Skip to main content
NCBI home page
Plant Physiology logoLink to Plant Physiology
. 2022 May 6;189(4):2015–2028. doi: 10.1093/plphys/kiac210

H-lignin can be deposited independently of CINNAMYL ALCOHOL DEHYDROGENASE C and D in Arabidopsis

Fabiola Muro-Villanueva 1,2, Hoon Kim 3, John Ralph 4,5, Clint Chapple 6,7,✉,b
PMCID: PMC9342963  PMID: 35522042

Abstract

Lignin contributes substantially to the recalcitrance of biomass toward saccharification. To circumvent this problem, researchers have genetically altered lignin, although, in a number of cases, these efforts have resulted in an undesirable yield penalty. Recent findings have shown that by knocking out two subunits (MED5A and MED5B) of the transcriptional regulatory complex Mediator, the stunted growth phenotype of mutants in p-coumaroyl shikimate 3′-hydroxylase, reduced epidermal fluorescence 8-1 (ref8-1), can be alleviated. Furthermore, these plants synthesize a lignin polymer almost entirely derived from p-coumaryl alcohol. Plants deficient in cinnamyl alcohol dehydrogenase (CAD) are notable in that they primarily incorporate coniferaldehyde and sinapaldehyde into their lignin. We tested the hypothesis that by stacking mutations in the genes encoding for the CAD paralogs C and D on an Arabidopsis (Arabidopsis thaliana) med5a/5b ref8-1 genetic background, the biosynthesis of p-coumaryl alcohol would be blocked, making p-coumaraldehyde available for polymerization into a novel kind of lignin. The med5a/5b ref8-1 cadc cadd plants are viable, but lignin analysis demonstrated that they continue to synthesize p-hydroxyphenyl lignin despite being mutated for the CADs typically considered to be required for monolignol biosynthesis. In addition, enzyme activity tests showed that even in the absence of CADC and CADD, there is high CAD activity in stems. We tested the potential involvement of other CADs in p-coumaraldehyde biosynthesis in the quintuple mutant by mutating them using the CRISPR/Cas9 system. Lignin analysis demonstrated that the resulting hextuple mutant plants continue to deposit p-coumaryl alcohol-derived lignin, demonstrating a route for the synthesis of p-hydroxyphenyl lignin in Arabidopsis independent of four CAD isoforms.

The biosynthesis of p-hydroxyphenyl subunits of the cell wall polymer lignin is independent of the canonical cinnamyl alcohol dehydrogenases involved in guaiacyl and syringyl monomers.

Introduction

The global economy is based upon petroleum-based industries but this dependency on fossil fuels is associated with many environmental concerns. Biofuels represent a potential alternative to oil that would alleviate many of these problems. Although starch-derived bioethanol is produced at industrial scale, its production competes with that of crops and for arable land (Nonhebel, 2005) that could be used to feed the increasing world population. In contrast, lignocellulosic biomass is an abundant resource for the production of biofuels and commodity chemicals that avoids the food versus fuel dilemma (US Department of Energy, 2005; McCann and Carpita, 2015). However, in order to extract its fermentable sugars, expensive pre-treatments are required that add substantially to the cost of biofuel production (Himmel et al., 2007). Recalcitrance of biomass toward saccharification can be attributed in large part to lignin, a polymer found in the plant secondary cell wall (Zeng et al., 2014). Successful engineering examples in monocots and dicots have shown that even modest reductions in lignin content can result in an increase in saccharification (Wang et al., 2015).

Lignin is a polymer of monolignols that are products of the phenylpropanoid pathway. This pathway gives rise to a wide variety of plant secondary metabolites, but the main products are the monolignols p-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol (Figure 1). Polymerization of these alcohols by radical coupling occurs in the apoplast and results in the deposition of p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) lignin. Lignin polymerization in the vasculature is neither a template-based nor an enzymatically guided process (Ralph et al., 2008). This unique feature allows for high plasticity when it comes to monomer incorporation into the growing polymer. In Nature, the incorporation of non-canonical monolignols into lignin is a widespread phenomenon. For instance, in some monocots the flavonoid tricin and the hydroxycinnamates ferulate and p-coumarate can be found cross-linked to the polymer (Ralph et al., 2019). In loblolly pine (Pinus taeda) and mulberry tree (Morus alba) mutants, the monolignol aldehyde precursors coniferaldehyde and sinapaldehyde (Figure 1) form a substantial component of lignin (Ralph et al., 1997; Yamamoto et al., 2020). Lignification was a key feature that allowed plants to colonize terrestrial ecosystems by imparting the necessary rigidity and hydrophobicity to vascular tissues (Turner and Somerville, 1997), but why distinct lineages of plants evolved the ability to incorporate atypical monomers is still an open question (Renault et al., 2019). Although the evolutionary reason behind this flexibility in monomer incorporation remains unknown, it has allowed the introduction of non-canonical monomers into the polymer via metabolic engineering and the development of lignins with new functionalities (Vanholme et al., 2019).

Figure 1.

Figure 1

Open in a new tab

The phenylpropanoid pathway. PAL, PHENYLALANINE AMMONIA-LYASE; C4H, CINNAMATE 4-HYDROXYLASE; 4CL, 4-COUMARATE-CoA LIGASE; HCT, HYDROXYCINNAMOYL-CoA:SHIKIMATE HYDROXYCINNAMOYL TRANSFERASE; C3′H, p-COUMAROYL SHIKIMATE 3′-HYDROXYLASE; CSE, CAFFEOYL SHIKIMATE ESTERASE; CCoAOMT, CAFFEOYL-CoA O-METHYLTRANSFERASE; F5H, FERULATE 5-HYDROXYLASE; COMT, CAFFEIC ACID O-METHYLTRANSFERASE; and CAD, CINNAMYL ALCOHOL DEHYDROGENASE.

Modifying monolignol ratios and incorporation of atypical monomers have proven to be effective approaches for reducing biomass recalcitrance (Mottiar et al., 2016). For example, by modulating the expression of FERULATE 5-HYDROXYLASE (F5H), the native G-to-S monomer ratio can be dramatically altered (Meyer et al., 1998; Franke et al., 2000; Huntley et al., 2003; Reddy et al., 2005; Stewart et al., 2009). F5H mutants deposit lignins dominated by G-subunits, but plants overexpressing F5H are nearly devoid of G-lignin and instead deposit almost pure S-lignin (Meyer et al., 1998; Franke et al., 2000; Huntley et al., 2003; Reddy et al., 2005; Stewart et al., 2009; Shuai et al., 2016). The shift to S-lignin subunits in transgenic poplar (Populus tremula×Populus alba) overexpressing F5H leads to an increase in pulping efficiency (Huntley et al., 2003) and sugar release following enzymatic digestion (Yang et al., 2019). The diversity of natural and engineered lignin variants indicates that there is a wide range of compounds that can be accepted as lignin monomers, some of which might similarly reduce biomass recalcitrance. Only a few basic principles dictate whether a compound can be a substantial component of lignin, including its production level, availability at the site of lignification, and its ability to undergo radical coupling (Ralph et al., 2004, 2008; Vanholme et al., 2019). These basic requirements result in a large repertoire of possible compounds that could be engineered into lignin or may already be incorporated in Nature.

H-lignin is naturally a minor component of lignins, accounting for only 2% of the total in Arabidopsis thaliana. The reaction immediately downstream of the branchpoint between H- and G-lignin synthesis is catalyzed by the enzyme p-COUMAROYL SHIKIMATE 3′-HYDROXYLASE (C3’H) and a plant carrying a mutation in the corresponding gene, reduced epidermal fluorescence 8 (ref8-1), is dwarfed and deposits primarily H-lignin (Figure 1; Franke et al., 2002). Elimination of two paralogs of the MED5 subunit of the transcriptional co-regulator Mediator enables ref8-1 plants to attain near-normal stature by blocking the transcriptional reprogramming that leads to dwarfing in the mutant, while retaining their H-subunit-rich lignin composition (Bonawitz et al., 2014). Upon enzymatic treatment, the biomass from med5a/5b ref8-1 plants yields more than double the amount of glucose when compared with wild-type (Bonawitz et al., 2014).

