Introduction
Plants produce a broad spectrum ofsecondary metabolites which play critical roles in plant-environmentinteractions and provide protection against biotic and abiotic stressessuch as pathogens, herbivores, drought, and ultraviolet light (
Yang et al., 2012;
Afrin et al., 2015). Secondary metabolitesare produced along with the primary metabolite pathway during plantgrowth and development. Although plant secondary metabolites are notinvolved in energy production, growth, reproduction, or other plantprimary functions, they perform important functions such as protection,attraction, and signaling (
Bernhoftet al., 2010). These plant bioactive compounds exhibitpharmacological and/or toxicological effects that could be furtherdeveloped into drugs for a wide range of diseases such as cancer,malaria, and schizophrenia (
Hickset al., 2011;
Afrin etal., 2015).
Recent studies have also shown thatmany bioactive compounds from natural products contain sugar moietyin their molecular structures. Glycosylation could enhance the physiological,selectivity, stability, solubility, and pharmacological propertiesof compounds acting as functional food additives and cosmetic ingredients(Luley-Goedl and Nidetzky, 2011;
Xiao et al., 2015,
2016). C-glycosylated flavonoidshave received less attention compared to O-glycosides (
Xiao et al., 2016). However, C-glycosylatedflavonoids are more stable than O-glycosidic bonds because of theirhigh resistance to chemical or enzyme-catalyzed hydrolysis (
Bungaruang et al., 2013).
Vitexin (apigenin 8-C-glycoside)is one of best-known C-glycosides because of its remarkable pharmacologicalactivities, which include anti-inflammatory (
Zunoliza et al., 2009) and antioxidantproperties (
Farsi et al., 2011) and a-amylase and a-glucosidase inhibition that can reducepostprandial hyperglycemia and diabetic complications (
Farsi et al., 2011;
Choo et al., 2012). It is also regardedas a marker compound for
F. deltoidea, locally known as Mas Cotek (
Shafaeiet al., 2012;
Azeminet al., 2014;
Mohd etal., 2016).
F. deltoidea, which belongs to the family Moraceae, is a popular medicinal herbin Malaysia. It has traditionally been used by the locals to treatillnesses, including fever and headache, to regulate blood sugar andblood pressure, and to control cholesterol levels (
Misbah et al. 2013). Therefore, theobjective of this study was to predict the secondary mechanism ofvitexin production in
F. deltoidea based on the identified proteins as primary building blocks. Understandingthis mechanism will enhance the production of vitexin as a potentiallead compound in drug discovery.
Material and methods
Plant material
F. deltoidea was obtained from Nursery Herba Pak Ali (Skudai, Johor, Malaysia)and authenticated by the herbarium of Universiti Kebangsaan Malaysia(Bangi, Selangor, Malaysia) under specimen number 40213. The samplewas confirmed to be F. deltoidea Jack var. trengganuensis Corner.The plant was cultivated in a mixture of sand and compost soil (1:1)and allowed to grow in natural glasshouse at the Plant BiotechnologyLaboratory, Faculty of Bioscience and Medical Engineering, UniversitiTeknologi Malaysia, 81310 Skudai, Johor Darul Takzim, Malaysia.
Protein extraction
Plant proteins were extracted fromthe leaves of
F. deltoidea accordingto the method described by
Isaacsonet al. (2006). One gram of frozen leaf tissues was homogenizedin liquid nitrogen and extracted by 10 mL cold extraction buffer consistingof 0.7 M sucrose, 0.1 M KCl, 0.5 M Tris-HCl (pH 7.5), 50 mM EDTA,1 mM phenyl methyl sulfonyl fluoride (PMSF), and 2% b-mercaptoethanol. An equal volume of Tris-bufferedphenol was added to the mixture and then the mixture was incubatedon a shaker for 30 min at 4°C. After incubation, the mixture wascentrifuged for 30 min at 5000 ×
g at 4°C. The phenol phase, which was located at the top of thetube, was carefully harvested and the remaining mixture was re-extractedby the extraction buffer in an equal volume ratio. The mixture wasincubated and centrifuged again to collect the phenol phase. Fivevolumes of cold ammonium acetate (0.1 M) in methanol was added intothe recovered phenol solution and stored at -20°C overnight. The precipitate was obtained aftercentrifugation for 30 min at 5000 ×
g at 4°C. The protein pellet was gently mixed andrinsed twice with ice-cold methanol prior to centrifugation for 10min at 5000 ×
g at 4°C.Acetone was used to perform the final wash for the pellet, and thenthe pellet was air-dried under a vacuum for 3 min. Protein concentrationwas estimated by Bradford assay (
Bradford,1976), using a serial concentration of bovine serumalbumin (0–15 µg/mL) as the standard chemical for thecalibration curve.
