Open access peer-reviewed chapter

Biosynthesis and Regulation of Antioxidant Flavonolignans in Milk Thistle

Written By

Samantha Drouet and Christophe Hano

Submitted: September 24th, 2020 Reviewed: February 24th, 2021 Published: September 8th, 2021

DOI: 10.5772/intechopen.96783

Chapter metrics overview

327 Chapter Downloads

View Full Metrics


Mature fruits (achenes) of milk thistle (Silybum marianum (L.) Gaertner, Asteraceae) accumulate high amounts of silymarin, a complex mixture of bioactive antioxidant flavonolignans deriving from taxifolin. Their biological activities in relation with human health promotion and disease prevention have been well described. The conditions of their biosynthesis in planta, however, have long been elusive and thus tend to be a limiting factor for their future applications. Significant advances in understanding their biosynthesis and regulation have been made over the last decade and are outlined in the current chapter.


  • Silybum marianum
  • flavonolignans
  • silymarin
  • biosynthesis
  • regulation

1. Introduction

Plant products, as food or as an “herbal medicine” preparation have been used by humans throughout history with varying success to prevent and/or cure various diseases [1]. Neanderthals had already understood the importance of plants in their diet but also their medicinal capacity [2]. For centuries, medicine has relied heavily on the use of plants. In many countries, traditional medicines, based on medicinal plants, are an essential part of their health system [3]. Some of these medicinal plants are still today, either collected from the wild or cultivated to ensure their availability for industry or traditional medicine. Nowadays, the badly named “weeds” such as thistles, nettle or burdock, represent a significant part of agricultural production [4]. These weeds are considered as pests, yet they are medicinal plants traditionally used for their beneficial properties. There is indeed a strong potential for the use of these plants as sources of pharmaceutical or cosmetic antioxidants [4]. In particular, the craze for silymarin from milk thistle is directly linked to the biological properties of this mixture. Silymarin is a complex mixture of bioactive flavonolignans accumulated in mature fruits (achenes) of milk thistle. Numerous biological activities, in particular hepatoprotective, anti-proliferative, immunomodulatory, anti-inflammatory and antioxidant, have demonstrated the high potential of these compounds [5, 6].

Free radicals, including superoxide radical (O2−•), hydroxyl radical (OH−•), hydrogen peroxide (H2O2) and peroxide radical (ROO−•), are implicated in liver disease [7, 8]. These reactive oxygen species (ROS) are produced through biochemical processes (cell metabolism) or induced by inflammatory diseases, cancers, or treatments such as radiotherapy or chemotherapy [9]. ROS play an important role in many signaling pathways (proliferation, cell activation, migration, etc.). However, when they are produced in large quantities in certain cellular compartments, they can become harmful to the body. This phenomenon is called “oxidative stress”. To protect themselves, cells respond by regulating the production of cellular antioxidants such as glutathione (GSH) and/or the activity of antioxidant enzymes such as superoxide dismutase (SOD) and catalase (CAT), glutathione peroxidase (GPx) [9, 10].

The abilities of silibinin (mixture of silybin A and B), which are among the main flavonolignans accumulated in S. marianum fruits, to fight certain oxidative stresses or oxidants such as superoxide anion radicals, hydrogen peroxide, hydroxyl radical and hypochlorous acid (HOCl) was studied [11, 12, 13]. The mixture of silybins does not seem to be a good O2−• scavenger and no reaction with hydrogen peroxide has been shown. On the other hand, these flavonolignans react rapidly in free solution with OH−• radicals [11]. Besides, in lymphocytes of patients with cirrhosis, in vitro incubation with silymarin markedly increases the expression of superoxide dismutase (SOD) [10]. Silymarin also helps increase the glutathione content in the cell leading to inhibition of lipid peroxidation, which enhances membrane stability [14].

The silymarin mixture also reduces oxidative stress induced by UV in epidermal and dermal cells. In certain skin conditions, it therefore exerts on humans a chemo-preventive effect on oxidative stress induced by solar UV rays (photo-aging and photo-carcinogenesis) [15, 16, 17]. Systemic and local administration of silymarin to burns is effective against oxidative damage (and morphological alterations) induced by sunburns to rat skin [18]. The study of the individual effect of some silymarin compounds has shown some differences. Other flavonolignans from milk thistle, isosilybin and silydianin, protect against glutathione depletion, ROS generation and activation of caspase-3 which plays a central role in the activation of apoptotic cell death. Both silychristin and silydianin only seem to reduce the levels of caspase-3 [19].

Hemisynthetic derivatives of silybins were found to be the effective radical scavengers and lipid peroxidation inhibitors [20, 21, 22, 23].

The biological activities in relation with human health promotion and disease prevention of the antioxidant milk thistle flavonolignans are well described. However, the conditions of their biosynthesis in plants are still unclear and have been a limiting factor for their potential developments. During the last decade, studies have paved the way toward a better understanding of the biosynthetic regulation of these flavonolignans during the maturation of S. marianum fruit. Providing important insights to better control the production of these medicinally important antioxidant compounds. This chapter summarizes these major outcomes.


2. Botanical description of Silybum marianum

Milk Thistle, S. marianum (L. Gaertner) is one of the oldest known medicinal plants belonging to the Asteraceae family. It was described and named by Carl von Linné, then replaced in the current classification by Joseph Gaertner in 1791. The term Silybum designates, in Greek and Latin, an edible thistle, as for the qualifier marianum, it would be linked to the Virgin Mary. Legend has it that during her journey from Judea to Herod’s escape, the Virgin Mary sheltered herself under a grove of thistles with the infant Jesus, where she breastfed him. Her breast milk would then have fallen on the leaves of thistles, hence the characteristic white mottling of this species. This legend could be the origin of the use of milk thistle to promote lactation, although its effectiveness in this area has never been demonstrated. Within the plant kingdom, S. marianum is an angiosperm (Magnoliophytes) commonly a flowering, fruit-bearing plant or Magnoliopsida (formerly dicotyledonous). The biological aspects of milk thistle (phenotype, life cycle) will be discussed in this section, followed by its worldwide distribution and its characteristics. Although this Asteraceae is considered a weed, it is valuable because of its interesting biological properties and antioxidant activities, in particular for applications in medicine, cosmetics, or even in the food industry.

Milk thistle is an annual or biennial herbaceous plant native to the Mediterranean basin. Due to its ecological abundance, this plant is considered as a common, even invasive species, it is nevertheless valuable. S. marianum can grow over a meter in height and diameter (at its base). Its stem is grooved, with a slightly cottony internal pith and branched at its top. This plant can be identified by its green leaves, marbled with white stripes, with an elliptical, toothed and thorny blade (Figure 1).

Figure 1.

Silybum marianum, redrawn from Bonnier and Douin, 1990 [24].

There is a plethora of thistles in the world, possessing similar morphological characteristics (Figure 2). Depending on the stage of development, it will be more or less easy to determine precisely the plant species. Indeed, many specific traits are unique to the flower or inflorescence, such as color or size. It is important to respect the “keys” of morphological identification. These taxonomic keys, based on aspects of fruits, flowers, stems, are often used as the main elements for the identification of most genera of plants [25]. The number, size, hairiness and shape of the organs often distinguish closely related species. In order not to confuse milk thistle with other species, flavonolignans could be used as taxonomic markers. It is important to have a critical look at different botanical, molecular and chemical techniques for the authentication of plant material for cosmetic applications. Molecular or phytochemical criteria are sometimes necessary, particularly when plants arrive crushed, as is customary in the industry. Identification, as well as the authentication of species, requires a wide range of technical knowledge and skills. It is easy to confuse one species with another, if only certain morphological characters, assumed to be specific to a species, are observed [25].

Figure 2.

Example of plant species that can be confused with S. marianum, all from the Asteraceae family.

S. marianum is native to southern Europe, specifically the mountains of the Mediterranean region, western Asia, and Russia, from where it has spread to most temperate regions of the world (Figure 3). This plant has also been introduced widely outside its natural range, such as North America, Japan, Iran, Australia, and New Zealand, and is now found all over the world. In some parts of the world, it is considered an invasive species, notably in Israel, Australia, Tasmania and Kansas in the USA [26]. S. marianum was introduced to the United States as an ornamental/medicinal plant and is also believed to have been introduced to the Pacific Northwest through the importation of contaminated hay. It is probably this species that Darwin calls the “giant pampas thistle” in his journal of the Voyage of the Beagle, 1831–1836 [27].

Figure 3.

Distribution of S. marianum, in green the native species and in purple the introduced species.

