Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

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Chapter 3

TRICIN IN GRASS LIGNIN: BIOSYNTHESIS, CHARACTERIZATION, AND QUANTITATION

Wu Lan*, Jorge Rencoret, José Carlos del Río and John Ralph Chemistry and Chemical Engineering, École polytechnique fédérale de Lausanne, Switzerland Instituto de Recursos Naturales y Agrobiología de Sevilla, Consejo Superior de Investigaciones Científicas, Seville, Spain Department of Biochemistry, and the Department of Energy’s Great Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin-Madison, Madison, WI, US

ABSTRACT

Tricin [5,7-dihydroxy-2-(4-hydroxy-3,5-dimethoxyphenyl)-4H-chromen-4-one] is a member of the flavonoid family with significant biological roles in plant tissues. Even though tricin has been extensively studied as a flavonoid, the presence of tricin in the lignin polymer was only recently discovered. Differently from the other monomers, tricin is derived from a combination of the shikimate and polyketide biosynthetic pathways, and increasingly attracts attention from researchers. This chapter briefly introduces the occurrence of tricin in plants and the relevant biosynthetic pathway, discusses the identification and characterization of tricin that is incorporated into the lignin polymer, methods for its quantitation, as well as the implications of the tricin-lignin structure.

Keywords: flavonoid, flavonolignin, biosynthesis, NMR, HSQC, HMBC

* Corresponding Author Email: [email protected]. 52 Wu Lan, Jorge Rencoret, José Carlos del Río et al.

INTRODUCTION

Lignin is a complex phenylpropanoid polymer composed primarily of p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) units derived from the monolignols p-coumaryl, coniferyl, and sinapyl alcohols, respectively [1, 2]. But as our understanding of the biosynthesis and structure of lignin has escalated in the recent decade, several new lignin monomers have been discovered in wild-type and transgenic plants, such as monolignol acetate, p-hydroxybenzoate, p-coumarate, and ferulate ester conjugates that are now recognized as authentic lignin monomers [2-4], as well as components of incomplete monolignol biosynthesis such as caffeyl alcohol [5, 6], and hydroxycinnamaldehydes [7]. Recently, a special compound, tricin, was identified in the isolated lignin fraction from wheat straw [8]. Tricin [5,7-dihydroxy-2-(4-hydroxy-3,5-dimethoxyphenyl)-4H-chromen-4-one], containing benzoyl, cinnamoyl, and heterocyclic structures in its backbone, is a member of the flavone family. It has long been studied as an extractable compound (but was not known previously to be in the lignin polymer). As a secondary metabolite in cell walls, tricin is widely distributed in the leaves and stems of herbaceous and cereal plants including but not limited to bamboo, sugarcane, wheat, oat, and maize, and can be extracted from these plant tissues by organic solvents (methanol, ethanol, acetonitrile, methylene chloride) under ambient conditions or with heating [9-16].

TRICIN AS A SECONDARY METABOLITE

Biosynthesis of Tricin

Biosynthesis of flavones has been extensively studied, but the biosynthetic pathway of tricin remains mildly controversial. It is well established that tricin is derived from a combination of the shikimate and acetate/malonate-derived polyketide biosynthetic pathways [17] (Figure 1). The first committed step is catalyzed by chalcone synthase (CHS) using p-coumaroyl-CoA, a key intermediate on the general phenylpropanoid pathway, and malonyl-CoA (via the fatty acid pathway) as substrates. The generated naringenin chalcone is further converted to naringenin, a compound in the flavanone category, by chalcone isomerase (CHI). By dehydrogenation of the heterocyclic ring with the enzyme flavone synthase (FNSII), the flavanone is converted to a flavone, apigenin [17, 18]. Flavonoid 3',5'-hydroxylases (F3',5'H) were believed to generate tricetin from apigenin for 3',5'-O-methylation by an O-methyltransferase (FOMT) to form tricin [19-22]. However, another study suggested that chrysoeriol (a 3'-methoxylated flavone), instead of tricetin, was the intermediate for tricin synthesis in rice. Hence the biosynthetic pathway Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation 53 leading to tricin should be reconstructed as being naringenin → apigenin → luteolin → chrysoeriol → selgin → tricin [23].

Figure 1. Proposed tricin biosynthetic pathway. The gray color indicates the pathway previously proposed [20,21], but that has been challenged recently [23]. PAL, pheammonialyase; C4H, cinnamate 4-hydroxylase; 4CL, 4-coumarate CoA ligase; HCT, p-hydroxycinnamoyl-CoA: quinate/shikimate p-hydroxycinnamoyltransferase; C3'H, p-coumaroyl ester 3-hydroxylase; CSE, caffeoyl shikimate esterase; CHS, chalcone synthase; CHI, chalcone isomerase; FNSII, flavone synthase II; F3'H, flavonoid 3'-hydroxylase; F5'H, flavonoid 5'-hydroxylase; FOMT, flavonoid O-methyltransferase; C5'H, chrysoeriol 5'-hydroxylase [18,24]. 54 Wu Lan, Jorge Rencoret, José Carlos del Río et al.

Tricin and Its Derivatives

Tricin was first isolated in 1930 from the rust-resistant wheat (Triticum dicoccum) leaf [25]. In a later study, it was also identified in many monocotyledons species in families Poaceae [26, 27], Cyperaceae [28], Gramineae [29], and even in eudicotyledons such as alfalfa (Medicago sativa) [30]. The extractives from plants usually contained various conjugated forms, including tricin-glycosides, tricin-monolignols, and tricin-glycoside- monolignols. The natural presence of free tricin was mostly found in cereal plants, such as wheat, oat (Avena sativa), maize (Zea mays), rice (Oryza glaberrima), and barley (Hordeum vulgare). The isolated yield was significantly different from species to species. Njavara rice (Oryza sativa cv. Niavara) [31], a medicinal rice cultivated in India, contains the highest level of free tricin in the bran, with a concentration of 1931 mg/kg of whole cell wall. This content was considerably higher than in the non-medicinal rice cultivars Sujatha and Palakkadan Matta which contain only 49 and 120 mg/kg of free tricin [31]. Tricin primarily accumulates in the aerial part of the plant including straw, leaves, and husk, at different levels, and the accumulation of tricin in plants can be affected by season: a winter wheat variety contains higher level of free tricin in the husk than a spring variety [16]. Tricin-glycoside conjugates are the compounds with one or two carbohydrate units attached to tricin via either C–O ether bonds (mainly on 5-OH, 7-OH, and/or 4'-OH) or C– C bonds (on 6-C and/or 8-C) [32]. Similarly to free tricin, tricin–O–glycosides (Figure 2, compound b) are widespread across different grasses, whereas tricin-C-glycosides are not very common in plants [9]. The carbohydrates in tricin-glycosides are predominately glucose, but xylose [33], arabinose [34], rhamnose [9], and biovinose [35] are also found. Another form of tricin conjugates are the compounds that tricin links via its 4'-OH with a phenylpropanoid unit. The most common one is tricin 4'–O–(β-guaiacylglyceryl) ether (Figure 2, compound c) that is extracted with methanol along with tricin and tricin- glycoside from Hyparrhenia hirta [10], oat [11], and rice [31]. These conjugates can also be in the form of a tricin-(4'–O–β)-p-coumaryl alcohol adduct [36] that has been detected in Aegilops ovate L. and maize. Additionally, coniferyl alcohol with γ-acylation by acetate or p-coumarate has also been reported in conjugation with tricin [37]. It is important to point out that, in previous studies, these conjugates were determined to be optically active and therefore termed as “flavonolignans” [11] (like their component lignan moieties which would logically be optically active [38]). However, a later study [39] demonstrated the racemic nature of these compounds. Therefore, now that tricin in lignin is known, these should be regarded as oligomers that are destined for the fully racemic lignins and should be termed as “flavonolignols”. Tricin-glycoside-phenylpropanoids are conjugates containing both carbohydrate and phenylpropanoid moieties (Figure 2, compounds d and e). Compared to tricin-glycosides and flavonolignols, these compounds have only been reported in a few plants such as Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation 55 sugarcane (Saccharum officinarum) [12], rice (Zizania latifolia) [40], and Acacia nilotica [41], and only in trace amounts. The phenylpropanoid either links onto tricin directly via β–O–4' ether bonding, or onto a glucose moiety on a tricin-glycoside conjugate. Similarly, the carbohydrate etherifies with the 4'/7-OH on tricin.

