Metabolic Engineering of Lignan Biosynthesis Pathways for the Production of Transgenic Plant-Based Foods

  • Honoo SatakeEmail author
  • Tomotsugu KoyamaEmail author
  • Erika MatsumotoEmail author
  • Kinuyo MorimotoEmail author
  • Eiichiro OnoEmail author
  • Jun MurataEmail author
Living reference work entry
Part of the Reference Series in Phytochemistry book series (RSP)


Lignans are major phytochemicals biosynthesized in several plants including Sesamum, Linum, Forsythia, and Podophyllum genus, and a great variety of lignans have received wide attentions as leading compounds of novel drugs for tumor treatment and healthy diets to reduce of the risks of lifestyle-related diseases. Recent genome and transcriptome studies have characterized multiple novel lignan-biosynthetic enzymes, and thus have opened new avenues to transgenic metabolic engineering of various nonmodel dietary or medicinal plants. Forsythia and Linum are the most useful and prevalent natural and agricultural sources for the development of both transgenic foods and medicinal compounds. Over the past few years, transiently gene-transfected or transgenic Forsythia and Linum plants or cell cultures have been shown to be promising platforms for the sustainable and efficient production of beneficial lignans. In this chapter, we present the essential knowledge and recent advances regarding metabolic engineering of lignans based on their biosynthetic pathways and biological activities and the perspectives in lignan production via metabolic engineering.


Lignan Biosynthesis Metabolic engineering Transgenic plant, Forsythia, Linum 



Cinnamylalcohol dehydrogenase


Cinnamoyl-CoA reductase


Dirigent protein


Estrogen receptor


Mitogen-activated protein kinase


Methyl jasmonate


Matairesinol O-methyltransferase


Phenylalanine ammonialyase


Pinoresinol-lariciresinol/isoflavone/phenylcoumaran benzylic ether reductase


Pinoresinol-lariciresinol reductase




RNA interference


Salicylic acid


Secoisolariciresinol diglucoside


Secoisolariciresinol dehydrogenase

1 Introduction

Functional foods, dietary supplements, and drug compounds are largely derived from specialized metabolites, previously called secondary metabolites of plants, including alkaloids, flavonoids, isoflavonoids, and lignans. Recently, the rapid increase in the number of elderly individuals has required various medical costs, which may eventually cause a serious disruption in essential medical care systems and national financial burdens. To address these issues, the consistent and appropriate intake of dietary supplements and the efficient development of clinical drugs are the most promising and effective ways to increase the healthy life expectancy and prevent lifestyle-related diseases. In this context, intensive efforts should be made on the development of functional food s and supplements as well as of clinical agents.

Lignan s are naturally occurring phenylpropanoid dimers (C6-C3 unit; e.g., coniferyl alcohol), in which the phenylpropane units are linked by the central carbons of the side chains (Fig. 1). These specialized metabolites are classified into eight groups based on their structural patterns, including their carbon skeletons, the way in which oxygen is incorporated into the skeletons, and the cyclization pattern: furofuran, furan, dibenzylbutane, dibenzylbutyrolactone, aryltetralin, arylnaphthalene, dibenzocyclooctadiene, and dibenzylbutyrolactol [1, 2]. Lignans have been shown to exhibit not only various pharmaceutical activities but also preventive or reductive effects on extensive life-related diseases (see the following section) [3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15], indicating the prominent potentials as functional foods and supplements. Indeed, a sesame lignan, sesamin, is already commercially available as functional supplements for antihypertension and protection of the liver based on reduction of lipid oxidation. Unfortunately, plant sources of lignans are frequently limited because of the high cost of plant hunting and collection, poor cultivation systems, long growth phase, and the low lignan content. For instance, sesamin is extracted from sesame seed oil, the most abundant source of this compound. Nevertheless, sesamin at most constitutes 0.4–0.6 % (w/w) of sesame seed oil. Moreover, sesame seeds are cultivated only once per year, limiting the ability to obtain large amounts of this compound. Likewise, podophyllotoxin (PTOX), a precursor of semisynthetic antitumor drugs (Fig. 1), is isolated from the roots and rhizomes of Podophyllum hexandrum, which is distributed in very limited regions, and is now endangered due to overharvesting and environmental disruption [16]. Moreover, lignans and their precursors are not or faintly biosynthesized in prevalent model plants such as Arabidopsis thaliana and Nicotiana tabacum, given that lignan biosynthetic pathways for plant specialized metabolites involve multiple enzymatic steps that are absent in most plant species including these model plants. Thus, transformation of a whole set of the biosynthetic genes would be required for the generation of transgenic model plants that could produce phytochemicals, indicating the critical shortcomings of these plants as biological lignan producers [12, 13, 17]. Similarly, transgenic microbial or animal cells are also not suitable for any biological lignan producers. In addition, the complicated chemical structures of lignans and the related compounds (Fig. 1a) make stereoselective organic synthesis impractical and costly for lignan producing large supplies of these compounds. These drawbacks indicate the requirement for efficient, stable, and sustainable systems for lignan production using lignan-rich plants or its precursor compound-rich plants.
Fig. 1

Chemical structures of typical lignans in dietary and medicinal sources (a) and synthetic podophyllotoxin derivatives (b)

There has been a growing body of reports on the molecular characterization of the enzymes involved in the biosynthesis of lignans, lignan production using lignan-rich plants or cultured plant cells [18, 19, 20, 21, 22, 23]. These recent findings have allowed us to attempt the metabolic engineering of lignan biosynthesis using transgenic lignin-rich plants such as Linum and Forsythia . In this chapter, we provide current knowledge of lignan production via metabolic engineering and perspectives in the development of metabolic engineering-based lignan production.