Speaking of the abundance of these species and their invasiveness, Darwin says, “There were immense beds of the thistle, as well as of the cardoon [i.e., Cynaria cardunculus]; the whole country, indeed, may be called one great bed. The two sorts [i.e., S. marianum and Cynaria cardunculus (artichoke and cardoon, edible and cultivated)] grow separate, each plant in company with its own kind. The cardoon is as high as a horse’s back, but the Pampas thistle is often higher than the crown of the rider’s head. To leave the road for a yard is out of the question; and the road itself is partly, and in some cases entirely closed” reported in Bulletin of the Torrey Botanical Club [27]. Milk thistle is also cultivated for industry and the production of large-scale pharmaceutical raw materials, in Austria (Waldviertel region), Germany, Hungary, Poland, China and Argentina, but also in France in the Loire Valley region.

Most medicinal plants can be harvested from the wild, but their cultivation ensures their availability for industry. Both traditional and biotechnological breeding techniques can be applied to improve yield and uniformity [4]. Conventional methods of plant breeding can improve both agronomic and medicinal characteristics [28] and in particular increase the content of active compounds. The time of harvest depends on the maturity stage of the considered crop. Flowering in flower heads is spread over time, maturation is not simultaneous throughout the plan [4], which is the general case with non-domesticated plants. In Poland, Andrzejewska and Sadowska [29] reported that the harvest of S. marianum should take place around the last days of July or early August when 40–50% of the inflorescences were in flower. After harvesting, the achene pappus should be removed with a knife and the harvested achenes should be dried at 50 °C at 8% humidity [30]. Carrier et al., [31] found that the highest silymarin content came from the late stages of development when the achenes are dark brown or even black [4]. Harvesting should be done just before the flower heads start to tear, for this an ordinary combine can be used. The large thorns present on the stems, leaves and inflorescences of milk thistle make manual harvesting of flower heads an extremely unpleasant task. However, in terms of efficiency, it is much better to harvest thistle flower heads manually rather than by machine, otherwise the yield and quality of the seeds could be compromised.

Milk thistle has been called an invasive plant because of its rapid growth and low requirement in terms of nutrients or the environment. This plant tends to grow outside its range, potentially contaminating other crop fields. S. marianum grows in compact groups and makes access to some paths or roads difficult due to its thorns. This plant can reach over 1 meter in height. This creates shadows for nearby plants (especially fodder species), helps to reduce their development, causes native vegetation to move or, in the worst case, causes them to wither. Introduced non-native thistles can invade an area, substitute for native plants, reduce crop yield or create problems for animals, when these plants infest a field or pasture. One of the problems with controlling thistles is that they are difficult to distinguish and to eradicate. Milk thistle reproduces thanks to the numerous achenes scattered in the soil. They can remain viable for up to nine years. These fruits are carried by rain, water and the movements of animals. Milk thistle can quickly invade native vegetation in natural areas. Uncontrolled, it can produce four tons of plant material every 4,000m2, leaving little room for other plants. For example, in Australia, Milk thistle was introduced as a medicinal plant. It quickly proliferated across the country, as far as Tasmania. The situation quickly became worrying. The species was quickly recognized as an invasive plant, leading to its inclusion as noxious grass in Australian and English legislation of 1851 and 1856 [32].


3. Phytochemical considerations on Silybum marianum

Milk thistle fruit extract contains 65–80% silymarin and 20–35% fatty acids, including linoleic and palmitic acid [33], but also many flavonoids (quercetin, kaempferol, apigenin, taxifolin) and phenolic compounds. Besides, around 38% of carbohydrates (mainly starch) and up to 23% of protein can be found, as well as some amines (tyramine, histamine). Silymarin is a mixture of polyphenolic molecules comprising seven closely related flavonolignans (silybin A, silybin B, isosilybin A, isosilybin B, silychristin, isosilychristin, silydianin) and a flavonoid (taxifolin).

Pelter and Haensel were the first, in 1968, to propose the generic term “flavonolignan” to designate these hybrid molecules of natural origin [34]. According to the definition recommended by the “International Union of Pure and Applied Chemistry” (IUPAC), the structure of flavonolignans is defined by the condensation of a C6C3C6 flavonoid unit with one or more C6C3 lignan precursors [35]. The precursors of lignans can be found under the general name of the unit phenylpropane (also called monolignols) which are generally in the form of hydroxycinnamic alcohol (Figure 4).

Figure 4.

Structure of a phenylpropane unit (A) and a flavonoid (B).

Silibinin, a semi-purified fraction of silymarin, is mainly a mixture of 2 diastereomers, silybin A and silybin B, in an almost equimolar ratio (close to 1:1) [36]. The silymarin content in fruits strongly depends on the selected ecotype. The geographical location of the crop (physical properties of the soil, weather conditions), as well as the agronomic conditions (sowing period, fertilization, irrigation, harvest period, seed maturity), greatly influence the composition of the flavonolignans constituting silymarin [37, 38, 39]. It is thanks to its compounds that this plant has been used for over 2,000 years in the European Pharmacopeia, making it one of the oldest medicinal plants in the Pharmacopeia.

Milk thistle is the model plant for the study of flavonolignans. The silymarin contained in the extract of milk thistle fruit, is used in diverse pharmaceutical and nutraceutical preparations. For example, Legalon® is used in the treatment of acute and chronic liver conditions. This mixture of several flavonolignans and a flavonoid account for 65–80% of the extract from the fruit of S. marianum. The constituents of silymarin are silybin, isosilybin, silychristin and silydianin [34, 40, 41, 42, 43]. These are the best known and most studied flavonolignans in the literature. These compounds are produced by the condensation of taxifolin and coniferyl alcohol.

In 1968, when the German phytochemist Hildebert Wagner and his colleagues described the mixture of flavonolignans extracted from milk thistle, only silybins, isosilybins, silydianin and silychristin had been characterized [44]. The different isomers were described later. What was initially described as a “simple” mixture of 4 molecules, actually turned out to be much more complex. In this mixture, most of the silymarin compounds are (regio)isomers and therefore contain the same number of atoms, but linked in different ways. Silybins and isosilybins are stereoisomers. These isomeric molecules differ in the three-dimensional orientations of their atoms, they are therefore diastereomers [45]. To date, by taking into account the various isomeric forms, nearly 23 flavonolignans have been identified in the genus Silybum [46]. Their structural similarity made the identification of these isomers difficult, especially by NMR, where their 1H and 13C spectra are similar, and no characteristic signal makes it possible to distinguish the three-dimensional orientations [47]. The different diastereomers were not isolated and fully characterized until 2003. These compounds were separated by column chromatography on silica gel, reverse phase HPLC, followed by recrystallization [48]. After the improvement of the various techniques and equipment, the study of silybins and isosilybins was facilitated, in particular by approaches based on chemistry, the generation of analogues, X-ray crystallography (to verify the structures of the four main isomers) and the development of tools making it possible to discern and quantify flavonolignans by 1H NMR spectroscopy, despite almost identical spectra [49, 50] (Figure 5).

Figure 5.

Chemical structure of the diastereomers of silybin and isosilybin. Complete structural assignments in C-2, C-3, C-7 ‘and C-8’ of these flavonolignans have thus been possible. The stereochemistry of these diastereomers was determined in the form of silybin A: 2R, 3R 7'R, 8'R; silybin B: 2R, 3R, 7′S, 8′S; isosilybin A: 2R, 3R, 7′R, 8′R; and isosilybin B: 2R, 3R, 7’S, 8’S [48].

Isosilybin C and isosilybin D also exist (Figure 6). The regiochemistry of these two compounds is similar to that of isosilybin A and isosilybin B. The major structural difference between these compounds and the other flavonolignans lies in their 1,3,5 substitution profile in the aromatic ring derived from the lignan precursor [47]. These two compounds are present in small quantities in the mixture of flavonolignans of milk thistle. They are therefore more rarely studied or analyzed in extracts from this plant.

Figure 6.

Chemical structure of other isosilybins present in minor amounts in milk thistle extract.

The other three silymarin flavonolignans are silychristin, isosilychristin, and silydianin. These are the structural isomers of the compounds mentioned above. Among these 3 compounds, only silychristin was characterized as two diastereoisomers (silychristin A and silychristin B) (Figure 7).

Figure 7.

Chemical structure of silychristin diastereomers.

The coupling, considered as non-stereo-selective, of taxifolin and coniferyl alcohol gives the two pairs of diastereomers (silybins and isosilybins). It is therefore not surprising to observe the existence of two diastereomers of silychristin in silymarin [51]. However, unlike silibinin (silybin A + silybin B) and isosilibinin (isosilybin A + isosilybin B) whose ratio between diastereomers varies according to the considered ecotype [52, 53], the silychristin A/silychristin B diastereomeric ratio is largely unbalanced, about 95: 5 in favor of silychristin A [51]. To date, no diastereoisomers of silydianin or isosilychristin have been reported. However, due to the low content of silychristin B, it is highly likely that a diastereomer of isosilychristin is also present but at a trace level. The presence of silydianin diastereomers cannot be excluded. However, the cyclic and compact structure of this molecule places these stereocenters close to the pre-existing stereocenters of taxifolin, which could explain the high stereoselectivity of this compound [51].