Figure 2. Chemical structure of tricin a and examples of a tricin-O-glycoside b [13], tricin-O- monolignol c [10], and tricin-glycoside-phenylpropanoid d [12] and e [40]. (Modified from [39]).

Biological Functions and Potential Applications

Flavonoid compounds generally function as antioxidants, antimicrobial/ antiviral agents, allelochemicals, and photoreceptors that are involved in plant growth and development [22]. Similarly, as first isolated from rust-infected wheat leaves, tricin and its related conjugates also possess important bio-functions in plant growth. A study on flavones isolated from rice leaves showed that tricin acted as an allelochemical, defending the rice against weeds by inhibiting their growth. Tricin was able to act against fungal pathogens by inhibiting their spore germination [42]. It was also involved in plant-insect interactions showing high activity against insects [43] and mosquito larvae [9], and acting as an anti-feedant against boll weevils [44]. As it is found in most of our crop plants, tricin is an important part of the human diet and has several effects on human health. It is therefore an attractive candidate for pharmacological and medicinal studies [22]. One of the most prominent and well documented properties of tricin is its potential antitumor/anticancer activity. Tricin has 56 Wu Lan, Jorge Rencoret, José Carlos del Río et al. been shown to inhibit the proliferation of human hepatic stellate cells [45], breast tumor cells [46-48], colon cancer cells [49], and leukemia HL-60 cells [50]. An acylated tricin- glycoside isolated from sugarcane (Saccharum officinarum) juice exhibited antiproliferative activity against several human cancer cell lines, with higher selectivity toward cells of the breast-resistant NIC/ADR line [12]. Additionally, tricin is able to interfere with inflammatory-related mouse colon carcinogenesis, suggesting the potential of tricin for clinical trials of colorectal cancer chemoprevention [51]. A preliminary study on the safety of applying tricin as a chemopreventive agent reported that tricin lacked genotoxic properties in the liver, lung, heart, and kidney tissues of mice, indicating that tricin could be safe enough for clinical development [52]. A structure-activity relationship study of the flavonoids suggested that the O-methylation and glucuronidation significantly increase the cytotoxicity [53]. Similarly, a study of flavones as colorectal cancer preventive agents indicated that the rank order of cancer chemopreventive efficacy is pentamethoxyflavone > tricin > apigenin [54]. Tricin has long been credited for its health-beneficial effects as an antioxidant [55] due to its ability to suppress lipoperoxidation. A previous study showed that the antioxidant activity of tricin was lower than that of luteolin and quercetin according to the rate of lipid peroxidation [55]. However, a later study on the reaction between 2,2-diphenyl-1- picrylhydrazyl radical and tricin and its conjugates suggested that tricin exhibited higher radical scavenging activity than the commonly used compounds such as myricetin, quercetin, and catechin, but lower than the tricin-monolignol dimer [56]. An oxygen radical absorbance capacity assay showed that tricin-glycoside conjugates possessed high antioxidant capacity [13]. The antioxidant ability of tricin and its derivatives was believed to be one of the reasons for their potent anti-inflammatory activities [57, 58]. Shalini et al. extracted tricin from Njavara rice (a medicinal rice cultivated in India) bran and investigated its inflammatory suppression in human peripheral blood mononuclear cells. In this study tricin showed powerful anti-inflammatory activity [59, 60]. Another study also demonstrated the inhibitory activity of tricin on the generation of inflammatory mediators in a cell line of mouse macrophages stimulated with lipopolysaccharide [57].

Preparation of Tricin

Unlike most flavonoid compounds, tricin is not readily available commercially and its isolation from plants is usually in extremely low yield; a total of 8 g of tricin was isolated from 40,000 kg of the Sasa albomarginata leaves in Oyama’s study [51]. In general, the phenolic compounds including tricin and its derivatives in plant tissues are extracted using a methanol/H2O mix solvent (50:50, 80:20, or 100:0, v/v) with or without dilute acid [13, 14, 40, 61]. Some studies also apply ethanol [9], hot water [51], dichloromethane [11], butanol [43], and acetonitrile [62] as the solvent to extract crude tricin extractives. A Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation 57 dewaxing step removes lipids and chlorophyll pigments by diethyl ether or hexane and can be done on the whole cell wall before the tricin solvent extraction or on the crude tricin extractives after the solvent extraction. After distillation, the condensed products are extracted by different solvents (n-hexane, diethyl ether, chloroform, dichloromethane, and butanol) to remove the compounds unrelated to tricin and thereby enrich the concentration of tricin. Such a tricin-rich fraction is further purified mainly by chromatographic techniques to give the pure tricin. An alternative way to prepare tricin in suitable quantities for experimentation and pharmacological testing is through chemical synthesis [63-65] (Figure 3). Some of the methods were based on the formation of a 1,3-diketone and followed by an intra-molecular ketone-hydroxyl reaction to form a flavone backbone (Figure 3a). Another way to make the flavone backbone is the direct condensation of appropriately protected 2,4,6- trihydroxyacetophenone and 4-hydroxy-3,5-dimethoxybenzaldehyde followed by dehydrogenation and cyclization by iodine and sodium acetate [66, 67] (Figure 3b).

Figure 3. Chemical synthesis of tricin. (a) Method based on the formation of a 1,3-diketone [64]; (2) method based on the direct condensation of a ketone and an aldehyde to form the chalcone backbone [67].

OCCURRENCE OF TRICIN IN THE LIGNIN POLYMER

In 2012, tricin was disclosed to be present in the milled wood lignin isolated from wheat straw for the first time [8], primarily via its characteristic HSQC NMR correlations (Figure 4). The mechanism of tricin’s incorporation into grass lignin was further investigated in a later study (Figure 5) [67]. Radical coupling reactions catalyzed by 58 Wu Lan, Jorge Rencoret, José Carlos del Río et al. hydrogen peroxidase showed that even though reaction rate of cross coupling between tricin and monolignols was lower than those of simple dimerization of the monolignol, by limiting their concentrations, tricin was able to form a 4'–O–β linkage with monolignols. Furthermore, the long-range correlation of C4'–Hβ was identified in the HMBC spectrum of acetylated maize stover lignin, as shown in Figure 6. Tricin moieties were also found in the high molecular weight fraction of isolated lignin according to HSQC characterization [67]. All of these data together provided solid evidence that tricin is incorporated into lignin polymers via 4'–O–β coupling.

Figure 4. Aromatic region of the short range 1H–13C correlation (2D HSQC) NMR spectrum of isolated milled wood lignin (MWL) from wheat straw cell walls (A) and long range 1H–13C correlation (2D HMBC) NMR spectrum of MWL from wheat straw cell wall showing the main correlations of tricin units in lignin. (Modified from [8]).

Additional studies verified the presence of tricin in lignin preparations from various monocots, including Carex meyeriana [68], bamboo (Phyllostachys pubescens) [69], coconut (Cocos nucifera) coir [70], giant cane (Arundo donax) [71], rice [72], barley [73], sugarcane [74], and V. planifolia [6]. The pretreatment methods to isolate lignin could also affect the presence of tricin because tricin is not stable under harsh conditions.

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation 59

Figure 5. Radical coupling reaction between tricin and a monolignol.

Figure 6. 2D HMBC spectrum of acetylated maize lignin [67]. Occurrence of the C4'–Hβ correlation demonstrates that tricin is incorporated into the lignin polymer via radical coupling and forming a 4'–O–β aryl ether bond.

60 Wu Lan, Jorge Rencoret, José Carlos del Río et al.

Figure 7. Aromatic region of the 2D HSQC spectra of wheat straw lignin. Samples were pretreated with water at 160°C (A), 0.25 wt% H2SO4 aqueous solution at 160°C (B), and 1.0 wt% H2SO4 aqueous solution at 160°C (C) (modified from [75]). Tricin was completely depleted under higher acid concentration pretreatments (C).