In addition to their presence in natural mutants, hydroxycinnamaldehydes have been successfully incorporated into lignin by mutation or downregulation of CINNAMYL ALCOHOL DEHYDROGENASES (CADs) in species such as poplar, Arabidopsis, Medicago (Medicago truncatula), sorghum (Sorghum bicolor), tobacco (Nicotiana tabacum), and loblolly pine (Pillonel et al., 1991; Baucher et al., 1996; Ralph et al., 1997; Kim et al., 2003; Sibout et al., 2005; Zhao et al., 2013). CADs catalyze the last step in monolignol biosynthesis by reducing hydroxycinnamaldehydes to hydroxycinnamyl alcohols (Figure 1). By stacking CAD-deficiency with F5H downregulation or upregulation, we were able to generate Arabidopsis lines in which the lignins derived heavily from coniferaldehyde, sinapaldehyde, or both rather than the conventional monolignols (Anderson et al., 2015). In Arabidopsis, polysaccharide hydrolysis of cadc cadd biomass resulted in a doubling of glucose release (Anderson et al., 2015).

H-lignin and aldehyde-rich lignins are desirable traits because they lead to greater cell wall digestibility. Combination of these two characteristics to generate p-coumaraldehyde lignins could lead to an enhancement in cell wall digestibility beyond that of each manipulation individually. To test if this polymer could be made in muro, we used the high H-lignin background med5a/5b ref8-1 as a genetic background into which the cadc, cadd (Supplemental Table S1) and two other cad mutations were stacked. Surprisingly, we found that H-lignin biosynthesis in Arabidopsis is independent of the two major CADs involved in the biosynthesis of the canonical G- and S-substituted subunits in Arabidopsis, and is also independent of two additional CAD isoforms.

Results

p-Coumaryl alcohol is the main constituent of lignin in med5a/5b ref8-1 cadc cadd

To determine whether p-coumaraldehyde can be incorporated into lignin, we generated genetic material in which p-coumaraldehyde is predicted to accumulate at high levels. As mentioned above, med5a/5b ref8-1 synthesizes lignin primarily derived from p-coumaryl alcohol (Bonawitz et al., 2014). In an attempt to block the reduction of p-coumaraldehyde to p-coumaryl alcohol in med5a/5b ref8-1, we crossed it to cadc cadd and identified quadruple and quintuple mutants, namely med5a/5b ref8-1 cadc, med5a/5b ref8-1 cadd, and med5a/5b ref8-1 cadc cadd plants among the segregating population. The triple mutant med5a/5b ref8-1 has nearly wild-type growth and the addition of cadc, cadd, or cadc cadd mutations to this genetic background did not alter its growth substantially (Supplemental Figure S1).

We next determined whether stacking these multiple mutations led to an effect on total lignin content and composition, as measured by the thioglycolic acid (TGA) (Campbell and Ellis, 1992) and derivatization followed by reductive cleavage (DFRC) (Lu and Ralph, 1998) methods. TGA analysis revealed that the total lignin content of the quintuple mutant was the same as that of the med5a/5b ref8-1 mutant (Table 1). The DFRC method cleaves only normal β-O-4 linkages, so the total amount of lignin appears to be lower in plants with lignins with alternate linkages (Lu and Ralph, 1998). For example, cadc cadd plants, which deposit lignins derived from cinnamaldehydes cross-coupled by 8-O-4 linkages, appeared to have only very low levels of lignin when analyzed by DFRC (Figure 2; Anderson et al., 2015). In contrast, alternate methods revealed that these plants only have a 30% reduction in total lignin (Sibout et al., 2005; Anderson et al., 2015). Unexpectedly, DFRC analysis revealed that the quadruple and quintuple mutants here had a profile very similar to that of med5a/5b ref8-1, rather than cadc cadd, suggesting a lignin composition still comprised of p-coumaryl alcohol units (Figure 2). Cell wall analysis by 2D heteronuclear single-quantum correlation (HSQC) NMR provides a more substantive structural profile of the polymers, including displaying unique signatures for the different lignin monomers and linkages. These analyses showed that the aldehyde, aromatic, and aliphatic regions of the spectrum of med5a/5b ref8-1 cadc cadd were all but indistinguishable from those of med5a/5b ref8-1 (Figure 3). To further explore this observation, synthetic dimers and oligomers of hydroxycinnamalaldehydes, as were prepared in previous studies (Kim et al., 2000, 2003), were included in the sample set for NMR analysis but none of the characteristic signatures of these synthetic compounds were observed by NMR in the quintuple mutant. No new correlations appeared in the aldehyde region (Figure 3C). In conclusion, mutation of CADC and CADD in med5a/5b ref8-1 appeared to have no effect on total lignin content nor composition.

Table 1.

Total lignin quantification by the TGA method

Genotype A 280 mg cell wall−1 ± sd
med5a/5b ref8-1 1.24 ± 0.12
med5a/5b ref8-1 cada 1.24 ± 0.09
med5a/5b ref8-1 cadc cadd 1.19 ± 0.05
med5a/5b ref8-1 cadc cadd cada 1.23 ± 0.07
Open in a new tab

sd, standard deviation among biological replicates (n = 3).

Figure 2.

Figure 2

Open in a new tab

Lignin monomer composition determined by gas chromatography using the DFRC method. Error bars indicate the standard deviation among three biological replicates. The asterisk represents a difference in H-lignin content between CAD mutants in the med5a/5b ref8-1 background and med5a/5b ref8-1 plants determined by one-way ANOVA and Dunnet’s test (P < 0.05).

Figure 3.

Figure 3

Open in a new tab

2D 1H–13C correlation (HSQC) NMR spectra of med5 ref8-1 and med5a/5b ref8-1 cadc cadd Arabidopsis transgenic lines. Resonance signals arising from H, G, and S lignin subunits and side-chain compositions, including aldehydes, are color-coded to match the structures shown on the right. Quantification is from correlation peak volume integration. A, Aromatics are on an S + G + X2G′ = 100% basis. A phenylalanine peak (Phe2/6) overlapped with the p-hydroxyphenyl group’s H2/6 correlation, preventing accurate quantification of H-units that may be over-estimated. B, Side-chain units in the aliphatic region are on an A + B + C = 100% basis with X1 expressed on the A + B + C basis. C, The aldehyde region shows aldehyde end-group components.

Table 2.

p-coumaryl alcohol and p-coumaraldehyde quantification in mutant stems

Genotype p-Coumaraldehyde (nmoles g−1 FW) ± SD p-Coumaryl alcohol (nmoles g−1 FW) ± SD 
Wild-type 0.08 ± 0.03b 0.83 ± 0.78b
cadc cadd 0.27 ± 0.18b 0.08 ± 0.03b
med5a/5b 0.51 ± 0.22b 25.05 ± 8.99ab
med5a/5b ref8-1 6.67 ± 1.42a 47.92 ± 17.13a
med5a/5b ref8-1 cadc/d 7.78 ± 2.22a 31.41 ± 11.01a
med5a/5b ref8-1 cadc/d/a 6.37 ± 0.75a 32.07 ± 14.81a
med5a/5b ref8-1 cadc/d/g 6.25 ± 0.67a 32.20 ± 7.22a
Open in a new tab

SD, standard deviation among biological replicates (n = 3). Letters represent difference between genotypes determined by one-way ANOVA and Tukey’s honest significant difference test (P < 0.05).

Substantial levels of residual CAD activity remain in cadc cadd plants

Arabidopsis cadc cadd mutants deposit a lignin mainly comprised of hydroxycinnamaldehyde subunits, indicating a dominant role for these two CADs in the synthesis of coniferyl and sinapyl alcohols (Anderson et al., 2015). The fact that med5a/5b ref8-1 cadc cadd plants continue to incorporate p-coumaryl alcohol in their lignin suggests that there is another enzyme(s) that can catalyze the reduction of p-coumaraldehyde to p-coumaryl alcohol. To explore this possibility, we first performed CAD enzyme activity tests on cadc cadd and med5a/5b ref8-1 cadc cadd stem tissue extracts and found that they have lower activity toward p-coumaraldehyde, coniferaldehyde, and sinapaldehyde than wild-type and med5a/5b ref8-1 plants but continue to exhibit substantial CAD activity (Figure 4), which was unexpected based upon their high aldehyde lignin phenotype. Additionally, we detected high CAD activity in med5a/5b backgrounds which is consistent with the increase in transcript levels of phenylpropanoid genes in this mutant, including CADD (Bonawitz et al., 2014). These observations and the fact that the genome of A. thaliana encodes for nine CAD paralogs (Costa et al., 2003) suggest that another CAD may catalyze the reduction of p-coumaraldehyde to p-coumaryl alcohol.