2 Lignan Biological Activity on Mammals

Lignans exhibit a wide variety of bioactivities on plants, insects, and mammals [12, 13, 24, 25, 26, 27, 28, 29], but special attentions have been paid to their unique antitumor-associated activities and reduction of the risks of lifestyle-related diseases. The modes of actions of lignans on mammals are classified in two ways: the pharmacological actions of specific metabolites of lignans by intestinal microflora and those of intact lignans. Many of lignans and their glycosides, including pinoresiniol, sesamin, lariciresinol, secoisolariciresinol, and matairesinol, are metabolized by intestinal microflora to yield enterodiol and enterolactone, which are defined as enterolignan s or mammalian lignan s [30, 31, 32]. These lignan metabolites are believed to elicit the modest estrogen (mammalian female steroid hormone)-like activity in mammals. For example, enterolignans bind to the mammalian estrogen receptors (ER), ERα or ERβ, which are key regulatory nuclear receptors in the sexual maturation of genital organs [33, 34]. Consequently, enterolignans, combined with other intestinal flora generating metabolites of isoflavones and coumestans, are also called phytoestrogen s.

Intact lignans have also been detected in the sera of mammals fed with lignan-rich diets, suggesting that nonmetabolized lignans are also taken up by the mammalian digestive system, and exhibit ER-independent activities in vivo and in vitro, including tumor growth suppression , angiogenesis inhibition, and reduction of diabetes [6, 35, 36, 37, 38, 39, 40]. Furthermore, lignans have been shown to manifest positive effects on other lifestyle-related diseases. Administration of flaxseed lignan complexes improved hyperglycemia and reduced markers of type II diabetes in elderly patients and various animal models [41, 42]. In particular, secoisolariciresinol diglucoside s (SDG), secoisolariciresinol, enterodiol, and enterolactone inhibited pancreatic α-amidase activity in a noncompetitive manner [43]. Sesamin and its metabolites also exhibited antihypertensive activities [44, 45, 46]. Moreover, the antioxidative activity of sesamin is believed to be involved in protecting the liver from oxidation by alcohols, lipids, and oxygen radicals [44, 47, 48, 49]. In human intestinal Caco 2 cells, pinoresinol suppressed expression of Cox-2, an inducible prostaglandin synthase that is responsible for the synthesis of prostaglandin H, a precursor of any other prostaglandins, leading to the decrease in the production of inflammatory factors, such as interleukin-6 and prostaglandin E2 [35]. In contrast, matairesinol increased levels of prostaglandin E2 [35]. These findings proved that pinoresinol and matairesinol have opposite activities in these cells [35].

Of the most prominent epidemiological significance is that intake of lignan-rich foods, such as flaxseeds and sesame seeds , has been shown to reduce breast cancer risk and to improve the breast cancer-specific survival of postmenopausal women [39, 50, 51, 52, 53, 54, 55]. Moreover, serum enterolactone levels were positively correlated with the improvement of prognosis in postmenopausal women with breast cancer [56]. These epidemiological studies suggest the unique and beneficial suppressive activity of lignans against breast cancer risks in elderly women.

Dietary lariciresinol was found to suppress tumor growth and angiogenesis in nude mice implanted with human MCF-7 breast cancer via the induction of apoptosis and the upregulation of ERβ expression [40]. SDG elicited potent inhibition of cell proliferation and induction of the apoptosis of breast cancer cells via the downregulation of ER- and growth factor-mediated gene expression in athymic mice [57]. Sesamin was found to reduce signaling downstream of mitogen-activated protein kinase (MAPK) [58]. Additionally, the inhibitory effect of sesamin on breast tumor growth is likely to be more potent than SDG [58]. These pharmacological effects, combined with the abundance of lignans in flax or sesame seeds and oils, confirm that the seeds and oils are promising functional diets for the prevention of breast cancer.

PTOX and its structurally related natural derivatives exhibit the suppressive activity on mitotic spindle assembly by binding to tubulin, resulting in cell cycle arrest at metaphase [22]. The PTOX semi-synthetic derivatives, etoposide, teniposide, and etopophos (Fig. 1b), are clinically utilized to treat certain types of cancers, including testicular/small-cell lung cancer, acute leukemia, Hodgkin’s and non-Hodgkin’s lymphoma [58, 59]. These PTOX-derived antitumor drugs induce apoptosis of tumor cells by binding to topoisimease II, a key enzyme for cell division [58, 59]. In addition, other new synthetic PTOX derivatives, including GP-11, NK-611, TOP-53, GL-331, and NPF, are undergoing phase I or II clinical trials as novel cancer drugs [22, 59]. Consistent with the difficulty in efficient chemical synthesis of PTOX due to its complicated structure, these findings reinforce the importance of PTOX as a natural seed material for the production of various anticancer drug s.

Altogether, these epidemiological and physiological studies demonstrate that lignans exert beneficial effects as dietary compounds or medicinal agents for the prevention of lifestyle-related diseases, such as cancer, hypertension, and diabetes. Of particular interest is that respective lignans exhibit specific bioactivities in mammals, strongly suggesting the requirements for the efficient, stable, and sustainable production of these compounds of interest. In other words, these findings not only endorse the high usefulness of lignin-rich sesame and flax seeds as unique functional foods but also shed light on the importance of the development of novel lignan production systems using transgenic lignan-rich plants.