For example, during dilute acid pretreatment of wheat straw, tricin was mostly retained under 160°C with 0.25% H2SO4 in water, whereas all of the tricin vanished when the acid concentration was increased to 1%, or when the pretreatment temperature was increased to 190°C (even without any acid) [75] (Figure 7). Similarly, steam explosion treatment at 200°C for only 10 minutes removed tricin from the lignin polymer significantly [76]. Alkaline conditions also affected the present of tricin even at low temperature. The content of tricin was much lower in the lignin isolated from alkaline pretreated wheat straw than that from the untreated sample [77]. On the other hand, some processes such as mild acid γ-valerolactone pretreatment [78, 79] and extractive-ammonia pretreatment [80] largely preserved the tricin moiety in the lignin polymer from corn stover. However, most of the above studies did not mention the reason for the absence of tricin. It is unclear whether it was cleaved from the polymer as an intact moiety or was degraded into other products. It is reported that the distribution of tricin in lignin polymer chains varied according to the molecular weight: the lower molecular weight fractions contained higher amounts of tricin, whereas in higher molecular weight fractions the tricin level was lower, and even no tricin units were found in the highest molecular weight fraction [81]. In contrast, our study showed that tricin moieties were approximately equally distributed in the lignin fractions of different molecular weights from corn stover [39]. Such differences might be due to the different lignin isolation methods applied in these two studies. Another study showed that different lignin-carbohydrate complex (LCC) fractions contained different levels of tricin [77]. The LCC is a lignin-cell wall cross-linked fraction in which ferulate (FA) presumably functions as a nucleation site. The FA acylates arabinose side-chains of arabinoxylans and radical coupling with monolignols can initiate the growth of lignin polymer chains. Zikeli et al. isolated two different LCC fractions; one was predominantly associated with glucan and the other one was mainly bound with xylan. It is surprising to find that the glucan-rich Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation 61

LCC contained remarkably higher levels of tricin units than the xylan-rich LCC fraction did [77].

Characterization of Tricin

NMR spectroscopy has enormously facilitated the investigations into structural aspects of complex lignin polymers [82, 83]. 1H NMR is the most widely applied NMR technique. But lignin appears broad and featureless in 1H NMR spectra in most of cases, not only because of lignin’s high molecular weight, but also the irregular bonding between monomers and, most importantly, the complexity caused by its stereochemical diversity. However, simple 1H NMR spectroscopy is able to identify the presence of tricin in lignin polymer because the tricin moiety shows a characteristic peak at 13.0 ppm, corresponding to 5-OH, that is very different from anything else in the lignin structure [84]. Compared to the 1D NMR techniques (1H and 13C NMR), the heteronuclear single- quantum coherence (HSQC) technique (a 2D heteronuclear NMR method) provides much more comprehensive information of inter-unit linkages and therefore is frequently applied for lignin characterization. The tricin moiety in the lignin polymer can be easily identified in HSQC spectroscopy because of its four characteristic correlations in the aromatic region, which do not overlap with the peaks from other lignin structures. The two correlation peaks at δC/δH 99.0/6.21 and 94.4/6.57 correspond to the C6/H6 and C8/H8 on aromatic ring (ring

A) and the peak at δC/δH 104.9/7.05 is from the C3/H3 on the C = C double bond (heterogeneous ring C). The C2'/H2'(C6'/H6') correlation of the symmetric aromatic ring

(ring B) raises a peak at δC/δH 104.0/7.30. After acetylation the peaks of C3/H3, C6/H6, and C8/H8 move to δC/δH 107.8/7.05, 110.2/7.71, and 114.3/7.07, whereas the peak of C2'/H2'(C6'/H6') does not change [67]. The chemical shift of C3/H3 indicates whether the tricin moiety is free or incorporated into the polymer via a 4'-O aryl ether bond. When the

4'-O-β bond is cleaved, the peak of C3/H3 slightly moves to downfield (δC/δH 103.5/7.04,

Figure 8). All of the abovementioned chemical shifts are on the basis of using DMSO-d6 as solvent. When using CDCl3 or acetone-d6 as the solvent the chemical shifts of these peaks would be changed, especially in the 1H dimension. Table 1 summarizes the chemical shifts of the characteristic peaks of tricin (non-acetylated and acetylated) using different deuterated NMR solvents. 31P NMR is another technique applied to characterize lignin, mainly focusing on qualitatively and quantitatively analyzing the various hydroxyl groups. It was also applied to characterize the tricin unit in lignin. The three hydroxyl groups (4'-OH, 5-OH, and 7- 31 OH) in tricin generate three peaks in the P NMR spectrum at δP 142.0, 137.6, and 136.4 ppm, respectively [76]. In the case of the 31P NMR spectrum of tricin-integrated lignin (enzymatic mild acidolysis lignin from wheat straw), two peaks at 137.6 and 136.4 ppm 62 Wu Lan, Jorge Rencoret, José Carlos del Río et al. originated from the 5-OH and 7-OH with different intensities indicating an overlap of the peaks from –OH on tricin and an –OH from another structure.

Table 1. Selected diagnostic tricin unit chemical shifts from tricin-(4'–O–β)-

monolignol dimers in acetone-d6, CDCl3, and DMSO-d6. (Modified from [67])

Unacetylated (δC/δH) C3/H3 C6/H6 C8/H8

Acetone-d6 106.01/6.82 99.79/6.27 94.98/6.56

CDCl3 105.83/6.61 99.75/6.33 94.34/6.49

DMSO-d6 104.87/7.05 98.98/6.21 94.37/6.57

Acetylated (δC/δH)

Acetone-d6 108.67/6.78 110.29/7.49 114.80/6.95

CDCl3 108.36/6.60 109.08/7.39 113.65/6.84

DMSO-d6 107.82/7.05 110.18/7.71 114.33/7.07

Figure 8. Aromatic region of the 2D HSQC spectrum of enzymatic lignin from maize. The blue dashed ellipsoids indicate the chemical shifts of free tricin itself. The C3-H3 correlation is especially different between free tricin and 4'-O etherified tricin. (Modified from [67]).

There is no peak at 142.0 ppm from the 4'-OH on tricin because this hydroxyl group is etherified in the lignin. The spectrum of lignin isolated from a sample treated with steam explosion shows that the two sharp peaks from tricin disappear demonstrating that tricin was both cleaved and degraded during pretreatment [76]. Ragauskas et al. systematically investigated the application of 31P NMR to identify tricin. By comparing the 31P NMR spectrum of tricin-incorporated lignin from Zea mays and non-tricin-incorporated lignin from Populous trichocarpa with the spectra of flavonoids, it was found that 31P NMR did Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation 63 provide diagnostic peaks for tricin and other flavonoids. However, again, cautions should be taken when applying such methods for quantitative studies due to peak overlap issues [85]. Another method to characterize tricin in lignin is using wet chemical degradative methods followed by chromatographic characterization. It should be noted that tricin is not able to be detected in gas chromatography (GC) because it is not sufficiently volatile, even when derivatized, under the temperature conditions applicable. Although the volatility of the compound can be improved by trimethylsilylation, the per-trimethylsilyl (TMS) tricin (with TMS on the 5, 7, and 4' hydroxyl groups) was not stable under the GC conditions and partially degraded into di- and mono-TMS tricin. Therefore, liquid chromatography with mass spectrometric (LC-MS) detection is more suitable for tricin-integrated lignin characterization. Tricin is able to be protonated and deprotonated, so it can be detected in both positive- and negative-ion mode, with mass to charge ratios (m/z) of 331 [M+H]+ and 329 [M-H]-, respectively. In tandem MS spectrometry (positive-ion mode), 331 [M+H]+ generates different ion fragments under specific collision energies. The fragments m/z 315 and 300 correspond to cleavage of one and two methoxyl groups. The cleavage of C2–C1' bond forms two ion fragments with m/z as 177 and 153 originating from the ring AC (chalcone backbone) and ring B (syringyl unit).

Figure 9. 31P NMR spectra of wheat straw enzymatic mild acidolysis lignin (EMAL, A), steam explosion-treated wheat straw EMAL (B), and tricin itself showing its three hydroxyl groups (C). (Adapted from [76]).

