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. 2010 Apr 6;22(4):1033–1045. doi: 10.1105/tpc.109.073528

Convergent Evolution of Syringyl Lignin Biosynthesis via Distinct Pathways in the Lycophyte Selaginella and Flowering Plants[C],[W]

Jing-Ke Weng a,1, Takuya Akiyama b, Nicholas D Bonawitz a, Xu Li a, John Ralph b,c, Clint Chapple a,2
PMCID: PMC2879749  PMID: 20371642

This study shows that the independent origin of syringyl lignin in the lycophyte Selaginella involved the elaboration of a biochemical pathway that bypasses four steps of the canonical lignin biosynthetic pathway established in flowering plants.

Abstract

Phenotypic convergence in unrelated lineages arises when different organisms adapt similarly under comparable selective pressures. In an apparent example of this process, syringyl lignin, a fundamental building block of plant cell walls, occurs in two major plant lineages, lycophytes and angiosperms, which diverged from one another more than 400 million years ago. Here, we show that this convergence resulted from independent recruitment of lignin biosynthetic cytochrome P450-dependent monooxygenases that route cell wall monomers through related but distinct pathways in the two lineages. In contrast with angiosperms, in which syringyl lignin biosynthesis requires two phenylpropanoid meta-hydroxylases C3′H and F5H, the lycophyte Selaginella employs one phenylpropanoid dual meta-hydroxylase to bypass several steps of the canonical lignin biosynthetic pathway. Transgenic expression of the Selaginella hydroxylase in Arabidopsis thaliana dramatically reroutes its endogenous lignin biosynthetic pathway, yielding a novel lignin composition not previously identified in nature. Our findings demonstrate a unique case of convergent evolution via distinct biochemical strategies and suggest a new way to genetically reconstruct lignin biosynthesis in higher plants.

INTRODUCTION

In response to environmental pressures, phylogenetically unrelated species sometimes arrive at similar adaptive solutions through independent mechanisms (Tanaka et al., 2009). Although convergent evolution happens rarely in nature, it represents an important evolutionary phenomenon, which has been documented in action at multiple levels of biological processes (Conant and Wagner, 2003). For example, birds and bats independently evolved wings for powered flight (Hedenstrom et al., 2007), both higher plants and fungi developed the ability to synthesize growth regulator gibberellins via nonorthologous pathways (Hedden et al., 2001), and mammals and fungi recruited two distinct families of proteins to methylate Lys residues on histone tails (Cheng et al., 2005). Elucidation of the molecular mechanisms underlying phenotypic convergence is key to understanding how this process contributes to evolution.

Vascular plants arose in the Late Silurian period (∼420 million years ago). They diversified rapidly during the Early Devonian period (416 to 398 million years ago), when an early split in the history of land plant evolution gave rise to two major lineages: the lycophytes and euphyllophytes (Kenrick and Crane, 1997) (Figure 1A). These two lineages are united as a monophyletic group by the presence of specialized water-conducting tracheary elements, the cell walls of which are physically reinforced by the phenolic lignin heteropolymer (Kenrick and Crane, 1997). Lignin endows vascular plants with the rigidity to stand upright, prevents their tracheids and vessel elements from collapsing during long-distance water transport, and has greatly contributed to the dominance of these plants in terrestrial environments (Boerjan et al., 2003). Although lignin appears to be fundamental to the biochemistry of all vascular plants, its monomer composition exhibits an intriguing distribution pattern across the major lineages. Among the euphyllophytes, ferns and gymnosperms generally contain p-hydroxyphenyl (H) and guaiacyl (G) lignin units, derived from p-coumaryl alcohol and coniferyl alcohol, respectively, and angiosperms additionally contain syringyl (S) units derived from sinapyl alcohol (Boerjan et al., 2003) (Figure 1). This observation has led to the notion that S lignin is a recent invention restricted primarily to angiosperms. In fact, S lignin has also been found in some lycophyte species belonging to the order of Selaginellales but is absent from species belonging to its extant sister order Lycopodiales and is thought to have been absent from its extinct sister order Lepidodendrales, suggesting that S lignin might have arisen independently in lycophytes and euphyllophytes by evolutionary convergence (Towers and Gibbs, 1953; White and Towers, 1967; Logan and Thomas, 1985, 1987; Jin et al., 2005; Weng et al., 2008c) (Figure 1A).

Figure 1.

Figure 1.

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S Lignin Biosynthesis via Distinct Biosynthetic Routes in Angiosperms and the Lycophyte Selaginella.

(A) Plant phylogenetic tree showing the distribution of lignin monomer composition across major land plant lineages. Open circle, no lignin; yellow circle, presence of H and G lignin; red circle, presence of S lignin in addition to H and G lignin. †Lignin composition indicated by the analysis of fossil samples of extinct Sigillaria species (Logan and Thomas, 1987).

(B) Alternative pathways toward S lignin biosynthesis in Selaginella and angiosperms. The shared routes between Selaginella and angiosperms are shaded in purple, whereas the divergent branches specific to Selaginella or angiosperms are shaded in pink or light blue, respectively. The two aromatic ring meta-hydroxylation steps are catalyzed by one bifunctional enzyme, Sm F5H (circled in red) in Selaginella, instead of two enzymes, C3′H and F5H (circled in blue), in angiosperms. At, Arabidopsis thaliana; Sm, Selaginella moellendorffii. PAL, phenylalanine ammonia-lyase; 4CL, 4-hydroxycinnamoyl-CoA ligase; CCoAOMT, caffeoyl-CoA O-methyl transferase; HCALDH, hydroxycinnamaldehyde dehydrogenase; COMT, caffeic acid O-methyltransferase; CAD, (hydroxy)cinnamyl alcohol dehydrogenase.

Lignin monomer biosynthesis in angiosperms is largely dependent on the expression of three cytochrome P450 monooxygenases (P450s; Figure 1B) (Boerjan et al., 2003). Cinnamate 4-hydroxylase (C4H) catalyzes the phenylpropanoid ring para-hydroxylation reaction that is required for the formation of all three types of lignin units (Teutsch et al., 1993), but two distinct P450s carry out each of the two subsequent hydroxylations required for S lignin synthesis. p-Coumaroyl shikimate 3′-hydroxylase (C3′H) catalyzes the first ring meta-hydroxylation reaction, necessary for the biosynthesis of both G and S lignin units (Franke et al., 2002a), and ferulate 5-hydroxylase (F5H) catalyzes the second ring meta-hydroxylation reaction leading to the biosynthesis of S lignin units from G-substituted precursors (Meyer et al., 1998).

We recently isolated from the lycophyte Selaginella moellendorffii a novel P450 (CYP788A1) that, like angiosperm F5Hs, can divert G-substituted intermediates toward S lignin synthesis (Weng et al., 2008c). Selaginella F5H (Sm F5H) shares only 37% amino acid sequence identity with its angiosperm counterparts, a level of similarity that can be expected from any of two random plant P450 enzymes from families with unrelated functions, suggesting that the similar catalytic activities of F5Hs in the two lineages were derived from convergent evolution (Weng et al., 2008c). Here, we report the discovery of a novel activity of Sm F5H that reveals an unexpected metabolic distinction between Selaginella and angiosperms and suggests that S lignin is elaborated via distinct biosynthetic routes in these two groups of plants.

