wu lan , jorge rencoret, josé carlos del río and john ralph

In: Lignin ISBN: 978-1-53614-769-8

Editors: Fachuang Lu and Fengxia Yue © 2019 Nova Science Publishers, Inc.

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].

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Wu Lan, Jorge Rencoret, José Carlos del Río et al. 52

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], con-

taining 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].


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


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


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


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


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.


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.


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


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-

OH) in tricin generate three peaks in the 31P 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


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


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


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


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

tricin

Extractable

T-(4'–O–β)-G

Lignin-integrated

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

tricin

Extractable

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


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]).


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].


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


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).

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wu lan , jorge rencoret, josé carlos del río and john ralph

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