enzymes involved in disinapoyl hexose biosynthesis also recognize and metabolize 5-hydroxylated analogs (Fraser et al., 2007). On the other hand, UDP-glucose-dependent glycosyltransferases (UGTs) have been shown to be involved in the detoxification of xenobiotics (Welinder et al., 2002; Lim and Bowles, 2004; Brazier-Hicks et al., 2007). It is thus anticipated that they will also detoxify the accumulating 5-hydroxylated phenolics, thereby giving rise to the hexosylated compounds of classes II, III and IV.

The subcellular location of synthesis of these compounds remains obscure; oxidative coupling occurs in the cell wall and possibly also in the vacuole, given the high number of peroxidases in this compartment (Welinder et al., 2002). However, glycosylation occurs in the cytoplasm but not in the vacuole, and malate transferase activity is present in the vacuole. Unraveling the subcellular routes for biosynthesis of the class II–IV compounds requires a new research approach that combines subcellular fractionation and phenolic profiling.

Genetic interactions within phenylpropanoid biosynthesis

Although the comt and C4H:F5H1 parental lines have normal lignin levels, comt C4H:F5H1 plants deposit significantly less lignin. Moreover, comt C4H:F5H1 plants remain small and their xylem vessels have an irx phenotype. One hypothesis to explain the lower lignin levels is that 5-hydroxyconiferyl alcohol is not efficiently translocated towards the cell wall and/or not efficiently incorporated into lignin. The reduction in the lignin levels might be sufficient to explain the irx phenotype (Jones et al., 2001; Franke et al., 2002; Goujon et al., 2003; Hoffmann et al., 2004; Besseau et al., 2007; Mir Derikvand et al., 2008; Schilmiller et al., 2009; Li et al., 2010), but the altered structural and physico-chemical properties of the 5H lignin may also contribute to the irx phenotype. In addition to the collapsed xylem, accumulating phenolic precursors might cause aberrant growth, because they are toxic to cellular processes. Indeed, the phenolic profile of comt C4H:F5H1 plants revealed numerous hexosylated (i.e. detoxified) compounds. Accumulation of these compounds is logical given that the sink, lignin, is reduced. However, we also observed a massive up-regulation of multiple phenylpropanoid pathway transcripts in the comt C4H:F5H1 plants (Figure 4). It is tempting to believe that the reduction in cell-wall integrity, possibly due to the shortage of lignin in the cell wall, results in a greater demand for biosynthesis of coniferyl and sinapyl alcohols, and thus a higher flux through the phenylpropanoid pathway (Figure 4). Up-regulation of the phenylpropanoid pathway might thus be part of the plant’s response to an aberrant cell-wall structure (Caño-Delgado et al., 2003; Rogers and Campbell, 2004).

We also tested whether the growth abnormalities in the comt C4H:F5H1 plants could have been caused by the higher flavonoid levels, as suggested by Besseau et al. (2007). However, in contrast with their results, and in support of the recent data by Li et al. (2010), our data indicate that flavonoids are certainly not the sole cause for the growth defect in comt C4H:F5H1 plants, as crossing in a chs mutation was not able to fully restore the phenotype. Instead, the main reason for the partial recovery in growth observed in comt C4H:F5H1 chs plants was reduction of C4H-driven gene expression caused by the chs mutation, resulting in lower over-expression of the F5H1 gene and a consequent a drop in the frequency of benzodioxane units (resulting from a reduction in the proportion of 5-hydroxyconiferyl alcohol monomers) and an increase in total lignin content (Table 2). The reason for reduction of this C4H-driven gene expression may either be negative feedback on the C4H promoter, or positive feedback on the C4H promoter that subsequently triggered gene silencing. In either case, the data clearly demonstrate a genetic interaction between the flavonoid and phenylpropanoid pathways. Note that this interaction in inflorescence stems was dependent on the genetic background: the chs mutation reduced C4H-driven gene expression only in comt C4H:F5H1 chs plants, but not in the single chs mutant. In addition, the reduction of C4H-driven gene expression appeared to be dependent on the developmental stage, as it was observed in stems but not in seedling leaves (Figure 4). It is furthermore important to note that the C4H promoter is one of the promoters of choice to drive transgene expression in lignifying cells in order to modify lignin biosynthesis in crop species (Huntley et al., 2003; Stewart et al., 2009). The observation that the expression driven by this promoter is regulated, possibly in response to altered concentrations of pathway intermediates, has repercussions on its use in gene engineering strategies. Such poorly studied genetic interactions within the phenylpropanoid pathway, for which full elucidation requires a systems approach, need to be taken into account during rational design of plant cell walls with improved properties for agro-industrial applications.