4. Flavonolignans biosynthesis in Silybum marianum

Most of the studies on flavonolignans focus on their pharmacological use and the chemistry of silymarin. In comparison, their biosynthesis remains less. Two different branches of the phenylpropanoids biosynthetic pathway are involved in the biosynthesis of their precursors, those of monolignols and flavonoids. In the case of the flavonolignans of purple milk thistle, the oxidative coupling allowing these molecules to be obtained occurs via the condensation of taxifolin and coniferyl alcohol. The reaction would take place via a radical coupling mechanism by oxidizing enzymes forming radicals, such as peroxidases or laccases, in a similar way to the formation of monolignol radicals and polymerization, during lignification [38, 45]. The synthesis would therefore involve three pathways: that of monolignols, that of flavonoids, and finally that specific to flavonolignans, which begins with the coupling of a flavonoid and a monolignol, possibly followed by intramolecular rearrangements.

Flavonolignans are produced mainly by oxidative coupling between a monolignol (generally coniferyl alcohol and a flavonoid). Their biosynthetic pathway includes a common element, p-coumaroyl-CoA, a key metabolite and precursor of multiple compounds, such as stilbenes, lignans, but also many phenolic compounds. The shikimate pathway generates the main metabolic flow of the flavonoid pathway. It provides the systemic phenylpropanoids pathway with a link with the primary metabolism in the form of l-phenylalanine [54]. The first enzyme acting on l -phenylalanine is l-phenylalanine ammonia lyase (PAL), followed by cinnamate 4-hydroxylase (C4H) and then p-coumarate-CoA ligase (4CL). These enzymes represent the first steps in the phenylpropanoid pathway and convert l-phenylalanine into a variety of secondary metabolites: lignin, lignans, coumarins, stilbenes and flavonoids [54, 55, 56].

The monolignols pathway allows the formation of various precursors, necessary for the formation of multiple phenylpropanoid compounds, such as lignin, lignans, neolignans, coumarins, flavonoids, stilbenes, etc. [57, 58, 59, 60, 61]. This pathway plays a central role in the secondary metabolism of plants [62]. Monolignols are synthesized from l-phenylalanine by the general route of phenylpropanoids. Several enzymatic steps are involved to obtain coniferyl alcohol, used in the biosynthesis of flavonolignans from purple milk thistle. Coniferyl alcohol is derived from the reduction of cinnamic acid by a NADPH-dependent reaction with coenzyme A, followed by aromatic hydroxylation and methylation (OMT). Then, the feruloyl-CoA is reduced by the enzyme cinnamoyl-CoA reductase to coniferyl aldehyde. The final step in the synthesis of coniferyl alcohol is catalyzed by cinnamyl alcohol dehydrogenase (CAD). This enzyme also catalyzes the final step in monomer biosynthesis, also synthesizing p-coumaryl alcohol and sinapyl alcohol. Monolignols are mainly known to be the precursors of lignin. The main components of lignin are derived from hydroxycinnamyl alcohols (or monolignols), such as coniferyl alcohol, sinapyl alcohol and p-coumaryl alcohol (usually present in minor amounts). The existence of these three monolignols is responsible for the significant structural variety of lignin within plant species [63]. The units resulting from these monolignols, when incorporated into the lignin polymer, are called guaiacyl (G), syringyl (S) and p-hydroxyphenyl (H) units and play a crucial role in the process of lignification. Lignin is one of the most common biopolymers along with cellulose [45, 64]. These three alcohols are also involved in the synthesis of certain flavonolignans, at present, the contribution of p-coumaryl alcohol and sinapyl alcohol remains largely less widespread than that of coniferyl alcohol. However, interest in flavonolignans continues to increase and there is no indication that other flavonolignans, derived from these two alcohols, will not be discovered.

Flavonoids include more than 9,000 known structures [54] and are found in almost all plant tissue [65]. The flavonoid pathway is a major branch of the phenylpropanoid pathway and therefore has the same enzymes, PAL, C4H and 4CL, involved in the early stages of the phenylpropanoid pathway, to obtain monolignols. Generally, in plants, chalcones are not the end products. These compounds go through several enzymatic steps before giving rise to other classes of flavonoids, such as dihydroflavonols, flavanols or anthocyanins, the main water-soluble pigments of flowers and fruits [66]. Certain classes of flavonoids, (namely isoflavones, flavones, pro-anthocyanidins and flavonols) represent secondary branches of the flavonoid pathway and are derived from intermediates that allow the formation of anthocyanins [66]. Flavonoids play a large number of roles and perform certain functions [67, 68, 69, 70]: 1) pigmentation of flowers, fruits and seeds, 2) protection against ultraviolet rays, 3) defense of plants against pathogens, microorganisms, 4) plant fertility and pollen germination.

In S. marianum, the precursor of silymarin, the flavanonol taxifolin, belongs to the flavonoids. After being formed through the phenylpropanoid, p-coumaroyl-CoA serves as a starting precursor for the biosynthesis of flavonoids. The p-coumaroyl-CoA molecule is associated with 3 units of malonyl-CoA, thanks to chalcone synthase (CHS) which catalyzes the reaction to give the naringenin chalcone [45]. The latter is a chalcone containing two aromatic rings. Subsequent cyclization results in other flavonoids [71].

In the next step, chalcone isomerase (CHI) converts chalcones into flavanones (naringenin, eriodictyol and dihydrotricin) and vice versa. For the rest of the biosynthesis leading to the flavanonols, the naringenin obtained can be hydroxylated twice, (once in the C3 position and a second time in the C3’ position) to obtain taxifolin (2,3-dihydroquercetin). The synthesis of taxifolin involves two enzymes flavanones-3-hydroxylases (F3H and F3’H) which allow the two successive hydroxylation steps from naringerin. The F3H enzyme belongs to the family of 2-oxoglutarate-dependent dioxygenases. It is highly conserved in plant species [72, 73]. Hydroxylation at the C3 position of flavanones to dihydroflavonols has been demonstrated in a wide variety of plant species, including Petunia hybrida, Antirrhinum majus or Zea mays [74, 75].

Silymarin flavonolignans are supposed derived from taxifolin and coniferyl alcohol by one of the two proposed mechanisms: 1) the traditional Freudenburg hypothesis or 2) Althagafy’s proposal.

The Freudenburg hypothesis suggests an oxidative coupling reaction between taxifolin and coniferyl alcohol, induced by the formation of free radicals, and probably catalyzed by an enzyme of the peroxidase or laccase type, known to generate radicals [52]. Mechanically, the reaction assumes the formation of two distinct radicals, one at the phenoxyl group of taxifolin, and the other at the side chain of coniferyl alcohol, leading to a very reactive quinone methide intermediate [45]. The free radical of taxifolin therefore couples with the quinone methide radical, generated from coniferyl alcohol, to produce a molecule via a mechanism which is neither regio- nor enantioselective [76, 77]. The last step in biosynthesis is therefore a thermodynamically controlled nucleophilic attack, of a free intramolecular hydroxyl group, on the quinone methide nucleus of the monolignol part. Some rearrangement and cyclization are necessary to lead to the two molecules of diastereomeric flavonolignans, called silybins A and B. The O-β coupling step is neither regio- nor enantioselective [42, 45, 78]. Similar radical coupling could also result in the formation of regioisomers, isosilybin A and isosilybin B isolated from S. marianum [71].

Besides the regioisomer of silybin, two other isomers, silychristin and silydianin, having different binding modes between dihydroflavonol and coniferyl alcohol, have been described. In both cases, these two molecules are derived from a mesomer of the free radical derived from taxifolin [52].

Silychristin illustrates another structural variant of flavonolignans, with the formation of a furan nucleus [71]. This structure would be obtained via a mesomerism of the taxifolin radical which differs from that allowing the synthesis of silybins and isosilybins. There is a resonance form of taxifolin where the unpaired electron is located on the carbon of the B ring at position 4′ depending on the original 4-hydroxyl [76]. It is this difference in the resonance shape of taxifolin that allows the formation of the furan ring. In the case of silydianin, the mesomerism of taxifolin is analogous to that allowing the production of silybins and isosilybins, with the unpaired electron located on the carbon of the B ring at position 3′ [76]. But its unique and more complex structure suggests a more complicated mechanism [76]. Indeed, the only significant difference here lies in the formation of two new C-C bonds during the radical coupling during which the product cyclizes again, thus creating a bi-cyclo structure [79]. Subsequently, an intramolecular attack of an enolate on quinone methide (hemiacetalization) occurs, thus forming the original ketone-hemiketal structure of silydianin [76, 79].