Quantitation of Tricin

As tricin has been proven to be a monomer in lignin, quantitation of the tricin moiety in the lignin polymer became an important target. The proportions of tricin were roughly 64 Wu Lan, Jorge Rencoret, José Carlos del Río et al. estimated to be 10-15% of wheat straw lignin on the basis of volume integration of diagnostic contours in HSQC spectra. It was, however, thought to be excessive because it is well known that end-groups are over-quantified by such methods that are semi- quantitative at best [86]. A suitable way to quantify tricin is to cleave tricin from the polymer by wet chemical degradative methods followed by LC(-MS) quantitation. Our group evaluated the efficiencies of cleaving the tricin-(4'–O–β)-ether bonds and the degradation of tricin under acidolysis, thioacidolysis, and derivatization followed by reductive cleavage (DFRC) methods [39]. To further increase the accuracy, a deuterated tricin (tricin-d6) was synthesized and used as an internal standard to correct not only sample variations during reaction and work up process, but to compensate for the variability in chromatographic separation, ionization, and MS detection [87, 88]. Thioacidolysis was found to be the best method as it produced a 96 mol% yield of tricin from tricin-(4'–O–β)- coniferyl alcohol. We then screened various seed-plant species and, when present, quantified tricin content in lignin using the thioacidolysis method followed by LC-MS characterization. Oat, wheat, and brachypodium showed the highest amount of tricin as 33.1, 32.7, and 28.0 mg/g of lignin, respectively. All Poaceae samples examined in our study showed the presence of tricin. Some other species, such as Arecaceae, Orchidaceae, and even Fabaceae in Eudicotyledons contained some amount of tricin in their lignin polymers. Compared to the amount of extractable free tricin, the amounts of tricin in the lignins were higher. The bran of Njavara rice (O. sativa cv. Niavara), a medicinal rice cultivated in India, was reported to contain the highest extractable tricin level (1931 mg/kg) [31]. Wheat husk, a more abundant source, contained the second highest tricin level (777 mg/kg) among plants that have been examined [16]. The wheat sample used in our study contained only 376 mg/kg of extractable tricin, whereas the content of lignin-integrated tricin was 4841 mg/kg [39]. The difference was much more substantial when the tricin level was calculated on a lignin basis. Therefore, lignin residue from certain biomass pretreatment processes could be a potential source of large amounts of tricin if a feasible and industrially relevant method to liberate tricin is available.

Bioengineering of Tricin

Lignin biosynthesis and bioengineering have attracted significant attention from researchers because it may play an important role in economically improving applications of agroindustrial biomass. Lignin has long been treated as an impediment to chemical pulping, forage digestion by livestock, and is a side-product of cellulosic biorefinery plants. But it has been increasingly recognized as potential source of aromatic bulk commodity chemicals. As tricin has been disclosed as one of the monomers in grass lignin, manipulation of the biosynthetic pathway responsible for flavone biosynthesis and Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation 65 investigation of the consequences on cell wall composition, recalcitrance, and lignin structure has become an interesting research focus. Branching off from p-coumaroyl-CoA, flavonoids and monolignols are the two major downstream metabolite classes. The flux toward flavone biosynthesis and, thus, to the tricin monomer, is controlled by CHS. In maize, Colorless 2 (C2) is the gene encoding CHS and expressed in many parts of the plant including the pericarp, tassels, and vegetative organs. Therefore, disruption of C2 would cause the depletion of flavonoids, including tricin in stems and leaves of maize. In our previous study [89], CHS-deficient plants showed strikingly lower abundance of flavone metabolites compared to the C2 control plants and no tricin monomer was detected in the lignin polymer. The stems of control and mutant plants contained similar Klason lignin contents, whereas the leaf of C2 mutant maize showed higher lignin content than the control. Accordingly, the cell wall recalcitrance to enzymatic saccharification were more profound for the leaves of C2 mutants than that of controls, but quite similar for the stem samples.

Table 2. Comparison of extractable tricin and lignin-integrated tricin (mg/kg) from non-extracted plant material. (Modified from [39])

Extractable tricin vs Lignin-integrated tricin Plant sample Extractable Extractable Lignin-integrated tricin T-(4'–O–β)-G tricin Wheat straw 376.1 ± 69.5 1044.3 ± 149.9 4841.3 ± 217.8 Maize straw 90.9 ± 12.4 299.7 ± 39.9 1304.0 ± 39.6 Oat straw 601.4 ± 48.4 1270.0 ± 49.7 5250.3 ± 121.9 Rice stem 64.1 ± 6.9 215.3 ± 28.1 979.7 ± 0.1 Extractable tricin reported in previous studies Plant species Part Extractable Extractable tricin T-(4'–O–β)-G Njavara rice (Oryza sativa cv. Niavara) [31] bran 1930.5 ± 0.3 1217.7 ± 1.2 Rice Oryza sativa cv. Sujatha [31] bran 48.6 ± 0.1 45.9 ± 0.9 Rice Oryza sativa cv. Palakkadam Matta [31] bran 119.8 ± 0.1 Not detected Wheat (a winter cultivar) [16] husk 772 ± 31.8 Not reported leaves 253 ± 18.3 Not reported bran 33 ± 15.9 Not reported

Flavone synthase II (FNSII) is the enzyme that catalyzes the direct conversion of flavanones to flavones. It is indispensable for the biosynthesis of both extractable tricin- derived metabolites [90] and tricin monomers for lignification in rice vegetative tissues [24]. In the culm, sheath, and leaf tissue of rice, disrupting FNSII significantly altered the cell wall properties, such as reducing the lignin content and decreasing the S/G ratio in lignin. HSQC characterization of the lignin from FNSII mutant plants indicated the absence of tricin, and a small amount of naringenin units was integrated into the lignin polymer (Figure 11). Additionally, FNSII-knockout mutants exhibited better saccharification 66 Wu Lan, Jorge Rencoret, José Carlos del Río et al. efficiency than the control samples and did not display negative impacts on the growth and development of vegetative tissues [24].

Figure 10. Content of tricin in the lignins from different plant species. The error bars were calculated on the basis of standard deviation (Adapted from [39]).

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation 67

Figure 11. Aromatic regions of 2D HSQC spectra of cell wall lignins from culm tissues of wild-type (WT, A) and FNSII-knockout mutant (B) rice plants. Lignin samples were prepared by enzymatic removal of cell wall polysaccharides with cellulase. In the lignin from fnsII mutant plants, tricin was completely depleted and naringenin units were evident. (Modified from [24]).

After the conversion of naringenin by FNSII to apigenin, substitutions on the B ring of the C6-C3-C6 flavan skeleton are catalyzed by flavonoid 3'-hydroxylases (F3'H) and O- methyltransferase (OMT) to produce flavonoids. CYP75B3 and CYP75B4 are the two enzymes in rice tissue with flavonoid 3'-hydroxylase activities [23,91,92]. By studying the metabolite profiles, cell wall properties, and lignin structure of the cyp75b3, cyp75b4 mutant and cyp75b3 cyp75b4 double-mutant plants, CYP75B3 was proved to be solely responsible for the biosynthesis of 3'-subsituted flavone C-glycosides. Disrupting CYP75B3 did not affect the lignin structure and cell wall properties. On the other hand, CYP75B4 was demonstrated to process both apigenin 3'-hydroxylation and chrysoeriol 5'- hydroxylation activity corresponding to the production of lignin-bound tricin and tricin O- conjugates. Similarly to the FNSII mutant, a knockout in CYP75B4 decreased the S/G lignin unit composition and remarkably reduced the lignin levels, accordingly enhancing the carbohydrates digestibility. As expected, cyp75b4 mutant plants produced tricin- depleted lignin. In culm tissue, apigenin, instead of tricin, was found to be integrated in lignin polymer [93]. 68 Wu Lan, Jorge Rencoret, José Carlos del Río et al.

Caffeoyl coenzyme A 3-O-methyltransferase (CCoAOMT) and caffeic acid-O- methyltransferase (COMT) were well documented as the important enzymes for the biosynthesis of monolignols. Downregulation or knockout of COMT or both enzymes enriched the G units in lignins and reduced the total lignin content [94-97]. A recent study also demonstrated that a comt mutant maize produced tricin-depleted lignin in stem tissue, whereas the tricin level was not significantly affected in the midrib. This observation suggested that COMT was also involved in the tricin biosynthetic pathway, at least in the stem tissue [98].