RESULTS

Sm F5H Is a Phenylpropanoid Dual Meta-Hydroxylase in Vitro

In our previous study, we demonstrated that like angiosperm F5Hs, Sm F5H uses coniferaldehyde and coniferyl alcohol as substrates in preference to ferulic acid (Humphreys et al., 1999; Weng et al., 2008c). To test whether Sm F5H can use phenylpropanoids other than those G-substituted intermediates as substrates, we performed a series of kinetic assays using Sm F5H against a wider range of phenylpropanoid pathway intermediates. Parallel comparative assays were conducted using Arabidopsis thaliana F5H (At F5H) recombinant protein.

Surprisingly, we found that although both At F5H and Sm F5H can catalyze 5-hydroxylation reactions on G-substituted intermediates equally well, Sm F5H can also efficiently catalyze the 3-hydroxylation of p-coumaraldehyde and p-coumaryl alcohol (Table 1; see Supplemental Figure 1 online). In contrast with Sm F5H, At F5H shows little activity toward p-coumaraldehyde and p-coumaryl alcohol with Km values so high that these activities are not likely to be relevant in vivo (Table 1; see Supplemental Figure 1 online). The similarly ring-substituted acid, p-coumaric acid, is not an optimal substrate for Sm F5H, consistent with the high Km value observed for ferulic acid. Neither Sm F5H nor At F5H showed any detectable activity toward cinnamic acid, cinnamaldehyde, and cinnamyl alcohol, indicating the ring para-hydroxyl is required for the meta-hydroxylase activity. When p-coumaroyl shikimic acid, the substrate for angiosperm C3′H (Schoch et al., 2001), was tested, no activity was detected in either Sm F5H or At F5H assays, suggesting that Sm F5H does not display promiscuous activity toward any given p-coumaroyl derivatives. We also detected no activity of Sm F5H toward caffeic acid, caffealdehyde, or caffeyl alcohol, suggesting that the 3-hydroxyl O-methylation is required for the subsequent 5-hydroxylase activity of Sm F5H. Collectively, the kinetic data imply that Selaginella may have a pathway for S lignin biosynthesis via the H-substituted aldehyde and alcohol, a route that is thought to be absent in angiosperms (Figure 1B).

Table 1.

Kinetic Properties of Sm F5H and At F5H against Phenylpropanoid Intermediates

Sm F5H
 At F5H
Substrate Kma (μM) Vmaxa (pkat·mg−1) Vmax/Km Kma (μM) Vmaxa (pkat·mg−1) Vmax/Km
p-Coumaric acid 1.67 × 103 1.67 1.00 × 10−3 N.D.b N.D.b N.D.b
p-Coumaraldehyde 4.50 4.45 0.989 3.71 × 102 0.700 1.89 × 10−3
p-Coumaryl alcohol 5.11 3.58 0.701 5.95 × 102 0.606 1.02 × 10−3
Ferulic acid 1.09 × 103 1.35 1.24 × 10−3 1.85 × 103 2.03 1.10 × 10−3
Coniferaldehyde 2.80 3.40 1.21 3.06 2.18 0.712
Coniferyl alcohol 3.63 2.56 0.705 1.76 1.73 0.983
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No activity was detected using either Sm F5H or At F5H against the following substrates at a concentration of 200 μM: cinnamic acid, cinnamaldehyde, cinnamyl alcohol, p-coumaroyl shikimic acid, caffeic acid, caffealdehyde, and caffeyl alcohol.

a

Extrapolated using Lineweaver-Burk plots based on triplicate assays. Note that the enzyme assays were performed using yeast microsomes containing the heterologously expressed catalysts in which the amount of P450 was not quantified. The kinetic constants involving Vmax cannot be compared between assays conducted using Sm F5H and At F5H.

b

Kinetic constants not determined (N.D.; no detectable activity).

Sm F5H Partially Rescues the Growth Defects of the Arabidopsis C3′H-Deficient Mutants

In Arabidopsis, mutants defective in each of the two phenylpropanoid meta-hydroxylases, C3′H and F5H, have been isolated and characterized. Whereas various alleles of the C3′H-deficient reduced epidermal fluorescence8 (ref8) mutants exhibit severe dwarfism, female sterility, greatly reduced soluble sinapate ester and total lignin content, and a lignin composed of almost pure H units (Franke et al., 2002a, 2002b; Abdulrazzak et al., 2006), the F5H-deficient fah1-2 mutant shows a total loss of sinapate esters and S lignin but normal growth (Chapple et al., 1992). We have shown that the Sm F5H transgene can rescue fah1-2 mutant biochemical phenotypes (Weng et al., 2008c), but our revised kinetic analysis of the enzyme's substrate specificity suggested that this complementation experiment may have exploited only a portion of the catalytic repertoire of Sm F5H. We postulated that if the phenylpropanoid dual meta-hydroxylase activity of Sm F5H observed in vitro is relevant in vivo, transgenic expression of Sm F5H would also be able to rescue ref8 because the p-coumaraldehyde and p-coumaryl alcohol that the mutant employs for H lignin synthesis would be available as substrates for the enzyme.

To test this hypothesis, we generated Sm F5H transgenics in ref8 fah1-2 double mutant backgrounds by crossing plants carrying one of two ref8 alleles, ref8-1 (a slightly leaky allele that carries a point mutation as described by Franke et al. [2002b]) and ref8-2 (a T-DNA insertional null allele), with four independent transgenic lines previously generated in the fah1-2 background that harbor the Sm F5H transgene under the control of the Arabidopsis C4H promoter (AtC4H:SmF5H), a promoter that has been shown to efficiently target expression in vascular tissue (Weng et al., 2008c). In both cases, visual inspection of the resulting ref8/fah1-2/AtC4H:SmF5H plants from the F2 generation indicated a partial but substantial complementation of the growth phenotype (Figures 2 and 3). Compared with ref8 and ref8 fah1-2 plants, which are severe dwarfs with miniature rosettes, ref8/fah1-2/AtC4H:SmF5H plants are significantly larger in stature (Figures 2 and 3). The dark-green/purple color typically observed in ref8 or ref8 fah1-2 rosette leaves is also greatly alleviated in the Sm F5H transgenics, indicating a decrease in the accumulation of anthocyanins. Despite the considerable complementation in growth phenotype, the female sterility phenotype of ref8 is not rescued in the Sm F5H transgenics, suggesting that the alternative phenylpropanoid meta-hydroxylation pathway mediated by Sm F5H is not sufficient to compensate for the loss of C3′H activity in flower development in ref8 or that the At C4H promoter does not target Sm F5H expression to the necessary tissues or cells.

Figure 2.

Figure 2.

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Partial Complementation of the Growth Phenotype of the Arabidopsis ref8 fah1-2 Double Mutant by the Sm F5H Transgene.

(A) Columbia wild type.

(B) fah1-2.

(C) ref8-1.

(D) ref8-1 fah1-2.

(E) A representative line of ref8-1/fah1-2/AtC4H:SmF5H.

(F) ref8-2.

(G) ref8-2 fah1-2.

(H) A representative line of ref8-2/fah1-2/AtC4H:SmF5H.

All plants were photographed at 2 months of age, and all images are shown at an identical scale.

[See online article for color version of this figure.]

Figure 3.

Figure 3.

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Photographs of Rosette-Stage Arabidopsis Plants under Visible Light and UV Light.

Blue fluorescence under UV light indicates the presence of sinapoylmalate in the leaf epidermis, whereas red fluorescence indicates its absence and results from the UV-induced chlorophyll fluorescence. Top, visible light; bottom, UV light.