3 Lignan Biosynthesis Pathways

Two major lignan biosynthesis pathways have thus far been identified. Both of the pathways originate from the coupling of achiral E-coniferyl alcohol, leading to the generation of pinoresinol, a basal lignan (Fig. 2). A pinoresinol synthase has yet to be identified. However, a dirigent protein (DIR) was shown to participate in the stereo-specific dimerization of E-coniferyl alcohol [60]. In several plants including Sesamum, pinoresinol is metabolized into piperitol, followed by further conversion into (+)-sesamin by a cytochrome P450 family enzyme, CYP81Q1 , which is responsible for the formation of two methylenedioxy bridges [61]. The CYP81Q1 gene is expressed almost exclusively in sesame seeds, which is compatible with sesamin production at the highest level in sesame seeds [61]. Sesamin is anticipated to be further metabolized into sesaminol and sesamolin (Fig. 2), although the relevant enzymes remain to be characterized. Sesaminol is glucosylated at its 2-hydroxyl group by the homologous enzymes UGT71A8 (S. radiatum), 9 (S. indicium), and 10 (S. alatum) [62]. Moreover, the resultant sesaminol 2-O-monoglucoside is further glucosylated by UGT94D1 , which is specific to the glucosylation of sesaminol 2-O-monoglucoside at 6-position of the conjugated glucose conjugated by UGT71A18 [62].
Fig. 2

Biosynthesis pathways of major lignans. Chemical conversions at each step are indicated in red. Solid and broken lines represent identified and unidentified enzyme-catalyzed reactions, respectively

No genes homologous to CYP81Q1 have been detected in diverse lignan-rich plant species including Forsythia, Linum, or Podophyllum [63, 64, 65, 66, 67, 68, 69]. This is in good agreement to the findings that these plants fail to biosynthesize sesamin and its derivatives. Instead, pinoresinol is stepwisely reduced to lariciresinol and then secoisolariciresinol by pinoresinol-lariciresinol reductase (PLR ), a member of the pinoresinol-lariciresinol/isoflavone/phenylcoumaran benzylic ether reductase (PIP) family in extensive plant species including Forsythia, Linum, and Podophyllum [70, 71, 72, 73, 74, 75]. PLR converts pinoresinol to secoisolariciresinol via lariciresinol (Fig. 2). Pinoresinol also undergoes glucosylation by UGT71A18, a UDP-glucose-dependent glucosyltranferase [76]. Such glycosylation is highly likely to suppress the chemical reactivity of a phenolic hydroxyl group of pinoresinol and to potentiate high water solubility of pinoresinol aglycone, resulting in large and stable amounts of pinoresinol [1, 2, 11, 12]. Indeed, approximately 90 % of pinoresinol is accumulated in its glucosylated form in Forsythia spp. [77, 78]. Thus, PLR-catalyzed metabolism and UGT71A18-directed glucosylation are reciprocally competitive pathways (Fig. 2), given that both of them share pinoresinol as a substrate. PLR shows opposite seasonal alteration in gene expression against UGT71A18; in Forsythia leaves in Japan, PLR gene is intensely expressed from April to August but poorly from September to November, whereas gene expression of UGT71A18 is observed at high level from September to November but at faint or no level from April to August in Japan [78]. These findings indicate that PLR and UGT71A18 participate in the competitive regulation of lignan biosynthesis via pinoresinol metabolism. In A. thaliana, AtPrR1 and 2 are only responsible for the reduction of pinoresinol to lariciresinol [74], and lariciresinol and pinoresinol are glucosylated by another novel UDP-glucose-dependent glucosyltranferase, UGT71C1 [79], revealing the diversity of lignan metabolism among plant species.

Secoisolariciresinol, like pinoresinol and lariciresinol, undergoes two metabolic pathways (Fig. 2). First, Secoisolariciresinol is converted into matairesinol by secoisolariciresinol dehydrogenase (SIRD ) [80]. Second, a novel UDP-glucose-dependent glucosyltranferase in Linum, UGT74S1, generates secoisolariciresinol monoglucoside and SDG [81]. Matairesinol is metabolized to arctigenin (Fig. 2) by matairesinol O-methytransferase (MOMT ) via methylation of a phenolic hydroxyl group in various plants including F. koreana, Carthamus tinctorius, and Anthriscus sylvestris [82, 83]. Additionally, 70–90 % of matairesinol is glucosylated throughout the year in the Forsythia leaves [78], although characterization of matairesinol-glucosylating enzymes awaits further study. As shown in Fig. 2, the biosynthetic pathways downstream of matairesinol are complexed and relatively species-specific. In Linum, Anthriscus, and Podophyllum plants, matairesinol is also converted into hinokinin, yatein, or PTOX via multiple biosynthetic pathways [1, 2, 12, 13, 60]. In A. sylvestris, AsTJOMT exclusively methylates the 5-hydroxyl group of thujaplicatin, an intermediate of the PTOX biosynthesis pathway [84].