Implications of Tricin’s Presence in Lignin

In maize metabolite profiling, coniferyl and sinapyl alcohol and their acetate and p- coumarate conjugates were all found to couple with tricin. Furthermore, using a chiral column, we were able to separate two enantiomers of the tricin-(4'–O–β)-coniferyl alcohol and tricin-(4'–O–β)-p-coumaryl alcohol using LC-MS/MS with multiple reaction monitoring (MRM). The identical peak areas of the two enantiomers indicated the racemic nature of the tricin conjugates [37]. In the case of the tricin-(4'–O–β)-coniferyl alcohol-(4'– O–β)-coniferyl alcohol trimer, 6 of the 8 isomers were separated and identified in LC- MS/MS, suggesting the compounds were formed by simple radical reactions [37]. As in the established theory and as increasingly evidenced, lignins are the products of simple, but combinatorial, radical coupling chemistry [99,100]. However, such a concept was heatedly debated after notions of absolute proteinaceous control over lignin structure were championed for a period [101], but were eventually convincingly debunked [102]. The fact that tricin cross-couples with acetate and p-coumarate monolignol conjugates, the racemic nature of flavonolignols, and the diversity of the diastereomers support the combinatorial radical coupling theory, demonstrating that that lignins are racemic polymers, are characterized by being products with a huge number of possible isomers, and have no defined sequence or (repeating) structure. Biomimetic radical coupling reactions between tricin and monolignols and characterization of lignin polymer from nature products indicate that tricin only incorporated into the lignin polymer in the form of 4'–O–β-coupled products and their higher oligomers [67]. In this case, the tricin unit must be localized at one terminus of its lignin chain, and that terminus must be at the starting end of that chain. In other words, tricin acts as a nucleation site for lignin chain growth in monocots, a role that was proposed for ferulate on arabinoxylans [103]. Such an observation contributes to resolving a monocot lignin structural dilemma that has existed for decades: that monocot lignins (especially maize lignin), unlike other syringyl-guaiacyl lignins in dicots/hardwoods, have essentially no, or very low levels of resinols (β–β-coupled units) [67,96]. Such β–β units are produced only as the result of monolignol (sinapyl alcohol) dimerization and are the obvious Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation 69 mechanism for starting a lignin chain. One of the reasons is that sinapyl p-coumarate homodimerization, instead of sinapyl alcohol coupling, is more preponderant to function as the starting point. Also, when the lignin chain is nucleated by another unit, such as tricin or ferulate, lignification does not need to start with a dimerization reaction.

CONCLUSION

Tricin, a member in flavonoid family with a C6-C3-C6 backbone, is a novel monomer in lignin discovered in recent years. Not only does it arise from a different biosynthetic pathway from the traditional monolignols, it operates in a polymer chain nucleation function; its biological functions and potential pharmaceutical applications increasingly attract research interest. Studies related to the biosynthesis and bioengineering, identification, characterization, and quantification of tricin are providing new insight into the lignin structure. The lignin-bound tricin also provides a new and abundant source of such a high-market-priced chemical, encouraging research into its production methods from lignin polymers and its applications beyond its current roles.

ACKNOWLEDGMENTS

The authors thank the China Scholarship Council, State Education Department, for supporting living expenses for Wu Lan’s PhD Program in the Department of Biological System Engineering, University of Wisconsin, Madison, USA. WL, and JRa were funded by the DOE Great Lakes Bioenergy Research Center (DOE Office of Science BER DE- FC02-07ER64494 and DE-SC0018409). JRe and JdR was funded by the Spanish Project CTQ2014-60764-JIN (co-financed by FEDER funds).

REFERENCES

[1] Boerjan, W.; Ralph, J.; Baucher, M. 2003. “Lignin biosynthesis.” Annual Review of Plant Biology 54, 519-46. [2] Ralph, J. 2010. “Hydroxycinnamates in lignification.” Phytochemistry Reviews 9, 65-83. [3] Karlen, S. D.; Smith, R. A.; Kim, H.; Padmakshan, D.; Bartuce, A.; Mobley, J. K.; Free, H. C. A.; Smith, B. G.; Harris, P. J.; Ralph, J. 2017. “Highly decorated lignins in leaf tissues of the Canary Island date palm Phoenix canariensis.” Plant Physiology 175, 1058-67. 70 Wu Lan, Jorge Rencoret, José Carlos del Río et al.

[4] Karlen, S. D.; Zhang, C. C.; Peck, M. L.; Smith, R. A.; Padmakshan, D.; Helmich, K. E.; Free, H. C. A.; Lee, S.; Smith, B. G.; Lu, F. C. et al. 2016. “Monolignol ferulate conjugates are naturally incorporated into plant lignins.” Science Advances 2. [5] Chen, F.; Tobimatsu, Y.; Jackson, L.; Nakashima, J.; Ralph, J.; Dixon, R. A. 2013. “Novel seed coat lignins in the Cactaceae: structure, distribution and implications for the evolution of lignin diversity.” Plant Journal 73, 201-11. [6] Chen, F.; Tobimatsu, Y.; Havkin-Frenkel, D.; Dixon, R. A.; Ralph, J. 2012. “A polymer of caffeyl alcohol in plant seeds.” Proceedings of the National Academy of Sciences of the United States of America 109, 1772-77. [7] Zhao, Q.; Tobimatsu, Y.; Zhou, R.; Pattathil, S.; Gallego-Giraldo, L.; Fu, C.; Jackson, L. A.; Hahn, M. G.; Kim, H.; Chen, F. et al. 2013. “Loss of function of cinnamyl alcohol dehydrogenase 1 leads to unconventional lignin and a temperature- sensitive growth defect in Medicago truncatula.” Proceedings of the National Academy of Sciences of the United States of America 110, 13660-65. [8] del Río, J. C.; Rencoret, J.; Prinsen, P.; Martinez, A. T.; Ralph, J.; Gutierrez, A. 2012. “Structural characterization of wheat straw lignin as revealed by analytical pyrolysis, 2D-NMR, and reductive cleavage methods.” Journal of Agricultural and Food Chemistry 60, 5922-35. [9] Ju, Y.; Sacalis, J. N.; Still, C. C. 1998. “Bioactive flavonoids from endophyte- infected blue grass (Poa ampla).” Journal of Agricultural and Food Chemistry 46, 3785-88. [10] Bouaziz, M.; Veitch, N. C.; Grayer, R. J.; Simmonds, M. S. J.; Damak, M. 2002. “Flavonolignans from Hyparrhenia hirta.” Phytochemistry 60: 515–520. [11] Wenzig, E.; Kunert, O.; Ferreira, D.; Schmid, M.; Schuhly, W.; Bauer, R.; Hiermann, A. 2005. “Flavonolignans from Avena sativa.” Journal of Natural Products 68, 289- 92. [12] Duarte-Almeida, J. M.; Negri, G.; Salatino, A.; de Carvalho, J. E.; Lajolo, F. M. 2007. “Antiproliferative and antioxidant activities of a tricin acylated glycoside from sugarcane (Saccharum officinarum) juice.” Phytochemistry 68, 1165-71. [13] Van Hoyweghen, L.; Karalic, I.; Van Calenbergh, S.; Deforce, D.; Heyerick, A. 2010. “Antioxidant flavone glycosides from the leaves of Fargesia robusta.” Journal of Natural Products 73, 1573-77. [14] Nakano, H.; Kawada, N.; Yoshida, M.; Ono, H.; Iwaura, R.; Tonooka, T. 2011. “Isolation and identification of flavonoids accumulated in proanthocyanidin-free barley.” Journal of Agricultural and Food Chemistry 59, 9581-87. [15] Bottcher, A.; Cesarino, I.; dos Santos, A. B.; Vicentini, R.; Mayer, J. L. S.; Vanholme, R.; Morreel, K.; Goeminne, G.; Moura, J. C. M. S.; Nobile, P. M. et al. 2013. “Lignification in sugarcane: Biochemical characterization, gene discovery, Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation 71