SDS gel electrophoresis (SDS-PAGE)
One-dimensional SDS–PAGE wasperformed to separate the extracted proteins according to the methoddescribed by Laemmli (1970). Samples were treated with rehydrationbuffer (8 M urea, 20 mM DTT, 4% CHAPS, and 5 mM Tris-Base), followedby boiling for 5 min. After centrifugation, 25 µL supernatantwas loaded onto a 12% (w/v) polyacrylamide running gel. Electrophoresiswas conducted at a constant current (20 mA) in 50 mM Tris–glycine–SDS(pH 8.3) running buffer for 2 h. A mixture of protein marker (PrecisionPlus Protein TM Prestained Standard Dual Xtra marker, BIO-RAD) wasused to determine the molecular masses of the detected proteins, andCoomassie Brilliant Blue G-250 was used to stain for protein visualization.
In-gel digestion
The protein bands on the electrophoreticgel were cut and divided into three sections according to their molecularsizes (1–20, 21–50, and 51–250 kDa). They were thendiced into smaller pieces of approximately 1–2 mm
2 in size and transferred into 1.5 mL centrifuge tubes.In-gel digestion was performed following the method explained by
Shevchenko et al. (2006). The CoomassieBrilliant Blue G-250 dye on the excised protein bands was removedwith three cycles of dehydration and hydration steps using acetonitrileand 100 mM ammonium bicarbonate, respectively. The proteins in thegel were then subjected to in situ reduction, alkylation, and finallydigestion by trypsin overnight. The peptides eluted from the gel wereconcentrated and followed by purification using C18 Zip-Tips (Merck,Millipore, USA).
LC-MS/MS
The tryptic peptides were re-suspendedin 0.1% (v/v) formic acid, and then analyzed by a micro-capillaryUltiMate 3000 (Sunnyvale, CA) system integrated with a quadrupole-time-of-flight(QTOF) mass spectrometer (AB SCIEX QSTAR Elite; Foster City, CA) witha turbo spray ionization (TIS) source. A C18 reversed phase Zorbax300SB column (150 × 0.3 mm, 5 µm) with a flow rate of5 µL/min was used for separation. A binary gradient system consistingof solvent A (water with 0.1% formic acid) and solvent B (acetonitrilewith 0.1% formic acid) was used. The LC gradient was: 0–5 min,2% B; 5–15 min, 2%–45% B; 15–16 min, 45%–80%B; 80% B hold for 2 min; 18–19 min, 80– 2% B; and 2% Bhold for 10 min. The injection volume was 5 µL.
Protein identification
The mass spectrometric data was searchedfor protein matches using MS-Fit (
University of California, 2017). The parameters were:database, NCBInr.2013.6.17 against
Arabidopsisthaliana; digest used, trypsin; maximum number of missedcleavages, 2; constant modification, carbamidomethyl; minimum matches,2; sort type, score sort; minimum parent ion matches, 1; MOWSE On,1; MOWSE P-factor, 0.4 (
Jiménezet al., 2001).