To test whether overexpression of At F5H can rescue ref8, we generated similar transgenic plants carrying an At C4H promoter-driven At F5H transgene (AtC4H:AtF5H). None of the resulting ref8-1/fah1-2/AtC4H:AtF5H transgenic lines showed any sign of phenotypic complementation (data not shown), which is consistent with the in vitro observation that At F5H is not effective as a 3-hydroxylase. The above data suggest that the complementation of ref8 by Sm F5H is due to its specific 3-hydroxylase activity and not the 5-hydroxylase activity it shares with At F5H.

Sm F5H Restores S Lignin Biosynthesis in the Arabidopsis C3′H-Deficient Mutants

To examine whether the rescue of the growth phenotype is associated with a restoration of normal lignin deposition in the ref8/fah1-2/AtC4H:SmF5H transgenics, we first performed phloroglucinol-HCl and Mäule histochemical staining on stem cross sections (Figure 4). The phloroglucinol-HCl reagent gives a red reaction when it reacts with hydroxycinnamaldehyde end groups in the lignin polymer, whereas the Mäule reagent gives a qualitative indication of lignin monomer composition by staining G lignin brown and S lignin red. Compared with the wild type and fah1-2, ref8-1 and the ref8-1 fah1-2 double mutant show much weaker phloroglucinol-HCl staining and no Mäule staining, consistent with decreased total lignin and lack of G and S lignin units in these mutants. By contrast, the ref8-1/fah1-2/AtC4H:SmF5H transgenics exhibited strong red Mäule staining in both xylem and interfascicular fiber cells, indicating that S lignin deposition is restored in these cells. Little phloroglucinol-HCl staining was detected in sections of these plants, suggesting low quantities of aldehyde end groups in the transgenic lignin, a character reminiscent of high S lignin as previously reported (Franke et al., 2000).

Figure 4.

Figure 4.

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Impact of Sm F5H Transgenic Expression on Lignin Histochemical Staining in the Arabidopsis ref8 fah1-2 Double Mutant.

Phloroglucinol-HCl (top) and Mäule (bottom) histochemical staining of 2-month-old Arabidopsis stem cross sections. The phloroglucinol-HCl reagent detects aldehyde groups contained in lignin and results in red staining that is generally indicative of the presence of lignin. The Mäule reagent is diagnostic for the presence of S units in lignin, leading to a yellow staining of G lignins and red staining of lignins containing S subunits.

We then quantified the total lignin content of the transgenic plants together with the control plants by Klason analysis (Figure 5). Compared with ref8-1 and the ref8-1 fah1-2 double mutant, which contain only about one-third of the Klason lignin content found in the wild type, the ref8-1/fah1-2/AtC4H:SmF5H transgenics deposit about three-quarters of the wild-type level of lignin. Interestingly, the ref8-2/fah1-2/AtC4H:SmF5H transgenics also have a Klason total lignin content similar to the ref8-1/fah1-2/AtC4H:SmF5H transgenics, despite their phenotypic difference in growth (Figures 2E and 2H). This observation is consistent with the previous suggestion that deficiency in factors other than lignin (e.g., some unknown growth substance synthesized via C3′H) may contribute to the dwarfism in ref8 mutants (Abdulrazzak et al., 2006).

Figure 5.

Figure 5.

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Restoration of Total Acid-Insoluble Lignin Content by Sm F5H in the Arabidopsis ref8 fah1-2 Double Mutant Determined by Klason Analysis.

Error bars represent 1 sd of biological triplicates. N.D., not determined (no inflorescence stems could be harvested from these plants).

We further examined the impact of Sm F5H on lignin monomer composition in the ref8 fah1-2 background using the derivatization followed by reductive cleavage (DFRC) method, a procedure that specifically releases lignin monomers (as their peracetates) from β-O-4-linked lignin units (Figure 6, Table 2; see Supplemental Figure 2 online). As mentioned previously, compared with the wild type, which contains both G and S units with traces of H units, S units in fah1-2 are below detectable limits. The ref8-1 fah1-2 line resembles ref8-1 in that it deposits essentially only H subunits and only accumulates low levels of lignin. Both ref8-1/fah1-2/AtC4H:SmF5H and ref8-2/fah1-2/AtC4H:SmF5H transgenics show a significant recovery of the total DFRC-releasable lignin monomer yield. More surprisingly, these plants contain lignin with comparable amounts of releasable H and S units, but very few G units, a lignin composition not previously identified in nature.

Figure 6.

Figure 6.

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Impact of Sm F5H Transgenic Expression on Lignin Composition in the Arabidopsis ref8 fah1-2 Double Mutant Examined by DFRC–Gas Chromatography Analysis.

H/G/S are p-hydroxyphenyl/guaiacyl/syringyl lignin-derived hydroxycinnamyl alcohol peracetates. c/t, cis/trans; IS, internal standard.

[See online article for color version of this figure.]

Table 2.

Lignin Monomer Composition of Arabidopsis Plants Inferred from DFRC Lignin Analysis and the Contour Integration in HSQC NMR Analysis

DFRC Releasable Monomer (μmol g−1 Cell Wall ± sd)
 DFRC Inferred H:G:S (Mol %) NMR Inferred H:G:S (Mol %)
Sample H G S
Columbia wild type 3.1 ± 1.1 49 ± 3.8 20 ± 0.62 4.4:68.2:27.4 0.1:81.5:18.4
fah1-2 3.2 ± 0.52 64 ± 1.2 N.D. 4.8:95.2:0.0 0.1:99.9:0.0
ref8-1 4.0 ± 0.50 N.D. N.D. 100.0:0.0:0.0 98.9:0.0:1.1
ref8-1 ah1-2 4.3 ± 0.33 N.D. N.D. 100.0:0.0:0.0 N.A.
ref8-1/fah1-2/AtC4H:SmF5H 22 ± 1.2 3.7 ± 0.11 47 ± 1.6 29.9:5.1:65.0 23.7:2.1:74.2
ref8-2/fah1-2/AtC4H:SmF5H 22 ± 1.6 2.5 ± 0.52 22 ± 1.2 46.8:5.4:47.8 48.7:3.8:47.5
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Standard deviation was calculated based on biological triplicates. N.D., not detectable; N.A., not analyzed.

To independently evaluate the novel lignin composition in the ref8/fah1-2/AtC4H:SmF5H transgenics, we conducted NMR analysis on whole lignins released from cell walls by treatment with polysaccharide hydrolases. These data clearly show that the transgenic expression of Sm F5H can rescue S lignin biosynthesis in Arabidopsis ref8 fah1-2 double mutants and provide support for the observation of unique H-S lignins (Figure 7). Contour integration in two-dimensional 13C–1H-correlated (HSQC) NMR also allowed better estimates of the H:G:S distribution of the whole lignin (Ralph et al., 2006; Wagner et al., 2007) (Table 2). The lignins displayed distinctly different structural attributes resulting from the diverse distributions of lignin monomers (see Supplemental Figure 3 online). For example, 5-5–linked structures (in dibenzodioxocin units) were elevated in the G-rich fah1-2 mutant but greatly reduced in the S-elevated transgenics.

Figure 7.

Figure 7.

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Impact of Sm F5H Transgenic Expression on Lignin Composition in the Arabidopsis ref8 fah1-2 Double Mutant Examined by Partial (Aromatic Region) HSQC NMR Spectra.

Correlations from the aromatic rings can be categorized by the type of lignin units: H (red), G (blue), and S (green).

