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. 2022 Sep 19;191(1):70–86. doi: 10.1093/plphys/kiac432

Lignocellulose molecular assembly and deconstruction properties of lignin-altered rice mutants

Andri Fadillah Martin 1,2,#, Yuki Tobimatsu 3,✉,b,#, Pui Ying Lam 4,5, Naoyuki Matsumoto 6, Takuto Tanaka 7, Shiro Suzuki 8,9, Ryosuke Kusumi 10, Takuji Miyamoto 11,12, Yuri Takeda-Kimura 13,14, Masaomi Yamamura 15,16, Taichi Koshiba 17,18, Keishi Osakabe 19, Yuriko Osakabe 20, Masahiro Sakamoto 21, Toshiaki Umezawa 22,23,b
PMCID: PMC9806629  PMID: 36124989

Abstract

Bioengineering approaches to modify lignin content and structure in plant cell walls have shown promise for facilitating biochemical conversions of lignocellulosic biomass into valuable chemicals. Despite numerous research efforts, however, the effect of altered lignin chemistry on the supramolecular assembly of lignocellulose and consequently its deconstruction in lignin-modified transgenic and mutant plants is not fully understood. In this study, we aimed to close this gap by analyzing lignin-modified rice (Oryza sativa L.) mutants deficient in 5-HYDROXYCONIFERALDEHYDE O-METHYLTRANSFERASE (CAldOMT) and CINNAMYL ALCOHOL DEHYDROGENASE (CAD). A set of rice mutants harboring knockout mutations in either or both OsCAldOMT1 and OsCAD2 was generated in part by genome editing and subjected to comparative cell wall chemical and supramolecular structure analyses. In line with the proposed functions of CAldOMT and CAD in grass lignin biosynthesis, OsCAldOMT1-deficient mutant lines produced altered lignins depleted of syringyl and tricin units and incorporating noncanonical 5-hydroxyguaiacyl units, whereas OsCAD2-deficient mutant lines produced lignins incorporating noncanonical hydroxycinnamaldehyde-derived units. All tested OsCAldOMT1- and OsCAD2-deficient mutants, especially OsCAldOMT1-deficient lines, displayed enhanced cell wall saccharification efficiency. Solid-state nuclear magnetic resonance (NMR) and X-ray diffraction analyses of rice cell walls revealed that both OsCAldOMT1- and OsCAD2 deficiencies contributed to the disruptions of the cellulose crystalline network. Further, OsCAldOMT1 deficiency contributed to the increase of the cellulose molecular mobility more prominently than OsCAD2 deficiency, resulting in apparently more loosened lignocellulose molecular assembly. Such alterations in cell wall chemical and supramolecular structures may in part account for the variations of saccharification performance of the OsCAldOMT1- and OsCAD2-deficient rice mutants.

Analysis of lignin-altered rice mutants reveals how lignin structural modifications impose loosened cell wall molecular assembly and boost biomass deconstruction.

Introduction

Lignocellulosic biomass is an abundant, renewable carbon source that can be exploited for the sustainable production of bio-based energy and chemicals. Lignocellulose is a sophisticated biocomposite majorly produced in the plant secondary cell walls, in which three major structural polymers—cellulose, hemicelluloses, and lignin—intricately interact with each other through both noncovalent and covalent linkages (Simmons et al., 2016; Nishimura et al., 2018; Kang et al., 2019; Terrett and Dupree, 2019; Kirui et al., 2022). The development of plant biotechnology methods for optimizing the composition, chemical structures, and supramolecular assemblies of lignocellulose is an active area of research worldwide as such approaches may greatly facilitate the production of lignocellulose-derived biofuels and biochemicals in an economically and environmentally sound manner (Ragauskas et al., 2014; Marriott et al., 2016; Bhatia et al., 2017; Umezawa et al., 2018).

Lignin, a phenylpropanoid polymer typically accounting for 15%–30% of lignocellulosic materials, has long been recognized as a key negative factor in conventional polysaccharide-oriented lignocellulose utilization processes, such as those used in the production of pulp and paper as well as fermentable sugars for value-added downstream products (Boerjan et al., 2003; Chen and Dixon, 2007; Umezawa, 2010; Wang et al., 2015; Mottiar et al., 2016). More recently, lignin has been increasingly viewed as a viable source for bio-based aromatic chemicals (Rinaldi et al., 2016; Schutyser et al., 2018; Sun et al., 2018; Abu-Omar et al., 2021). Bioengineering approaches for the manipulation of lignin content and structure for both polysaccharide- and lignin-oriented lignocellulose utilization are accordingly becoming an important research focus (Umezawa, 2018; Halpin, 2019; Mahon and Mansfield, 2019; Ralph et al., 2019; Umezawa et al., 2020). To date, numerous mutants and transgenic plants producing cell walls with altered lignin contents and/or chemical structures have been generated via upregulation and/or downregulation of various lignin biosynthetic genes in different plant species. Some of these lignin-modified plants indeed display desired, improved biomass characteristics, apparently without compromising plant growth (Umezawa, 2018; Halpin, 2019; Mahon and Mansfield, 2019; Ralph et al., 2019). Nevertheless, however, the underlying relationships between altered lignin chemistry and lignocellulose properties, including those associated with biomass utilization and also plant development and growth, are still only partially understood.

In this study, we conducted a comparative and integrative analysis of cell wall chemical structures, supramolecular structures, and digestibility in lignin-modified mutant lines of rice (Oryza sativa L.), a model crop species representative of various grasses, including numerous biomass crops showing great potential as lignocellulose feedstocks (Umezawa, 2018; Halpin, 2019; Coomey et al., 2020; Umezawa et al., 2020). In particular, we focused on a series of rice mutant lines deficient in 5-HYDROXYCONIFERALDEHYDE O-METHYLTRANSFERASE (CAldOMT; also known as CAFFEIC ACID O-METHYLTRANSFERASE, COMT) and/or CINNAMYL ALCOHOL DEHYDROGENASE (CAD), both of which encode key enzymes involved in lignin biosynthesis and represent major bioengineering targets for improving lignin characteristics toward better biomass utilization.

Because CAldOMT and CAD both play key roles in the metabolic pathways leading to diverse lignin monomers (Supplemental Figure S1), manipulations of their gene expression result in prominent shifts in subunit composition of lignin polymers, occasionally accompanied by changes in total lignin contents of cell walls. CAldOMT catalyzes the O-methylation of the 5-hydroxyconiferaldehyde (and/or 5-hydroxyconiferyl alcohol) intermediate in the syringyl (S) lignin biosynthetic pathway diverging from the guaiacyl (G) lignin biosynthetic pathway (Supplemental Figure S1) (Osakabe et al., 1999; Li et al., 2000; Koshiba et al., 2013a). Consistent with this notion, downregulation of CAldOMT typically results in a decrease in S units and an increase in G units, which is often accompanied by the appearance of atypical 5-hydroxyguaiacyl (5H) units in lignin polymers (Jouanin et al., 2000; Guo et al., 2001; Ralph et al., 2001; Nakatsubo et al., 2008; Vanholme et al., 2010; Weng et al., 2010; Koshiba et al., 2013a; Daly et al., 2019; Lam et al., 2019a; Wu et al., 2019). In addition, recent studies have provided evidence that grasses utilize CAldOMT in the parallel flavonoid biosynthetic pathway dedicated to the production of tricin, a grass-specific lignin monomer (Lam et al., 2021) (Supplemental Figure S1). Downregulation of CAldOMT in grasses therefore results in concomitant reductions of S and tricin (T) units in lignin polymers (Eudes et al., 2017; Fornalé et al., 2017; Lam et al., 2019a). On the other hand, CAD is responsible for the final step in the biosynthesis of monolignols by reducing hydroxycinnamaldehyde precursors into their corresponding monolignols (Supplemental Figure S1) (Umezawa, 2010). Accordingly, downregulation of CAD usually leads to augmentation of atypical γ-aldehyde functionalities in lignin polymers through incorporation of noncanonical hydroxycinnamaldehydes (Higuchi et al., 1994; Lapierre et al., 1999; Kim et al., 2000, 2003; Ralph et al., 2001; Sibout et al., 2005; Zhao et al., 2013; Koshiba et al., 2013b; Anderson et al., 2015; Van Acker et al., 2017; Martin et al., 2019).