The homologous enzymes, CYP719A23 (from P. hexandrum) and CYP719A24 (from P. peltatum) participate in the conversion of matairesinol into pluviatolide, a more downstream intermediate of PTOX (Fig. 2), via methylenedioxy bridge formation [63, 64]. Quite recently, six novel genes, which were also detected by NGS-based transcriptome, have been characterized from P. hexandrum and shown to be responsible for the PTOX biosynthesis [23]. CYP71CU1 was found to hydroxydise (−)-5′-desmethoxy-yatein into (−)-5′-desmethyl-yatein followed by O–methylation by OMT1 to (−)-yatein (Fig. 2). (−)- yatein is converted into (−)-deoxy-podophyllotoxin, which is demethylated to (−)-4′-desmethyl-deoxy-podophyllotoxin by CYP71BE54 (Fig. 2). CYP82D61 was shown to participate in the production of (−)-4′-desmethyl-epipodophyllotoxin via hydroxylation of (−)-4′-desmethyl-deoxy-podophyllotoxin (Fig. 2). Notably, (−)-4′-desmethyl-epipodophyllotoxin, which is an aglycone of an antitumor drug, etoposide, was detected in transgenic tobacco transected with these six genes [23]. Taken into account that (−)-4′-desmethyl-epipodophyllotoxin is synthesized from PTOX in the industrial production of etoposide, this study leads to the development of the novel procedure for the production etoposide using transgenic tobacco as well as explored total biosynthesis pathways of PTOX and its related lignans [23].

Over the past few years, the genomes or transcriptomes of lignan-rich plants including Linum [68, 85, 86], Sesamum [65, 66, 67], and Podophyllum [23, 63, 64, 69] have been documented, followed by in silico detection of functional genes. Particularly, next-generation sequencers (NGS)-based de novo transcriptome has been shown to be a powerful procedure for molecular characterization of lignan-biosynthetic genes at the first step, as described above. These findings are highly likely to remarkably enhance the molecular and functional characterization of unknown lignan biosynthetic enzymes. In addition, it is suggested that a Podophyllum endophyte may produce PTOX [87]. NGS analyses of the genome, metagenome, and transcriptome of Podophyllum and its endophytes are expected to provide crucial clues to understanding the PTOX biosynthesis pathways.

4 Metabolic Engineering of Lignan Biosynthesis

A growing body of studies has revealed that lignan biosynthesis is altered by genetic modification, light, and elicitors. This section presents an overview and discussion of recent progress in major lignan metabolic engineering using plants, plant cells, and organ cultures.

4.1 Metabolic Engineering of Transgenic Plants and Cells

Stable transfection or gene silencing , namely authentic transgenic metabolic engineering of a lignan biosynthetic enzyme gene is expected to directly alter the lignan production cascades in host plants, organs, and cells. To date, Forsythia and Linum cell, organ cultures, and plants are attempted to generate the transgenic plants among lignan-rich plants. Agrobacterium-based gene introduction was employed for transformation of both Forsythia and Linum, which is also essentially common to generation of transgenic model plants [17, 77, 88, 89, 90, 91, 92, 93, 94]. Figure 3 demonstrates the typical procedure for Agrobacterium -based transformation of Forsythia. In the subsection, we present the recent progress in transformation of Forsythia and Linum and metabolic engineering of lignan biosynthetic pathways using these plants.
Fig. 3

Scheme for generation of transgenic Forsythia mediated by Agrobacterium. This process is common for the generation of transgenic Forsythia plants and suspension cultures

4.1.1 Transgenic Forsythia Cells

Forsythia is a perennial plant commonly known as the golden bell flower and is used for a variety of Chinese medicines and health diets [1, 2, 5, 7, 12, 13]. As shown in Fig. 2, Forsythia biosynthesizes pinoresinol, lariciresinol, secoisolariciresinol, matairesinol, and arctigenin, with >90 % of pinoresinol, >80 % of matairesinol, and 40–80 % of arctigenin accumulated in glucosylated forms [12, 13, 17, 77, 78]. Seasonal changes in amounts of major Forsythia lignans and the relevant gene expression were also reported. All of the lignans in the leaf continuously increased from April to June, reached the maximal level in June, and then decreased [78]. PLR was stably expressed from April to August, whereas the PLR expression was not detected from September to November [78]. In contrast, the UGT71A18 expression was detected from August to November but not from April to July. The SIRD expression was prominent from April to May, not detected in June to July, and then increased again from September to November [78]. These expression profiles of the lignan-synthetic enzymes are essentially correlated with the alteration in lignan amounts.

Several transgenic Forsythia plants and cells have been documented for the past 5 years [17, 77, 89]. It is noteworthy that the regeneration efficiency of callus (shoot formation and rooting) and optimal condition for them differ among the variety of Forsythia species (F. koreana, F. intermedia, and F. suspense) (Fig. 4). For instance, F. koreana , F. intermedia , and F. suspense explants regenerated more than 100, 36, and 4 shoots per leaf, respectively [89]. Likewise, F. intermedia calli, unlike F. koreana, calli, were much more effectively regenerated on the F0 medium than on the FM0 medium [89]. Two transgenic F. intermedia and one transgenic F. koreana have acquired hygromycin resistance, but none of them have exhibited metabolic alteration in lignan biosynthesis [89]. Moreover, the greatest drawback in generation of transgenic Forsythia lies in extremely low transformation efficiencies. These findings strongly suggest the potential requirement for innovation of transgenic Forsythia plant generation. In other words, a high-efficient transgenic method for Forsythia is expected to remarkably enhance transgenic metabolic engineering-based lignan production using transgenic Forsythia.
Fig. 4

Different conditions in regeneration between Forsythia varieties (F. koreana and F. intermedia). Note that different media is used for regeneration of the respective Forsythia species. Culturing periods also vary between these species