Sm F5H Rescues the Arabidopsis HCT-Deficient Mutant

Taken together, the above results suggest that Sm F5H can mediate a novel lignin biosynthetic route that bypasses three enzymes, hydroxycinnamoyl-CoA:shikimate hydroxycinnamoyl transferase (HCT), C3′H, and caffeoyl-CoA O-methyl transferase (CCoAOMT), which catalyze four consecutive steps in the canonical pathway in angiosperms (Figure 1B). This hypothesis predicts that Sm F5H should also be able to rescue the phenotypes exhibited by an Arabidopsis HCT-deficient mutant, just as it rescues the ref8 mutant. To test this prediction, we generated AtC4H:SmF5H transgenics in a genetic background where HCT is downregulated by RNA interference (RNAi). As has been reported previously, HCT downregulated plants phenocopy ref8 in that they show severe dwarfism and a dark-green/purple coloration of their rosette leaves, although in general they are not female-sterile like ref8 (Figure 8A) (Hoffmann et al., 2004). While quantitative RT-PCR shows that HCT transcript levels in HCTRNAi/AtC4H:SmF5H plants are similar to those of HCTRNAi plants (Figure 8B), the plants are significantly rescued in growth and exhibit decreased leaf pigmentation (Figure 8A). DFRC lignin analysis revealed that HCTRNAi plants deposit trace amounts of H lignin, whereas the lignin of HCTRNAi/AtC4H:SmF5H plants contains comparable amount of H and S subunits similar to the lignin of ref8/fah1-2/AtC4H:SmF5H plants (Figure 8C). These observations further support the rerouted lignin biosynthetic pathway mediated by Sm F5H, which bifurcates from the canonical pathway upstream of HCT.

Figure 8.

Figure 8.

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Complementation of the Arabidopsis HCTRNAi Plant by Sm F5H.

(A) Four-week-old Arabidopsis plants showing the partial rescue of growth phenotype of an HCTRNAi plant by the Sm F5H transgene.

(B) Relative HCT transcript levels in plants quantified by quantitative RT-PCR. Error bars represent standard deviations based on assays of biological duplicates.

(C) DFRC–gas chromatography analysis of lignin monomer composition. H/G/S are p-hydroxyphenyl/guaiacyl/syringyl lignin-derived hydroxycinnamyl alcohol peracetates. c/t, cis/trans; IS, internal standard.

[See online article for color version of this figure.]

Impact of Sm F5H Transgenic Expression on Soluble Phenylpropanoid Metabolism in the Arabidopsis C3′H-Deficient Background

We have previously shown that Sm F5H can rescue sinapoylmalate accumulation in fah1-2 leaves, thus complementing its ref phenotype under UV light (Weng et al., 2008c). To test whether Sm F5H could also rescue the ref phenotype in the ref8 background, we first inspected the rosette-stage ref8/fah1-2/AtC4H:SmF5H transgenics under UV light (Figure 3). Compared with ref8 mutants, the blue epidermal fluorescence is increased in the corresponding Sm F5H transgenics, but clearly not to the level of the wild type. We further analyzed the leaf soluble phenylpropanoid metabolites by HPLC (Figure 9). Consistent with the visual phenotyping result, we found that ref8-1/fah1-2/AtC4H:SmF5H plants showed only a modest increase in the sinapoylmalate levels compared with ref8-1, and ref8-2/fah1-2/AtC4H:SmF5H plants showed no increase compared with ref8-2. HPLC profiling of leaf soluble phenylpropanoids also revealed a significant decrease in the level of three major flavonol glycosides in the Sm F5H transgenics, compared with ref8 and ref8/fah1-2 where these flavonoids are hyperaccumulated. This observation indicates that either stress induced by the ref8 mutation is generally alleviated by the expression of Sm F5H transgene or that metabolites that are otherwise channeled into flavonoid biosynthesis when C3′H is blocked are redirected into lignin biosynthesis via Sm F5H.

Figure 9.

Figure 9.

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HPLC Quantification of Major Phenylpropanoid Metabolites in Leaves of 3-Week-Old Arabidopsis.

Sinapoylmalate can be detected in ref8-1 and even the null mutant ref8-2, which is consistent with a previous report (Abdulrazzak et al., 2006) and suggests the presence of a C3′H independent 3-hydroxylation pathway in ref8 mutant background. We observed that At F5H possesses a level of 3-hydroxylase activity toward p-coumaraldehyde and p-coumaryl alcohol (Table 1). Although the kinetic constants suggest such reactions cannot take place in vivo, under ref8 mutant background, the levels of relevant substrates might reach local concentrations that could permit some flux through At F5H. KG1, kaempferol 3-O-β-[β-d-glucopyranosyl(1→6)- d-glucopyranoside]-7-O-α-l-rhamnopyranoside; KG2, kaempferol 3-O-β-d-glucopyranoside-7-O-α-l-rhamnopyranoside; KG3, kaempferol 3-O-α-l-rhamnopyranoside-7-O-α-l-rhamnopyranoside; SM, sinapoylmalate.

[See online article for color version of this figure.]

Coexistence of H and S Lignin in Selaginella Stem Cortex

To gain insight into whether the Sm F5H-mediated S lignin biosynthetic pathway is operating in Selaginella, we took the advantage of the unique anatomy of Selaginella stems and separated the stem xylem and cortical tissue for DFRC lignin analysis (Figure 10A). The results confirmed the presence of a high percentage of S lignin in cortex and revealed that a significant amount of H lignin coexists in the same tissue (Figure 10B). By contrast, xylem tissue is dominated by G lignin with only trace amounts of S lignin and no detectable level of H lignin (Figure 10B). Consistent with what we have observed in the Arabidopsis transgenics, the coexistence of H and S lignin in Selaginella stem cortex suggests that the novel S lignin biosynthetic pathway derived from H lignin precursors may be active in Selaginella.

Figure 10.

Figure 10.

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Tissue-Specific Lignin Analysis in Selaginella Stems.

(A) A longitudinal section of mature Selaginella stem showing that xylem and cortical tissue can be separated. Bar = 1 mm.

(B) Lignin monomer composition (H, G, and S) in separated stem xylem and cortical tissue quantified by DFRC lignin analysis. N.D., not detectable.

[See online article for color version of this figure.]

DISCUSSION

F5H Mediates a C3′H-Independent S Lignin Biosynthetic Pathway in Selaginella

We have provided both in vitro and in vivo evidence to show that the convergent evolution of S lignin in lycophytes and angiosperms is the result of the elaboration of alternative biochemical pathways. Whereas in angiosperms two phenylpropanoid meta-hydroxylases, C3′H and F5H, are required for S lignin biosynthesis from para-hydroxylated intermediates, a unique phenylpropanoid dual meta-hydroxylase, Sm F5H, evolved in the lycophyte Selaginella where it mediates S lignin biosynthesis via a different pathway (Figure 1B). Our data suggest that G and S lignin can be biosynthesized independently of one another through two distinct sets of enzymes in Selaginella, in comparison to in angiosperms where S lignin subunits are derived from G lignin precursors.

We postulate that the C3′H-independent S lignin biosynthetic pathway in Selaginella requires two independent or one bifunctional COMT to catalyze the methylation step after the addition of each ring meta-hydroxyl by Sm F5H (Figure 1B). Bifunctional COMTs with activities toward 3- and 5-hydroxylated phenylpropanoid aldehydes and alcohols are known in angiosperms (Parvathi et al., 2001; Do et al., 2007). Such bifunctional COMTs could be ubiquitous in vascular plants or, like the phenylpropanoid 5-hydroxylases, may have emerged independently in Selaginella and angiosperms.