However, Althagafy suggests, based on a biomimetic synthesis of the 4 major flavonolignans of silymarin (silybins and isosilybins), that the latter would be produced by a different mechanism, via the coupling of coniferyl alcohol and taxifolin [80]. There would be the formation of an intermediate quinone methide, via an oxidation causing the loss of an electron on the coniferyl alcohol. This intermediate would then add to taxifolin, via one of its hydroxyl groups, at the level of its catechol part. A second oxidation would allow the synthesis of silybins and isosilybins constituting silymarin. This is contrary to the mechanism proposed by Freudenburg for this process. Althagafy et al. [80], exclusively studied oxidative couplings carried out using silver (I) oxide (Ag2O) instead of enzyme, to form flavonolignans. It is postulated that a similar reaction mechanism could occur in the biosynthesis of flavonolignans in vivo.

Although the Althagafy hypothesis is shown to be functional in vitro, it does not offer any hypothesis on the mechanism of silydianin or silychristin biosynthesis. This second hypothesis focuses exclusively on silybins and isosilybins, which are the major components of silymarin.

The protein(s) responsible for the formation of flavonolignans, during the oxidative coupling step between taxifolin and coniferyl alcohol, remains to be identified. As stated above, the reaction could take place via a radical coupling mechanism by the intervention of oxidative radical-forming enzymes, such as peroxidases or laccases [38]. These enzymes are associated with the formation of monolignol radicals, with polymerization during lignification and lignan synthesis [38]. These two types of enzymes cause electron loss by catalyzing the oxidation of phenolic substrates [61], especially monolignols such as coniferyl alcohol, p-coumaryl alcohol and alcohol. Sinapylic thus leading to the formation of radicals with a view to their polymerization [81]. Peroxidases are among the most common and widespread enzymes. Many peroxidases incorporate an iron-porphyrin (heme) derivative at the heart of their active site. They can catalyze the oxidation of a wide variety of organic compounds using hydrogen peroxide (H2O2) [82]. The most studied function of peroxidases is their role in the polymerization of lignin (polymer constituent of the plant cell wall). Peroxidases have been shown to catalyze the polymerization of lignin monomers (p-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol) in vitro to form a lignin-like polymer [83]. Peroxidases are very widespread, especially in Arabidopsis where around 73 different peroxidase genes have been identified [84, 85]. So far, the expression of 23 peroxidases has been studied in different organs of this plant [85]. Numerous examples of functional characterization in vivo confirm this role in planta [86]. A peroxidase potentially involved in the formation of flavonolignans could be structured in the same way as class III peroxidases [45, 87]. Extracellular peroxidases are believed to be also responsible for the oxidation of taxifolin [88, 89]. Laccases are widely distributed oxidases, harboring a multi-copper center, among plants, insects and fungi [90]. They are mainly monomeric, dimeric or tetrameric glycoproteins [91]. These enzymes catalyze oxidations on a wide variety of organic and inorganic substrates, in particular mono-, di- and polyphenols, with simultaneous reduction of oxygen to water by 4 electrons from their multi-copper center [92]. These oxidases play a role in the degradation, but also in the formation of polymers of lignin, by promoting the oxidative coupling of monolignol units [45]. In comparison with peroxidases, besides the different structural aspects, laccases are generally considered to have a lower oxidative power than that of peroxidases [93].

The proposed biosynthetic sequence leading to flavonolignans accumulation in S. marianum fruit is summarized in Figure 8.

Figure 8.

Partial scheme of the silymarin (SILM) flavonolignans biosynthesis pathway in S. marianum. Flavonolignans are mainly accumulated in mature achenes (yellow box). In red are presented genes potentially involved in this pathway: PAL (l-phenylalanine ammonia-lyase), CAD (cinnamyl alcohol dehydrogenase), CHS (chalcone synthase), flavanone 3-dioxygenase (F3H), flavone 3′-hydroxylase (F3’H), LAC (laccases) and POX (peroxidases). The dotted arrows indicate a single step, while dashed arrows indicate several steps in the metabolic pathway while the full arrows indicate direct synthesis of the compound through an enzyme.

A recent study on the spatial organization of silybin biosynthesis in S. marianum demonstrated, through biomimetic synthesis, that peroxidase and laccase can both catalyze silybin synthesis [94]. This study was further developed by identifying, through an RNAseq sequencing study, five peroxidase candidates involved in the production of silybins and isolybins. Among these, APX1 (ascorbate peroxidase 1) exhibits distinct peroxidase activity and the ability to synthesize silybins [94]. This first step provides a better understanding of the biosynthesis of silybin and other flavonolignans. Despite these encouraging results, it is still unknown, at this time, whether peroxidases are the only enzymes involved in the final stage of flavonolignan biosynthesis [88, 89]. Based on these results, two different coupling modes can be suggested, depending on the type of flavonolignan obtained. One would involve a classic, simple coupling, while the other would be a more complex coupling, with complementary proteins depending on the flavonolignans synthesized, suggesting the possible intervention of dirigent proteins (DIRs).

The two mechanisms proposed above are radical types. They are therefore, a priori, non-stereospecific and non-regioselective, which implies a wide variety of possibilities at the coupling level, and therefore of the isomers formed. Based on this information, the diastereomers of silybin and isosilybin are described as equimolar mixtures reflecting the lack of stereospecificity of the initial radical couplings [76, 95]. Among the available experimental data, none confirms the 1: 1 ratio between these diastereoisomeric couples [31, 37, 88, 96, 97]. Also, the theory of random radical coupling cannot explain the proportions obtained in the mixture of silymarin, since in certain genotypes, silydianin is practically non-existent [37, 88, 97]. An important question then arises on the biosynthetic pathway of flavonolignans [45] is how to explain the strong disparities observed during the quantification of the constituents of silymarin in the fruits of various origins of milk thistle since 3 distinct chemotypes have already been described in S. marianum based on their phytochemical diversity in silymarin [37, 38, 39, 52].

The in-depth study of these chemotypes could provide important information on the biosynthetic pathways leading to the different flavonolignans of silymarin. This could allow relational models to be established, based on the levels of metabolites in chemotypes, to infer the presence of steps in a metabolic pathway [88]. To study flavonolignans from a scientific point of view, it would be interesting to understand the underlying mechanism allowing to obtain the different chemotypes and positional isomers in S. marianum, and more generally. Despite the hypotheses made and studied about the biosynthesis of flavonolignans, little investigation has been carried out on how the proportional differences of isomers arise, and how the plant differentiates them [45]. In 1995, the discovery of DIRs by Davin and Lewis opened up new perspectives on how free radical coupling of plant phenols is controlled in the production of lignans and lignin [98]. DIRs, by their abundance, would be the first essential elements of phenoxy-radical couplings. The first action hypotheses of these proteins concerned the stereoselective biosynthesis of (+)-pinoresinol from coniferyl alcohol monomers. Coniferyl alcohol (or the resulting radical) was oriented and directed by a DIR and allowed specific synthesis of (+)-pinoresinol during bimolecular coupling [99]. Until recently, DIRs were not considered as enzymes, due to their lack of oxidative activity. The proper binding and orientation of the substrate radicals appeared sufficient for DIR activity. In 2016, the structure of the AtDIR6 from Arabidopsis thaliana confirms that DIRs are actively involved and that they have a catalytic function in the cyclization step of the quinone methide intermediate [45, 100, 101]. This has been attributed to the active donation of protons by the formation of hydrogen bonds or by acid catalysis [100]. Other DIRs have been identified, notably in Arabidopsis thaliana, with DIR responsible for the enantioselective synthesis of (−)-pinoresinol [102]. Other DIRs have been described in other plant species including flax [103], Schisandra [102, 104] or Isatis indigotica [105]. In Gossypium hirsutum (cotton), a leader protein has been characterized. It is involved in the stereoselective coupling of hemigossypol in the formation of the terpene (+)-gossypol [106, 107]. In 2018, the study of the leader proteins of Linum usitatissimum made it possible to highlight around forty DIRs in the flax genome [103] and to identify new conserved motifs linked to functions specific biochemicals. The mechanism of reactions by oxidative coupling leading to the accumulation of different steroisomers in the synthesis of flavonolignans and lignans, in S. marianum, makes possible the hypothesis of the involvement of DIRs [53, 108].


5. Regulation of flavonolignans biosynthesis in Silybum marianum

The genes, the expression of which has been studied in milk thistle, are the genes involved in the formation of precursors to taxifolin and coniferyl alcohol. The mixture of flavonolignans from Milk Thistle mainly accumulates in the pericarp of the fruit of milk thistle. It is therefore necessary to wait several months for the plant to produce its fruits and the molecules of interest. Experiments have shown that the accumulation of silymarin in the fruits of S. marianum is associated with the ripening process. The study of gene expression in milk thistle fruit is very recent. So far, it has only concerned 5 genes, involved in the pathway of flavonoids biosynthesis, CHS [109, 110], CHI, F3H, F3’H and CAD [108, 111]. The objective was to verify their roles in the biosynthesis of flavonolignans. These first functional studies showed the potential association of these genes in the synthesis of silymarin, because their expression coincides with the accumulation of taxifolin in fruits. In addition, induction of the CAD gene appears to be necessary for the accumulation of silymarin in ripe fruits [108, 111].