The transgenic metabolic engineering of Forsythia culture cells was originally reported in 2009. F. koreana suspension cells stably transfected with a PLR-RNA interference (RNAi) sequence ( PLR-RNAi ) showed complete loss of matairesinol and an approximately 20-fold increase in total pinoresinol (pinoresinol aglycone and glucoside), compared with the wild type cells [77]. Furthermore, Forsythia transgenic cells CPi-Fk, which are stably double transfected with PLR-RNAi and the sesamin-producing enzyme CYP81Q1 (Fig. 2), produced sesamin (0.01 mg/g dry weight of the cell [DW]) (Fig. 5), although sesamin is not biosynthesized in native Forsythia [77]. This is the first success in the metabolic engineering leading to an exogenous lignan using transgenic plant cells, demonstrating that the Forsythia cell culture system is an efficient and promising platform for producing both endogenous and exogenous lignans by transgenic metabolic engineering. A striking feature is that light irradiation has been shown to improve the production of both endogenous and exogenous lignans by CPi-Fk cells. Irradiation of CPi-Fk cells for 2 weeks with white fluorescent, blue LED , and red LED light increased sesamin production 2.3-fold, 2.7-fold, and 1.6-fold, respectively, compared with cells cultured in the dark [94]. Likewise, irradiation of CPi-Fk cells increased pinoresinol (aglycone and glucoside) production 1.5- to 3.0-fold [94]. Intriguingly, expression of the pinoresinol-glucosylating enzyme UGT71A18 was also downregulated in CPi-Fk cells under blue LED or red LED light [94]. This reduction of the expression of UGT71A18 is also likely to contribute to the increase of sesamin production [94], given that pinoresinol glucoside is not metabolized into sesamin by CYP81Q1 [12, 75], and 90 % of pinoresinol is glucosylated in Forsythia wildtype cells [12, 13, 17, 77, 78]. In other words, these findings suggested that suppression of UGT71A18 by RNAi might contribute to an increase in productivity of pinoresinol and sesamin in CPi-Fk. This presumption was substantiated in our subsequent study. Quite recently, we have created more efficient, stable, and sustainable sesamin production system using triple-transgenic Forsythia koreana cell suspension cultures, U18i-CPi-Fk , compared to CPi-Fk [17]. These transgenic cells were generated by stable transfection of CPi-Fk with an RNAi sequence against the pinoresinol-glucosylating enzyme UGT71A18. UGT71A18 expression was not detected in the triple-transgenic Forsythia cells [17]. Moreover, U18i-CPi-Fk accumulated approximately fivefold higher amounts of pinoresinol aglycone than CPi-Fk, and the ratio of pinoresinol aglycone to total pinoresinol in U18i-CPi-Fk is 81.81 ± 6.43 %, which is approximately 6.5-fold greater than that in CPi-Fk (13.19 ± 2.35 %). These results proved that UGT71A18-RNAi contributed a great deal to the increase in the ratio of pinoresinol aglycone to total pinoresinol. Notably, U18i-CPi-Fk has also been shown to display 1.4-fold higher production of sesamin than CPi-Fk [17], confirming that the suppression of UGT71A18 gene expression is effective for improvement of the sesamin production. Furthermore, pinoresinol aglycone was 3.4-fold and 2.8-fold greater produced under white fluorescent and red LED, respectively, than under the dark condition. Consistently, sesamin production in U18i-CPi-Fk was approximately threefold (31.02 ± 3.45 μg/g DW) upregulated specifically under red LED, whereas white fluorescent or blue light failed to affect sesamin production [17]. It should be noteworthy that the light types effective for the sesamin production differed between CPi-Fk and U18i-CPi-Fk; the sesamin production was potentiated exclusively by blue LED light in CPi-Fk [94], whereas red LED light upregulated the sesamin production in U18i-CPi-Fk [17]. The molecular mechanism underlying such different sensitivity of these transgenic Forsythia cells remains unclear, but the suppression of UGT71A18 gene is likely to alter other biosynthetic pathways than pinoresinol glucosylation, which ultimately may affect light sensitivity of the sesamin production. In addition, upregulation of lignan production by light was also observed in other natural plants or cells. In Linum species, suspension of L. album cells produced twofold more PTOX under red light than those in the dark [95]. Irradiation of S. indicum leaves 3–5 weeks after sowing with blue LED light increased sesamin content twofold, compared with white fluorescent light, whereas irradiation with red LED light reduced sesamin content twofold [9, 96]. In combination, these findings highlight the different specificity of light types to lignan production among lignan compounds and host plant species.
Fig. 5

Metabolic engineering of Forsythia suspension cell cultures. The double-transgenic Forsythia suspension cells, CPi-Fk, acquired the ability to produce sesamin by stable transfection of PLR-RNAi and an exogenous (Sesamum) CYP81Q1 gene. The triple-transgenic cells, U18i-CPi-Fk, were generated by the introduction of UGT71A18-RNAi into CPi-Fk and exhibit higher productivity of pinoresinol and sesamin than CPI-Fk. The lignan productivity is approximately three- to fivefold upregulated under red LED. U18i-CPi-Fk can also be stocked in liquid nitrogen for a long period, and re-thawed U18i-CPi-Fk exhibit as high productivity of sesamin as noncryopreserved U18i-CPi-Fk

U18i-CPi-Fk has also been found to possess another prominent advantage over CPi-Fk, that is, long-term and reproducible storage [17]. Universal procedures for long-term stock of plant cell cultures, unlike those of seeds, bacteria, or animal cells, have not been well established, and cryopreservation procedures for a particular plant species are not always applicable to other plant cells [97, 98, 99]. Indeed, we failed to establish any procedure for long-term stock of CPi-Fk, and thus observed a decrease in the growth rate of CPi-Fk cells after 2 years of culture and, eventually, proliferation loss. Nevertheless, we have developed a procedure for sodium alginate-based long-term storage of U18i-CPi-Fk in liquid nitrogen [17]. Moreover, production of sesamin in U18i-CPi-Fk re-thawed after 6-month cryopreservation was equivalent to that of noncryopreserved U18i-CPi-Fk, proving the reproducible functionalities of U18i-CPi-Fk [17]. Altogether, the high lignan (pinoresinol and sesamin) productivity and establishment of the freeze stocks of U18i-CPi-Fk endorses the marked usefulness of U18i-CPi-Fk as a stable and sustainable platform of lignan production (Fig. 5).