Such lignin modifications induced by CAldOMT and CAD deficiencies may substantially mitigate lignin-associated biomass recalcitrance in polysaccharide-oriented biorefinery contexts. In several grass species, defects in CAldOMT and CAD expressions are known to be associated with the brown midrib (bm) phenotype that often enhances forage digestibility (Sattler et al., 2010; Umezawa et al., 2018; Halpin, 2019). Similar to bm mutants, CAldOMT- and CAD-deficient transgenic plants, including nongrass species, frequently exhibit improved biomass deconstruction characteristics related to forage digestibility (Chen et al., 2003; Tu et al., 2010) and/or performance in chemical pulping (Lapierre et al., 1999; O’Connell et al., 2002; Pilate et al., 2002) and enzymatic saccharification (Chen and Dixon, 2007; Fu et al., 2011; Saathoff et al., 2011; Fornalé et al., 2012, 2017; Jung et al., 2012; Bouvier D’Yvoire et al., 2013; Van Acker et al., 2013, 2017; Koshiba et al., 2013a, 2013b; Anderson et al., 2015; Ho-Yue-Kuang et al., 2016; Kannan et al., 2018; Daly et al., 2019; Kim et al., 2019; Martin et al., 2019; Wu et al., 2019; Lam et al., 2019a).

Despite the many studies on CAldOMT- and CAD-deficient plants, the molecular-level relationship between CAldOMT- and CAD-associated lignin modifications and concurrent improvements in cell wall digestibility remains largely unknown. Recently, we demonstrated that lignin modifications imposed by CAD deficiency in rice lead to notably disordered polysaccharide assembly in lignocellulose, which may account for improved saccharification performance of the CAD-deficient mutant (Martin et al., 2019). In this study, we extend our investigations to rice mutant lines deficient in CAldOMT. Accordingly, a set of rice mutants deficient in either or both CAldOMT and CAD in a same genetic background were generated in part using the Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/CRISPR-associated protein 9 (CRISPR/Cas9) system (Osakabe and Osakabe, 2015). These mutants were subjected to comparative, in-depth analyses of lignocellulose chemical and supramolecular structures and biomass saccharification performance. Lignin profiles of the rice mutants were obtained by solution-state two-dimensional (2D) 1H-13C heteronuclear single-quantum coherence (HSQC) nuclear magnetic resonance (NMR) and complementary wet-chemical methods. The profiles were then analyzed in conjunction with concurrent improvements in biomass saccharification efficiency and changes in lignocellulose supramolecular structures as revealed by solid-state 13C magic-angle-spinning (MAS) NMR spectroscopy, nuclear magnetic relaxation, wide-angle X-ray diffraction (WAXD), and Simon’s staining analyses. We highlight how differently CAldOMT- and CAD-associated lignin modifications impose “loosened” lignocellulose molecular assembly and thereby boost biomass deconstruction.

Results

OsCAldOMT1- and OsCAD2-deficient rice mutant lines

We previously demonstrated that OsCAldOMT1 (Koshiba et al., 2013a; Lam et al., 2019a) and OsCAD2 (Koshiba et al., 2013b; Martin et al., 2019), respectively, function as the major CAldOMT and CAD genes responsible for cell wall lignification in rice. In this study, we comparatively investigated the impacts of CAldOMT- and CAD-induced lignin modifications on rice cell wall structure and saccharification performance. To this end, we first prepared a series of Nipponbare rice (O. sativa L. spp. japonica cv. Nipponbare) mutant lines deficient in either or both OsCAldOMT1 and OsCAD2. The previously isolated cad2 Nipponbare rice mutant, which harbors a homozygous Tos17 retrotransposon insertion in OsCAD2 (Figure 1A) (Miyao et al., 2003; Koshiba et al., 2013a; Martin et al., 2019), was included as an OsCAD2 single-knockout line for testing in this study. In addition, OsCAldOMT1 single-knockout and OsCAldOMT1 and OsCAD2 double-knockout mutant lines of Nipponbare rice were developed here via CRISPR/Cas9-mediated targeted mutagenesis.

Figure 1.

Figure 1

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Generation and phenotypic characterization of OsCAldOMT1- and OsCAD2-knockout rice mutant lines. A, gene structures and mutations in the mutant lines. CRISPR/Cas9-induced mutation sites in OsCAldOMT1-knockout lines (caldomt1, cad2 caldomt1-a, and cad2 caldomt1-b) and Tos17-retrotransposon insertion sites in OsCAD2-knockout lines (cad2, cad2 caldomt1-a, and cad2 caldomt1-b) are shown. PAM sites (navy) and deletion or insertion mutation sites (blue) are indicated. B and C, growth characteristics (B) and morphological phenotype (C) of wild-type and homozygous mutant lines at the ripening stage. Bars in (C) = 10 cm. Values refer to mean ± standard deviation (sd) from individually analyzed plants (n = 5). Different letters indicate significant differences (ANOVA with post-hoc Tukey–Kramer’s test, P < 0.05). WT, wild-type control line; caldomt1, OsCAldOMT1 single-knockout line; cad2, OsCAD2 single-knockout line; cad2 caldomt1-a and cad2 caldomt1-b; OsCAldOMT1 and OsCAD2 double-knockout lines.

A single-guide RNA (sgRNA) targeting the first exon of OsCAldOMT1 (Figure 1A) with minimum off-target potential was designed using the CRISPR-P program (Lei et al., 2014; Liu et al., 2017). The CRISPR/Cas9 binary vector (Mikami et al., 2015) harboring the designed sgRNA sequence was then transformed into embryogenic calli derived from wild-type and cad2 Nipponbare mutant rice seeds to generate OsCAldOMT1 single-knockout and OsCAldOMT1 and OsCAD2 double-knockout mutant lines, respectively. Numerous T0 transformants containing various insertion–deletion mutations (indels) in the targeted OsCAldOMT1 site were identified based on genotyping by direct sequencing (Takeda et al., 2018, 2019). Consequently, we successfully isolated T1 individuals for one OsCAldOMT1 single-knockout line (caldomt1 containing a homozygous 1-nt insertion in the targeted site) and two OsCAldOMT1 and OsCAD2 double-knockout mutant lines (cad2 caldomt1-a and cad2 caldomt1-b, respectively, containing a homozygous 1-nt deletion and a 1-nt insertion in the targeted site) (Figure 1A). No off-target mutations had occurred in any of the selected T1 individuals in at least the five potential off-target sites (Supplemental Table S1). Multiple sequence alignment suggested that the CRISPR/Cas9-induced indels in caldomt1 and cad2 caldomt1-a/b lines caused frame-shift mutations; the resulting premature stop codons in turn lead to truncated OsCAldOMT1 mutant proteins that lacked conserved residues essential for CAldOMT catalytic activity (Zubieta et al., 2002; Kim et al., 2006) (Supplemental Figure S2). Loss-of-function mutations of OsCAldOMT1 were thus expected in these CRISPR/Cas9 mutant lines.