Experimental Procedures

Plant material

All experiments were performed with Arabidopsis ecotype Columbia (Col-0). The comt mutant (At5g54160, SALK_002373) and chs mutant (At5g13930, SALK_020583) were obtained from the SALK collection (Alonso et al., 2003). Zygosity was analyzed using primers 1 (5′-ATGCCAAAGAGTCATGTTTTA-3′) and 2 (5′-TGGTGTCTCTGGAAGTATAC-3′) to amplify the endogenous COMT gene, and primers 1 and 3 (5′-GCCCTTTGACGTTGGAGTC-3′) to amplify the comt T-DNA left-border insertion region. For CHS, primers 4 (5′-CTCGTCGGTCAGGCTCTTT-3′) and 5 (5′-TTCCATACTCGCTCAACACG-3′) were used for the endogenous gene, and primers 3 and 4 for the chs T-DNA left-border insertion region. The C4H:F5H1 transgenic line (C4H-F5H) in the fah1-2 mutant background (Meyer et al., 1998) was kindly provided by Clint Chapple (Department of Biochemistry, Purdue University, IN). This line over-expresses F5H1 (At4g36220) under the control of the C4H (At2g30490) promoter. The presence of the C4H:F5H1 construct was analyzed using primers 5′-CGTCGTGGGATCTGATAGTT-3′ and 5′-CTCCCTCTTATAAAATCCTCTC-3′. To confirm the presence of the fah1-2 point mutation, primers 5′-ATGGAGAACTTTGCCTCCTG-3′ and 5′-GGATAAACAAGCGGCTCGT-3′ were used. The C4H:F5H1 transgene is not amplified by these primers. The 907 bp PCR product was further digested using MseI (New England Biolabs Inc.,, resulting in DNA fragments of 398, 225, 221, 56 and 7 bp for the wild-type F5H1 and 267, 225, 221, 131, 56 and 7 bp for the mutant fah1-2 allele. The comt C4H:F5H1 fah1-2, comt C4H:F5H1 and C4H:F5H1 lines were isolated from the F2 progeny of a comt x C4H-F5H1 fah1-2 cross. Because comt C4H:F5H1 fah1-2 and comt C4H:F5H1 plants are sterile, these genotypes were generated from the parental plants heterozygous for comt and homozygous for C4H:F5H1 and fah1-2. All experiments described were performed on comt C4H:F5H1 plants. However, comt C4H:F5H1 fah1-2 plants had similar phenotypes.

Plant growth conditions

Agar-grown plants were grown at 23°C, with 16 h of light per day, on the following medium: 0.43% Murashige and Skoog vitamin medium/basal salt mixture (M0221.0050, Duchefa Biochemie,, 1% sucrose (177140010, Acros,, 0.05% MES (M1503.0100, Duchefa Biochemie), pH 5.7, 0.725% agar (A1296, Sigma, and 0.1% 1000× Murashige and Skoog vitamin solution (M3900, Sigma). Soil-grown plants were germinated directly in Saniflor soil (Van Israel; supplemented with 10% v/v vermiculite (16 h light, 22°C, 55% humidity).