It is also important to note that the in vitro kinetics data showed that Sm F5H does not produce 3,4,5-trihydroxycinnamyl aldehyde/alcohol from p-coumaryl- or caffeyl-substituted substrates. These data suggest that after the first meta-hydroxylation step, the dihydroxy-substituted product is released from the active site of Sm F5H, and subsequent 3-O-methylation is required for the reentry of the guaiacyl-substituted substrate into the enzyme active site for the 5-hydroxylation. These biochemical characteristics resemble those of the recently characterized Arabidopsis CYP98A8, a phenylpropanoid dual meta-hydroxylase involved in the biosynthesis of N1,N5-di(hydroxyferuloyl)-N10-sinapoylspermidine (Matsuno et al., 2009). Interestingly, the phenylpropanoid dual meta-hydroxylase activities of Sm F5H and CYP98A8 are distinct from that of the petunia flavonoid 3′,5′-hydroxylase, which is capable of catalyzing 3′- and 5′-hydroxylations of 4′-hydroxylated flavonone substrates without meta-O-methylation after the addition of the first meta-hydroxyl group (Menting et al., 1994).

The C3′H-independent S lignin biosynthetic pathway in Selaginella also requires a form of hydroxycinnamoyl-CoA reductase (CCR) that can efficiently convert p-coumaroyl-CoA to p-coumaraldehyde. It is possible that, like Sm F5H, this enzyme is also specifically expressed in the cortex (Weng et al., 2008c), where it provides H-substituted precursors for Sm F5H, some of which escape hydroxylation and lead to the deposition of H lignin in this tissue.

Unlike Selaginella, angiosperms usually contain only trace amount of H lignin (Boerjan et al., 2003), an observation that may be explained by previous studies that have shown that for a number of angiosperm CCR isoforms, p-coumaroyl-CoA is a less favorable substrate than the G lignin precursor, feruloyl-CoA (Wengenmayer et al., 1976; Sarni et al., 1984; Baltas et al., 2005). That carbon flux is not stoichiometrically redirected into H lignin when G and S lignin is eliminated in the Arabidopsis ref8 mutant further supports this suggestion (Franke et al., 2002a; Abdulrazzak et al., 2006). Surprisingly, we found that transgenic expression of the bifunctional Sm F5H in the ref8 background not only restored S lignin biosynthesis but also significantly enhanced H lignin deposition. This result suggests that at least in the lignifying tissue, Arabidopsis contains CCR isoforms that can support the biosynthesis of substantial amounts of H-substituted lignin precursors. Thus, in contrast with what has been thought previously, it seems that CCR is unlikely to be the factor that limits H lignin biosynthesis, at least in wild-type Arabidopsis. Although it is unclear why Sm F5H is more effective in restoring S lignin synthesis than it is in complementing the sinapoylmalate-deficient phenotype of the fah1 ref8 double mutant, it is possible that a CCR that catalyzes the conversion of p-coumaroyl-CoA to p-coumaraldehyde may be limiting or absent in Arabidopsis leaf tissue, limiting the availability of substrate for Sm F5H in leaves.

Despite the presence of a C3′H-independent S lignin biosynthetic pathway mediated by Sm F5H, the Selaginella genome contains a C3′H ortholog that is >60% identical to angiosperm C3′H at amino acid level (Weng et al., 2008c), which suggests that Selaginella may also contain a C3′H-dependent pathway, analogous to the one defined in angiosperms (Franke et al., 2002b). The fact that Selaginella xylem, where Sm F5H is not highly expressed (Weng et al., 2008c), contains lignin composed of almost pure G units, is consistent with this notion, although the exact biochemical function of C3′H ortholog in Selaginella is still to be determined.

Sm F5H Provides a Valuable Tool for Genetic Engineering of Lignin Biosynthesis in Higher Plants

Lignin is a ubiquitous component of the cell wall of vascular plants and is essential to normal plant development, and plants in which the lignin biosynthetic pathway has been downregulated often suffer from significantly reduced growth (Weng et al., 2008b). The discovery of the phenylpropanoid dual meta-hydroxylase activity of Sm F5H and its role in the unique C3′H-independent S lignin biosynthetic pathway suggests it may be a valuable tool for fundamentally rerouting lignin biosynthesis in higher plants. In the presence of Sm F5H, several essential lignin biosynthetic genes in the canonical pathway, including C3′H and HCT as demonstrated in this study, could be downregulated without causing deleterious effects and resulted in a unique H-S lignin composition in the transgenic plants, which may not occur in nature. The mechanism that leads to the unique H-S lignin is currently unknown but deserves additional investigation in the future.

S Lignin Could Have Emerged Multiple Times during Plant Evolution

Some older literature reported that S lignin could be detected in some fern and gymnosperm species (e.g., cuplet fern, yew plum pine, sandarac-cypress, and gnetophytes) (reviewed in Weng et al., 2008a), which suggests that S lignin might have evolved multiple times during plant evolution, but additional F5H analogs and/or alternative pathways toward S lignin biosynthesis have yet to be discovered. The apparently independent occurrence of S lignin in distantly related plant lineages implies that it may have an important role in plants’ adaptation to their environment. S lignin is present in the anatomically analogous fiber cells in stems of angiosperms and Selaginella, suggesting that S lignin may function similarly in both lineages for mechanical support and/or defense against pathogens and herbivores (Li et al., 2001; Weng et al., 2008c). For example, the evolution of S-lignified fibers in angiosperms, Selaginella, and some members of the Gnetales may indicate that the presence of these strengthening cell types permitted the development of relatively weaker vessel elements in the vasculature of these plants (Logan and Thomas, 1985; Carlquist, 1996).

METHODS

Plant Material

Arabidopsis thaliana was grown under a 16-h-light/8-h-dark photoperiod at 100 μE·m−2·s−1 at 22°C. Columbia-0 was used as the wild type. The T-DNA insertional ref8-2 null allele was obtained from the ABRC under the accession of SALK_036132 (Alonso et al., 2003). Selaginella moellendorffii was obtained from Plant Delights Nursery (Raleigh, NC) and grown in a local greenhouse under 50% shade cloth.

Transgenic Plants

To generate ref8/fah1-2/AtC4H:SmF5H plants, ref8-1 and ref8-2 heterozygous plants were used as the female parent in a cross with four independent lines of previously described fah1-2/AtC4H:SmF5H plants (lines 2, 6, 7, and 8) (Weng et al., 2008c). In the F2 generation, plants with the genotype REF8/ref8 fah1-2/fah1-2 that were homozygous for the Sm F5H transgene were selected and allowed to self. All subsequent analyses were performed in the F3 generation on homozygous Sm F5H transgenics that were genotyped as being ref8 fah1-2 double homozygotes. All the independent transgenic lines showed partial phenotypic complementation compared with the corresponding ref8 fah1-2 plants. Transgenics that derived from fah1-2/AtC4H:SmF5H line 6 were used for further detailed analysis. To generate ref8-1/fah1-2/AtC4H:AtF5H plants, a similar approach was adopted as previously described using the fah1-2/AtC4H:AtF5H transgenic plants as the male parent in the initial cross.