and expression analysis in two genotypes contrasting for lignin content.” Plant Physiology 163, 1539-57. [16] Moheb, A.; Grondin, M.; Ibrahim, R. K.; Roy, R.; Sarhan, F. 2013. “Winter wheat hull (husk) is a valuable source for tricin, a potential selective cytotoxic agent.” Food Chemistry 138, 931-37. [17] Winkel-Shirley, B. 2001. “Flavonoid biosynthesis. A colorful model for genetics, biochemistry, cell biology, and biotechnology.” Plant Physiology 126, 485-93. [18] Morreel, K.; Goeminne, G.; Storme, V.; Sterck, L.; Ralph, J.; Coppieters, W.; Breyne, P.; Steenackers, M.; Georges, M.; Messens, E. et al. 2006. “Genetical metabolomics of flavonoid biosynthesis in Populus: a case study.” Plant Journal 47, 224-37. [19] Zhou, J. M.; Fukushi, Y.; Wang, X. F.; Ibrahim, R. K. 2006. “Characterization of a novel flavone O-methyltransferase gene in rice.” Natural Product Communications 1, 981-84. [20] Zhou, J. M.; Gold, N. D.; Martin, V. J. J.; Wollenweber, E.; Ibrahim, R. K. 2006. “Sequential O-methylation of tricetin by a single gene product in wheat.” Biochimica Et Biophysica Acta-General Subjects 1760, 1115-24. [21] Zhou, J. M.; Fukushi, Y.; Wollenweber, E.; Ibrahim, R. K. 2008. “Characterization of two O-methyltransferase-like genes in barley and maize.” Pharmaceutical Biology 46, 26-34. [22] Zhou, J.-M.; Ibrahim, R. K. 2010. “Tricin—a potential multifunctional neutraceutical.” Phytochemistry Reviews 9, 413-24. [23] Lam, P. Y.; Liu, H. J.; Lo, C. 2015. “Completion of tricin biosynthesis pathway in rice: Cytochrome P450 75B4 is a unique chrysoeriol 5'-hydroxylase.” Plant Physiology 168, 1527-U760. [24] Lam, P. Y.; Tobimatsu, Y.; Takeda, Y.; Suzuki, S.; Yamamura, M.; Umezawa, T.; Lo, C. 2017. “Disrupting flavone synthase II alters lignin and improves biomass digestibility.” Plant Physiology 174, 972-85. [25] Anderson, J. A.; Perkin, A. G. 1931. “The yellow colouring matter of Khapli wheat, Triticum dicoccum.” Journal of Chemical Society [London] 1931, 2624-25. [26] Harborne, J. B.; Hall, E. 1964. “Plant polyphenols. XII. The occurrence of tricin and of glycoflavones in grasses.” Phytochemistry 3, 421-28. [27] Kuwazuka, S.; Oshima, Y. 1964. “Studies on polyphenols in rice plant. III. Isolation and determination of tricin-glycosides “glucotricin” and “tricinin”.” Agricultural and Biological Chemistry 28, A31-A31. [28] Harborne, J. B. 1971. “Comparative biochemistry of flavonoids. 16. distribution and taxonomic significance of flavonoids in leaves of Cyperaceae.” Phytochemistry 10, 1569-&. [29] Harborne, J. B.; Williams, C. A. 1976. “Flavonoid patterns in leaves of the Gramineae.” Biochemical Systematics and Ecology 4, 267-80. 72 Wu Lan, Jorge Rencoret, José Carlos del Río et al.

[30] Bickoff, E. M.; Livingston, A. L.; Booth, A. N. 1964. “Tricin from alfalfa: Isolation and physiological activity.” Journal of Pharmaceutical Sciences 53, 1411-12. [31] Mohanlal, S.; Parvathy, R.; Shalini, V.; Helen, A.; Jayalekshmy, A. 2011. “Isolation, characterization and quantification of tricin and flavonolignans in the medicinal rice Njavara (Oryza sativa L.), as compared to staple varieties.” Plant Foods for Human Nutrition 66, 91-96. [32] Ferreres, F.; Gil-Izquierdo, A.; Andrade, P. B.; Valentao, P.; Tomas-Barberan, F. A. 2007. “Characterization of C-glycosyl flavones O-glycosylated by liquid chromatography-tandem mass spectrometry.” Journal of Chromatography A 1161, 214-23. [33] Theodor, R.; Zinsmeister, H. D.; Mues, R.; Markham, K. R. 1981. “Flavone C- glycosides of 2 Metzgeria species.” Phytochemistry 20, 1851-52. [34] Parveen, M.; Khanam, Z.; Ali, A.; Ahmad, S. M. 2010. “A novel antimicrobial flavonoidic glycoside from the leaves of Alstonia macrophylla Wall ex A. DC (Apocynaceae).” Chinese Chemical Letters 21, 593-95. [35] Sun, J.; Yue, Y. D.; Tang, F.; Guo, X. F.; Wang, J.; Yao, X. 2013. “Flavonoids from the Leaves of Neosinocalamus affinis.” Chemistry of Natural Compounds 49, 822- 25. [36] Cooper, R.; Gottlieb, H. E.; Lavie, D. 1977. “New flavolignan of biogenetic interest from Aegilop Ovata L. 1.” Israel Journal of Chemistry 16, 12-15. [37] Lan, W.; Morreel, K.; Lu, F.; Rencoret, J.; del Río, J. C.; Voorend, W.; Vermerris, W.; Boerjan, W.; Ralph, J. 2016. “Maize tricin-oligolignol metabolites and their implications for monocot lignification.” Plant Physiology 171, 810-20. [38] Umezawa, T. 2004. “Diversity in lignan biosynthesis.” Phytochemistry Reviews 2, 371-90. [39] Lan, W.; Rencoret, J.; Lu, F. C.; Karlen, S. D.; Smith, B. G.; Harris, P. J.; del Río, J. C.; Ralph, J. 2016. “Tricin-lignins: occurrence and quantitation of tricin in relation to phylogeny.” Plant Journal 88, 1046-57. [40] Lee, S. S.; Baek, N. I.; Baek, Y. S.; Chung, D. K.; Song, M. C.; Bang, M. H. 2015. “New flavonolignan glycosides from the aerial parts of Zizania latifolia.” Molecules 20, 5616-24. [41] Khanam, Z.; Adam, F.; Singh, O.; Ahmad, J. 2011. “A novel acylated flavonoidic glycoside from the wood of cultivated Acacia nilotica (L.) Willd. Ex. Delile.” Bioresources 6, 2932-40. [42] Kong, C. H.; Xu, X. H.; Zhou, B.; Hu, F.; Zhang, C. X.; Zhang, M. X. 2004. “Two compounds from allelopathic rice accession and their inhibitory activity on weeds and fungal pathogens.” Phytochemistry 65, 1123-28. [43] Adjei-Afriyie, F.; Kim, C. S.; Takemura, M.; Ishikawa, M.; Horiike, M. 2000. “Isolation and identification of the probing stimulants in the rice plant for the white- Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation 73

back planthopper, Sogatella furcifera (Homoptera: Delphacidae).” Bioscience Biotechnology and Biochemistry 64, 443-46. [44] Miles, D. H.; Tunsuwan, K.; Chittawong, V.; Kokpol, U.; Choudhary, M. I.; Clardy, J. 1993. “Boll weevil antifeedants from arundo donax.” Phytochemistry 34, 1277- 79. [45] Seki, N.; Toh, U.; Kawaguchi, K.; Ninomiya, M.; Koketsu, M.; Watanabe, K.; Aoki, M.; Fujii, T.; Nakamura, A.; Akagi, Y. et al. 2012. “Tricin inhibits proliferation of human hepatic stellate cells in vitro by blocking tyrosine phosphorylation of PDGF receptor and its signaling pathways.” Journal of Cellular Biochemistry 113, 2346- 55. [46] Cai, H.; Hudson, E. A.; Mann, P.; Verschoyle, R. D.; Greaves, P.; Manson, M. M.; Steward, W. P.; Gescher, A. J. 2004. “Growth-inhibitory and cell cycle-arresting properties of the rice bran constituent tricin in human-derived breast cancer cells in vitro and in nude mice in vivo.” British Journal of Cancer 91, 1364-71. [47] Yan, J.; Sun, L. R.; Zhang, X. M.; Qiu, M. H. 2005. “A new flavone from Lycopodium japonicum.” Heterocycles 65, 661-66. [48] Jeong, Y. H.; Chung, S. Y.; Han, A. R.; Sung, M. K.; Jang, D. S.; Lee, J.; Kwon, Y.; Lee, H. J.; Seo, E. K. 2007. “P-glycoprotein inhibitory activity of two phenolic compounds, (-)-syringaresinol and tricin from Sasa borealis.” Chemistry & Biodiversity 4, 12-16. [49] Hudson, E. A.; Dinh, P. A.; Kokubun, T.; Simmonds, M. S. J.; Gescher, A. 2000. “Characterization of potentially chemopreventive phenols in extracts of brown rice that inhibit the growth of human breast and colon cancer cells.” Cancer Epidemiology Biomarkers & Prevention 9, 1163-70. [50] Ninomiya, M.; Nishida, K.; Tanaka, K.; Watanabe, K.; Koketsu, M. 2013. “Structure-activity relationship studies of 5,7-dihydroxyflavones as naturally occurring inhibitors of cell proliferation in human leukemia HL-60 cells.” Journal of Natural Medicines 67, 460-67. [51] Oyama, T.; Yasui, Y.; Sugie, S.; Koketsu, M.; Watanabe, K.; Tanaka, T. 2009. “Dietary tricin suppresses inflammation-related colon carcinogenesis in male Crj: CD-1 mice.” Cancer Prevention Research 2, 1031-38. [52] Verschoyle, R. E.; Greaves, P.; Cai, H.; Arndt, B.; Broggini, M.; D'Incalci, M.; Riccio, E.; Doppalapudi, R.; Kapetanovic, I. M.; Steward, W. P. et al. 2006. “Preliminary safety evaluation of the putative cancer chemopreventive agent tricin, a naturally occurring flavone.” Cancer Chemotherapy and Pharmacology 57, 1-6. [53] Plochmann, K.; Korte, G.; Koutsilieri, E.; Richling, E.; Riederer, P.; Rethwilm, A.; Schreier, P.; Scheller, C. 2007. “Structure-activity relationships of flavonoid- induced cytotoxicity on human leukemia cells.” Archives of Biochemistry and Biophysics 460, 1-9. 74 Wu Lan, Jorge Rencoret, José Carlos del Río et al.