For biomass measurements, plants were grown in soil until senescence. Cauline leaves and siliques were removed from the inflorescence before weighing. Statistical analysis was performed using anova followed by the Scheffe post hoc test.


Electron microscopy and lignin staining were performed essentially as described previously (Rohde et al., 2004). Stem segments were cut into small pieces and fixed with 4% paraformaldehyde and 2.5% glutaraldehyde, and post-fixed in 1% OsO4 with 1.5% K3Fe(CN)6 in 0.1 m sodium cacodylate buffer, pH 7.2. Samples were dehydrated through a graded ethanol series, including bulk staining with 2% uranyl acetate at the 50% ethanol step, followed by embedding in Spurr’s resin. For tissue morphology, semi-thin sections (0.5 μm) were cut using an Ultracut E microtome (Reichert-Jung,, stained in a 1% toluidine blue and 2% borate solution in distilled water, and viewed using a Leica DM LB microscope (

For electron microscopy, ultra-thin sections of a gold interference color were produced on an Ultracut E microtome (Reichert-Jung), and then post-stained in an ultrastainer (Leica) using uranyl acetate and lead citrate. They were viewed using a JEOL 1010 transmission electron microscope (JEOL,

Lignin histochemistry was performed by Mäule staining as previously described (Rohde et al., 2004). Briefly, agar-imbedded stem segments were sectioned with a vibroslicer, incubated in 4% KMnO4 for 5 min, rinsed with water, incubated in 37% HCl/H2O (1:1) for 2 min, and observed after addition of a drop of aqueous NH3. Sections were viewed under a binocular microscope (Leica MZ16).

Phenolic profiling and MS analysis

Single inflorescences, lacking cauline leaves, flowers and siliques, were sampled and immediately frozen in liquid nitrogen. Ten whole tissue-culture grown stems between 2.2 and 6.0 cm long were obtained from comt C4H:F5H1 plants and compared with five wild-type whole stems of approximately 8.0 cm, as well as five apical and five basal stem segments of 5.0 cm from approximately 20 cm tall wild-type plants. Samples were transferred to 1.5 ml Eppendorf tubes. After grinding, the samples were extracted in 600 μl THF/isopropanol/methanol (15:15:70). The extract (550 μl) was freeze-dried in a SAVANT Speedvac (SC21A SpeedVac Concentrator, Thermo,, and subjected to extraction using water/cyclohexane (100 μl of each). Each individual sample (60 μl) was injected by means of a SpectraSystem AS1000 autosampler (Thermo Separation Products) onto a reverse-phase Luna C18(2) column (150 × 2.1 mm, 3 μm; Phenomenex,

Gradient separation was performed on a SpectraSystem P1000XR HPLC pump (Thermo Separation Products), using a gradient of two buffers: buffer A (water/acetonitrile (ACN)/acetic acid, 100:1:0.1, pH 2) and buffer B (ACN/water/acetic acid, 100:1:0.1, pH 2). A flow rate of 0.25 ml min−1 was used, with a buffer composition starting at 5% B, linearly increasing to 17% at 1 min, 77% at 33 min, and finally 100% at 36 min. A SpectraSystem UV6000LP detector (Thermo Separation Products) was used to measure UV/vis absorption between 200 and 450 nm with a scan rate of 2 Hz. Atmospheric pressure chemical ionization (APCI) operated in the negative-ion mode, was used as an ion source to couple HPLC to LCQ Classic MS instrument (ThermoQuest) (vaporizer temperature 350°C, capillary temperature 140°C, source current 5 mA, sheath gas flow set at 27, auxiliary gas flow set at 3, m/z range 140–1000, normalized collision energy for MS2 35).