Fully genotyped caldomt1, cad2 caldomt1-a, and cad2 caldomt1-b mutant lines (T2 generation) were grown side-by-side with the previously isolated cad2 mutant (Miyao et al., 2003; Koshiba et al., 2013a; Martin et al., 2019) and wild-type control lines under controlled greenhouse conditions. These plants were used for phenotypic characterization and preparation of extractive-free cell wall residue (CWR) samples from dried culm tissues for further cell wall analyses. In line with our earlier observations (Koshiba et al., 2013a, 2013b; Martin et al., 2019; Lam et al., 2019a), the overall growth performance of OsCAldOMT1 and OsCAD2 single-knockout lines, caldomt1 and cad2, was similar to that of wild-type rice in terms of plant height, culm length, and biomass and culm CWR yields (Figure 1, B and C). The OsCAldOMT1 and OsCAD2 double-knockout lines, cad2 caldomt1-a and cad2 caldomt1-b, had significantly reduced plant heights and culm lengths. Nevertheless, their overall biomass and culm CWR yields were comparable to those of caldomt1, cad2, and the wild-type at least in the current greenhouse conditions, although there was a tendency for both of the two double-knockout lines to have lower biomass compared to others albeit with no statistical significance (Figure 1, B and C).

Chemical analyses of OsCAldOMT1 and OsCAD2 mutant rice cell walls

Cell wall chemotypes of the OsCAldOMT1- and OsCAD2-deficient rice mutants were first characterized by a series of chemical analyses. As determined by the Klason lignin assay, lignin content of caldomt1 cell walls was reduced by ca. 30% compared with that of the wild-type, consistent with our previous observations of other OsCAldOMT1-deficient rice lines (Koshiba et al., 2013a; Lam et al., 2019a), whereas the lignin content of cad2 cell walls was not significantly different compared with that of the wild-type control (Figure 2A). Similar to the caldomt1 single-knockout line, both cad2 caldomt1-a and cad2 caldomt1-b double-knockout lines had lignin contents that were significantly reduced, by ca. 30%, compared with the wild-type line (Figure 2A).

Figure 2.

Figure 2

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Lignin content and composition analyses of OsCAldOMT1 and OsCAD2-knockout rice mutant lines based on chemical analyses. A, lignin content determined by a Klason lignin assay. B–F, thioacidolysis-derived lignin composition data. Total yield of p-hydroxyphenyl (H)-, guaiacyl (G)-, and syringyl (S)-type monomers (B), H/G/S monomer composition (C), S/G monomer ratio (D), normalized GC/MS peak area of 5H-type monomers (E) and structures of H-, G-, S-, and 5H-type monomers (trimethylsilylated) released upon thioacidolysis (F) are shown. Values refer to mean ±  sd from individually analyzed plants (n = 3). Different letters indicate significant differences (ANOVA with post-hoc Tukey–Kramer’s test, P < 0.05).

To examine changes in lignin composition in OsCAldOMT1- and OsCAD2-deficient mutant cell walls, we carried out analytical thioacidolysis, which quantifies lignin-derived monomeric compounds released by the chemical cleavage of the major β–O–4 linkages in lignin polymers (Lapierre et al., 1986). When subjected to thioacidolytic degradation, both caldomt1 and cad2 single-knockout mutant cell walls released considerably fewer monomeric compounds, ca. 61% and 66% fewer, respectively, than did wild-type cell walls (Figure 2B). The large decrease in the yield of monomers from caldomt1 cell walls can be attributed to the reduced lignin content and also the replacement of the normal β–O–4 linkages in lignin polymers by the cyclic β–O–4/α–O–5 (benzodioxane) linkages, which are relatively tolerant to thioacidolysis, via incorporation of 5-hydroxyconiferyl alcohol (Lam et al., 2019a), whereas that from cad2 cell walls can be primarily due to the replacement of the normal β–O–4 linkages by the hydroxycinnamaldehyde-derived linkages (Martin et al., 2019). Monomer yields from cad2 caldomt1-a and cad2 caldomt1-b double-knockout mutant cell walls were even more drastically decreased—by ca. 84% and 81%, respectively, compared with wild-type cell walls (Figure 2B). This result can be attributed to the concomitant effects of reduced lignin content and changes in the types of β–O–4 linkages in lignin polymers produced by these CAldOMT and CAD double-knockout mutants.

Lignin compositional analysis based on proportional changes in thioacidolysis-derived S-, G-, and p-hydroxyphenyl (H)-type monomers (Figure 2F) revealed that S-type monomers were heavily depleted in the OsCAldOMT1-deficient mutant lines (caldomt1, cad2 caldomt1-a, and cad2 caldomt1-b). As a result, S/G monomer ratios were decreased considerably, by ca. 80%, compared with wild-type levels (Figure 2, C and D). In addition, cell walls from the three OsCAldOMT1-deficient mutant lines released significantly increased levels of 5H-type monomers upon thioacidolysis (Koshiba et al., 2013a) (Figure 2, E and F), thereby indicating the incorporation of noncanonical 5H lignin units in cell walls. These results are very consistent with the notion that OsCAldOMT1 is the major CAldOMT involved in S lignin biosynthesis in rice (Supplemental Figure S1) (Koshiba et al., 2013a; Lam et al., 2019a). In contrast, the S/G monomer ratio of cad2 mutant cell walls was increased by ca. 15% compared with wild-type levels (Figure 2, C and D). This result suggests that OsCAD2 is preferentially involved in the biosynthesis of G lignin relative to that of S lignin, a prediction based on our previous study (Martin et al., 2019).

Neutral sugar compositions in OsCAldOMT1- and OsCAD2-knockout mutant cell walls were generally similar to those in wild-type cell walls, except that we observed tendencies toward decreased hemicellulosic glucose and increased mannose contents in cad2 caldomt1-a and cad2 caldomt1-b cell walls (Supplemental Table S2). We also quantified cell wall-bound p-coumarates (pCAs) and ferulates (FAs) by measuring their corresponding free acids released by mild alkaline hydrolysis of CWRs (Yamamura et al., 2011). All OsCAldOMT1- and OsCAD2-knockout mutant cell walls had greatly decreased pCA levels, with reductions of ca. 60% compared with the wild-type in caldomt1 and cad2 cell walls and more drastic reductions of ca. 85% in cad2 caldomt1-a and cad2 caldomt1-b cell walls. In contrast, overall FA levels were similar to those of wild-type cell walls (Supplemental Table S2). Considering that most cell wall-associated pCAs and FAs are bound to lignins and hemicellulosic arabinoxylans, respectively, in cell walls of rice (Takeda et al., 2018, 2019) and generally also other grasses (Karlen et al., 2016, 2018), the observed variation (or lack thereof) in pCA and FA levels of OsCAldOMT1- and OsCAD2-knockout mutant cell walls can be at least partly attributed to the variation (or lack thereof) in their lignin and polysaccharide contents (Figure 2A; Supplemental Table S2).