Recently, Drouet et al. [108] precisely described development stages of fruit to study the kinetics of accumulation of silymarin constituents during fruit ripening (Figure 9). During fruit maturation, the accumulation profiles of the silymarin components were evaluated by LC–MS analysis at each of the development stages identified [108]. Reference genes have been identified, selected and validated to allow accurate gene expression profiling of candidate biosynthetic genes [108]. Enzyme activity and biosynthetic gene expression indicated a possible in situ biosynthesis of silymarin from l-Phenylalanine during fruit ripening [108]. The gene expression profiles were well correlated with silymarin kinetic accumulation and preferential location in pericarp during S. marianum fruit maturation, reaching maximum biosynthesis when desiccation occurs [108]. This observation led us to consider the possible involvement of abscisic acid (ABA), a key phytohormone in fruit ripening control, for which accumulation timing and location during fruit ripening were consistent with the potential regulation of the silymarin accumulation. This possible transcriptional regulation of silymarin biosynthesis by ABA was further supported by the presence of ABA-responsive cis-acting elements in the silymarin biosynthetic gene promoter regions studied [108].

Figure 9.

Development stages of the S. marianum achene defined according to their morphological characteristic. (A) Six achene developmental stages were defined. For the developmental stages 4, 5 and 6 achenes were manually dissected to allow the visualization of both seed and pericarp. The capitula morphology corresponding to each defined achene developmental stage is presented. Note that the capitula morphology is not predictive of the fruit developmental stage. (B) Morphological features of achene maturation during time such as achene, seed, and pericarp length, dry weight (DW) and day of flowering (DAF). Each value represents means ± SD of n = 10 independent sampling.

Biotechnological approaches have been used to increase the production of these molecules in vitro. The in vitro culture of a plant is a possible source of secondary metabolites. The use of in vitro models makes it possible to overcome the constraints of plant growth and makes it possible to have a biomass containing the molecules of interest more quickly. The cell models used aim to increase the yields of flavonolignans, by optimizing their biosynthesis. The main objective is therefore to modify the expression of one or more genes, or/and to elicit cultures, to overcome the steps limiting the accumulation of these compounds in the biosynthetic pathway [111]. However, one of the major constraints is the limited information available on the coupling of flavonolignans, for the genes and proteins involved.

S. marianum in vitro cultures can constitute an alternative and renewable source of this natural product. However, the concentration of silymarin in this type of crop is often lower than that found naturally in fruit [111, 112]. A higher accumulation was observed in cell suspensions obtained from calli [113]. To obtain silymarin in sufficiently high concentrations for commercial purposes, techniques have been proposed to stimulate its production in calli and cultured cells of S. marianum, via the use of elicitors or by the addition of precursors. The elicited systems are a starting point in improving the production of silymarin for industrial purposes. Elicitation is one of the most effective approaches to increase the yield of secondary metabolites in in vitro cultures [114]. It modulates the rates of biosynthesis and accumulation [112, 115]. The use of stress mediators such as methyl jasmonate, exposure to certain lights [116, 117], auxin, [118], yeast extracts, chitosan [119], cyclodextrin, strongly induce the extracellular accumulation of coniferyl alcohol and Silymarin [111, 120]. The “feeding” experiments consisting in providing the initial substrates to increase the content of flavonolignans are less conclusive. Coniferyl alcohol added to cultures allows for a significant and more surprisingly increase in silydianin, but no other component of silymarin has been detected [121].

Hairy root cultures have multiple advantages, in particular their genetic/biochemical stability and the ability to rapidly produce large biomasses [112]. Hairy roots are roots of plant tissue obtained by genetic transformation using a bacteria naturally present in the soil, Agrobacterium rhizogenes. Hairy root cultures of S. marianum have been established [122, 123, 124]. Elicitation experiments have been undertaken on this model. Salicylic acid is effective in increasing the content of flavonolignans, the content of linoleic acid and the activity of lipoxygenase (an important enzyme in jasmonate biosynthesis) in hairy root crops [125]. It is therefore likely that elicitation by salicylic acid regulates the jasmonate pathway which in turn mediates the accumulation of silymarin [125].

A recent and comprehensive review summarized these different biotechnological strategies [126].


6. Conclusions

Many experiments have provided a deeper understanding of the biosynthesis and control of silymarin flavonolignans from S. marianum over the last decade. However, to completely take advantage of their multiple biological activities, including antioxidant activity, for pharmaceutical and/or cosmeceutical uses, this view is still partial and further study is needed. To allow a better understanding of the biosynthetic steps leading to these flavonolignans, more detailed enzymatic and genetic studies are therefore needed. Only a thorough and comprehensive understanding of the metabolic regulation of these pathways in S. marianum can make it possible to identify andpromote their accumulations, the limiting steps and key points toward their regulation on which it will be possible to act.



S.D. acknowledges research fellowships of Loire Valley Region. This research was part of the MariLine project (grant number ICO16029) focusing on the cosmetic application of the milk thistle flavonolignans, supported by Cosmetosciences, a global training and research program dedicated to the cosmetic industry. Located in the heart of the cosmetic valley, this program led by University of Orléans is funded by the Region Centre-Val de Loire.


Conflict of interest

The authors declare no conflict of interest.