4.1.2 Transgenic Linum

Linum spp. (flax, Linaceae) are annual flowering plants comprised of approximately 200 species. This genus has received pharmaceutical and medicinal attention due to the presence of various lignans, including PTOX and its related compounds, which are practically applied for the semisynthesis of antitumor drugs for breast and testicular cancers described above. Since Linum is also known to biosynthesize PTOX and its derivatives, and the procedures for tissue and cell culture are well established, optimal conditions and stimulating factors for production of various lignans, including (−)-podophyllotoxin, by Linum calli, suspension cell cultures, and roots have been extensively investigated as described in the followings. Recently, flax seeds and oils have also received attentions as functional foods due to the contents of lignans beneficial for human health [14, 15]. Accordingly, metabolic engineering of Linum is also highly likely to contribute a great deal to the development of novel lignan production and transgenic foods.

In various Linum species, the effects of RNAi on production of endogenous lignans via metabolic engineering were investigated. PLR-RNAi- transgenic plants of L. usitatissimum showed the high accumulation of pinoresinol diglucoside and loss of SDG in the seed coat [90]. Intriguingly, these PLR-RNAi-transgenic L. usitatissimum plants produced the 8–5′ linked neolignans, dehydrodiconifnyl alcohol and dihydro-dehydrodiconifnyl alcohol, while these neolignan s were not biosynthesized in the wildtype plants [90]. These findings indicate that RNAi occasionally affects some biosynthetic pathways in an indirect fashion.

4.1.3 Transient Transformation of Linum

Hairy roots of L. perenne transiently transfected with PLR-RNAi reduced the production of the major endogenous lignan, justicidin B, to 25 %, compared with the untreated hairy roots [72]. Likewise, transient transfection of L. corymbulosum hairy roots with PLR-RNAi resulted in a marked reduction of hinokinin [73]. Combined with the justicidin B and hinokinin biosynthetic pathways, in which PLR converts pinoresinol into secoisolariciresinol (Fig. 2), these findings indicate that PLR-directed conversion of pinoresinol into secoisolariciresinol is a rate-limiting step of justicidin B and hinokinin biosynthesis, at least in the hairy roots of L. perenn and L. corymbulosum, respectively. Therefore, identification and genetic manipulation of justicidin B and hinokinin synthase will contribute a great deal to the establishment of procedures for the direct metabolic engineering of these lignans. Taken together, these findings reinforce the potential of Forsythia and Linum transgenic or transiently gene-transfected cells and plants as the metabolic engineering-based platforms for on-demand production of both endogenous and exogenous lignans. The draft genome and transcriptome of L. usitatissimum [68, 85, 86] will also accelerate the identification of the enzymes involved in the biosynthesis of Linum lignans, leading to the efficient lignan production using gene-modified plant sources.

4.1.4 What Should Be Considered for Lignan Production via Metabolic Engineering Using Gene-Modified Plants?

To establish gene-modified plant platforms for lignan production, we should consider two crucial factors: the type of host and the use of transgenic or transiently transfected hosts. Host types can be classified into plants, organs, and cell cultures. For example, although the amount of sesamin produced by U18i-CPi-Fk cells is ~100-fold lower than that by native sesame seeds, U18i-CPi-Fk-based lignan metabolic engineering has several advantages. Furthermore, the Forsythia transgenic cells are propagated tenfold in 2 weeks in standard culture medium [17] and can be cultivated at all times and locations which is also endorsed by the fact that U18i-CPi-Fk can be stocked for a long time [17]. In contrast, sesame seeds are cultivated in limited regions only once a year. Moreover, the conditions used in the culturing of U18i-CPi-Fk cells, including temperature, light wavelength and intensity, and medium components, can be altered to optimize sesamin production [17, 77]. Forsythia plants have much greater biomass, with higher amount of lignans, than suspension cell cultures, and these plants can grow and propagate from small explants without flowering or seed formation [89]. However, efficient generation of transgenic Forsythia plants still requires further basic research due to the markedly low transformation efficiency by any known gene transfection methods [89]. In addition, as mentioned above, the optimal culturing and regeneration conditions were found to vary among Forsythia species [89]. In other words, the development of a procedure for efficient generation of transgenic Forsythia would surely enhance novel lignan production system.

The generation of both stable (namely transgenic) and transient transfectants of Linum species are well established [72, 73, 90, 93], and thus the amounts of precursors or intermediates of targeted lignans are major determinants for the employment of cell cultures, organ cultures, or plants as host platforms. Additionally, gene-modified host plants may fail to normally grow or to produce lignans of interest due to cytotoxicity of lignans, although the underlying molecular mechanisms have not fully been elucidated. Therefore, generation of lignan-producing plants using multiple plant species is occasionally required.