2D HSQC NMR analysis of OsCAldOMT1 and OsCAD2 mutant lignins

To further investigate changes in the chemical structure of lignin in OsCAldOMT1- and OsCAD2-knockout mutant rice, we performed 2D HSQC NMR analysis on dioxane-soluble lignin samples prepared from CWRs via enzymatic removal of cell wall polysaccharides followed by extraction with a dioxane-water (96:4, v/v) solvent system for further lignin purification (Björkman, 1954; Tobimatsu et al., 2013; Martin et al., 2019). For the analysis of the OsCAldOMT1 and OsCAD2 double-knockout mutant lines, we focused on the analysis of cad2 caldomt1-a as the chemical analyses described above determined similar lignin characteristics between the two mutant line with different mutation types (cad2 caldomt1-a and cad2 caldomt1-b). The aromatic sub-regions of HSQC spectra of wild-type and mutant dioxane-soluble lignin samples displayed monolignol-derived lignin aromatic signals from G and S units (G and S, respectively) as well as signals from other typical components of grass lignins, such as pCA (pCA) and T (T) units (Figure 3A; Supplemental Table S3); we also detected minor signals from H lignin units. In addition, oxygenated-aliphatic and aldehyde sub-regions displayed signals from various inter-monomeric (I–III, I′, and I″) and end-unit linkage types (IV, IV′, and IV″) in rice lignin polymers (Figure 4, A and B; Supplemental Table S4). Volume integration analysis of these major lignin signals allowed us to estimate the shifts of lignin substructure distributions in OsCAldOMT1- and OsCAD2-knockout mutant cell walls (Figures 3, B and 4, C). The reported aromatic and inter-monomeric/end-unit linkage signal intensities were relative intensities normalized by the total of the major lignin aromatic unit (G2 + ½S2/6 + Gʹ2 + ½Sʹ2/6 = 100%) and inter-monomeric unit (Iα + IIα +½IIIα + Iʹα + I″9 = 100%) signals, respectively, reflecting the proportional amount of each substructure in the lignin polymer.

Figure 3.

Figure 3

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Lignin composition analysis of OsCAldOMT1- and OsCAD2-knockout rice mutant lines based on solution-state 2D NMR. A, aromatic sub-regions of HSQC NMR spectra of dioxane-soluble lignin samples from OsCAldOMT1- and OsCAD2-knockout rice mutant lines. Contour coloration matches the substructures shown. B, normalized signal intensity values and ratio of major lignin aromatic units. Data are expressed on a G2 + ½S2/6 + Gʹ2 + ½Sʹ2/6 = 1 basis. The dioxane-soluble lignin samples were prepared from culm cell wall residues pooled from three independent plants for each mutant and wild-type line. n.d., not detected.

Figure 4.

Figure 4

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Lignin inter-monomeric and end-unit linkage analysis of OsCAldOMT1- and OsCAD2-knockout rice mutant lines based on solution-state 2D NMR. A and B, oxygenated-aliphatic (A) and aldehyde (B) sub-regions of HSQC NMR spectra of dioxane-soluble lignin samples from OsCAldOMT1- and OsCAD2-knockout rice mutant lines. Contour coloration matches the substructures shown in each panel. C, normalized signal intensity values of major lignin inter-monomeric linkage types. Data are expressed on Iα + IIα + ½IIIα + Iʹα + Iʹʹ9 = 1 basis. The dioxane-soluble lignin samples were prepared from culm cell wall residues pooled from three independent plants for each mutant and wild-type line.

Our NMR data demonstrated that caldomt1 produced altered lignin polymers that were largely depleted of S and T units. This result is consistent with the proposed role of OsCAldOMT1 as a bifunctional OMT functioning in both S and T lignin biosynthetic pathways in rice (Supplemental Figure S1) (Lam et al., 2019a). According to volume integration of the HSQC signals, the decrease in the S/G signal ratio was ca. 96% in the caldomt1 lignin spectrum compared with the wild-type control, with the proportional decrease in the T signal likewise estimated as ca. 96% (Figure 3B); the S unit could be overestimated in the caldomt1 (and also cad2 caldomt1-a) lignin spectra because the residual S aromatic signals (S2/6) could be overlapped with the new 5H aromatic signals (5H2/6) (Vanholme et al., 2010; Lam et al., 2019a). The incorporation of noncanonical 5H lignin units in caldomt1 as more clearly demonstrated above by thioacidolysis (Figure 2) was further corroborated by the appearance of HSQC signals from cyclic β–O–4/α–O–5 (benzodioxane) units (I′) (Figure 4A), which could have been derived from the incorporation of catechol-type aromatic units, such as 5H and catechyl (C) units, in lignin polymers (Ralph et al., 2001; Chen et al., 2012). According to volume integration estimates, I′ signals accounted for ca. 22% of the total inter-monomeric linkage-type signals detected in the caldomt1 lignin spectrum (Figure 4C). Further, cad2 produced lignins incorporating noncanonical hydroxycinnamaldehyde-derived G′ and S′ aromatic units. G′ and S′ signals accounted for ca. 13% and ca. 6%, respectively, of the major lignin aromatic signals in the cad2 lignin spectrum compared with ca. 4% and none (not detected), respectively, in the wild-type control (Figure 3B). In addition, the aldehyde C9–H9 signals from hydroxycinnamaldehyde-derived 8–O–4 units (I″, ca. 12% of detected inter-monomeric linkage types), along with those from cinnamaldehyde (IV′, ca. 6%) and benzaldehyde (IV″, ca. 8%) end units, were clearly visible in the cad2 spectrum but were much smaller or practically absent in wild-type control spectra (Figure 4, B and C). This result provides further evidence of the incorporation of hydroxycinnamaldehyde precursors into lignin through radical coupling (Kim et al., 2000).

The OsCAldOMT1 and OsCAD2 double-knockout mutant lines produced lignins displaying a combination of the structural features of the two modified lignins produced by the OsCAldOMT1 and OsCAD2 single-knockout lines. Similar to the lignin produced by caldomt1, lignin from cad2 caldomt1-a was largely depleted of both S (S, ca. 24% of wild-type levels) and T (T, ca. 36% of wild-type levels) aromatic units (Figure 3) and incorporated at least some atypical 5H units bearing benzodioxane linkages (I′, ca. 19% of detected inter-monomeric linkage signals) (Figure 4). Reflecting structural features of the lignin produced by cad2, they also contained substantially augmented hydroxycinnamaldehyde-derived G′ and S′ aromatic units (G′ and S′, ca. 20% and ca. 8%, respectively, of detected aromatic unit signals) (Figure 3B) and 8–O–4 units (I″, ca. 12% of detected inter-monomeric linkage signals) (Figure 4C), and cinnamaldehyde (IV′, ca. 8% of detected inter-monomeric linkage signals) and benzaldehyde (IV″, ca. 9% of detected inter-monomeric linkage signals) end-units (Figure 4C).