To generate the HCTRNAi binary vector, a 356-bp HCT cDNA fragment was PCR amplified by the primer pair 5′-GGGGACAAGTTTGTACAAAAAAGCAGGCTAACATCAGAGATTCCACCA-3′ and 5′-GGGGACCACTTTGTACAAGAAAGCTGGGTCAACTTCGGGAATAAGC-3′. The resulting PCR product was recombined with the modified version of pDONR221 (Invitrogen) using BP clonase (Invitrogen) to generate the entry clone. The entry clone was then recombined with a destination binary vector (modified from pBI121; Clontech) that harbors a cauliflower mosaic virus 35S promoter-driven Gateway RNAi cassette to generate the final construct. The HCTRNAi construct was introduced into Arabidopsis via Agrobacterium tumefaciens–mediated transformation (Clough and Bent, 1998). To generate HCTRNAi/AtC4H:SmF5H plants, plants with the genotype REF8/ref8 fah1-2/fah1-2 that were homozygous for the Sm F5H transgene were used as the female parent to cross with the HCTRNAi plants. In the F1 generation, the plants with the genotype REF8/REF8 FAH1/fah1-2 that were hemizygous for both the Sm F5H transgene and the HCTRNAi transgene were analyzed. In parallel, HCTRNAi plants were crossed to fah1-2 to generate plants with the genotype REF8/REF8 FAH1/fah1-2 that were hemizygous for the HCTRNAi transgene in the F1 generation, which were used as control plants.

To genotype ref8-1, the primer pair 5′-CGAGCTATCATGGAGGAGCATA-3′ and 5′-CAACAAGAGCATGAGCAGCAG-3′ was used in combination with EcoRV digestion, exploiting the cleaved amplified polymorphic sequence marker resulting from the point mutation in ref8-1. Similarly, fah1-2 was genotyped using the primer pair 5′-TGGTGTGTACATATATGGATGAAGAA-3′ and 5′-TAGCAAGAGTGGTGAATATGTGAAGT-3′ in combination with MseI digestion. To genotype ref8-2, LP primer 5′-TCGTGGTTTCTAATAGCGGTG-3′ and RP primer 5′-TGTTAAGAAAAACAATTAGGGTTTTTG-3′, together with the T-DNA left border BP primer 5′-TGGTTCACGTAGTGGGCCATCG-3′ were used according to the previously described method (Alonso et al., 2003). To genotype the presence of the Sm F5H transgene, the gene-specific primer pair 5′-CAAGGTCCTCCACAAGAAGC-3′ and 5′-CAGTCGAAGCACTGGATGAA-3′ was used. To genotype the presence of the HCTRNAi transgene, a primer to the 35S promoter 5′-GACCTAACAGAACTCGCCGTAAAGA-3′ and an HCT gene-specific primer 5′-TAAGGGTAGGAGCAAAATCACCAAA-3′ were used.

Yeast Expression of P450s and Enzyme Assays

The construction of Sm F5H and At F5H yeast expression vectors is described by Weng et al. (2008c) and Humphreys et al. (1999), respectively. Constructs were transformed into the WAT11 yeast strain, and yeast growth, induction, and preparation of yeast microsomal extracts were conducted as previously described (Humphreys et al., 1999). Enzyme kinetic assays were performed essentially as described (Humphreys et al., 1999). In brief, a reaction system containing 1 mM NADP+, 10 mM glucose-6-phosphate, and 4 units of glucose-6-phosphate dehydrogenase was incubated for 5 min at 30°C in the presence of substrate to allow the generation of NADPH. The reaction was incubated for 20 min, at 30°C after adding the yeast microsomal extract, and was terminated by adding glacial acetic acid. All the assays except those involving hydroxycinnamyl alcohols were extracted with ethyl acetate, dried in vacuo, resuspended in 50% methanol, and analyzed by HPLC. The assays testing hydroxycinnamyl alcohols were analyzed by HPLC directly. For assays testing Sm F5H against p-coumaric acid, a substrate concentration series of 10, 14, 18, 25, 50, and 200 μM was used. For assays testing Sm F5H against p-coumaraldehyde/p-coumaryl alcohol or Sm F5H/At F5H against coniferaldehyde/coniferyl alcohol, a substrate concentration series of 0.7, 0.8, 1, 1.3, 2, and 10 μM was used. For assays testing Sm F5H/At F5H against ferulic acid, a substrate concentration series of 40, 50, 70, 100, 200, and 1000 μM was used. The same amount of microsomal extract from a single prep was used for each enzyme toward various substrates.

Histochemical Staining

For phloroglucinol-HCl staining, hand sections of 2-month-old Arabidopsis stems were stained with 1% phloroglucinol (w/v) in 12% HCl for 5 min and observed under light microscope. For Mäule staining, hand sections of 2-month-old Arabidopsis stems were fixed in 4% glutaraldehyde, rinsed in water, and treated for 10 min with 0.5% KMnO4. Sections were then rinsed with water, treated for 5 min with 10% HCl, rinsed in water, mounted in concentrated NH4OH, and examined under the light microscope.

Lignin Analysis

Cell wall samples free of soluble metabolites were prepared as previously described (Meyer et al., 1998). For Klason lignin analysis, 100 mg of cell wall sample was swelled with 3 mL of 72% H2SO4 for 30 min at 30°C and then diluted with water to 4% H2SO4 and autoclaved at 120°C for 1 h. The residue was filtered through a preweighed glass filter. The residue was dried in an 80°C oven on the filter overnight before being weighed at room temperature. The DFRC lignin analysis was performed essentially as previously reported (Lu and Ralph, 1998). Briefly, cell wall samples were dissolved in acetyl bromide/acetic acid solution, containing 4,4′-ethylidenebisphenol as an internal standard. The reaction products were dried down using nitrogen gas, dissolved in dioxane/acetic acid/water (5/4/1, v/v/v), reacted with Zn dust, purified with C-18 SPE columns (SUPELCO), and acetylated with pyridine/acetic anhydride (2/3, v/v). The lignin derivatives were analyzed by gas chromatography/flame ionization detection using response factors relative to the internal standard of 1.26 for p-coumaryl alcohol peracetate, 1.30 for coniferyl alcohol peracetate, and 1.44 for sinapyl alcohol peracetate. The same samples were run through gas chromatography–mass spectrometry in parallel to confirm the identity of the derived hydroxycinnamyl alcohol peracetates.

Lignin Preparation for NMR Spectroscopy

Dried Arabidopsis stems (0.5 to 1.5 g) were ground in a Retsch MM301 shaker mill for 3 min at 30 s−1 and extracted sequentially with water (sonication, 20 min, four times), 80% methanol (four times), acetone (two times), chloroform-acetone (1/1, v/v, two times), and acetone (one time) again. The obtained isolated cell walls (0.3 to 1 g) were ball-milled for 5 h per 1 g of sample weight (in 20 min on/10 min off cycles) using a Retsch PM100 ball mill vibrating at 600 rpm with zirconium dioxide vessels (50 mL) containing ZrO2 ball bearings (10 × 10 mm). Ball milled walls were transferred to centrifuge tubes (50 mL) and digested at 30°C with crude cellulases (Cellulysin; Calbiochem; 30 mg g−1 of sample, in pH 5.0 sodium acetate buffer, 3 × 2 d, fresh buffer and enzyme each time), leaving all of the lignin and residual polysaccharides totaling 0.202 g (20.1% of the original cell wall after extractions, wild type), 0.232 g (23.1%, fah1-2), 0.0452 g (15.3%, ref8-1), 0.0749 g (11.9%, ref8-1/fah1-2/AtC4H:SmF5H), and 0.137 g (16.1%, ref8-2/fah1-2/AtC4H:SmF5H). The polysaccharidase-digested cell wall fractions (70 mg each except for ref8-1, 45 mg) were subjected to solubilization in DMSO/N-methylimidazole (2/1, v/v). Following acetic anhydride addition (0.5 mL, 1.5 h), the polysaccharidase-digested cell walls gave acetylated samples for NMR measurement (Lu and Ralph, 2003).