Figure 4.

Figure 4

Open in a new tab

CAD enzyme activity in 4-week-old stems. CAD activity using p-coumaraldehyde (top) and coniferaldehyde (bottom) as substrates reported as picokatal per mcrogram of protein. Error bars represent standard deviation among biological replicates (n = 4). Letters represent difference between genotypes determined by one-way ANOVA and Tukey’s honest significant difference test (P < 0.05).

Knocking out three CAD paralogs does not have an effect on lignin composition

To identify candidate CAD paralogs that might have a role in H-lignin synthesis, we first considered whether relevant CADs might be upregulated in genetic backgrounds that synthesize high levels of p-coumaryl alcohol. Accordingly, we mined an RNA-seq data set generated from ref8-1 and med5a/5b ref8-1 rosettes (Bonawitz et al., 2014), searched for non-canonical CADs that were upregulated in both of these backgrounds (Supplemental Table S3), and identified CADA as a candidate. In addition, because the RNA-seq data available was from rosettes that are not highly lignified tissues, we performed qPCR using cDNA from med5a/5b ref8-1 cadc cadd stems and found that the CADG transcript was upregulated in this background (Figure 5), as was previously reported in the Wassilevskija ecotype (Sibout et al., 2005). CADA was not upregulated in these stem analyses (Figure 5). CADB1 and CADB2 transcripts were not detected in any of the genotypes analyzed.

Figure 5.

Figure 5

Open in a new tab

Expression of CAD genes in 4-week-old Arabidopsis stems. Relative expression measured by RT-qPCR, normalized to the reference gene ACTIN2 (At1g18780). Error bars represent standard deviation among biological replicates (n = 3). Transcript levels significantly different from wild-type (P < 0.05), determined by one-way ANOVA and Dunnet’s test (P < 0.05) are marked with (*).

To test the hypothesis that these CADs are catalyzing the reduction of p-coumaraldehyde in the quintuple mutant, we generated CADA and CADG knockouts in med5a/5b ref8-1 cadc cadd using the CRISPR/Cas9 system (Mao et al., 2013). Independent single-strand guide RNAs (sgRNAs) were designed to target two regions upstream of the CAD active site in CADA and CADG. Plants with a single base pair insertion upstream of the active site were selected for both CADs. For CADA, a line carrying a homozygous “T” insertion in the third exon was used for further experiments. For CADG, one of the transformants carried a single “A” insertion that generated a new BsmAI restriction site in the fourth exon. The CRISPR/Cas9-mediated edits in the coding sequence resulted in frame shifts and early stop codons in cada and cadg upstream of the conserved cofactor binding GX(X)GXXG motif; therefore, protein function is almost certainly disrupted.

To determine whether CADA and/or CADG contribute to the quintuple mutant’s total CAD activity, we performed CAD activity assays in the resulting hextuple mutant stems and found that there was no statistical difference in CAD activity between med5a/5b ref8-1 cadc cadd, med5a/5b ref8-1 cadc cadd cada, and med5a/5b ref8-1 cadc cadd cadg (Figure 4). In addition, to determine if mutation of CADA and/or CADG had an effect on lignin quantity or composition we analyzed the hextuple mutant’s lignin by DFRC and TGA analysis. Neither lignin quantity nor composition changed in med5a/5b ref8-1 cadc cadd cada nor med5a/5b ref8-1 cadc cadd cadg when compared with med5a/5b ref8-1 cadc cadd (Figures 6 and 7 and Table 1). In conclusion, neither CADA nor CADG alone are responsible for p-coumaryl alcohol synthesis in lignifying tissue.

Figure 6.

Figure 6

Open in a new tab

Lignin monomer composition of CADA knockouts determined by gas chromatography using the DFRC method. Error bars indicate the standard deviation among three biological replicates (n = 3). Statistical differences in H-lignin content were determined by Student’s t test, not significant (n.s.).

Figure 7.

Figure 7

Open in a new tab

Lignin monomer composition of CADG knockouts determined by gas chromatography using the DFRC method. Error bars indicate the standard deviation among three biological replicates (n = 3). Products not detected (n.d.). Statistical differences in H-lignin content were determined by Student’s t test, not significant (n.s.).

We also tested by liquid chromatography tandem mass spectrometry whether these mutants were accumulating higher levels of p-coumaraldehyde that was not incorporated into the lignin polymer. As seen in Table 2, even in the absence of CADC, CADD, CADA, and CADG, these plants continued to accumulate the same level of both the aldehyde and alcohol precursors as med5a/5b ref8-1.

Neither CCR1 nor CCR2 can catalyze the reduction of p-coumaraldehyde

Knocking out CADC, CADD, CADA, and/or CADG was not sufficient to prevent H-lignin synthesis and, although five other CADs remain untested, we also considered the possibility that an enzyme other than typical CADs might be involved. The enzyme 3-hydroxy-3-methyl-glutaryl-coenzyme A (HMG-CoA) reductase catalyzes the two-step reduction of HMG-CoA to mevalonate through a mevaldehyde intermediate (Haines et al., 2013). Based on this precedent, we tested the possibility that CINNAMOYL-CoA REDUCTASE (CCR), in a reaction analogous to that catalyzed by HMG-CoA reductase, can catalyze the two-step reduction of p-coumaroyl-CoA directly to p-coumaryl alcohol, obviating the need for an alcohol dehydrogenase comparable to CADC and CADD. Arabidopsis thaliana CCR1 and CCR2 were heterologously expressed with an N-terminal His-tag and purified, rendering active proteins in vitro. Both CCRs catalyzed the reduction of p-coumaroyl-CoA to p-coumaraldehyde, but p-coumaryl alcohol synthesis could not be detected under our assay conditions (Supplemental Figure S2) even when p-coumaraldehyde was provided directly as a substrate.

CADD is transcribed in med5a/5b cadd despite having a disrupted promoter

In the course of verifying the genotypes of several mutants with testcrosses, we conducted a backcross of med5a/5b ref8-1 cadc cadd to med5a/5b ref8-1. The F1 showed the expected ref phenotype and normal growth of med5a/5b ref8-1. On the contrary, the F1 progeny of a backcross of med5a/5b ref8-1 cadc cadd to cadc cadd did not exhibit the orange pigmentation in the interfascicular fibers as expected for a cadc cadd mutant suggesting that CAD activity is elevated in med5a/+ med5b/+ ref8-1/+ cadc cadd plants (Figure 8A). SAIL_776_B06 is a T-DNA line with an insertion in the CADD promoter that reduces transcript accumulation to levels close to the limit of detection (Figure 8B). Transcript analysis showed that although not statistically significant, CADD transcripts accumulate to nearly 20% of wild-type levels in med5a/5b ref8-1 cadc cadd and med5a/5b cadc cadd plants, consistent with the lack of xylem pigmentation in the F1 plants described earlier (Figure 8A). These findings suggest that the activation of CADD expression in the med5a/5b background may be responsible for the reduction of p-coumaraldehyde to p-coumaryl alcohol and the continued deposition of H-lignin in med5a/5b ref8-1 cadc cadd and the higher-order CAD mutants developed in this study.

Figure 8.