The second factor involves construction of either transgenic or transiently transfected hosts. Transgenic plant s and cell cultures, once generated, are sustainably used for lignan production and readily upscaled, whereas generation of transgenic plants, in particular nonmodel plants, may be time- and cost-consuming. Moreover, cultivation of transgenic plants in general requires a closed facility for transgenic plants. Transiently transfected plants require repeated transfections, and transient transfection of multiple genes may dramatically decrease the transfection efficiency. Furthermore, massive transient transfection methods for industrial use remains to be fully developed [100]. Further research on lignan metabolic engineering, using transgenic or transiently gene-transfect ed plants, organ cultures, and cell cultures, is expected to lead to the establishment of both universal and molecular species-specific strategies for gene-regulated metabolic engineering of lignan biosynthesis pathways.

4.2 Metabolic Engineering by Elicitation

Plant defense systems are triggered upon injury or infection via signaling by the phytohormones, methyl jasmonate (MeJA) and salicylic acid (SA), and treatment with elicitors, including fungi, their extracts, and the glycan components, MeJA and SA, also mimic such activation. Moreover, lignans, at least in part, are likely to be implicated with host defense systems [12, 13, 22, 101]. In combination, elicitor s are expected to enhance lignan biosynthesis [13, 22, 102]. As summarized in Table 1, the effects of various elicitors on lignan production have been examined in a wide variety of cell cultures and hairy roots of Forsythia, Juniperus, and Podophyllum (Table 1).
Table 1

List of major elicitors and their effects on lignan biosynthesis





Chito-oligosaccharides (1 mg)

Juniperus chinensis callus culture

Increased PTOX production


Methyl jasmonate (MeJA) (100 μM)

Forsythia intermedia cell suspension culture

Increased pinoresinol and matairesinol production


Mannan (0.1 mg mL−1)

β-1,3-glucan (0.1 mg mL−1)

Ancymidol (10−7 M)

L. austriacum callus culture

Enhanced activity of tyrosine ammonia-lyase (TAL), coumarate 3-hydroxylase (C3H), polyphenoloxidase (PPO) and PAL

Increased PTOX,



α- and β-peltatins production

Increased PTOX and α-peltatins production

Increased PTOX, 6-MPTOX,

dPTOX and α- peltatins production


Indanoyl-isoleucine (5–100 μM)

Coronalon, (10–50 μM)

MeJA(100 μM)

L. nodiflorum cell suspension culture

Increased deoxypodophyllotoxin production

Enhanced activity of 6-hydroxylase and β-peltatin 6-O-methyltransferase,

Increased 6-MPTOX and 5′-d-6-MPTOX production


MeJA (100 μM)

L. album cell suspension culture

Increased PTOX production


Botrytis cinerea extract (3 % v/v)

Phoma exigua extract (3 % v/v)

Fusarium oxysporum extract (3 % v/v)

L. usitatissimum cell suspension culture

Rapid stimulation of the monolignol pathway, enhanced PAL activity, and expression of genes encoding PAL, CCR, and CAD


MeJA(50–200 μM)

L. tauricum hairy root culture

Increased 6MPTOX and 4′-DM6MPTOX production


Salicylic acid (SA)(10 μM)

L. album cell suspension culture

Enhanced PAL, CCR, and CAD gene expression and PTOX production


Chitin (100 mg l−1)

Chitosan (100–200 mg l−1)

MeJA(100–200 μM)

L. album cell suspension culture

Increased lariciresinol and/or PTOX production


Fusarium graminearum extract(1 %v/v)

Sclerotinia sclerotiorum extract(1 %v/v)

Rhizopus stolonifer extract(1 % v/v)

Rhizoctonia solani extract(1 % v/v)

L. album cell suspension culture

Enhanced PAL, CCR, CAD, and PLR gene expression

Increased PTOX and lariciresinol production

[105, 109]

MeJA(10–100 μM)

Podophyllum hexandrum cell suspension culture

Changes in cell proteome Increased PTOX production


Fusarium graminearum extract (1 %v/v)

Sclerotinia sclerotiorum extract(1 %v/v)

Trichoderma viride extract (1 %v/v)

Chitosan (100 mg l−1)

L. album hairy root culture

Enhanced PAL, CCR, CAD, and PLR gene expression

Increased PTOX, 6MPTOX, and lariciresinol production


Chitosan and chitin oligomers (100 mg l−1)

L. album cell suspension culture

Enhanced PAL, CCR, CAD, and PLR gene expression

Increased PTOX, 6MPTOX, and lariciresinol production


Fusarium graminearum culture filtrate (1 % v/v)

L. album cell suspension culture

Increased phenolic compound, PTOX, and lariciresinol production

Enhanced PAL activity


MeJA and SA were found to increase the production of PTOX and the structurally related lignan production or the gene expression of lignan biosynthetic enzymes responsible for biosynthesis of conifenyl alcohol, phenylalanine ammonialyase (PAL ), cinnamoyl-CoA reductase (CCR ), and cinnamylalcohol dehydrogenase (CAD ) in cell suspension cultures of L. album [103, 104] and L. nodiflorum [103], Podophyllum hexandrum [105], and callus of L. austriacum callus culture [106]. These phytohormones also increased the PTOX production or the relevant gene expression in hairy roots of L. tauricum [107]. Additionally, an increase in production of pinoresinol and matairesinol by MeJA was observed in Forsythia intermedia cell suspension culture [108].