NMR Spectroscopy

The NMR spectra were acquired on a Bruker Biospin DMX-500 instrument fitted with a sensitive cryogenically cooled 5-mm DCH 1H/13C gradient probe with inverse geometry (proton coils closest to the sample). Acetylated lignin preparations (5 to 80 mg) were dissolved in 0.5 mL CDCl3; the central chloroform solvent peak was used as internal reference (δC 77.0, δH 7.26 ppm). HSQC experimental conditions were as described previously (Wagner et al., 2007). Volume integration of contours in HSQC plots was accomplished by Bruker's TopSpin 2.0 software as described (Ralph et al., 2006).

Quantitative RT-PCR

Total RNA was extracted from 3-week-old rosette leaves of Arabidopsis plants using the RNeasy plant mini kit (Qiagen). Single-strand cDNAs were synthesized via reverse transcription using the High Capacity cDNA reverse transcription kit (Applied Biosystems). The cDNAs was treated with RNase and used as template for real-time PCR. Quantitative real-time PCR was performed on the StepOne Real-Time PCR system (Applied Biosystems) using the ΔΔCT method with default cycling program. HCT was amplified using the primer pair 5′-GAATTCCATACGAGGGTTTGTCTT-3′ and 5′-GGGCAATGGCAACGGATA-3′, whereas At1g13320, as an internal standard (Czechowski et al., 2005), was amplified using the primer pair 5′-TAACGTGGCCAAAATGATGC-3′ and 5′-GTTCTCCACAACCGCTTGGT-3′. Both the primer pairs were tested in a standard curve analysis beforehand, which showed amplification efficiency higher than 90%.

Leaf Sinapoylmalate and Flavonoid Analysis

Three-week-old Arabidopsis rosette leaves were harvested, ground in liquid nitrogen, and extracted with 50% methanol (1 mL per 100 mg fresh weight) for 2 h at 65°C, and the extracts were then analyzed by HPLC. Metabolites were separated on a C18 column, using a gradient from 1.5% acetic acid to 35% acetonitrile in 1.5% acetic acid at a flow rate of 1 mL min−1. Sinapoylmalate content was quantified using sinapic acid as standard. The identification of the three major kaempferol glycosides was according to a previous report (Veit and Pauli, 1999); quantification used kaempferol as standard.

Accession Numbers

Sequence data from this article can be found in the Arabidopsis Genome Initiative or GenBank/EMBL databases under the following accession numbers: Sm F5H, EU032589; At F5H, At4g36220; and HCT, At5g48930.

Supplemental Data

The following materials are available in the online version of this article.

  • Supplemental Figure 1. Lineweaver-Burk Representation of the Kinetic Analysis of Phenylpropanoid Meta-Hydroxylation Reactions Catalyzed by Yeast-Expressed Sm F5H or At F5H.

  • Supplemental Figure 2. GC-MS Confirmation of the Identity of Peracetates of Monolignols Derived from the DFRC Lignin Analysis Shown in Figure 6.

  • Supplemental Figure 3. Partial (Sidechain Region) HSQC Spectra Showing the Structural Changes Resulting from Lignification Using Altered Monolignol Supplies.

Supplementary Material

[Supplemental Data]
tpc.109.073528_index.html (756B, html)

Acknowledgments

This work is funded by the National Science Foundation (Grant IOB-0450289). Partial funding to J.R. was via the Department of Energy (DOE) Office of Science (Grant DE-AI02-06ER64299) and the DOE Great Lakes Bioenergy Research Center (DOE Office of Science BER DE-FC02-07ER64494). N.D.B. is supported by a fellowship from the Life Sciences Research Foundation. We thank J.A. Banks for providing Selaginella moellendorffii plant materials.