NMR sample preparation

NMR samples were prepared essentially as described previously (Weng et al., 2010). Senesced inflorescence stems of Arabidopsis (approximately 200 mg) from tissue culture- and soil-grown plants (Figures S1 and S2) were pre-ground in a Retsch MM400 mixer mill (, equipped with a 10 ml ZrO2 vessel and two ZrO2 ball-bearings (12 mm diameter), for 1 min at 30 Hz. The ground material was extracted three times with water and subsequently three times with 80% ethanol by sonicating in an ultrasonic bath for 30 min each time. The freeze-dried extractive-free Arabidopsis samples (approximately 100 mg) were ball-milled using a Retsch PM 100 planetary ball mill in a 50 ml ZrO2 vessel with 10 ZrO2 ball-bearings (10 mm diameter) at 600 rpm, with 5 min breaks after every 5 min of milling. The total ball-milling time for the samples was 16.5 min. Cellulolytic enzyme lignins (CEL) were then isolated by enzymatically saccharifying (most of the) polysaccharides as described by Chang et al. (1975). Cellulysin cellulase (EC, Calbiochem,, a crude cellulase preparation containing hemicellulase activities, from Trichoderma viride was used. Its activity was 10 000 units g−1 dry weight. The ball-milled material was suspended in 20 mm NaOAc buffer (30 ml, pH 5.0) in a 50 ml centrifuge tube, 8 mg of Cellulysin was added, and the reaction slurry was incubated at 35°C for 48 h. The solids were pelleted by centrifugation (1500 g, 21°C, 40 min), and the process was repeated with fresh buffer and enzyme. CEL was recovered by filtration through a nylon membrane (0.2 μm). The product was washed with water and dried under vacuum at room temperature, yielding 16–28 mg CEL, depending on the sample. CEL (12 mg) was acetylated using 2 ml DMSO, 1 ml N-methylimidazole and 0.6 ml acetic anhydride, stirring for 1.5 h at room temperature, before quenching the reaction by pouring the solution into 1 L of distilled water. The precipitated material was recovered by filtration through a nylon membrane (0.2 mm). Acetylated CEL, typically approximately 15–18 mg from 12 mg CEL, was dried under vacuum and dissolved in 0.5 ml CDCl3 for NMR analysis. For the comt C4H:F5H1 line, only approximately 3–4 mg of acetylated CEL was obtained after filtering out unidentified gelatinous CDCl3-insoluble material. Nevertheless, the 2D HSQC spectrum was of excellent quality, and it was even possible to obtain a 3D TOCSY-HSQC spectrum from this sample.

NMR experiments

NMR spectra were acquired on a Bruker-Biospin AVANCE 500 MHz spectrometer (Bruker-Biospin, fitted with a cryogenically cooled 5 mm Bruker TCI (triple resonance cryoprobe optimized for 1H and 13C observation) gradient probe with inverse geometry (proton coils closest to the sample). The 2D 13C–1H correlation spectra were acquired using an adiabatic HSQC pulse program (Bruker standard pulse sequence ‘hsqcetgpsisp2.2′) (Kupče and Freeman, 2007) and the following parameters: spectra were acquired from 10 to 0 ppm (5000 Hz) in F2 (1H) using 1000 data points for an acquisition time (AQ) of 200 msec, an interscan delay (D1) of 500 msec, and from 196 to −23 ppm (25 154 Hz) in F1 (13C) using 400 increments (F1 acquisition time 8 msec) of 90 scans, for a total acquisition time of 17 h 17 min. The number of scans can be adjusted depending on the signal-to-noise required from a sample. The 1JCH used was 145 Hz. Processing used typical matched Gaussian apodization in the 1H dimension and squared cosine-bell apodization in the 13C dimension. Prior to Fourier transformation, the data matrices were zero-filled to 1024 points in the 13C dimension. The central chloroform solvent peak at a δCH of 77.0/7.26 ppm was used as an internal reference. The 3D TOCSY-HSQC spectrum was acquired and processed exactly as described previously, providing similar evidence for the benzodioxane units (Marita et al., 2001).