Enzymatic saccharification efficiency of OsCAldOMT1 and OsCAD2 mutant cell walls

Given that both CAldOMT and CAD are potent lignin bioengineering targets for improving cell wall deconstruction properties, we comparatively evaluated the enzymatic saccharification efficiency of our OsCAldOMT1- and OsCAD2-knockout mutant cell walls. As anticipated from previous research (Koshiba et al., 2013a, 2013b; Martin et al., 2019; Lam et al., 2019a), both caldomt1 and cad2 single-knockout lines had substantially enhanced saccharification efficiencies compared with the wild-type control. Under identical saccharification conditions, caldomt1 clearly surpassed cad2 in terms of saccharification efficiency. After 48 h of incubation with a cocktail of cellulases, the yield of released glucose (per CWR and per glucan content, respectively) from caldomt1 cell walls was 67% and 68% higher than the wild-type control, whereas cad2 cell walls had a released glucose yield that was 23% and 34% higher than the control (Figure 5). The glucose release profiles of cad2 caldomt1-a and cad2 caldomt1-b cell walls were very similar to each other: their saccharification efficiencies appeared to be higher than those of wild-type and cad2 mutant cell walls but were similar or lower than that of caldomt1 mutant cell walls. Thus, our data suggested that single OsCAldOMT1 mutation enhanced cell wall digestibility better than single OsCAD2 mutation and even better than the effect of two simultaneous mutations of OsCAldOMT1 and OsCAD2.

Figure 5.

Figure 5

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Enzymatic saccharification efficiency of culm cell walls from OsCAldOMT1- and OsCAD2-knockout rice mutant lines. Data are expressed as glucose yield per cell wall residue, CWR (upper) and glucose yield per total glucan (lower). Values refer to mean ±  sd from individually analyzed plants (n = 3). Different letters indicate significant differences (ANOVA with post-hoc Tukey–Kramer’s test, P < 0.05).

Cellulose assembly analyses based on solid-state NMR and WAXD

The variation in saccharification performance of the current OsCAldOMT1 and OsCAD2 mutant set suggested differently altered lignocellulose supramolecular structures in their cell wall samples. To further investigate this aspect, we performed a comparative analysis of polysaccharide molecular alignment (more specifically, cellulose crystallinity) and mobility in the mutant cell wall samples. The comparison was based on the solid-state NMR (13C MAS spectra and relaxation time) and WAXD patterns of culm CWR samples prepared from caldomt1, cad2, and cad2 caldomt1-a mutant lines along with the wild-type control. For the characterizations of the OsCAldOMT1 and OsCAD2 double-knockout mutant lines using these solid-state NMR and WAXD analyses as well as the Simon’s staining assay described below, only cad2 caldomt1-a out of the two mutant lines (cad2 caldomt1-a and cad2 caldomt1-b), was used because of the similarities in cell wall chemical structure and saccharification performance of the two mutant lines.

The 13C MAS NMR spectra were collected using 1H–13C cross-polarization (CP) for the initial 13C magnetization to preferentially detect relatively rigid polymer components, mainly cellulose, over other relatively mobile components such as lignin and hemicelluloses (Wang et al., 2014; Dupree et al., 2015; Simmons et al., 2016; Kang et al., 2019; Martin et al., 2019). As expected, all CP-MAS spectra of rice cell wall samples were dominated by cellulose signals from glucose residues in two distinct cellulose environments: “crystalline and/or internal” and “amorphous and/or surface” cellulose domains (Supplemental Table S5; Supplemental Figure S3). Volume integration analysis of cellulose C4 signals, that is, C4a at 89 ppm from crystalline/internal cellulose and C4b at 84 ppm from amorphous/surface cellulose (Simmons et al., 2016; Martin et al., 2019), revealed that the relative abundance of crystalline/internal over amorphous/surface cellulose domains was notably decreased, by ca. 13%–19% compared with the wild-type, in all OsCAldOMT1 and OsCAD2 mutant cell wall samples (Figure 6A). Consistent with this observation, the relative crystallinity index, determined from WAXD profiles based on the ratio of crystalline and amorphous scattering (Supplemental Figure S4), similarly decreased, by ca. 7%–12%, in all OsCAldOMT1 and OsCAD2 mutant cell walls compared with the wild-type (Figure 6B). These data suggest that the crystalline network of cellulose was disrupted in all the tested OsCAldOMT1 and OsCAD2 mutant cell wall samples.

Figure 6.

Figure 6

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Polysaccharide assembly and mobility analyses of culm cell walls from OsCAldOMT1- and OsCAD2-knockout rice mutant lines. A, expanded 13C CP–MAS NMR spectra showing crystalline/internal (C4a) and amorphous/surface (C4b) cellulose C4 signals. Inserted values are volume integrals for indicated peak areas (C4a + C4b = 100). B, apparent cellulose crystallinity index based on WAXD. Values refer to mean ±  sd from individually analyzed plants (n = 3). Different letters indicate significant differences (ANOVA with post-hoc Tukey–Kramer’s test, P < 0.05). C, CP 13C spin-lattice relaxation time (T1) data for major cellulose carbon sites. Delay time-dependent signal decay data were fitted using a double exponential function to determine two independent T1 for slower and faster relaxing components (see Supplemental Methods). T1 values (left) and fraction (slower + faster-relaxing components = 100%) data (right) for slower-relaxing components are shown. All data including T1 and fraction data for faster-relaxing components are listed in Supplemental Table S6. The error bars indicate sds of the fitting coefficients.

We also assessed the molecular mobility of cellulose in mutant and wild-type cell walls based on site-specific 13C T1 data which are sensitive to nanosecond timescale motions of the cell wall components (García et al., 2011; Wang et al., 2014; Kang et al., 2019). Because we used Torchia pulse sequences using CP for the 13C magnetization (Torchia, 1978), the collected T1 data mainly reflect the molecular motions of CP-enhanced rigid cellulose components (Martin et al., 2019). Similar to the Torchia-CP relaxation profiles of powdery biomass and cellulose samples reported in other studies (Horii et al., 1984; Focher et al., 2001; Zuckerstätter et al., 2013; Ghosh et al., 2019; Martin et al., 2019), the obtained relaxation decay curves were well fitted by a double exponential function with two distinctively slow- and fast-relaxing components, each representing relatively rigid and mobile components, respectively, both in crystalline/internal and amorphous/surface cellulose domains (Figure 6C; Supplemental Table S6). Consistent with the data obtained in our previous study (Martin et al., 2019), cellulose T1 values in cad2 cell walls were notably increased relative to the wild-type at most cellulose carbon sites, whereas the fractional weighing of the slower-relaxing components was generally similar or slightly lower than that in the wild-type control (Figure 6C; Supplemental Table S6). In contrast, cellulose T1 was markedly decreased at all cellulose carbon sites in both caldomt1 and cad2 caldomt1-a cell walls (Figure 6C; Supplemental Table S6). Although the fractional weighing of faster- and slower-relaxing components in the caldomt1 single-knockout mutant cell wall was generally similar to that in the wild-type control, the fractional weighing of the slower-relaxing components relative to that of the faster-relaxing components were notably decreased at most cellulose carbon sites (except at C6b) in the cad2 caldomt1-a double-knockout mutant cell walls compared with the wild-type and caldomt1 mutant cell walls. Collectively, our data suggest that not only the crystallinity but also the molecular mobility of cellulose were notably affected in cell wall samples of the OsCAldOMT1 and OsCAD2 rice mutants. Apparently, disruption of OsCAldOMT1 increased cellulose mobility (lower T1) and stacking OsCAldOMT1 and OsCAD2 mutations further increased cellulose mobility (lower T1 and increased proportion of faster- over slow-relaxing components), whereas disruption of OsCAD2 alone contributed to decreased cellulose mobility (higher T1).