[54] Cai, H.; Sale, S.; Schmid, R.; Britton, R. G.; Brown, K.; Steward, W. P.; Gescher, A. J. 2009. “Flavones as colorectal cancer chemopreventive agents-phenol-O- methylation enhances efficacy.” Cancer Prevention Research 2, 743-50. [55] Watanabe, M. 1999. “Antioxidative phenolic compounds from Japanese barnyard millet (Echinochloa utilis) grains.” Journal of Agricultural and Food Chemistry 47, 4500-05. [56] Ajitha, M. J.; Mohanlal, S.; Suresh, C. H.; Jayalekshmy, A. 2012. “DPPH radical scavenging activity of tricin and its conjugates isolated from “Njavara” rice bran: A density functional theory study.” Journal of Agricultural and Food Chemistry 60, 3693-99. [57] Moscatelli, V.; Hnatyszyn, O.; Acevedo, C.; Megias, J.; Alcaraz, M. J.; Ferraro, G. 2006. “Flavonoids from Artemisia copa with anti-inflammatory activity.” Planta Medica 72, 72-74. [58] Lee, S. S.; Baek, Y. S.; Eun, C. S.; Yu, M. H.; Baek, N. I.; Chung, D. K.; Bang, M. H.; Yang, S. A. 2015. “Tricin derivatives as anti-inflammatory and anti-allergic constituents from the aerial part of Zizania latifolia.” Bioscience Biotechnology and Biochemistry 79, 700-06. [59] Shalini, V.; Bhaskar, S.; Kumar, K. S.; Mohanlal, S.; Jayalekshmy, A.; Helen, A. 2012. “Molecular mechanisms of anti-inflammatory action of the flavonoid, tricin from Njavara rice (Oryza sativa L.) in human peripheral blood mononuclear cells: Possible role in the inflammatory signaling.” International Immunopharmacology 14, 32-38. [60] Shalini, V.; Jayalekshmi, A.; Helen, A. 2015. “Mechanism of anti-inflammatory effect of tricin, a flavonoid isolated from Njavara rice bran in LPS induced hPBMCs and carrageenan induced rats.” Molecular Immunology 66, 229-39. [61] Ogo, Y.; Ozawa, K.; Ishimaru, T.; Murayama, T.; Takaiwa, F. 2013. “Transgenic rice seed synthesizing diverse flavonoids at high levels: a new platform for flavonoid production with associated health benefits.” Plant Biotechnology Journal 11, 734- 46. [62] Norbaek, R.; Aaboer, D. B. F.; Bleeg, I. S.; Christensen, B. T.; Kondo, T.; Brandt, K. 2003. “Flavone C-glycoside, phenolic acid, and nitrogen contents in leaves of barley subject to organic fertilization treatments.” Journal of Agricultural and Food Chemistry 51, 809-13. [63] Owada, E.; Mieno, M. 1970. “Synthetic studies of flavonoid compounds. 2. improved synthesis of tricin.” Nippon Kagaku Zasshi 91, 1002-&. [64] Nagarathnam, D.; Cushman, M. 1991. “A short and facile synthetic route to hydroxylated flavones - new syntheses of apigenin, tricin, and luteolin.” Journal of Organic Chemistry 56, 4884-87. Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation 75

[65] Pandurangan, N. 2014. “A new synthesis for acacetin, chrysoeriol, diosmetin, tricin and other hydroxylated flavones by modified Baker-Venkataraman transformation.” Letters in Organic Chemistry 11, 225-29. [66] Wang, J.; Zhou, R. G.; Wu, T.; Yang, T.; Qin, Q. X.; Li, I.; Yang, B.; Yang, J. 2012. “Total synthesis of apigenin.” Journal of Chemical Research, 121-22. [67] Lan, W.; Lu, F. C.; Regner, M.; Zhu, Y. M.; Rencoret, J.; Ralph, S. A.; Zakai, U. I.; Morreel, K.; Boerjan, W.; Ralph, J. 2015. “Tricin, a flavonoid monomer in monocot lignification.” Plant Physiology 167, 1284-95. [68] Mao, J. Z.; Zhang, X.; Li, M.-F.; Xu, F. 2013. “Effect of biological pretreatment with white-rot fungus trametes hirsuta C7784 on lignin structure in Carex meyeriana Kunth.” Bioresources 8, 3869-83. [69] Wen, J.; Sun, S.; Xue, B.; Sun, R. 2013. “Quantitative structural characterization of the lignins from the stem and pith of bamboo (Phyllostachys pubescens).” Holzforschung 67, 613-27. [70] Rencoret, J.; Ralph, J.; Marques, G.; Gutierrez, A.; Martinez, A. T.; del Río, J. C. 2013. “Structural characterization of lignin isolated from coconut (Cocos nucifera) coir fibers.” Journal of Agricultural and Food Chemistry 61, 2434-45. [71] You, T. T.; Mao, J. Z.; Yuan, T. Q.; Wen, J. L.; Xu, F. 2013. “Structural elucidation of the lignins from stems and foliage of Arundo donax Linn.” Journal of Agricultural and Food Chemistry 61, 5361-70. [72] Wu, M.; Pang, J.; Lu, F.; Zhang, X.; Che, L.; Xu, F.; Sun, R. 2013. “Application of new expansion pretreatment method on agricultural waste. Part I: Influence of pretreatment on the properties of lignin.” Industrial Crops and Products 50, 887-95. [73] Rencoret, J.; Prinsen, P.; Gutierrez, A.; Martinez, A. T.; del Río, J. C. 2015. “Isolation and structural characterization of the milled wood lignin, dioxane lignin, and cellulolytic lignin preparations from Brewer’s spent grain.” Journal of Agricultural and Food Chemistry 63, 603-13. [74] del Río, J. C.; Lino, A. G.; Colodette, J. L.; Lima, C. F.; Gutierrez, A.; Martinez, A. T.; Lu, F. C.; Ralph, J.; Rencoret, J. 2015. “Differences in the chemical structure of the lignins from sugarcane bagasse and straw.” Biomass & Bioenergy 81, 322-38. [75] Jensen, A.; Cabrera, Y.; Hsieh, C. W.; Nielsen, J.; Ralph, J.; Felby, C. 2017. “2D NMR characterization of wheat straw residual lignin after dilute acid pretreatment with different severities.” Holzforschung 71, 461-69. [76] Heikkinen, H.; Elder, T.; Maaheimo, H.; Rovio, S.; Rahikainen, J.; Kruus, K.; Tamminen, T. 2014. “Impact of steam explosion on the wheat straw lignin structure studied by solution-state nuclear magnetic resonance and density functional methods.” Journal of Agricultural and Food Chemistry 62, 10437-44. 76 Wu Lan, Jorge Rencoret, José Carlos del Río et al.