G:5H:S integral ratios were obtained by integrating the contours from the 2 or 2,6 positions of each type of aromatic unit (with S integrals being logically halved as the 2 and 6 positions are identical in the symmetrical S units). In samples with 5H correlations, where the 5H6 correlations overlap with the S2/6 correlations, the S integral was calculated by subtracting the 5H2 integral (which should be the same as the 5H6 integral) from the integral of the overlapping S2/6 + 5H6 contours (Figure 3b–e).

Lignin quantification

Cell-wall material was prepared from 5 mg of senescent inflorescence stems of soil-grown plants, washed sequentially with 0.5 ml water, ethanol, chloroform and acetone for 30 min each at 98, 76, 59 and 45°C, respectively. Acetyl bromide lignin extraction was performed according a downscaled method similar to that described previously (Dence, 1992). Briefly, 0.1 ml acetyl bromide (25% in acetic acid) and 4 μl 60% perchloric acid were added to the dry cell wall and the mixture was incubated at 70°C for 30 min. NaOH (2 m, 0.2 ml) and 0.5 ml acetic acid were then added. After centrifugation (15 000 g, room temperature, 15 min), the supernatant was separated from the pellet. The pellet was further washed by adding 0.5 ml acetic acid (vortex, 5 s), followed by centrifugation (15 000 g, room temperature, 15 min). The second supernatant was combined with the first. Acetic acid was added to the combined supernatants to a volume of 2 ml, and absorption was measured at 280 nm using a NanoDrop ND-1000 spectrophotometer (Nanodrop, An extinction coefficient for lignin at 280 nm of 23.35 L g−1 cm−1 was used (Chang et al., 2008). As for every lignin quantification method, the response varies with the composition of the lignin; the acetyl bromide protocol gives only an estimate of the total lignin amount (Hatfield and Fukushima, 2005) and we used the same lignin UV280 extinction coefficient for all samples here. Statistical tests were performed as indicated for biomass measurements.

RNA extraction, cDNA preparation and quantitative RT-PCR

Total RNA was extracted from frozen soil-grown inflorescence stems and tissue-cultured seedling leaves using a plant RNeasy extraction kit (Qiagen,, and quantified using a NanoDrop ND-1000 spectrophotometer. Total RNA (4.5 μg) was reverse-transcribed in a final volume of 190 μl using a Superscript II reverse transcription kit (Invitrogen, Quantitative RT-PCR was performed using a SYBR Green I master kit (Roche, on a Lightcycler 480 (Roche). The primer sequences used for quantitative RT-PCR are given in Table S1. Gene expression was normalized on the basis of ACTIN expression. Statistical tests were performed using a procglm nested model with unequal variances (SAS 9.2; SAS Institute Inc.,


The authors thank Clint Chapple (Department of Biochemistry, Purdue University, West Lafayette, IN) for kindly providing the C4H:F5H1 fah1-2 line, Bart Ivens and David Casini for practical assistance, and Martine De Cock for help in preparing the manuscript. We gratefully acknowledge partial funding through the United States Department of Energy (DOE) Energy Biosciences program (grant number DE-AI02-00ER15067) and the DOE Great Lakes Bioenergy Research Center (DOE Office of Science, grant number BER DE-FC02-07ER64494) to J.R., the Research Foundation-Flanders (grant number G.0352.05N), the European Community's 7th Framework Programme (FP7/2007) under grant agreement no. 211982 (RENEWALL), the Global Climate and Energy Project (GCEP) (grants to W.B. for ‘Towards New Degradable Lignin Types’ and to J.R. for ‘Efficient Biomass Conversion: Delineating the Best Lignin Monomer Substitutes’), and the Multidisciplinary Research Partnership ‘Biotechnology for a Sustainable Economy’ (01MRB510W) of Ghent University. Some of the NMR experiments on the Bruker DMX-500 cryoprobe system made use of the National Magnetic Resonance Facility at Madison ( R.V. is indebted to the Agency for Innovation by Science and Technology for a pre-doctoral fellowship.