Figure 8

Open in a new tab

Stem cross-sections and expression of CADC and CADD in stems. A, Stem cross-sections of 7-week-old plants. Left, cadc cadd. Right, F1 of a cross between cadc cadd and med5a/5b ref8-1 cadc cadd. Scale bar = 100 µm. B, Expression of CADC and CADD genes in 4-week-old stems. Relative expression of CADC (top) and CADD (bottom) measured by RT-qPCR, normalized to the reference gene ACTIN2 (At1g18780). Error bars represent standard deviation among biological replicates (n = 3). The letters represent differences between genotypes determined by one-way ANOVA and Tukey’s honest significant difference test (P < 0.05).

med5a/5b ref8-1 cadc/d CRISPR-CADD plants still deposit lignin derived from p-coumaryl alcohol

In order to test the hypothesis that the low-level expression of CADD is associated with the synthesis of p-coumaryl alcohol in med5a/5b ref8-1 cadc cadd, an sgRNA was designed to target a region upstream of the CAD active site. The sgRNA targeted a BsaAI site located in the fourth exon of CADD that allowed screening for homozygous mutants by genotyping. Putative homozygous mutants were identified in the T1 by the loss of the BsaAI site. A trans-heterozygous T2 line, segregating for an “A” insertion and an “AC” deletion was used to test the effect of mutating CADD in med5a/5b ref8-1 cadc cadd. Elimination of CADD in med5a/5b ref8-1 cadc cadd resulted in plants with compromised growth (Figure 9A) and decreased fertility; however, extracts of med5a/5b ref8-1 cadc cadd CRISPR-cadd plants had the same activity toward coniferaldehyde and p-coumaraldehyde as the med5a/5b ref8-1 cadc cadd mutant (Figure 4). Lignin analysis (Figure 9B) revealed that the CRISPR-cadd plants have the same lignin composition as the med5a/5b ref8-1 cadc cadd mutant.

Figure 9.

Figure 9

Open in a new tab

Height and lignin composition of CRISPR-CADD knockouts. A, Height measurements of 5-week-old stems. Box plots depicting the median, first, and third quartiles. Dots represent biological replicates (n = 17) and (****) marks a significant difference between genotypes (P < 0.0001) by Student’s t test. B, Lignin monomer composition of CRISPR-CADD knockouts determined by gas chromatography using the DFRC method. Error bars indicate the standard deviation among three biological replicates (n = 3). Statistical differences in H-lignin content were determined by Student’s t test, not significant (n.s.).

Discussion

The saccharification potential and the absence of a growth deficit of biomass containing aldehyde-derived lignins make them an attractive substitute for biomass with conventional monolignol-derived lignins. Aldehyde-rich lignocellulosic material may not only reduce future biofuel production costs but has already been demonstrated to be a superior animal feedstock (Sattler et al., 2010). Hydroxycinnamaldehyde-derived lignins exist in Nature and they have also been generated in many plant species through metabolic engineering (Pillonel et al., 1991; Baucher et al., 1996; Ralph et al., 1997; Sibout et al., 2005; Zhao et al., 2013; Yamamoto et al., 2020). The discovery that p-coumaryl alcohol can be incorporated into lignin as the primary component in med5a/5b ref8-1 plants (Bonawitz et al., 2014) and the availability of a strategy to generate aldehyde-derived lignins (Anderson et al., 2015) appeared to provide the opportunity to generate plants that would deposit lignin derived from p-coumaraldehyde. To our knowledge, this compound has only been associated with lignin in cucumber (Cucumis sativa) under biotic stress (Varbanova et al., 2011) or as the end units of H-lignin, but not as a substantial component of lignin. In an attempt to generate this novel lignin in planta, we knocked out CADC and CADD in med5a/5b ref8-1 plants. Surprisingly, these experiments instead indicated that H-lignin can be synthesized independently of the two CADs that are known to participate in the synthesis of G and S lignin.

cadc cadd plants continue to exhibit CAD activity

There are nine CAD genes in A. thaliana: CADC, CADD, CAD1, CADA, CADB1, CADB2, CADE, CADF, and CADG (Costa et al., 2003). The involvement of all of these CADs in stem lignification has been tested using individual T-DNA mutant lines, with the exception of CADF, which lacks detectable transcript accumulation in wild-type plants (Eudes et al., 2006). CADC and CADD are the catalysts that contribute most substantially to lignification in wild-type mature stems (Sibout et al., 2005) as indicated by the fact that cadc cadd mutants deposit lignin, 99% of which is derived from coniferaldehyde and sinapaldehyde (Sibout et al., 2005; Anderson et al., 2015). Although both CADC and CADD exhibit activity toward p-coumaraldehyde in vitro, the p-coumaryl alcohol-derived lignin in med5a/5b ref8-1 cadc cadd plants (Figure 3) indicates that the activities of these two enzymes are not required for H-lignin synthesis, at least in this mutant context, and suggest that another CAD or related oxidoreductase may be involved. Consistent with this hypothesis, cadc cadd and med5a/5b ref8-1 cadc cadd plants exhibited substantial remaining CAD activity toward p-coumaraldehyde and coniferaldehyde (Figure 4). Previous in vitro studies revealed that CADB1, CADB2, CADE, and CADF exhibit activity toward p-coumaraldehyde, coniferaldehyde, and sinapaldehyde, although for the most part with lower catalytic efficiency than CADC and CADD (Kim et al., 2004). Based on their expression patterns derived from promoter reporter gene fusions and/or transcript analysis, CAD1, CADB1, CADB2, CADG, CADE, CADF, and/or CADA (Eudes et al., 2006; Kim et al., 2007) may contribute to the remaining CAD activity measured in cadc cadd stems. Given that cadc cadd mutants deposit aldehyde-derived lignins, the CAD activity in cadc cadd stems (Figure 4) must be located in nonlignifying tissues or in subcellular compartments that do not have access to the coniferaldehyde and sinapaldehyde that is directed into lignin biosynthesis, otherwise cadc cadd would incorporate conventional monolignols into its lignin instead of aldehydes. Underlining the importance of compartmentalization/tissue-specific expression, CAD1 can partially complement cadc cadd when its expression is driven by the CADD promoter (Eudes et al., 2006). Nevertheless, how these alternate CADs are inaccessible to coniferaldehyde and sinapaldehyde pools but can still function in the reduction of p-coumaraldehyde to p-coumaryl alcohol is unclear but presumably involves compartmentalization of substrate, enzyme, or both. Although published results using reporter gene constructs are somewhat variable, probably due to the nature of the promoter::reporter gene fusions employed, in general CADB1 is unlikely to play a major role in the reduction of p-coumaraldehyde because it is not expressed in lignifying tissues (Eudes et al., 2006; Kim et al., 2007). Furthermore, CADB1 and CADB2 were expressed at low levels in all the genotypes tested such that they were not detected by RT-qPCR (Figure 5). RNA-seq data from med5a/5b ref8-1 stems corroborated this finding (P. Wang, L. Guo, J. Morgan, N. Dudareva, and C. Chapple, unpublished data). According to promoter::reporter gene fusions, CADE and CADF are not expressed in stems (Kim et al., 2007), but we were able to amplify the transcript of at least one of these nearly identical genes by RT-qPCR leaving the localization of their expression and involvement in lignification an open question. Previous reports on the promoter activity of CAD1 are inconsistent (Eudes et al., 2006; Kim et al., 2007) and, although its role in lignification in young stems has been explored (Eudes et al., 2006), it is as yet unknown whether CAD1 plays a role in p-coumaryl alcohol synthesis. Alternatively, H-lignin synthesis in med5a/5b ref8-1 cadc cadd plants may depend on an as yet unidentified p-coumaraldehyde-specific form of the enzyme.

Because the bulk of monolignol synthesis in wild-type plants goes toward G and S monomers, little attention has been paid to H-lignin synthesis. Furthermore, the low levels of H-lignin in plants would have made quantifying changes in the deposition of these subunits challenging. In previous studies, CADA and CADG single mutants did not show a reduction in total CAD activity or lignin content (Eudes et al., 2006). Similarly, neither of these manipulations had an effect in stem total CAD activity (Figure 4) or H-lignin content in our studies (Figures 6 and 7), suggesting that these CADs are not involved in lignin monomer production, or are redundant with other CADs that can also catalyze p-coumaraldehyde reduction. Quantification of free p-coumaraldehyde from mutant stems also suggests that these two CADs do not play dominant roles in the reduction of p-coumaraldehyde, as it revealed that adding cada and cadg mutations to the med5a/5b ref8-1 cadc cadd background did not further enhance p-coumaraldehyde levels.