Cellulose surface accessibility assessed by Simon’s staining assay

Finally, we assessed the relative porosity and accessibility of the cellulose surface in OsCAldOMT1 and OsCAD2 mutant and wild-type cell wall samples by performing Simon’s staining assay. In this technique, pore surface area amplitudes and cellulose substrate accessibility in biomass are semi-quantitatively determined according to the adsorption of two cellulose binding dyes: direct orange (DO) and direct blue (DB). In particular, DO better resembles the molecular size of typical cellulases compared with DB; the ratio of adsorbed DO to DB dyes has therefore been used to evaluate the physical accessibility of cellulolytic enzymes to cellulose substrate (Chandra et al., 2008). As shown in Figure 7, cellulose dye adsorptions were overall similar between the wild-type and tested OsCAldOMT1 and OsCAD2 mutant lines. The DO adsorption of caldomt1 was slightly higher than that of the other genotypes (Figure 7A), and a higher DO/DB ratio, which was not statistically different from that of the wild-type, was detected in caldomt1 compared with cad2 and cad2 caldomt1-a (Figure 7B). No statistical difference was observed in DO adsorption or the DO/DB ratio among wild-type, cad2 and cad2 caldomt1-a lines. Given the amplitude of data variations using this assay reported in the literature (Chandra et al., 2008; Arantes and Saddler, 2011; Biswal et al., 2018; Wang et al., 2018), however, we interpret these data as indicating that no drastic changes occurred in the porosity and physical accessibility of cellulose in the cell walls of the OsCAldOMT1 and OsCAD2 mutants.

Figure 7.

Figure 7

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Simon’s staining assay of culm cell walls from OsCAldOMT1- and OsCAD2-knockout rice mutant lines. Amounts of absorbed DO and DB dyes (A) and ratio of absorbed DO to DB dyes (B) are shown. Values are means ±  sd from individually analyzed plants (n = 3). Different letters indicate significant differences (ANOVA with post-hoc Tukey–Kramer’s test, P < 0.05).

Discussion

Lignin content and structure are the two major factors contributing to biomass recalcitrance in polysaccharide-oriented biomass utilizations, possibly by affecting the lignocellulose supramolecular assembly (Li et al., 2016). Previous efforts to control these parameters by manipulating CAldOMT and CAD gene expressions have improved the deconstruction properties of cell walls in many plant species to varying extents. Nevertheless, much remains to be learned about how changes in lignin chemistry affect supramolecular assembly and consequently the deconstruction properties of lignocellulose. In this study, we comparatively analyzed a series of CAldOMT- and CAD-deficient rice mutant lines with respect to their altered lignin chemistry, lignocellulose supramolecular structures, and biomass digestibility. Our goal was to provide further insights into the contribution of lignin to lignocellulose properties and to inform the development of approaches to overcome biomass recalcitrance.

This study further corroborates the notion that manipulations of CAldOMT and CAD gene expressions are effective for the modification of lignin composition and functionality in grass cell walls (Figures 2–4). The single-knockout rice mutant deficient in CAldOMT, caldomt1, produced altered lignin that was largely depleted of canonical S and T units and incorporated at least some noncanonical 5H units derived from the polymerization of 5-hydroxyconiferyl alcohol (Supplemental Figure S1). The mutant deficient in CAD, cad2, generated another type of lignin incorporating noncanonical G′ and S′ units derived from the polymerization of hydroxycinnamaldehydes (Supplemental Figure S1). Our data, which are consistent with our previously obtained results for other CAldOMT- and CAD-deficient rice transgenic plants (Koshiba et al., 2013a, 2013b; Martin et al., 2019; Lam et al., 2019a), further establish the proposed functions of the two enzyme genes in the grass lignin biosynthetic pathways. The double-knockout rice mutants deficient in both CAldOMT and CAD, cad2 caldomt1-a and cad2 caldomt1-b, produced even more drastically altered lignins that displayed a combination of the structural features of the two modified lignins produced by caldomt1 and cad2 (Figures 2–4). The two lignin modification strategies targeting CAldOMT and CAD can thus be combined for simultaneous introduction of the two unique lignin features.

In line with our previous research (Koshiba et al., 2013a, 2013b; Martin et al., 2019; Lam et al., 2019a), the caldomt1 and cad2 single-knockout lines both had improved cell wall saccharification efficiencies compared with the wild-type control (Figure 5). We also determined that caldomt1 surpasses cad2 with respect to cell wall saccharification performance under identical testing conditions (Figure 5). Given that the saccharification efficiency of cad2, although less increased than that of caldomt1, was substantially higher than that of the wild-type, cad2 caldomt1 double-knockout lines would be expected to be synergistically improved or to at least have a saccharification performance similar to the corresponding caldomt1 single-knockout lines. However, although the saccharification efficiencies of cad2 caldomt1-a and cad2 caldomt1-b double-knockout mutant cell walls were improved compared with the performance of the wild-type control cell walls, the efficiencies appeared to be intermediate between those of the caldomt1 and cad2 single-knockout mutant cell walls (Figure 5). This result suggests that stacking the downregulation of CAD in the CAldOMT-downregulated background had little or no synergistic effect at least within the current rice mutant set under the tested conditions (Figure 5).

According to our solid-state NMR and WAXD data, the CAldOMT- and CAD-deficient rice mutant cell walls both had notably altered lignocellulose assemblies compared to that in the wild-type cell walls (Figure 6), which may, at least in part, account for their improved saccharification performances. In particular, the depletion of the apparent cellulose crystallinity—as gauged by the decreased peak areas of crystalline/internal carbons relative to amorphous/surface carbons of cellulose in the 13C CP–MAS spectra (Figure 6A) and the decreased ratio of crystalline to amorphous scattering in the WAXD profiles (Figure 6B)—can improve saccharification performance of biomass. In fact, several earlier studies detected negative associations between cellulose crystallinity and biomass susceptibility to enzymatic saccharification (Laureano-Perez et al., 2005; Hall et al., 2010; Vandenbrink et al., 2012; Marriott et al., 2016; Martin et al., 2019). In addition, our cellulose T1 analysis suggested that the cellulose mobility of cell wall samples from CAldOMT-deficient caldomt1 and cad2 caldomt1-a mutants was notably increased over those observed in the corresponding wild-type and cad2 mutant cell wall samples (Figure 6; Supplemental Table S6). The disruption of CAldOMT may therefore contribute to a more “loosened” lignocellulose assembly, and thereby a more enhanced susceptibility of cellulose to enzymatic saccharification than does the disruption of CAD. Nevertheless, despite our cellulose T1 analysis determined that cad2 caldomt1-a cell walls had an apparently more increased cellulose mobility, that is, lower T1 values and decreased proportions of slower- over faster-relaxing components (Figure 6; Supplemental Table S6), their saccharification performance did not surpass that of caldomt1 single-knockout mutant cell walls; it is currently unclear why stacking OsCAD2 mutation on OsCAldOMT1 mutation background contributed to increased cellulose mobility (in cad2 caldomt1-a), whereas OsCAD2 mutation alone on wild-type background conversely contributed to decreased cellulose mobility (in cad2) (Figure 5). Overall, neither our current lignocellulose supramolecular structural data based on solid-state NMR and X-ray approaches (Figure 6) nor the cellulose surface accessibility data from Simon’s staining (Figure 7) provide a clear explanation for why the mutation of CAD in the CAldOMT-mutated (but not in wild-type) background had little positive impact on the enzymatic saccharification of rice biomass. Further studies are needed to clarify this issue.