  1. 1. Raskin I, Ribnicky DM, Komarnytsky S, Ilic N, Poulev A, Borisjuk N, et al. Plants and human health in the 21st century. Trends Biotechnol [Internet]. 2002;20(12):522-531. Available from:
  2. 2. Shipley GP, Kindscher K. Evidence for the Paleoethnobotany of the Neanderthal: A Review of the Literature. Scientifica (Cairo). 2016;2016:1-12.
  3. 3. Ma JKC, Chikwamba R, Sparrow P, Fischer R, Mahoney R, Twyman RM. Plant-derived pharmaceuticals - The road forward. Trends Plant Sci. 2005;10(12):580-585.
  4. 4. Karkanis A, Bilalis D, Efthimiadou A. Cultivation of milk thistle (Silybum marianum L. Gaertn.), a medicinal weed. Ind Crops Prod [Internet]. 2011;34(1):825-30. Available from:
  5. 5. Shaker E, Mahmoud H, Mnaa S. Silymarin, the antioxidant component and Silybum marianum extracts prevent liver damage. Food Chem Toxicol [Internet]. 2010;48(3):803-806. Available from:
  6. 6. Bijak M. Silybin, a Major Bioactive Component of Milk Thistle (Silybum marianum L. Gaernt.)—Chemistry, Bioavailability, and Metabolism. Molecules. 2017;22(11):1942.
  7. 7. Abenavoli L, Capasso R, Milic N, Capasso F. Milk Thistle in Liver Diseases: Past, Present, Future. Phyther Res. 2010;24(June):1423-1432.
  8. 8. Urtasun R, Rosa LC de la, Nieto N. Oxidative and Nitrosative Stress and Fibrogenic Response. Clin Livers Dis. 2008;12(4).
  9. 9. Wu D, Yotnda P. Production and Detection of Reactive Oxygen Species (ROS) in Cancers. J Vis Exp. 2011;(57):2-5.
  10. 10. Pradhan S, Girish C. Hepatoprotective herbal drug, silymarin from experimental pharmacology to clinical medicine. Indian J Med Res. 2006;124:491-504.
  11. 11. Detaille D, Sanchez C, Sanz N, Lopez-Novoa JM, Leverve X, El-Mir MY. Interrelation between the inhibition of glycolytic flux by silibinin and the lowering of mitochondrial ROS production in perifused rat hepatocytes. Life Sci. 2008;82(21-22):1070-1076.
  12. 12. Polyak SJ, Morishima C, Lohmann V, Pal S, Lee DYW, Liu Y, et al. Identi fi cation of hepatoprotective fl avonolignans from silymarin. 2010;107(13).
  13. 13. Drouet S, Leclerc EA, Garros L, Tungmunnithum D, Kabra A, Abbasi BH, et al. A Green Ultrasound-Assisted Extraction Optimization of the Natural Antioxidant and Anti-Aging Flavonolignans from Milk Thistle Silybum marianum ( L .) Gaertn . Fruits for Cosmetic Applications. Antioxidants. 2019;8(304):1-19.
  14. 14. Karimi G, Vahabzadeh M, Lari P, Rashedinia M, Moshiri M. Silymarin, a promising pharmacological agent for treatment of diseases. Iran J Basic Med Sci. 2011;14(4):308-317.
  15. 15. Katiyar SK. Silymarin and skin cancer prevention: Anti-inflammatory, antioxidant and immunomodulatory effects (Review). Int J Oncol. 2005;26:169-176.
  16. 16. Katiyar SK, Meleth S, Sharma SD. Silymarin, a flavonoid from milk thistle (Silybum marianum L.), inhibits UV-induced oxidative stress through targeting infiltrating CD11b+cells in mouse skin. Photochem Photobiol. 2008;84(2):266-271.
  17. 17. Meeran SM, Katiyar S, Elmets C a, Katiyar SK. Silymarin inhibits UV radiation-induced immunosuppression through augmentation of interleukin-12 in mice. Mol Cancer Ther. 2006;5(7):1660-1668.
  18. 18. Toklu HZ, Tunali-Akbay T, Erkanli G, Yüksel M, Ercan F, Şener G. Silymarin, the antioxidant component of Silybum marianum, protects against burn-induced oxidative skin injury. Burns. 2007;33(7):908-916.
  19. 19. Rajnochová A, Eva S, Jitka G, Bohumil U, David Z. A pilot study of the UVA - photoprotective potential of dehydrosilybin , isosilybin , silychristin , and silydianin on human dermal fibroblasts. Arch Dermatol Res [Internet]. 2019; Available from:
  20. 20. Gažák R, Purchartová K, Marhol P, Živná L, Petr Sedmera, Valentová K, et al. Antioxidant and antiviral activities of silybin fatty acid conjugates. Eur J Med Chem. 2010;45:1059-1067.
  21. 21. Zarrelli A, Romanucci V, Greca M Della, De Napoli L, Previtera L, Di Fabio G. New silybin scaffold for chemical diversification: Synthesis of novel 23-phosphodiester silybin conjugates. SYNLETT. 2013;24(1):45-48.
  22. 22. Zarrelli A, Romanucci V, Tuccillo C, Federico A, Loguercio C, Gravante R, et al. New silibinin glyco-conjugates: Synthesis and evaluation of antioxidant properties. Bioorganic Med Chem Lett [Internet]. 2014;24(22):5147-5149. Available from:
  23. 23. Drouet S, Doussot J, Garros L, Mathiron D, Bassard S, Favre-Réguillon A, et al. Selective Synthesis of 3-O-Palmitoyl-Silybin, a New-to-Nature Flavonolignan with Increased Protective Action against Oxidative Damages in Lipophilic Media. Molecules. 2018;23(10):2594.
  24. 24. Bonnier G, Douin R. La grande flore. Belin, editor. 1990.
  25. 25. Drouet S, Garros L, Hano C, Lainé É. A Critical View of Different Botanical , Molecular , and Chemical Techniques Used in Authentication of Plant Materials for Cosmetic Applications. 2018;
  26. 26. International C. CABI, 2018. Invasive species compendium [Internet]. 2018. Available from:
  27. 27. W.E Safford. The Flora of Banda Oriental. In: Bulletin of the Torrey Botanical Club [Internet]. New York : Torrey Botanical Club,; 1887 [cited 2019 Apr 12]. p. 159-64. Available from:
  28. 28. Canter PH, Thomas H, Ernst E. Bringing medicinal plants into cultivation: Opportunities and challenges for biotechnology. Trends Biotechnol. 2005;23(4):180-185.
  29. 29. Andrzejewska J, Sadowska K. Effect of cultivation conditions on the variability and interrelation of yield and raw material quality in Milk thistle (Silybum marianum (L.) GAERTN.). Acta Sci Pol Agric. 2008;7(3):3-11.
  30. 30. Andrzejewska J, Sadowska K, Mielcarek S. Effect of sowing date and rate on the yield and flavonolignan content of the fruits of milk thistle (Silybum marianum L. Gaertn.) grown on light soil in a moderate climate. Ind Crops Prod [Internet]. 2011;33(2):462-468. Available from:
  31. 31. Carrier DJ, Crowe T, Sokhansanj S, Wahab J, Branka B. Milk Thistle, Silybum marianum (L.) Gaertn., flower head development and associated marker compound profile. J Herbs, Spices Med Plants. 2003;10(1):65-74.
  32. 32. Parsons WT, Cuthbertson EG. Noxious Weeds of Australia. Inkata Pre. 1992.
  33. 33. Greenlee H, Abascal K, Yarnell E, Ladas E. Clinical applications of Silybum marianum in oncology. Integr Cancer Ther. 2007;6(2):158-165.
  34. 34. Pelter A, Hansel R. The structure of silybin (Silybum substance E6), the first flavonolignan. Tetrahedron Lett. 1968;1(25):2911-2916.
  35. 35. Rauter AP, Ennis M, Hellwich K-H, Herold BJ, Horton D, Moss GP, et al. Nomenclature of flavonoids (IUPAC Recommendations 2017). Vol. 90, Pure and Applied Chemistry. 2018. 1429-1486 p.
  36. 36. Kroll DJ, Shaw HS, Oberlies NH, Kroll BDJ, Shaw HS, Oberlies NH. Milk thistle nomenclature: why it matters in cancer research and pharmacokinetic studies. Integr Cancer Ther. 2007;6(2):110-119.
  37. 37. Martin RJ, Lauren DR, Smith WA, Jensen DJ, Deo B, Douglas JA, et al. Factors influencing silymarin content and composition in variegated thistle (Silybum marianum). New Zeal J Crop Hortic Sci. 2006;34(3):239-245.
  38. 38. Poppe L, Petersen M. Variation in the flavonolignan composition of fruits from different Silybum marianum chemotypes and suspension cultures derived therefrom. Phytochemistry [Internet]. 2016;131:68-75. Available from:
  39. 39. Radjabian T, Rezazadeh SH, Huseini HF. Analysis of silymarin components in the seed extracts of some milk thistle ecotypes from Iran by HPLc. Iran J Sci Technol. 2008;32:6-8.
  40. 40. Delfini M, Cocco MED, Gaggelli E, Lepri A, Valensin G. H-1 delineation of conformational parameters of constituents of silybum marianum in solution. Spectrosc Lett. 1990;23(8):1007-1025.
  41. 41. Hansel R, Janiak VB. Phytochemisch-pharmakognostische untersuchungen über fructus Cardui mariae. Planta Med. 1960;8:71-84.
  42. 42. Nam-Cheol K, Graf TN, Sparacino CM, Wani MC, Wall ME. Complete isolation and structure identification of hepatoprotective flavonolignans silybins and isosilybins from the medicinal herb milk thistle (Silybum marianum). Org Biomol Chem. 2003;1:1684-1689.
  43. 43. Wagner H, Diesel P, Seitz M. The chemistry and analysis of silymarin from Silybum marianum Gaertn. Arzneimittelforschung [Internet]. 1974;24(4):466—471. Available from:
  44. 44. Wagner H, Hörhammer L, Münster R. On the chemistry of silymarin (silybin), the active principle of the fruits from Silybum marianum (L.) Gaertn. (Carduus marianus L.). Naturwissenschaften [Internet]. 1965;18(6):688—696. Available from:
  45. 45. Poppe L. Thesis: Flavonolignan Biosynthesis in Silybum marianum – Potential Regulatory Mechanisms and Candidate Genes. 2017.
  46. 46. Csupor D, Csorba A, Hohmann J, Cuspor DD, Csorba A, Hohmann J. Recent advances in the analysis of flavonolignans of Silybum marianum. J Pharm Biomed Anal. 2016;130:301-317.
  47. 47. Abouzid S, Ahmed OM. Silymarin flavonolignans: Structure-activity relationship and biosynthesis. 1st ed. Vol. 40, Studies in Natural Products Chemistry. © 2013 Elsevier B.V. All rights reserved.; 2013. 469-484 p.
  48. 48. Lee DY, Liu Y. Molecular structure and stereochemistry of Silybin A, Silybin B , Isosilybin , and Isosilybin B, isolated from Silybum marianum (Milk Thistle). J Nat Prod. 2003;66:1171-1174.
  49. 49. Althagafy HS, Meza-Aviña ME, Oberlies NH, Croatt MP. Mechanistic study of the biomimetic synthesis of flavonolignan diastereoisomers in milk thistle. J Org Chem. 2013;78(15):7594-7600.
  50. 50. Napolitano JG, Lankin DC, Graf TN, Brent Friesen J, Chen SN, McAlpine JB, et al. HiFSA fingerprinting applied to isomers with near-identical NMR Spectra: The silybin/isosilybin case. J Org Chem. 2013;78(7):2827-2839.
  51. 51. Smith WA, Lauren DR, Burgess EJ, Perry NB, Martin RJ. A silychristin isomer and variation of flavonolignan levels in milk thistle (Silybum marianum) fruits. Planta Med. 2005;71(9):877-880.
  52. 52. Chambers CS, Hole V, Petrásková L, Biedermann D, Buchta M, Vladimír K. The silymarin composition … and why does it matter??? Food Res Int. 2017;100(July):339-353.
  53. 53. Drouet S, Abbasi BH, Falguières A, Ahmad W, Sumaira, Ferroud C, et al. Single Laboratory Validation of a Quantitative Core Shell-Based LC Separation for the Evaluation of Silymarin Variability and Associated Antioxidant Activity of Pakistani Ecotypes of Milk Thistle (Silybum marianum L.). Molecules. 2018;23(904).
  54. 54. Bashandy H. Flavonoid metabolomics in Gerbera hybrida and elucidation of complexity in the flavonoid biosynthetic pathway. 2016.
  55. 55. Tohge T, Watanabe M, Hoefgen R, Fernie AR. The evolution of phenylpropanoid metabolism in the green lineage. Crit Rev Biochem Mol Biol. 2013;48(2):123-152.
  56. 56. Winkel-shirley B. Flavonoid Biosynthesis. A Colorful Model for Genetics, Biochemistry, Cell Biology, and Biotechnology. Plant Physiol. 2001;126(June):485-493.
  57. 57. Attoumbre J, Hano C, Mesnard F, Lamblin F, Bensaddek L, Raynaud-Le Grandic S, et al. Identification by NMR and accumulation of a neolignan, the dehydrodiconiferyl alcohol-4-$β$-D-glucoside, in Linum usitatissimum cell cultures. Comptes Rendus Chim. 2006;9(3-4).
  58. 58. Hano C, Addi M, Bensaddek L, Crônier D, Baltora-Rosset S, Doussot J,et al. Differential accumulation of monolignol-derived compounds in elicited flax (Linum usitatissimum) cell suspension cultures. Planta. 2006 Apr;223(5):975-989.
  59. 59. Martins V, Garcia A, Costa C, Sottomayor M, Gerós H. Calcium- and hormone-driven regulation of secondary metabolism and cell wall enzymes in grape berry cells. J Plant Physiol [Internet]. 2018;231(August):57-67. Available from:
  60. 60. Umezawa T. Diversity in lignan biosynthesis. Phytochem Rev. 2003;2(3):371-390.
  61. 61. Vanholme R, Demedts B, Morreel K, Ralph J, Boerjan W. Lignin Biosynthesis and Structure. Plant Physiol. 2010;153(3):895-905.
  62. 62. Umezawa T. The cinnamate/monolignol pathway. Phytochem Rev. 2010;9(1):1-17.
  63. 63. Rodrigo CP, James WH, Zwier TS. Single-Conformation ultraviolet and infrared spectra of jet-cooled monolignols: P-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol. J Am Chem Soc. 2011;133(8):2632-2641.
  64. 64. Freudenberg K, Neish AC. Constitution and biosynthesis of lignin. Springer. Science. Berlin; 1968.
  65. 65. Martens S, Mithöfer A. Flavones and flavone synthases. Phytochemistry. 2005;66(20):2399-2407.
  66. 66. Schijlen EGWM, Ric de Vos C, Tunen AJ Van, Bovy AG. Modification of flavonoid biosynthesis in crop plants. Phytochemistry. 2004;65:2631-2648.
  67. 67. Dooner HK, Robbins TP, Jorgensen RA. Genetic and Developmental Control of Anthocy an in. Most. 1991;
  68. 68. Spangenberg G. Flavonoids : a metabolic network mediating plants adaptation to their real estate. Front Plant Sci. 2014;5(November):1-16.
  69. 69. Vasilev N, Schmitz C, Dong L, Ritala A, Imseng N, Häkkinen S, et al. Comparison of plant-based expression platforms for the heterologous production of geraniol. Plant Cell, Tissue Organ Cult [Internet]. 2014;117(3):373-80. Available from:
  70. 70. Winkel-shirley B. Biosynthesis of flavonoids and effects of stress. 2002;218-23.
  71. 71. Begum S., Sahai M, Ray A. Non-conventional Lignans: Coumarinolignans, Flavonolignans, and Stilbenolignans. Kinghorn A., Falk H, Kobayashi J, editors. SpringerWienNewYork; 2010. 28-49 p.
  72. 72. Britsch L, Dedio J, Saedler H, Forkmann G. Molecular characterization of flayanone 3β-hydroxylases: Consensus sequence, comparison with related enzymes and the role of conserved histidine residues. Eur J Biochem. 1993;217(2):745-754.
  73. 73. Hall J, Ananga A, Georgiev V, Ochieng J, Cebert E, Tsolova V. Molecular Cloning, Characterization, and Expression Analysis of Flavanone 3-Hydroxylase (F3H) Gene during Muscadine Grape Berry Development. J Biotechnol Biomater [Internet]. 2015;05(02). Available from:
  74. 74. Weiss D, van der Luit AH, Kroon JTM, Mol JNM, Kooter JM. The petunia homologue of the Antirrhinum majus candi and Zea mays A2 flavonoid genes; homology to flavanone 3-hydroxylase and ethylene-forming enzyme. Plant Mol Biol. 1993;22(5):893-897.
  75. 75. Deboo GB, Albertsen MC, P.Taylor L. Flavanone 3-hydroxylase transcripts and flavonol accumulation are temporally coordinate in maize anthers. plant J. 1995;7(5):703-713.
  76. 76. Dewick PM. The shikimate pathway: aromatic amino acids and phenylpropanoids. In: Medicinal Natural Products: A Biosynthetic Approach. John Wiley. 2007. p. 137-184.
  77. 77. Merlini L, Zanarotti A, Pelter A, Rochefort MP, Hänsel R. Benzodioxans by oxidative phenol coupling. Synthesis of silybin. J Chem Soc, Perkin Trans 1. 1980;(0):775-778.
  78. 78. Mascher H, Kikuta C, Weyhenmeyer R. Diastereomeric separation of free and conjugated silibinin in plasma by reversed phase hplc after specific extraction. J Liq Chromatogr. 1993;16(13):2777-2789.
  79. 79. Biedermann D, Moravcová V, Valentová K, Kuzma M, Petrásková L, Císařová I, et al. Oxidation of flavonolignan silydianin to unexpected lactone-acid derivative. Phytochem Lett [Internet]. 2019;30(October 2018):14-20. Available from:
  80. 80. Althagafy HS, Meza-Aviña ME, Oberlies NH, Croatt MP. Mechanistic study of the biomimetic synthesis of flavonolignan diastereoisomers in milk thistle. J Org Chem. 2013;78(15):7594-7600.
  81. 81. Kersten PJ, Kalyanaraman B, Hammel KE, Reinhammar B, Kirk TK. Comparison of lignin peroxidase, horseradish peroxidase and laccase in the oxidation of methoxybenzenes. Biochem J. 1990;268(2):475-480.
  82. 82. Watanabe Y, Nakajima H. Creation of a Thermally Tolerant Peroxidase [Internet]. 1st ed. Vol. 580, Methods in Enzymology. Elsevier Inc.; 2016. 455-470 p. Available from:
  83. 83. Christensen JH, Montagu M Van, Bauw G, Boerjan W. Xylem peroxidases: Purification and altered expression. Prog Biotechnol. 2001;18(C):171-176.
  84. 84. Tognolli M, Overney S, Penel C, Greppin H, Simon P. A genetic and enzymatic survey of Arabidopsis thaliana peroxidases. Plant Perox. 2000;
  85. 85. Tognolli M, Penel C, Greppin H, Simon P. Analysis and expression of the class III peroxidase large gene family in Arabidopsis thaliana. Gene. 2002;288(1-2):129-138.
  86. 86. Boerjan W, Ralph J, Baucher M. Lignin biosynthesis. Annu Rev Plant Biol [Internet]. 2003;54(1):519-546. Available from:
  87. 87. Veitch NC. Horseradish peroxidase: a modern view of a classic enzyme. Phytochemistry [Internet]. 2004;65(3):249-59. Available from:
  88. 88. Martinelli T, Whittaker A, Benedettelli S, Carboni A. The study of flavonolignan association patterns in fruits of diverging Silybum marianum (L.) Gaertn. chemotypes provides new insights into the silymarin biosynthetic pathway. Phytochemistry [Internet]. 2017;144:9-18. Available from:
  89. 89. Sánchez-Sampedro A, Kim HK, Choi YH, Verpoorte R, Corchete P. Metabolomic alterations in elicitor treated Silybum marianum suspension cultures monitored by nuclear magnetic resonance spectroscopy. J Biotechnol. 2007;130(2):133-142.
  90. 90. Yaropolov AI, Skorobogat’ko O V., Vartanov SS, Varfolomeyev S. Laccase: Properties, Catalytic Mechanism, and Applicability. Appl Biochem Biotechnol. 1994;49:257-280.
  91. 91. Shraddha, Shekher R, Sehgal S, Kamthania M, Kumar A. Laccase: microbial sources, production, purification, and potential biotechnological applications. Enzyme Res [Internet]. 2011;2011:217861. Available from:
  92. 92. Madhavi V, Lele SS. Laccase: Properties and applications. BioResources. 2009;4(4):1694-1717.
  93. 93. Heinzkill M, Bech L, Halkier T, Schneider P, Anke T. Characterization of laccases and peroxidases from wood-rotting fungi (family Coprinaceae). Appl Environ Microbiol. 1998;64(5):1601-1606.
  94. 94. Lv Y, Gao S, Xu S, Du G, Zhou J, Chen J, et al. Spatial organization of silybin biosynthesis in milk thistle (Silybum marianum (L.) Gaertn.). Plant J. 2017;92:995-1004.
  95. 95. Nyiredy S, Szücs Z, Antus S, Samu Z. New Components from Silybum marianum L.Fruits: A Theory Comes True. Chromatographia. 2008;68:5-11.
  96. 96. Abouzid SF, Ahmed HS, Moawad AS, Owis AI, Chen S, Nachtergael A, et al. Chemotaxonomic and biosynthetic relationships between flavonolignans produced by Silybum marianum populations. Fitoterapia [Internet]. 2017;119(April):175-184. Available from:
  97. 97. Martinelli T, Potenza E, Moschella A, Zaccheria F, Benedettelli S, Andrzejewska J. Phenotypic Evaluation of a Milk Thistle Phenotypic Evaluation of a Milk Thistle Germplasm Collection: Fruit Morphology and Chemical Composition. Crop Sci. 2016;56(September).
  98. 98. Davin LB, Lewis NG. Dirigent Proteins and Dirigent Sites Explain the Mystery of Specificity of Radical Precursor Coupling in Lignan and Lignin Biosynthesis. Plant Physiol. 2000;123:453-461.
  99. 99. Davin LB, Wang H, Crowell AL, Bedgar DL, Martin DM, Sarkanen S, et al. Stereoselective Bimolecular Phenoxy Radical Coupling by Auxialiary (Dirigent) Protein Without an Active Center. Reports. 1997;275(16).
  100. 100. Gasper R, Effenberger I, Kolesinski P, Terlecka B, Hofmann E, Schaller A. Dirigent Protein Mode of Action Revealed by the Crystal Structure of AtDIR6. Plant Physiol. 2016;172(4):2165-2175.
  101. 101. Kim KW, Smith CA, Daily MD, Cort JR, Davin LB, Lewis NG. Trimeric structure of (+)-pinoresinol-forming dirigent protein at 1.95 A° resolution with three isolated active sites. J Biol Chem. 2015;290(3):1308-1318.
  102. 102. Pickel B, Constantin MA, Pfannstiel J, Conrad J, Beifuss U, Schaller A. An enantiocomplementary dirigent protein for the enantioselective lacease-catalyzed oxidative coupling of phenols. Angew Chemie - Int Ed. 2010;49(1):202-204.
  103. 103. Corbin C, Drouet S, Markulin L, Auguin D, Lainé É, Davin LB, et al. A genome-wide analysis of the flax (Linum usitatissimum L .) dirigent protein family: from gene identification and evolution to differential regulation. Plant Mol Biol [Internet]. 2018;97(1-2):73-101. Available from:
  104. 104. Kim K-W, Moinuddin SGA, Atwell KM, Costa MA, Davin LB, Lewis NG. Opposite Stereoselectivities of Dirigent Proteins in Arabidopsis and Schizandra Species. J Biol Chem [Internet]. 2012 Oct 5;287(41):33957-33972. Available from:
  105. 105. Li Q, Chen J, Xiao Y, Di P, Zhang L, Chen W. The dirigent multigene family in Isatis indigotica: Gene discovery and differential transcript abundance. BMC Genomics. 2014;15(1):1-13.
  106. 106. Benedict CR, Liu J, Stipanovic RD, Ho O, C H, Oh HO, et al. The peroxidative coupling of hemigossypol to (+)- and (−_)-gossypol in cottonseed extracts. Contents / Phytochem. 2006;67(4):323-326.
  107. 107. Liu J, Stipanovic RD, Bell AA, Puckhaber LS, Magill CW. Stereoselective coupling of hemigossypol to form (+)-gossypol in moco cotton is mediated by a dirigent protein. Phytochemistry. 2008;69(18):3038-3042.
  108. 108. Drouet S, Tungmunnithum D, Lainé É, Hano C. Gene Expression Analysis and Metabolite Profiling of Silymarin Biosynthesis during Milk Thistle (Silybum marianum (L.) Gaertn.) Fruit Ripening. Int J Mol Sci. 2020;21(13):4730.
  109. 109. El-garhy HAS, Khattab S, Moustafa MMA, Abou R, Abdel AZ, Elhalwagi A, et al. Silybin content and overexpression of chalcone synthase genes in Silybum marianum L. plants under abiotic elicitation. Plant Physiol Biochem [Internet]. 2016;108:191-202. Available from:
  110. 110. Sepideh S, Shobbar ZS, Ebrahimi M, Hasanloo T, Sadat-Noori S-A, Tirnaz S. Chalcone synthase genes from milk thistle (Silybum marianum): isolation and expression. J Genet. 2015;94(4):611-617.
  111. 111. Torres M, Corchete P. Gene expression and flavonolignan production in fruits and cell cultures of Silybum marianum. J Plant Physiol [Internet]. 2016;192:111-117. Available from:
  112. 112. Abouzid S. Silymarin , Natural Flavonolignans from Milk Thistle. Rao DV, editor. A Global Perspective of Their Role in Nutrition and Health. 2012. 255-272 p.
  113. 113. Cacho M, Mora M, Ferna J. Influence of medium composition on the accumulation of flavonolignans in cultured cells of Silybum marianum (L.) Gaertn. Plant Sci. 1999;144:63-68.
  114. 114. Namdeo A. Plant Cell Elicitation for Production of Secondary Metabolites: A Review. Pharmacogn Rev. 2007;1(1):69-79.
  115. 115. Barz W, Beimen A, Drager B, Jaques U. Turnover and storage of secondary products in cell cultures. Second Prod from Plant Tissue Cult [Internet]. 1990 [cited 2019 May 15];79-102. Available from:
  116. 116. Shah M, Ullah MA, Drouet S, Younas M, Tungmunnithum D, Giglioli-Guivarc’h N, et al. Interactive effects of light and melatonin on biosynthesis of silymarin and anti-inflammatory potential in callus cultures of Silybum marianum (L.) gaertn. Molecules. 2019;24(7).
  117. 117. Younas M, Drouet S, Nadeem M, Giglioli-Guivarc’h N, Hano C, Abbasi BH. Differential accumulation of silymarin induced by exposure of Silybum marianum L. callus cultures to several spectres of monochromatic lights. J Photochem Photobiol B Biol [Internet]. 2018;184(February):61-70. Available from:
  118. 118. Hasanloo T, Majidi E, Ardakani MRS. Flavonolignan Production in Cell Suspension Culture of Silybum marianum. 2008;46(12):876-882.
  119. 119. Shah M, Jan H, Drouet S, Tungmunnithum D, Shirazi JH, Hano C, et al. Chitosan Elicitation Impacts Flavonolignan Biosynthesis in Silybum marianum (L.) Gaertn Cell Suspension and Enhances Antioxidant and Anti-Inflammatory Activities of Cell Extracts. Molecules [Internet]. 2021;26(4). Available from:
  120. 120. Sánchez-Sampedro MA, Fernández-Tárrago J, Corchete P. Yeast extract and methyl jasmonate-induced silymarin production in cell cultures of Silybum marianum (L.) Gaertn. J Biotechnol. 2005;119(1):60-69.
  121. 121. Tůmová L, Řimáková J, Tůma J, Dušek J. Silybum marianum in vitro-flavonolignan production. 2006;2006(10):454-458.
  122. 122. Alikaridis FU, Papadakis D, Pantelia K, Kephalas T. Flavonolignan production from Silybum marianum transfored and untransfromed root cultures. Fitoterapia. 2000;71:379-384.
  123. 123. Hasanloo T, Rahnama H, Sepehrifar R, Shams M. Silymarin production in hairy root cultures of Silybum marianum L. Gaertn. Planta Med. 2008;74(09):109-114.
  124. 124. Rahnama H, Hasanloo T, Shams MR, Sepehrifar R. Silymarin production by hairy root culture of Silybum marianum (L.) Gaertn. Iran J Biotechnol. 2008;6(2):113-118.
  125. 125. Khalili M, Hasanloo T, Tabar KSK, Rahnama H. Influence of exogenous salicylic acid on flavonolignans and lipoxygenase activity in the hairy root cultures of Silybum marianum. Cell Biol Int. 2009;33(9):988-994.
  126. 126. Elateeq AA, Sun Y, Nxumalo W, Gabr AMM. Biotechnological production of silymarin in Silybum marianum L.: A review. Biocatal Agric Biotechnol [Internet]. 2020;29:101775. Available from:

Written By

Samantha Drouet and Christophe Hano

Submitted: September 24th, 2020 Reviewed: February 24th, 2021 Published: September 8th, 2021