It is also possible that the reduction of p-coumaraldehyde is catalyzed by non-CAD oxidoreductases. We tested whether A. thaliana CCR1 and/or CCR2 could catalyze the reduction of p-coumaroyl-CoA to p-coumaraldehyde and subsequently to p-coumaryl alcohol, but only catalyze the reduction of feruloyl-CoA to coniferaldehyde. Our results indicated that CCR1 and CCR2 can catalyze the reduction of the CoA esters to their corresponding aldehydes (Supplemental Figure S2), as has been previously described in literature (Zhou et al., 2010), but cannot facilitate the two-step reduction of the CoA esters to the corresponding alcohols in a manner similar to HMG-CoA reductase (Haines et al., 2013).

Leaky expression of CADD in med5a/5b plants

By crossing med5a/5b ref8-1 cadc cadd to cadc cadd, we made the serendipitous finding that CADD is expressed in med5a/5b backgrounds carrying a homozygous T-DNA insertion in CADD (Figure 8B), even though it is expressed at levels close to the limit of detection in an otherwise wild-type background. An increase in CADC transcript levels was also detected in med5a/5b ref8-1 cadc cadd; however, the cadc T-DNA line has an insertion in the coding region that would result in a truncated enzyme with an altered active site (Youn et al., 2006). Unlike cadc, the T-DNA insertion in CADD disrupts the promoter region leaving the coding region intact, which if transcribed and translated, would lead to a functional enzyme. MED5A and MED5B negatively regulate phenylpropanoid metabolism by directly interacting with transcription factors and RNA polymerase II (Bonawitz et al., 2012). The increase in CADC, CADG, and CADD transcripts together with the increase in CAD activity in med5a/5b backgrounds exemplifies the role of MED5 as a negative regulator of phenylpropanoid biosynthesis. The absence of MED5 in the cadd T-DNA line leads to expression of CADD, probably explaining the lack of vascular pigmentation in med5a/5b cadc cadd and med5a/5b ref8-1 cadc cadd (Figure 8A). Indeed, even med5a/+ med5b/+ ref8-1/+ cadc cadd plants lacked the characteristic orange interfascicular fibers of cadc cadd mutants, which suggests that either MED5A or MED5B or both are haploinsufficient for suppression of the CADD allele in the T-DNA line. Despite the low expression of CADD in the med5a/5b background, we tested whether the leaky expression of CADD was leading to the synthesis of p-coumaryl alcohol by using the CRISPR/Cas9 system to target an exon in CADD in med5a/5b ref8-1 cadc cadd plants. Our results showed that these mutations had no effect on the lignin composition of med5a/5b ref8-1 cadc cadd (Figure 9B) but did have a negative effect on growth (Figure 9A) and fertility. It has yet to be determined whether the effects on growth and fertility arise from reduced CADD activity or they are the result of an off-target effect from the CRISPR/Cas9 system. Future users of SAIL_776_B06 should proceed with caution when interpreting their results, because it is possible that other conditions, such as abiotic or biotic stresses, might turn on the expression of this gene.

The enzymes involved in the lignin biosynthetic pathway are relatively well delineated in A.thaliana, although additional players have been identified in recent years (Vanholme et al., 2013; Barros et al., 2019). CADC and CADD had been established as the only reductases involved in H, G, and S monolignol biosynthesis in mature stems; however, this study has shown that the biosynthetic pathway for H-lignin synthesis remains to be elucidated.

Materials and methods

Plant materials

Arabidopsis (A.thaliana) ecotype Columbia-0 plants were grown in Propagation Mix (Sun Gro Horticulture, Agawam, MA, USA) supplemented with Osmocote Plus (ICL Fertilizers, Dublin OH, USA) at 22°C under long-day conditions (16-h light/8-h dark). The med5a/5b ref8-1 triple mutant and its genotyping information has been described previously (Bonawitz et al., 2014). The cadc (SAIL_1265_A06) and cadd (SAIL_776_B06) T-DNA lines were obtained from the Arabidopsis Biological Resource Center (The Ohio State University, Columbus, OH) and crossed to generate the cadc cadd mutant. Homozygous double mutants were identified by PCR using the primers listed in Supplemental Table S2.

RNA extraction and RT-qPCR

RNA from 4-week-old stems was extracted using the QIAGEN RNeasy Plant Kit and DNAse-treated using the Invitrogen TURBO DNA-free kit following the manufacturer’s instructions. Complementary DNA was synthesized using 1 µg of RNA and the Invitrogen MultiScribe Reverse Transcriptase. Quantitative PCR was performed using Applied Biosystems SYBR Green PCR Master Mix, gene-specific primers and ACTIN2 (At3g18780) as the internal standard (Supplemental Table S2). Fold change expression was determined by the ΔΔCt method. CADE and CADF were targeted with the same primers because they are 98% identical at the nucleotide level. Fold change expression was determined by the ΔΔCt method.

Lignin analysis

Lignin monomers were quantified by gas chromatography after DFRC (Lu and Ralph, 1998). Total lignin content was measured by the TGA method (Campbell and Ellis, 1992). Briefly, 30 mg of dried extractive-free cell wall residue was resuspended in 750 µL of distilled water, 250 µL of concentrated HCl, and 100 µL of TGA and incubated at 80°C for 3 h. Samples were centrifuged at 15,500 × g for 5 min and the supernatant was discarded. The samples were washed with distilled water and resuspended in 1 mL of 1 M NaOH, followed by incubation overnight on a rocking plate. Samples were centrifuged at 15,500 × g for 10 min and the supernatant was transferred to a new tube and 200 µL of concentrated HCl were added. Tubes were vortexed and incubated at 4°C for 4 h. The precipitate was collected by centrifugation and the supernatant was discarded. The pellet was dissolved in 1 mL of 1 M NaOH. Absorbance at 280 nm was measured with a spectrophotometer using 15 µL of sample diluted with 1 mL of 1 M NaOH.

Protein extraction and CAD activity assays

Four biological replicates consisting of two 5-week-old stems each, were harvested for each genotype. Stems without axillary stems, cauline leaves, and siliques were flash frozen in liquid nitrogen and ground with mortar and pestle. Tissue was suspended in four volumes of 100 mM Tris–HCl (pH 7.5) containing 5% (v/v) glycerol and 15 µM β-mercaptoethanol. Samples were centrifuged 10 min at 16,000 × g at 4°C and the supernatant was transferred to a new tube and re-centrifuged. A 250-µL sample of the supernatant was desalted in a disposable column packed with Sephadex G-50 (medium) pre-equilibrated in extraction buffer. Protein was quantified using the Bradford assay (Bradford, 1976) using bovine serum albumin as a standard.

CAD enzyme assays were performed using a modified version from Wyrambik and Grisebach (1975). Forward assays were carried out using an NADPH regenerating system that consisted of 500 µM NADP+, 5 mM glucose-6-phosphate and 0.5 units of glucose-6-phosphate dehydrogenase in 200 mM potassium phosphate (pH 6.5) in a final volume of 200 µL. Aliquots (180 µL) containing the NADPH regenerating system and 340 µM of one of the hydroxycinnamaldehyde substrates were preincubated for 5 min at 30°C. Assays were started by the addition of 20 µL of desalted protein and were incubated for 30 min at 30°C. Assays were stopped by the addition of 24 µL of glacial acetic acid. Samples were centrifuged for 1 h at 16,000 × g at 4°C prior to analysis by HPLC.

CCR heterologous expression and enzyme assays

Arabidopsis thaliana CCR1 and CCR2 were cloned into the Gateway pDEST17 vector and expressed in Escherichia coli BL21 (DE3) cells as fusion proteins with an N-terminal 6X-His tag. To generate the substrate p-coumaroyl-CoA, 4-COUMARATE-CoA LIGASE 1 (4CL1) was expressed in E. coli BL21 (DE3) cells as a fusion protein with a 6X-His tag as previously described in Li et al. (2010). For all proteins, cells were grown to an optical density of 0.6, expression was induced by the addition of 2.5 mM isopropyl β-d-thiogalactoside, and cells were incubated overnight at 18°C. Cells were harvested and lysed with lysozyme and DNAseI as described by Weng et al. (2011). The three fusion proteins were purified using Ni-NTA agarose beads. PD-10 (GE-Healthcare, Chicago, IL, USA) columns were used for desalting the proteins and buffer exchange; the column was equilibrated with 100 mM Tris–HCl containing 10% glycerol (v/v) for 4CL and 100 mM HEPES-NaOH (pH 7.5) containing 10% glycerol (v/v) for CCR1/2 (Zhou et al., 2010). Protein was quantified as described above.