Our data support the notion that genetic manipulation of lignin content and structure can substantially affect the organization of lignocellulose polymers in cell walls (Ruel et al., 2009; Carmona et al., 2015; Liu et al., 2016; Martin et al., 2019). In particular, reduced lignin contents, as are prominent in the CAldOMT-deficient lines (Figure 2, A and B), may be one of the major factors causing the observed disintegration of lignocellulose in rice mutants tested in this study. In line with this idea, previous cell wall structural studies using X-ray diffraction approaches also detected considerable decreases in the proportion of orientated cellulose microfibrils in cell walls from Arabidopsis mutants having reduced lignin contents with little or minor changes in lignin composition (Ruel et al., 2009; Liu et al., 2016). In addition to the effect of reduced lignin content, lignocellulose assembly might be affected by altered lignin chemical structures. Given that lignin and cellulose may interact with each other indirectly via hemicelluloses through independent lignin–hemicellulose and cellulose–hemicellulose associations (Simmons et al., 2016; Kang et al., 2019; Terrett and Dupree, 2019; Kirui et al., 2022), some types of changes in lignin composition and functionality may lead to a looser lignocellulose assembly primarily by disrupting lignin–hemicellulose associations in lignocellulose. This situation may apply to the disrupted cellulose assembly observed in the cell wall samples from our CAD-deficient rice mutants as well as the CAD-deficient Arabidopsis cadc cadd mutants reported earlier by Liu et al. (2016). Indeed, a previous study using computational molecular modeling has suggested that CAD-associated structural modifications of lignin polymers, for example, augmentations of aldehyde functionalities, may reduce noncovalent lignin–hemicellulose interactions in cell walls (Carmona et al., 2015). In addition, it has been proposed that introducing hydroxycinnamaldehydes (by disrupting CAD) or catechol-type monolignols such as 5-hydroxyconiferyl alcohol (by disrupting CAldOMT) in lignin polymerization may avert the formations of covalent lignin–polysaccharide cross-linking by trapping quinone methide intermediates which can react with hemicelluloses to form covalent lignin–polysaccharide linkages during the β–O–4 coupling in lignin polymerization (Ralph, 2006; Tobimatsu et al., 2012; Vanholme et al., 2012; Anderson et al., 2015; Martin et al., 2019).

In conclusion, we have provided a molecular basis for understanding the relationship between altered lignin chemistry and changes in the supramolecular structure and deconstruction properties of lignocellulose in CAldOMT- and CAD-deficient plants. Both CAldOMT and CAD hold promise as potent bioengineering targets for improving biomass conversions in biochemical productions. Although this study failed to detect any synergistic effect due to simultaneous disruption of CAldOMT and CAD on rice biomass saccharification at least within the current testing conditions, it is still tempting to test whether the CAldOMT and CAD double-knockout mutant lines are improved over their corresponding single-knockout mutants in regards to other biorefinery contexts; for example, in biomass saccharification combined with various chemical pretreatments and/or in lignin valorization chemistries for production of useful aromatic chemicals. Future studies may also extend comparative analyses of chemical and supramolecular structures of lignocellulose to other lignin mutant and transgenic plants—ideally those with more systematically altered lignin contents and structures, such as plants with altered lignin composition but with constant lignin content and vice versa, to further investigate independent effects of lignin content and structure on the lignocellulose supramolecular structure. It is also important to further investigate the growth characteristics and biomass productivity of the developed rice mutants in field conditions and/or under various abiotic and biotic stresses for their potential biorefinery applications. Such studies should promote our understanding of the still-elusive architectures and functions of plant cell walls and ultimately increase our ability to manipulate these characteristics for better biomass utilization.

Materials and methods

Plant materials

The cad2 (gh2; accession: NE4246) rice (O. sativa L.) mutant was originally identified from a Tos17-insertional mutant population derived from O. sativa L. spp. japonica cv. Nipponbare (Miyao et al., 2003; Koshiba et al., 2013a). The caldomt1, cad2 caldomt1-a, and cad2 caldomt1-b mutant lines were generated via CRISPR/Cas9-mediated targeted mutagenesis (Mikami et al., 2015) as described in Supplemental Methods. Regenerated plants and isolated T1 mutant lines were grown under growth chamber conditions as described previously (Lam et al., 2017). Oligonucleotides used for vector construction, genotyping, and off-target analysis are listed in Supplemental Table S7. Fully genotyped caldomt1, cad2 caldomt1-a, and cad2 caldomt1-b T2 mutant lines were grown side-by-side with cad2 and wild-type rice in a greenhouse (Takeda et al., 2017) and subjected to phenotypic characterization and cell wall analyses. Rice culm CWR samples used for cell wall chemical and supramolecular structural analyses and enzymatic saccharification assay were prepared as described in Supplemental Methods in Supporting Information.

Cell wall chemical and supramolecular structural analyses

Klason lignin assay (Hatfield et al., 1994), sugar analysis (Lam et al., 2017), quantitation of alkaline-releasable cell-wall-bound pCAs and FAs (Yamamura et al., 2011), and the modified Simon’s staining assay (Chandra et al., 2008; Martin et al., 2019) were performed according to the methods described in literature. Analytical thioacidolysis was performed according to the method described previously (Yamamura et al., 2012), and the released lignin monomers were derivatized with N,O-bis(trimethylsilyl)acetamide and quantified by gas chromatography/mass spectrometry (GC/MS) using 4,4′-ethylenebisphenol as an internal standard (Yue et al., 2012). The GC/MS detection of the 5H-lignin-derived monomeric compounds was based on MS detection of their characteristic base ion peaks (m/z, 357) (Koshiba et al., 2013a) and the peak intensity data reported in Figure 2E are normalized intensities relative to the base ion peak of the internal standard (4,4′-ethylenebisphenol, m/z, 343) (Yue et al., 2012). Solution-state 2D HSQC NMR, solid-state 13C CP–MAS NMR and WAXD measurements were performed as described in Supplemental Methods. Peak assignments for solution-state HSQC spectra (Supplemental Tables S3 and S4) (Kim and Ralph, 2010; Mansfield et al., 2012; Tarmadi et al., 2018; Martin et al., 2019; Lam et al., 2019a) and solid-state CP–MAS spectra (Supplemental Table S5) (Simmons et al., 2016; Martin et al., 2019) were based on comparison with literature data.