References

  1. Abdulrazzak N., et al. (2006). A coumaroyl-ester-3-hydroxylase insertion mutant reveals the existence of nonredundant meta-hydroxylation pathways and essential roles for phenolic precursors in cell expansion and plant growth. Plant Physiol. 140: 30–48 [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Alonso J.M., et al. (2003). Genome-wide insertional mutagenesis of Arabidopsis thaliana. Science 301: 653–657 [DOI] [PubMed] [Google Scholar]
  3. Baltas M., Lapeyre C., Bedos-Belval F., Maturano M., Saint-Aguet P., Roussel L., Duran H., Grima-Pettenati J. (2005). Kinetic and inhibition studies of cinnamoyl-CoA reductase 1 from Arabidopsis thaliana. Plant Physiol. Biochem. 43: 746–753 [DOI] [PubMed] [Google Scholar]
  4. Boerjan W., Ralph J., Baucher M. (2003). Lignin biosynthesis. Annu. Rev. Plant Physiol. Plant Mol. Biol. 54: 519–546 [DOI] [PubMed] [Google Scholar]
  5. Carlquist S. (1996). Wood, bark, and stem anatomy of gnetales: A summary. Int. J. Plant Sci. 157: S58–S76 [Google Scholar]
  6. Chapple C.C., Vogt T., Ellis B.E., Somerville C.R. (1992). An Arabidopsis mutant defective in the general phenylpropanoid pathway. Plant Cell 4: 1413–1424 [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Cheng X., Collins R.E., Zhang X. (2005). Structural and sequence motifs of protein (histone) methylation enzymes. Annu. Rev. Biophys. Biomol. Struct. 34: 267–294 [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Clough S.J., Bent A.F. (1998). Floral dip: A simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J. 16: 735–743 [DOI] [PubMed] [Google Scholar]
  9. Conant G.C., Wagner A. (2003). Convergent evolution of gene circuits. Nat. Genet. 34: 264–266 [DOI] [PubMed] [Google Scholar]
  10. Czechowski T., Stitt M., Altmann T., Udvardi M.K., Scheible W.R. (2005). Genome-wide identification and testing of superior reference genes for transcript normalization in Arabidopsis. Plant Physiol. 139: 5–17 [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Do C.T., Pollet B., Thevenin J., Sibout R., Denoue D., Barriere Y., Lapierre C., Jouanin L. (2007). Both caffeoyl Coenzyme A 3-O-methyltransferase 1 and caffeic acid O-methyltransferase 1 are involved in redundant functions for lignin, flavonoids and sinapoyl malate biosynthesis in Arabidopsis. Planta 226: 1117–1129 [DOI] [PubMed] [Google Scholar]
  12. Franke R., Hemm M.R., Denault J.W., Ruegger M.O., Humphreys J.M., Chapple C. (2002a). Changes in secondary metabolism and deposition of an unusual lignin in the ref8 mutant of Arabidopsis. Plant J. 30: 47–59 [DOI] [PubMed] [Google Scholar]
  13. Franke R., Humphreys J.M., Hemm M.R., Denault J.W., Ruegger M.O., Cusumano J.C., Chapple C. (2002b). The Arabidopsis REF8 gene encodes the 3-hydroxylase of phenylpropanoid metabolism. Plant J. 30: 33–45 [DOI] [PubMed] [Google Scholar]
  14. Franke R., McMichael C.M., Meyer K., Shirley A.M., Cusumano J.C., Chapple C. (2000). Modified lignin in tobacco and poplar plants over-expressing the Arabidopsis gene encoding ferulate 5-hydroxylase. Plant J. 22: 223–234 [DOI] [PubMed] [Google Scholar]
  15. Hedden P., Phillips A.L., Rojas M.C., Carrera E., Tudzynski B. (2001). Gibberellin biosynthesis in plants and fungi: A case of convergent evolution? J. Plant Growth Regul. 20: 319–331 [DOI] [PubMed] [Google Scholar]
  16. Hedenstrom A., Johansson L.C., Wolf M., von Busse R., Winter Y., Spedding G.R. (2007). Bat flight generates complex aerodynamic tracks. Science 316: 894–897 [DOI] [PubMed] [Google Scholar]
  17. Hoffmann L., Besseau S., Geoffroy P., Ritzenthaler C., Meyer D., Lapierre C., Pollet B., Legrand M. (2004). Silencing of hydroxycinnamoyl-coenzyme a shikimate/quinate hydroxycinnamoyltransferase affects phenylpropanoid biosynthesis. Plant Cell 16: 1446–1465 [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Humphreys J.M., Hemm M.R., Chapple C. (1999). New routes for lignin biosynthesis defined by biochemical characterization of recombinant ferulate 5-hydroxylase, a multifunctional cytochrome P450-dependent monooxygenase. Proc. Natl. Acad. Sci. USA 96: 10045–10050 [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Jin Z.F., Matsumoto Y., Tange T., Akiyama T., Higuchi M., Ishii T., Iiyama K. (2005). Proof of the presence of guaiacyl-syringyl lignin in Selaginella tamariscina. J. Wood Sci. 51: 424–426 [Google Scholar]
  20. Kenrick P., Crane P.R. (1997). The origin and early evolution of plants on land. Nature 389: 33–39 [Google Scholar]
  21. Li L., Cheng X.F., Leshkevich J., Umezawa T., Harding S.A., Chiang V.L. (2001). The last step of syringyl monolignol biosynthesis in angiosperms is regulated by a novel gene encoding sinapyl alcohol dehydrogenase. Plant Cell 13: 1567–1586 [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Logan K.J., Thomas B.A. (1985). Distribution of lignin derivatives in plants. New Phytol. 99: 571–585 [DOI] [PubMed] [Google Scholar]
  23. Logan K.J., Thomas B.A. (1987). The distribution of lignin derivatives in fossil plants. New Phytol. 105: 157–173 [DOI] [PubMed] [Google Scholar]
  24. Lu F., Ralph J. (1998). The DFRC method for lignin analysis. 2. Monomers from isolated lignins. J. Agric. Food Chem. 46: 547–552 [DOI] [PubMed] [Google Scholar]
  25. Lu F., Ralph J. (2003). Non-degradative dissolution and acetylation of ball-milled plant cell walls: high-resolution solution-state NMR. Plant J. 35: 535–544 [DOI] [PubMed] [Google Scholar]
  26. Matsuno M., et al. (2009). Evolution of a novel phenolic pathway for pollen development. Science 325: 1688–1692 [DOI] [PubMed] [Google Scholar]
  27. Menting J., Scopes R.K., Stevenson T.W. (1994). Characterization of flavonoid 3[prime],5[prime]-hydroxylase in microsomal membrane fraction of Petunia hybrida flowers. Plant Physiol. 106: 633–642 [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Meyer K., Shirley A.M., Cusumano J.C., Bell-Lelong D.A., Chapple C. (1998). Lignin monomer composition is determined by the expression of a cytochrome P450-dependent monooxygenase in Arabidopsis. Proc. Natl. Acad. Sci. USA 95: 6619–6623 [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Parvathi K., Chen F., Guo D., Blount J.W., Dixon R.A. (2001). Substrate preferences of O-methyltransferases in alfalfa suggest new pathways for 3-O-methylation of monolignols. Plant J. 25: 193–202 [DOI] [PubMed] [Google Scholar]
  30. Ralph J., Akiyama T., Kim H., Lu F., Schatz P.F., Marita J.M., Ralph S.A., Reddy M.S., Chen F., Dixon R.A. (2006). Effects of coumarate 3-hydroxylase down-regulation on lignin structure. J. Biol. Chem. 281: 8843–8853 [DOI] [PubMed] [Google Scholar]
  31. Sarni F., Grand C., Boudet A.M. (1984). Purification and properties of cinnamoyl-CoA reductase and cinnamyl alcohol dehydrogenase from poplar stems (Populus X euramericana). Eur. J. Biochem. 139: 259–265 [DOI] [PubMed] [Google Scholar]
  32. Schoch G., Goepfert S., Morant M., Hehn A., Meyer D., Ullmann P., Werck-Reichhart D. (2001). CYP98A3 from Arabidopsis thaliana is a 3′-hydroxylase of phenolic esters, a missing link in the phenylpropanoid pathway. J. Biol. Chem. 276: 36566–36574 [DOI] [PubMed] [Google Scholar]
  33. Tanaka K., Barmina O., Kopp A. (2009). Distinct developmental mechanisms underlie the evolutionary diversification of Drosophila sex combs. Proc. Natl. Acad. Sci. USA 106: 4764–4769 [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Teutsch H.G., Hasenfratz M.P., Lesot A., Stoltz C., Garnier J.M., Jeltsch J.M., Durst F., Werck-Reichhart D. (1993). Isolation and sequence of a cDNA encoding the Jerusalem artichoke cinnamate 4-hydroxylase, a major plant cytochrome P450 involved in the general phenylpropanoid pathway. Proc. Natl. Acad. Sci. USA 90: 4102–4106 [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Towers G.H.N., Gibbs R.D. (1953). Lignin chemistry and the taxonomy of higher plants. Nature 172: 25–26 [DOI] [PubMed] [Google Scholar]
  36. Veit M., Pauli G.F. (1999). Major flavonoids from Arabidopsis thaliana leaves. J. Nat. Prod. 62: 1301–1303 [DOI] [PubMed] [Google Scholar]
  37. Wagner A., Ralph J., Akiyama T., Flint H., Phillips L., Torr K., Nanayakkara B., Te Kiri L. (2007). Exploring lignification in conifers by silencing hydroxycinnamoyl-CoA:shikimate hydroxycinnamoyltransferase in Pinus radiata. Proc. Natl. Acad. Sci. USA 104: 11856–11861 [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Weng J.K., Banks J.A., Chapple C. (2008a). Parallels in lignin biosynthesis: A study in Selaginella moellendorffii reveals convergence across 400 million years of evolution. Commun. Integr. Biol. 1: 20–22 [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Weng J.K., Li X., Bonawitz N.D., Chapple C. (2008b). Emerging strategies of lignin engineering and degradation for cellulosic biofuel production. Curr. Opin. Biotechnol. 19: 166–172 [DOI] [PubMed] [Google Scholar]
  40. Weng J.K., Li X., Stout J., Chapple C. (2008c). Independent origins of syringyl lignin in vascular plants. Proc. Natl. Acad. Sci. USA 105: 7887–7892 [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Wengenmayer H., Ebel J., Grisebach H. (1976). Enzymic synthesis of lignin precursors. Purification and properties of a cinnamoyl-CoA: NADPH reductase from cell suspension cultures of soybean (Glycine max). Eur. J. Biochem. 65: 529–536 [DOI] [PubMed] [Google Scholar]
  42. White E., Towers G.H.N. (1967). Comparative biochemistry of lycopods. Phytochemistry 6: 663–667 [Google Scholar]

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