For CCR assays, recombinant 4CL1 was first used to generate p-coumaroyl-CoA by incubating it with 200 µM p-coumarate, 220 µM Coenzyme A, 1 mM ATP, and 1 mM MgCl2 in 100 mM Tris–HCl (pH 8.0) for 30 min at 30°C. Recombinant CCR was then added to the reaction in the presence of 1 mM NADPH and incubated at 30°C for 30 min. The reactions were terminated by acidification with glacial acetic acid and the products analyzed by HPLC.

p-Coumaryl alcohol and p-coumaraldehyde quantification

Soluble metabolites were extracted from 36-day-old stems and analyzed by LC–MS/MS as previously described (Jaini et al., 2017).

Generation of cada, cadg, and cadd using CRISPR/Cas9

The cada mutant was generated by constructing two sgRNAs targeting CADA. The first sgRNA was generated by cloning the complementary primers CC4942 and CC4943 into the BbsI-digested psgR-Cas9-At plasmid (Mao et al., 2013) to generate psgR-Cas9-At-target1. The second sgRNA was generated in a similar manner using primers CC4944 and CC4945 and resulted in psgR-Cas9-At-target2. The A.thaliana small nucleolar RNA U6 and the sgRNA from psgR-Cas9-At-target2 was PCR amplified with primers containing KpnI and EcoRI overhangs as described in Mao et al. (2013) and cloned into psgR-Cas9-At-target1 to generate psgR-Cas9-At-target1&2. The cassette contains both sgRNAs and the Cas9 gene were excised from psgR-Cas9-At-target1&2 by digesting with EcoRI and HindIII and cloned into the binary vector pCAMBIA1300 resulting in psgR-Cas9-At-CADA (pCC 2436). The construct was introduced into med5a/5b ref8-1 cadc cadd plants via Agrobacterium tumefaciens-mediated transformation (Clough and Bent, 1998). Transformants were selected on plates with hygromycin and transferred to soil. Genomic DNA was extracted from the transformants and PCR-amplified with primer pair CC4949/CC5009 for target 1 and CC5012/CC5007 for target 2. Amplicons were sequenced with primers CC5010 and CC5011 for target 1 and C4950 and CC5013 for target 2.

The cadg mutant was generated by transforming med5a/5b ref8-1 cadc cadd plants with psgR-Cas9-At-CADG. The psgR-Cas9-At-CADG plasmid (pCC 2437) was constructed as described above using primers CC5494 and CC5495 for target 1 and CC5496 and CC5497 for target 2. After selection, genomic DNA was extracted from the transformants and both targets were PCR-amplified with primers CC5418 and CC5236. The amplicon was sequenced using primer CC5417 for target 1 and CC5236 for target 2. One of the transformants carried a single “A” insertion in target 1 that generated a new BsmAI site. A homozygous line for this edit was selected by genotyping the next generations using the primer pair CC5416 and CC5417 and digesting the amplicon with BsmAI.

CADD was targeted with a single sgRNA which was generated by cloning the complementary primers CC5847 and CC5848 into psgR-Cas9-At. The plasmid was digested with KpnI and HindIII to excise the sgRNA and Cas9 cassette and cloned into pCAMBIA1300 to generate psgR-Cas9-At-CADD (pCC 2423). The sgRNA was designed to target an existing BsaAI site, therefore after transformation of med5a/5b ref8-1 cadc cadd, genomic DNA from T1 plants was PCR amplified with primers CC5861 and CC5862 and digested with BsaAI. DNA from plants that generated amplicons that did not digest with BsaAI were used for amplification of the corresponding genomic region and were sequenced with primer CC5863.

CADG and CADD targets were designed using the online tool E-CRISP (http://www.e-crisp.org/E-CRISP/) (Heigwer et al., 2014). Primer sequence information can be found on Supplemental Table S2.

NMR analysis of enzyme lignin

The enzyme lignins (ELs) were prepared from whole cell walls after solvent-extraction (water ×3; 80% EtOH [v/v] ×3) as described in a previous publication (Kim et al., 2017). The extractive-free cell walls were ball-milled and treated with cellulases (Cellulysin, EC 3.2.1.4, activity >10,000 units/g, Calbiochem) from Trichoderma viride. The ground cell walls were suspended in NaOAc buffer (pH 5), and Cellulysin was added. The reaction mixture was shaken on a rotary incubator shaker at 35°C for 72 h. The residue was collected by centrifugation (8,000 rpm, 30 min), and the digestion process was repeated 3 times. The collected lignin residue was sonicated and washed with deionized water 3 times and lyophilized.

NMR experiments were performed and the data were analyzed as previously reported (Kim et al., 2010, 2017). NMR spectra of the ELs were acquired on a Bruker Biospin (Billerica, MA, USA) Avance 700 MHz spectrometer equipped with a 5-mm QCI 1H/31P/13C/15N cryoprobe with inverse geometry (proton coils closest to the sample). DMSO-d6:pyridine-d5 (4:1, v/v) was the solvent, and the central DMSO solvent peak (δC 39.5, δH 2.49 ppm) was used as the internal reference. Isotopically enriched pyridine-d5 (“100”; min. 99.94 atom% D) was used to minimize interference between the peaks from aromatic moieties and residual solvent. All NMR experiments used Bruker’s standard pulse programs; an adiabatic 1H–13C 2D HSQC experiment (hsqcetgpsisp2.2; phase-sensitive gradient-edited 2D HSQC using adiabatic pulse sequences for inversion and refocusing) was used to collect the main data (Kupče and Freeman, 2007). The HSQC experiments were acquired from 11.5 to −0.5 ppm (12 ppm spectral width) in F2 (1H) with 3,366 data points (acquisition time, 200 ms) and 215 to −5 ppm (220 ppm spectral width) in F1 (13C) with 620 increments (F1 acquisition time, 8.0 ms) of 32 scans with a 1 s interscan delay (D1); the d24 delay was 0.86 ms (1/8J, J = 145 Hz). The total acquisition time for a sample was 5.5 h. The spectra were processed using Gaussian apodization (GB = 0.001, LB = −0.5) in F2 and squared cosine-bell and 32 coefficients of linear prediction in F1. Volume integration of contours in HSQC plots was carried out using TopSpin version 4.1.3 (Mac version) software.

Statistical analysis

All statistical analyses were performed using GraphPad Prism version 9.3.1 for macOS, GraphPad Software, San Diego, CA, USA, www.graphpad.com. The statistical analyses performed are indicated in the figure legends.

Accession numbers

Sequence data from this article can be found in the GenBank/EMBL data libraries under the accession numbers listed in Supplemental Table S1.

Supplemental data

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

Supplemental Figure S1. Height measurements of 6-week-old stems.

Supplemental Figure S2 . CCR assays with heterologously expressed CCR1 and CCR2 from A. thaliana.

Supplemental Table S1. CAD genes in Arabidopsis and the nomenclatures used for them.

Supplemental Table S2. List of primers used.

Supplemental Table S3. Fold change expression of AtCAD genes relative to wild-type determined by RNA-seq.

Supplementary Material

kiac210_Supplementary_Data
Click here for additional data file. (891.9KB, pdf)

Acknowledgments

NMR data were acquired on a Bruker NEO 700 MHz instrument at the NMR facility in the Wisconsin Energy Institute of the University of Wisconsin-Madison.

Funding

This work was supported by the Center for Direct Catalytic Conversion of Biomass to Biofuels, an Energy Frontier Research Center funded by the US Department of Energy, Office of Science, Office of Basic Energy Sciences (DE-SC0000997), and the US Department of Energy, Office of Science, Office of Basic Energy Sciences, Chemical Sciences, Geosciences, and Biosciences Division (DE-FG02-07ER15905). J.R. and H.K. were funded by the Great Lakes Bioenergy Research Center (DOE BER Office of Science DE-SC0018409).

Conflict of interest statement: The authors declare no conflict of interest.