[77] Zikeli, F.; Ters, T.; Fackler, K.; Srebotnik, E.; Li, J. B. 2016. “Wheat straw lignin fractionation and characterization as lignin-carbohydrate complexes.” Industrial Crops and Products 85, 309-17. [78] Luterbacher, J. S.; Rand, J. M.; Alonso, D. M.; Han, J.; Youngquist, J. T.; Maravelias, C. T.; Pfleger, B. F.; Dumesic, J. A. 2014. “Nonenzymatic sugar production from biomass using biomass-derived γ-valerolactone.” Science 343, 277- 80. [79] Luterbacher, J. S.; Azarpira, A.; Motagamwala, A. H.; Lu, F.; Ralph, J.; Dumesic, J. A. 2015. “Aromatic monomer production integrated into the γ-valerolactone sugar platform.” Energy and Environmental Science 8, 2657-63. [80] Sousa, L. D.; Foston, M.; Bokade, V.; Azarpira, A.; Lu, F. C.; Ragauskas, A. J.; Ralph, J.; Dale, B.; Balan, V. 2016. “Isolation and characterization of new lignin streams derived from extractive-ammonia (EA) pretreatment.” Green Chemistry 18, 4205-15. [81] Zikeli, F.; Ters, T.; Fackler, K.; Srebotnik, E.; Li, J. B. 2016. “Fractionation of wheat straw Dioxane lignin reveals molar mass dependent structural differences.” Industrial Crops and Products 91, 186-93. [82] Lu, F. C.; Ralph, J. 2011. “Solution-state NMR of lignocellulosic biomass.” Journal of Biobased Materials and Bioenergy 5, 169-80. [83] Ralph, J.; Landucci, L. L. In Lignin and Lignans; Advances in Chemistry; Heitner, C.; Dimmel, D. R.; Schmidt, J. A., Eds.; CRC Press (Taylor & Francis Group): Boca Raton, FL, 2010, doi: org/10.1201/EBK1574444865 org/10.1201/EBK1574444865. [84] Kwon, Y. S.; Kim, C. M. 2003. “Antioxidant constituents from the stem of Sorghum bicolor.” Archives of Pharmaceutical Research 26, 535-39. [85] Li, M.; Pu, Y. Q.; Tschaplinski, T. J.; Ragauskas, A. J. 2017. “31P NMR Characterization of tricin and its structurally similar flavonoids.” Chemistry Select 2, 3557-61. [86] Mansfield, S. D.; Kim, H.; Lu, F. C.; Ralph, J. 2012. “Whole plant cell wall characterization using solution-state 2D NMR.” Nature Protocols 7, 1579-89. [87] Stokvis, E.; Rosing, H.; Beijnen, J. H. 2005. “Stable isotopically labeled internal standards in quantitative bioanalysis using liquid chromatography/mass spectrometry: necessity or not?” Rapid Communications in Mass Spectrometry 19, 401-07. [88] Schafer, J.; Urbat, F.; Rund, K.; Bunzel, M. 2015. “A stable-isotope dilution GC-MS approach for the analysis of DFRC (Derivatization Followed by Reductive Cleavage) monomers from low-lignin plant materials.” Journal of Agricultural and Food Chemistry 63, 2668-73. [89] Eloy, N. B.; Voorend, W.; Lan, W.; Saleme, M. D. S.; Cesarino, I.; Vanholme, R.; Smith, R. A.; Goeminne, G.; Pallidis, A.; Morreel, K. et al. 2017. “Silencing Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation 77

CHALCONE SYNTHASE in maize impedes the incorporation of tricin into lignin and increases lignin content.” Plant Physiology 173, 998-1016. [90] Lam, P. Y.; Zhu, F. Y.; Chan, W. L.; Liu, H. J.; Lo, C. 2014. “Cytochrome P450 93G1 is a flavone synthase II that channels flavanones to the biosynthesis of tricin O-linked conjugates in rice.” Plant Physiology 165, 1315-27. [91] Park, S.; Choi, M. J.; Lee, J. Y.; Kim, J. K.; Ha, S. H.; Lim, S. H. 2016. “Molecular and biochemical analysis of two rice flavonoid 3'-hydroxylase to evaluate their roles in flavonoid biosynthesis in rice grain.” International Journal of Molecular Sciences 17. [92] Shih, C. H.; Chu, H.; Tang, L. K.; Sakamoto, W.; Maekawa, M.; Chu, I. K.; Wang, M.; Lo, C. 2008. “Functional characterization of key structural genes in rice flavonoid biosynthesis.” Planta 228, 1043-54. [93] Lam, P. Y.; Tobimatsu, Y.; Lui, A. C.; Yamamura, M.; Wang, L.; Takeda, Y.; Suzuki, S.; Liu, H.; Zhu, F.; Chen, M. et al. 2018. “Recruitment of specific flavonoid B-ring hydroxylases for two independent biosynthesis pathways of flavone-derived metabolites in grasses.” New Phytologist, under revision. [94] Do, C. T.; Pollet, B.; Thevenin, J.; Sibout, R.; Denoue, D.; Barriere, Y.; Lapierre, C.; Jouanin, L. 2007. “Both caffeoyl Coenzyme A 3-O-methyltransferase 1 and caffeic acid O-methyltransferase 1 are involved in redundant functions for lignin, flavonoids and sinapoyl malate biosynthesis in Arabidopsis.” Planta 226, 1117-29. [95] Marita, J. M.; Ralph, J.; Hatfield, R. D.; Guo, D. J.; Chen, F.; Dixon, R. A. 2003. “Structural and compositional modifications in lignin of transgenic alfalfa down- regulated in caffeic acid 3-O-methyltransferase and caffeoyl coenzyme A 3-O- methyltransferase.” Phytochemistry 62, 53-65. [96] Marita, J. M.; Vermerris, W.; Ralph, J.; Hatfield, R. D. 2003. “Variations in the cell wall composition of maize brown midrib mutants.” Journal of Agricultural and Food Chemistry 51, 1313-21. [97] Palmer, N. A.; Sattler, S. E.; Saathoff, A. J.; Funnell, D.; Pedersen, J. F.; Sarath, G. 2008. “Genetic background impacts soluble and cell wall-bound aromatics in brown midrib mutants of sorghum.” Planta 229, 115-27. [98] Fornalé, S.; Rencoret, J.; García-Calvo, L.; Encina, A.; Rigau, J.; Gutiérrez, A.; del Río, J. C.; Caparros-Ruiz, D. 2017. “Changes in cell wall polymers and degradability in maize mutants lacking 3' -and 5'-O-methyltransferases involved in lignin biosynthesis.” Plant and Cell Physiology 58, 240-55. [99] Ralph, J.; lundquist, K.; Brunow, G.; Lu, F.; Kim, H.; Schatz, P. F.; Marita, J. M.; Hatfield, R. D.; Ralph, S. A.; Christensen, J. H. 2004. “Lignins: natural polymers from oxidative coupling of 4-hydroxyphenylpropanoids.” Phytochemistry Reviews 3, 29-60. [100] Vanholme, R.; Demedts, B.; Morreel, K.; Ralph, J.; Boerjan, W. 2010. “Lignin biosynthesis and structure.” Plant Physiology 153, 895-905. 78 Wu Lan, Jorge Rencoret, José Carlos del Río et al.

[101] Davin, L. B.; Lewis, N. G. 2005. “Lignin primary structures and dirigent sites.” Current Opinion in Biotechnology 16, 407-15. [102] Ralph, J.; Brunow, G.; Harris, P. J.; Dixon, R. A.; Schatz, P. F.; Boerjan, W. In Recent Advances in Polyphenol Research; Daayf, F.; El Hadrami, A.; Adam, L.; Ballance, G. M., Eds.; Wiley-Blackwell Publishing: Oxford, UK, 2008; Vol. 1. [103] Ralph, J.; Grabber, J. H.; Hatfield, R. D. 1995. “Lignin-ferulate cross-links in grasses: Active incorporation of ferulate polysaccharide esters into ryegrass lignins.” Carbohydrate Research 275, 167-78.