Chitosan, chitin oligomers, and other glycans also enhanced PTOX production or gene expression of lignan biosynthetic enzymes in Juniperus chinensis callus culture [109], L. austriacum callus culture [106], and L. album cell suspension culture and hairy roots [110, 111, 112]. In particular, comparisons of chitin tetramer, pentamer, and hexamer and chitosan tetramer and pentamer showed that treatment of L. album hairy roots with chitosan hexamer for 5 days most potently enhanced PTOX and lariciresinol production, as well as upregulating the expression of PAL, CCR, CAD, and PLR genes [112]. In summary, treatment with these elicitors resulted in two- to sevenfold increase in PTOX synthesis and expression of genes encoding enzymes involved in the early steps of lignan biosynthesis in various plant cells and hairy roots.

Fungal co-culturing, extracts, and filtrate exhibited unique effects on the metabolic engineering of lignan production (Table 1). Botrytis cinerea, Phoma exigua, and Fusarium oxysporum extracts triggered the accumulation of monolignols and enhanced PAL activity and gene expression of PAL, CCR, and CAD in L. usitatissimum cell suspension cultures [113]. Treatment of L. album cell cultures with Fusarium graminearum extract for 5 days increased PTOX sevenfold and PAL, CCR, and CAD mRNAs > tenfold compared with untreated cells. These results confirmed that this extract is a more potent elicitor of PTOX production and PAL, CCR, and CAD expression than treatment with chitosan, chitin, or MeJA treatment for 3 days [110, 111, 114].

Rhizopus stolonifer and Rhizoctonia solani extract stimulated 8.8-fold and 6.7-fold greater accumulation of lariciresinol, instead of PTOX, in L. album cell cultures after 5-days treatment as compared with untreated cells, and the highest (6.5-fold) PLR gene induction was observed in L. album cell cultures treated with Rhizopus stolonifer extract for 2 days [114]. Similar data were obtained in L. album hairy roots with the same fungal extracts [111] or L. album cell suspension culture with Fusarium graminearum culture filtrate [115], but the latter manifested less lignan production. These studies revealed that fungal extract exhibited the species-specific effects on the lignan biosynthesis pathways, although investigation of the molecular basis awaits further study.

As described above, the regulation of gene expression has thus far been restricted to enzymes responsible for the upstream of lignan biosynthesis pathways. Therefore, the effects of these elicitors on lignans and the relevant biosynthetic genes downstream of PLR, such as SIRD or 719A23 (Fig. 2), would provide a clue to understanding the molecular mechanisms underlying upregulation of PTOX production and to identifying more effective elicitors for lignan production.

5 Conclusion

In this chapter, we have provided diverse recent advances in metabolic engineering for lignan production by plants, including: (i) the molecular characterization of novel genes encoding enzymes for biosynthesis pathways of dietary and medicinal lignans; (ii) the production of both endogenous and exogenous lignans by transient or stable transfection of lignan biosynthetic genes into cultured cells, tissues, and plants; (iii) the long-term stock and following reproduction of the cell functionality of a transgenic Forsythia lignan producing cells, U18i-CPI-Fk ; and (iv) the upregulation of productivity of lignans in cells and plants by exogenous stimuli such as light and elicitors in a plant species- and lignan-specific fashion. Taken together, combination of transgenesis, light, and elicitors will be a promising strategy for further improvement of the lignan productivity. For example, elicitation of U18i-CPi-Fk under red LED light is expected to increase the amounts of sesamin and/or pinoresinol. Moreover, bioinformatic integration of the aforementioned experimental data is likely to enable the systematic prediction of optimal lignan production strategy: hosts (cells, organ cultures, plants), light conditions, elicitor types, and transfection types. For example, three Forsythia varieties, F. koreana, F. intermedia, and F. suspensa, displayed differential growth and regeneration in a medium component-dependent fashion or selection marker antibiotics-dependent fashions [89], and Linum spp. showed genus-specific sensitivities to different elicitors (Table 1).

Public acceptance of transgenic dietary products is not yet sufficient all over the world. Nevertheless, it should be noted that lignans produced by transgenic hosts are chemically identical to natural ones and free from any recombinant genes or proteins. Thus, public acceptance for lignans produced by transgenic plants should also be more easily garnered than that for transgenic foods themselves. In this context, we will pay more attention to the establishment of scaling-up and following industrialization of the lignan production systems as well as the development of efficient generation of transgenic plant s in the near future [116, 117, 118]. Large-scale lignan production by transgenic plants must be carried out in a closed cultivation system to prevent contamination of the environment by transgenic plants. Recently, various closed plant factories have been emerging, which completely shut off a gene flow into the outer environment and enables the transgenic plants-based molecular breeding of genes or compounds of interest under optimal and sterile conditions [116, 117, 118]. Such advances in the metabolic engineering of lignan biosynthesis will surely pave the way for the conversion of conventional agricultural lignan production to innovative industrial production of various beneficial lignans and, ultimately, contribute a great deal to the improvement of quality of life and national financial burdens for medical care via extension of our healthy life expectancy owing to the preventive effects of lignans on diverse diseases.



This work was, in part, supported by the Plant Factory Project of the Ministry of Economy, Technology, and Industry of Japan.


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Copyright information

© Springer International Publishing Switzerland 2016

Authors and Affiliations

  1. 1.Bioorganic Research InstituteSuntory Foundation for Life SciencesSeika, KyotoJapan
  2. 2.Research InstituteSuntory Global Innovation Center (SIC) Ltd.Seika, KyotoJapan

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