Contributor Information

Fabiola Muro-Villanueva, Department of Biochemistry, Purdue University, West Lafayette, Indiana 47907, USA; Center for Plant Biology, Purdue University, West Lafayette, Indiana 47907, USA.

Hoon Kim, US Department of Energy’s Great Lakes Bioenergy Research Center (GLBRC), Wisconsin Energy Institute (WEI), Madison, Wisconsin 53726, USA.

John Ralph, US Department of Energy’s Great Lakes Bioenergy Research Center (GLBRC), Wisconsin Energy Institute (WEI), Madison, Wisconsin 53726, USA; Department of Biochemistry, University of Wisconsin–Madison, Madison, Wisconsin 53706, USA.

Clint Chapple, Department of Biochemistry, Purdue University, West Lafayette, Indiana 47907, USA; Center for Plant Biology, Purdue University, West Lafayette, Indiana 47907, USA.

 

C.C. and F.M.-V. conceived the project and designed and executed the experiments with the exception of the NMR which was designed and executed by J.R. and H.K. F.M.-V., C.C., H.K., and J.R. wrote the manuscript.

The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (https://academic.oup.com/plphys/pages/general-instructions) is: Clint Chapple (chapple@purdue.edu).

References

  1. Anderson NA, Tobimatsu Y, Ciesielski PN, Ximenes E, Ralph J, Donohoe BS, Ladisch M, Chapple C (2015) Manipulation of guaiacyl and syringyl monomer biosynthesis in an Arabidopsis cinnamyl alcohol dehydrogenase mutant results in atypical lignin biosynthesis and modified cell wall structure. Plant Cell 27: 2195–2209 [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Barros J, Escamilla-Treviño L, Song L, Rao X, Serrani-Yarce JC, Palacios MD, Engle N, Choudhury FK, Tschaplinski TJ, Venables BJ, et al. (2019) 4-Coumarate 3-hydroxylase in the lignin biosynthesis pathway is a cytosolic ascorbate peroxidase. Nat Commun 10: 1994. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Baucher M, Chabbert B, Pilate G, Van Doorsselaere J, Tollier MT, Petit-Conil M, Cornu D, Monties B, Van Montagu M, Inzé D, et al. (1996) Red xylem and higher lignin extractability by down-regulating a cinnamyl alcohol dehydrogenase in poplar. Plant Physiol 112: 1479–1490 [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Bonawitz ND, Soltau WL, Blatchley MR, Powers BL, Hurlock AK, Seals LA, Weng JK, Stout J, Chapple C (2012) REF4 and RFR1, subunits of the transcriptional coregulatory complex mediator, are required for phenylpropanoid homeostasis in Arabidopsis. J Biol Chem 287: 5434–5445 [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bonawitz ND, Kim JI, Tobimatsu Y, Ciesielski PN, Anderson NA, Ximenes E, Maeda J, Ralph J, Donohoe BS, Ladisch M, et al. (2014) Disruption of Mediator rescues the stunted growth of a lignin-deficient Arabidopsis mutant. Nature 509: 376–380 [DOI] [PubMed] [Google Scholar]
  6. Bradford M (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 172: 248–254 [DOI] [PubMed] [Google Scholar]
  7. Campbell MM, Ellis BE (1992) Fungal elicitor-mediated responses in pine cell cultures. I. Induction of phenylpropanoid metabolism. Planta 186: 409–417 [DOI] [PubMed] [Google Scholar]
  8. Clough SJ, Bent AF (1998) Floral dip: a simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J 16: 735–743 [DOI] [PubMed] [Google Scholar]
  9. Costa MA, Collins RE, Anterola AM, Cochrane FC, Davin LB, Lewis NG (2003) An in silico assessment of gene function and organization of the phenylpropanoid pathway metabolic networks in Arabidopsis thaliana and limitations thereof. Phytochemistry 64: 1097–1112 [DOI] [PubMed] [Google Scholar]
  10. Eudes A, Pollet B, Sibout R, Do CT, Séguin A, Lapierre C, Jouanin L (2006) Evidence for a role of AtCAD 1 in lignification of elongating stems of Arabidopsis thaliana. Planta 225: 23–39 [DOI] [PubMed] [Google Scholar]
  11. Franke R, McMichael CM, Meyer K, Shirley AM, Cusumano JC, 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]
  12. Franke R, Hemm MR, Denault JW, Ruegger MO, Humphreys JM, Chapple C (2002) 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. Haines BE, Wiest O, Stauffacher CV (2013) The increasingly complex mechanism of HMG-CoA reductase. Acc Chem Res 46: 2416–2426 [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Heigwer F, Kerr G, Boutros M (2014) E-CRISP: fast CRISPR target site identification. Nat Methods 11: 122–123 [DOI] [PubMed] [Google Scholar]
  15. Himmel ME, Ding S, Johnson DK, Adney WS, Nimlos MR, Brady JW, Foust TD (2007) Biomass recalcitrance: engineering plants and enzymes for biofuels production. Science 315: 804–807 [DOI] [PubMed] [Google Scholar]
  16. Huntley SK, Ellis D, Gilbert M, Chapple C, Mansfield SD (2003) Significant increases in pulping efficiency in C4H-F5H-transformed poplars: improved chemical savings and reduced environmental toxins. J Agric Food Chem 51: 6178–6183 [DOI] [PubMed] [Google Scholar]
  17. Jaini R, Wang P, Dudareva N, Chapple C, Morgan JA (2017) Targeted metabolomics of the phenylpropanoid pathway in Arabidopsis thaliana using reversed phase liquid chromatography coupled with tandem mass spectrometry. Phytochem Anal 28: 267–276 [DOI] [PubMed] [Google Scholar]
  18. Kim H, Ralph J, Yahiaoui N, Pean M, Boudet AM (2000) Cross-Coupling of hydroxycinnamyl aldehydes into lignins. Organ Lett 2: 2197–2200 [DOI] [PubMed] [Google Scholar]
  19. Kim H, Ralph J, Lu F, Ralph SA, Boudet AM, MacKay JJ, Sederoff RR, Ito T, Kawai S, Ohashi H, Higuchi T (2003) NMR analysis of lignins in CAD-deficient plants. Part 1. Incorporation of hydroxycinnamaldehydes and hydroxybenzaldehydes into lignins. Org Biomol Chem 1: 268–281 [DOI] [PubMed] [Google Scholar]
  20. Kim SJ, Kim MR, Bedgar DL, Moinuddin SGA, Cardenas CL, Davin LB, Kang CH, Lewis NG (2004) Functional reclassification of the putative cinnamyl alcohol dehydrogenase multigene family in Arabidopsis. Proc Natl Acad Sci USA 101: 1455–1460 [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Kim SJ, Kim KW, Cho MH, Franceschi VR, Davin LB, Lewis NG (2007) Expression of cinnamyl alcohol dehydrogenases and their putative homologues during Arabidopsis thaliana growth and development: lessons for database annotations? Phytochemistry 68: 1957–1974 [DOI] [PubMed] [Google Scholar]
  22. Kim H, Ralph J (2010) Solution-state 2D NMR of ball-milled plant cell wall gels in DMSO-d6/pyridine-d5. Organic Biomol Chem 8: 576–591 [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Kim H, Padmakshan D, Li Y, Rencoret J, Hatfield RD, Ralph J (2017) Characterization and elimination of undesirable protein residues in plant cell wall materials for enhancing lignin analysis by solution-state NMR. Biomacromolecules 18: 4184–4195 [DOI] [PubMed] [Google Scholar]
  24. Kupče E, Freeman R (2007) Compensated adiabatic inversion pulses: broadband INEPT and HSQC. J Magnet Resonance 187: 258–265 [DOI] [PubMed] [Google Scholar]
  25. Li X, Bonawitz ND, Weng JK, Chapple C (2010) The growth reduction associated with repressed lignin biosynthesis in Arabidopsis thaliana is independent of flavonoids. Plant Cell 22: 1620–1632 [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. 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]