Enzymatic saccharification assay

Enzymatic saccharification efficiency was determined as described previously (Hattori et al., 2012; Martin et al., 2019). Briefly, destarched CWRs were digested using an enzyme cocktail composed of Celluclast 1.5 L (Novozymes, Bagsvaerd, Denmark) (1.1 FPU), Novozyme 188 (Novozymes) (2.5 CbU), and Ultraflo L (Novozymes) (65 μg) in 50-mM sodium citrate buffer (pH 4.8) at 50°C for 1, 6, 12 and 48 h. Released glucose was measured using a Glucose CII test kit (Wako Pure Chemicals, Osaka, Japan). Total glucan content for calculation of glucose yield per glucan was independently determined by complete hydrolysis of destarched CWRs with 72% (w/w) sulfuric acid (Hattori et al., 2012).

Statistical analysis

One-way analysis of variance (ANOVA) with a post-hoc Tukey–Kramer’s test (P < 0.05) was performed using R statistical software (R Core Team, 2019) with the agricolae package (de Mendiburu, 2019).

Accession numbers

Sequence data from this study can be found in the GenBank/EMBL data libraries under accession numbers AK064768 (OsCAldOMT1; LOC_Os08g06100) and AK105011 (OsCAD2; LOC_Os02g09490).

Supplemental data

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

Supplemental Methods. Additional experimental procedures.

Supplemental Table S1. Potential off-target sites in OsCAldOMT1-knockout lines.

Supplemental Table S2. Polysaccharide and cinnamate analyses of rice cell walls.

Supplemental Table S3. Aromatic peak assignments in 2D NMR spectra.

Supplemental Table S4. Aliphatic and aldehyde peak assignments in 2D NMR spectra.

Supplemental Table S5. Peak assignments in 13C CP–MAS NMR spectra.

Supplemental Table S6. 13C spin-lattice relaxation time data of rice cell walls.

Supplemental Table S7. Primers used in this study.

Supplemental Figure S1. Proposed lignin biosynthetic pathways in grasses.

Supplemental Figure S2. Predicted effects of CRISPR/Cas9-mediated mutations.

Supplemental Figure S3. 13C CP–MAS NMR spectra of rice cell walls.

Supplemental Figure S4. WAXD profiles of rice cell walls.

Supplementary Material

kiac432_Supplementary_Data
Click here for additional data file. (1.2MB, pdf)

Acknowledgments

We thank Dr. Hirohiko Hirochika and Dr. Akio Miyao of the National Agriculture and Food Research Organization (NARO) for providing cad2 mutant rice seeds, and Dr. Seiichi Toki, Dr. Masaki Endo, and Mr. Masafumi Mikami of NARO for providing pZH_OsU6gRNA_MMCas9 vector. We also thank Dr. Hironori Kaji, Ms. Ayaka Maeno, and Ms. Kyoko Yamada for their assistance and helpful suggestions in NMR analysis, and Dr. Tomoya Imai for his helpful suggestions in X-ray analysis. A part of this study was conducted using the DASH/FBAS facilities at RISH, Kyoto University, and the NMR spectrometer in the JURC at ICR, Kyoto University.

Funding

This work was supported in part by grants from the Japan Science and Technology Agency/Japan International Cooperation Agency (Science and Technology Research Partnership for Sustainable Development, SATREPS), the Japan Society for the Promotion of Science (JSPS, grant nos. #JP16K14958, #JP16H06198, #JP17F17103, #JP17K05882, and #JP20H03044) and RISH Kyoto University (Mission-linked Research Funding for Mission 2 and Mission 5-2). A.F.M. acknowledges the support of the RISET-Pro PhD fellowship program of the Ministry of Research, Technology, and Higher Education of Republic of Indonesia (World Bank Loan no. 8245-ID).

Conflict of interest statement. None declared.

Contributor Information

Andri Fadillah Martin, Research Institute for Sustainable Humanosphere, Kyoto University, Gokasho, Uji 611-0011, Japan; Research Center for Genetic Engineering, National Research and Innovation Agency (BRIN), Bogor, 16911, Indonesia.

Yuki Tobimatsu, Research Institute for Sustainable Humanosphere, Kyoto University, Gokasho, Uji 611-0011, Japan.

Pui Ying Lam, Research Institute for Sustainable Humanosphere, Kyoto University, Gokasho, Uji 611-0011, Japan; Center for Crossover Education, Graduate School of Engineering Science, Akita University, Akita, 010-8502, Japan.

Naoyuki Matsumoto, Research Institute for Sustainable Humanosphere, Kyoto University, Gokasho, Uji 611-0011, Japan.

Takuto Tanaka, Research Institute for Sustainable Humanosphere, Kyoto University, Gokasho, Uji 611-0011, Japan.

Shiro Suzuki, Research Institute for Sustainable Humanosphere, Kyoto University, Gokasho, Uji 611-0011, Japan; Faculty of Applied Biological Sciences, Gifu University, Gifu, 501-1193, Japan.

Ryosuke Kusumi, Graduate School of Agriculture, Kyoto University, Kyoto, 606-8502, Japan.

Takuji Miyamoto, Research Institute for Sustainable Humanosphere, Kyoto University, Gokasho, Uji 611-0011, Japan; Sakeology Center, Niigata University, Niigata, 950-2181, Japan.

Yuri Takeda-Kimura, Research Institute for Sustainable Humanosphere, Kyoto University, Gokasho, Uji 611-0011, Japan; Department of Botany, University of Wisconsin-Madison, Madison, Wisconsin, 53706, USA.

Masaomi Yamamura, Research Institute for Sustainable Humanosphere, Kyoto University, Gokasho, Uji 611-0011, Japan; Faculty of Bioscience and Bioindustry, Tokushima University, Tokushima, 770-8503, Japan.

Taichi Koshiba, Research Institute for Sustainable Humanosphere, Kyoto University, Gokasho, Uji 611-0011, Japan; National Agriculture and Food Research Organization, Tsukuba, 305-8517, Japan.

Keishi Osakabe, Faculty of Bioscience and Bioindustry, Tokushima University, Tokushima, 770-8503, Japan.

Yuriko Osakabe, School of Life Science and Technology, Tokyo Institute of Technology, Tokyo, 152-8550, Japan.

Masahiro Sakamoto, Graduate School of Agriculture, Kyoto University, Kyoto, 606-8502, Japan.

Toshiaki Umezawa, Research Institute for Sustainable Humanosphere, Kyoto University, Gokasho, Uji 611-0011, Japan; Research Unit for Realization of Sustainable Society (RURSS), Kyoto University, Uji, 611-0011, Japan.

 

A.F.M., Y.T., and T.U. conceived the research. Y.T., S.S., R.K., K.O., Y.O., M.S., and T.U. designed the experiments. A.F.M., Y.T., P.Y.L., N.M., T.T., S.S., T.M., Y.T.K., M.Y., and T.K. performed the experiments and analyzed the data. A.F.M., Y.T., and T.U. wrote the manuscript with help from all the others. A.F.M. and Y.T. contributed equally to this work.

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 Yuki Tobimatsu (ytobimatsu@rish.kyoto-u.ac.jp).

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