  27. Mao Y, Zhang H, Xu N, Zhang B, Gou F, Zhu JK (2013) Application of the CRISPR-Cas system for efficient genome engineering in plants. Mol Plant 6: 2008–2011 [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. McCann MC, Carpita NC (2015) Biomass recalcitrance: a multi-scale, multi-factor, and conversion-specific property. J Exp Bot 66: 4109–4118 [DOI] [PubMed] [Google Scholar]
  29. Mottiar Y, Vanholme R, Boerjan W, Ralph J, Mansfield SD (2016) Designer lignins: harnessing the plasticity of lignification. Curr Opin Biotechnol 37: 190–200 [DOI] [PubMed] [Google Scholar]
  30. Meyer K, Shirley A M, Cusumano JC, Bell-Lelong DA, 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]
  31. Nonhebel S (2005) Renewable energy and food supply: will there be enough land? Renew Sustain Energy Rev 9: 191–201 [Google Scholar]
  32. Pillonel C, Mulder MM, Boon JJ, Forster B, Binder A (1991) Involvement of cinnamyl-alcohol dehydrogenase in the control of lignin formation in Sorghum bicolor L. Moench. Planta 185: 538–544 [DOI] [PubMed] [Google Scholar]
  33. Ralph J, Mackay JJ, Hatfield RD, O’Malley DM, Whetten RW, Sederoff RR (1997) Abnormal lignin in a loblolly pine mutant. Science 277: 235–239 [DOI] [PubMed] [Google Scholar]
  34. Ralph J, Lundquist K, Brunow G, Lu F, Kim H, Schatz PF, Marita JM, Hatfield RD, Ralph SA, Christensen JH, et al. (2004) Lignins: natural polymers from oxidative coupling of 4-hydroxyphenyl-propanoids. Phytochem Rev 3: 29–60 [Google Scholar]
  35. Ralph J, Brunow G, Harris PJ, Dixon RA, Schatz PF, Boerjan W (2008) Lignification: are lignins biosynthesized via simple combinatorial chemistry or via proteinaceous control and template replication? InDaayf F, Lattanzio V, eds, Recent Advances in Polyphenol Research, Vol 1. Willey-Blackwell Publishing, Oxford, pp 36–66 [Google Scholar]
  36. Ralph J, Lapierre C, Boerjan W (2019) Lignin structure and its engineering. Curr Opin Biotechnol 56: 240–249 [DOI] [PubMed] [Google Scholar]
  37. Reddy MSS, Chen F, Shadle G, Jackson L, Aljoe H, Dixon RA (2005) Targeted down-regulation of cytochrome P450 enzymes for forage quality improvement in alfalfa (Medicago sativa L.). Proc Natl Acad Sci USA 102: 16573–16578 [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Renault H, Werck-Reichhart D, Weng JK (2019) Harnessing lignin evolution for biotechnological applications. Curr Opin Biotechnol 56: 105–111 [DOI] [PubMed] [Google Scholar]
  39. Sattler SE, Funnell-Harris DL, Pedersen JF (2010) Brown midrib mutations and their importance to the utilization of maize, sorghum, and pearl millet lignocellulosic tissues. Plant Sci 178: 229–238 [Google Scholar]
  40. Shuai L, Amiri MT, Questell-Santiago YM, Héroguel F, Li Y, Kim H, Meilan R, Chapple C, Ralph J, Luterbacher JS (2016) Formaldehyde stabilization facilitates lignin monomer production during biomass depolymerization. Science 354: 329–333 [DOI] [PubMed] [Google Scholar]
  41. Sibout R, Eudes A, Mouille G, Pollet B, Lapierre C, Jouanin L, Séguin A (2005) CINNAMYL ALCOHOL DEHYDROGENASE-C and -D are the primary genes involved in lignin biosynthesis in the floral stem of Arabidopsis. Plant Cell 17: 2059–2076 [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Stewart JJ, Akiyama T, Chapple C, Ralph J, Mansfield SD (2009) The effects on lignin structure of overexpression of ferulate 5-hydroxylase in hybrid poplar. Plant Physiol 150: 621–635 [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Tavares R, Aubourg S, Lecharny A, Kreis M (2000) Organization and structural evolution of four multigene families in Arabidopsis thaliana: AtLCAD, AtLGT, AtMYST and AtHD-GL2. Plant Mol Biol 42: 703–717 [DOI] [PubMed] [Google Scholar]
  44. Turner SR, Somerville CR (1997) Collapsed xylem phenotype of Arabidopsis identifies mutants deficient in cellulose deposition in the secondary cell wall. Plant Cell 9: 689–701 [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. US Department of Energy, US Department of Agriculture (2005) Biomass as Feedstock for a Bioenergy and Bioproducts Industry: The Technical Feasibility of a Billion-Ton Annual Supply. https://www1.eere.energy.gov/bioenergy/pdfs/final_billionton_vision_report2.pdf
  46. Vanholme R, Cesarino I, Rataj K, Xiao Y, Sundin L, Goeminne G, Kim H, Cross J, Morreel K, Araujo P, et al. (2013) Caffeoyl shikimate esterase (CSE) is an enzyme in the lignin biosynthetic pathway in Arabidopsis. Science 341: 1103–1106 [DOI] [PubMed] [Google Scholar]
  47. Vanholme R, De Meester B, Ralph J, Boerjan W (2019) Lignin biosynthesis and its integration into metabolism. Curr Opin Biotechnol 56: 230–239 [DOI] [PubMed] [Google Scholar]
  48. Varbanova M, Porter K, Lu F, Ralph J, Hammerschmidt R, Daniel Jones A, Day B (2011) Molecular and biochemical basis for stress-induced accumulation of free and bound p-coumaraldehyde in cucumber. Plant Physiol 157: 1056–1066 [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Wang P, Dudareva N, Morgan JA, Chapple C (2015) Genetic manipulation of lignocellulosic biomass for bioenergy. Curr Opin Chem Biol 29: 32–39 [DOI] [PubMed] [Google Scholar]
  50. Weng JK, Akiyama T, Ralph J, Chapple C (2011) Independent recruitment of an O-methyltransferase for syringyl lignin biosynthesis in Selaginella moellendorffii. Plant Cell 23: 2708–2724 [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Wyrambik D, Grisebach H (1975) Purification and properties of isoenzymes of cinnamyl-alcohol dehydrogenase from soybean-cell-suspension cultures. Eur J Biochem 59: 9–15 [DOI] [PubMed] [Google Scholar]
  52. Yamamoto M, Tomiyama H, Koyama A, Okuizumi H, Liu S, Vanholme R, Goeminne G, Hirai Y, Shi H, Takata N, et al. (2020) A century-old mystery unveiled: sekizaisou is a natural lignin mutant. Plant Physiol 182: 1821–1828 [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Yang H, Zhang X, Luo H, Liu B, Shiga TM, Li X, Kim JI, Rubinelli P, Overton JC, Subramanyam V, et al. (2019) Overcoming cellulose recalcitrance in woody biomass for the lignin-first biorefinery. Biotechnol Biofuels 12: 1–18 [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Youn B, Camacho R, Moinuddin SGA, Lee C, Davin LB, Lewis NG, Kang C (2006) Crystal structures and catalytic mechanism of the Arabidopsis cinnamyl alcohol dehydrogenases AtCAD5 and AtCAD4. Org Biomol Chem 4: 1687–1697 [DOI] [PubMed] [Google Scholar]
  55. Zeng Y, Zhao S, Yang S, Ding SY (2014) Lignin plays a negative role in the biochemical process for producing lignocellulosic biofuels. Curr Opin Biotechnol 27: 38–45 [DOI] [PubMed] [Google Scholar]
  56. Zhao Q, Tobimatsu Y, Zhou R, Pattathil S, Gallego-Giraldo L, Fu C (2013) Loss of function of cinnamyl alcohol dehydrogenase 1 leads to unconventional lignin and a temperature-sensitive growth defect in Medicago truncatula. Proc Natl Acad Sci USA 110: 13660–13665 [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Zhou R, Jackson L, Shadle G, Nakashima J, Temple S, Chen F, Dixon RA (2010) Distinct cinnamoyl-CoA reductases involved in parallel routes to lignin in Medicago truncatula. Proc Natl Acad Sci USA 107: 17803–17808 [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

kiac210_Supplementary_Data
Click here for additional data file. (891.9KB, pdf)

Articles from Plant Physiology are provided here courtesy of Oxford University Press

RESOURCES