Results and discussion
Plant protein extraction
Phenol-based extraction was carriedout to extract the plant proteins, which were then recovered by phenol.The extraction buffer was composed of PMSF to inhibit proteases thatmight be released upon cell rupture during extraction. EDTA was addedinto the buffer to hinder the activities of metalloproteases and oxidasesby chelating metal ions. b-mercaptoethanol,a reducing agent, was used to protect proteins from oxidation, whilepotassium chloride was used to facilitate extraction via its saltingeffect. The addition of sucrose assisted phase separation betweenthe extraction buffer and phenol phases so that phenol could be harvestedfor high recovery of the proteins.
In the present study, phenol-basedextraction recovered 459.24 µg/g plant proteins from the leavesof F. deltoidea. The protein pelletwas white in color, indicating that few plant contaminants such aschlorophylls and pigments were co-extracted by this method. The electrophoreticgel of the extracted protein mixture is presented in Fig. 1. Thirteenintact proteins were separated on the 12% polyacrylamide gel. Themolecular size of the intact proteins ranged from 10 to 245 kDa, withmost of them being in the intermediate range of 20–135 kDa.The protein bands were excised and digested with trypsin into peptidesfor mass spectrometric analysis.
Plant protein identification
The mass spectra were analyzed andmatched to the protein database of the National Center for BiotechnologyInformation (NCBI) using MS-Fit. A total of 229 proteins were found.The identified proteins were categorized into several classes, suchas proteins involved in secondary metabolism, hormone metabolism,and signaling, according to their functional groups as stated in thegene ontology of the GoMapMan database (http://www.gomapman.org/ontology).Only proteins related to secondary metabolite production were selected,and these are listed together with their matched peptide sequencesin Table 1. Some of the proteins perform more than one function, andtherefore are grouped into the miscellaneous category.
Biosynthesis of plant secondary metabolites
The biosynthesis of secondary metabolitesis controlled by a complex network of many regulatory proteins knownas transcription factors (TFs). The transcription factors usuallybind to specific regions of promoters at the target genes, followedby activation or repression of their expressions to regulate secondarymetabolism. Members of the TF family, including putative WRKY transcriptionfactor 2, putative c-myb-like transcription factor MYB3R-4, ethylene-responsivefactor 2 (ERF2) and 10 (ERF 10), auxin response factor 10 (ARF 10),and ABRE binding factor 4, were identified in the present study. TFssuch as MYC, MYB, WRKY, and APETALA2/Ethylene-Responsive Factor (AP2/ERF)have been shown to be involved in the regulation of secondary metabolismin medicinal plants (
Afrin et al.,2015;
Dey and Corina,2015). The expression of MYB-like gene encoding enzyme,which is involved in the biosynthesis of secondary metabolites, iscontrolled by TFs. For example, PbMYB9 is an activator of the proanthocyanidin,anthocyanin, and flavonol pathways, and its function is essentialfor flavonoid biosynthesis in pear fruits (
Zhai et al., 2016).
In this study, 12-oxophytodienoatereductase 1 (OPR1) was identified. This protein is involved in thefinal step of the b-oxidation cycleto yield the end product of jasmonic acid (
Schaller, 2001). The oxylipin-typemolecule of jasmonic acid is synthesized from a-linolenic acid via the octadecanoid pathway (
Schaller, 2001;
Pauwels et al., 2009). Jasmonic acidand its derivatives are important signaling molecules for the productionof secondary metabolites in the plant kingdom (
Zhao et al., 2005;
Pauwels et al., 2009). Jasmonic acid,which also acts as a phytohormone, can synthesize a wide variety ofplant secondary metabolites, primarily terpenoids, flavonoids, alkaloids,glucosinolates, anthocyanins, and isoprenoids (
Pauwels et al., 2009). Jasmonic acidand jasmonates (methyl jasmonate) are key signaling compounds thatcontrol the expression of specific genes such as jasmonate-responsivegene 1 (jrg1), and are followed by the synthesis of jasmonate-inducedproteins (JIPs) whenever they are exposed to external stimuli suchas biotic and abiotic stress (
Kramellet al., 2000). Jasmonates induce transcription of thegene encoding phenylalanine ammonia lyase (PAL) as the key enzymeof the phenylpropanoid pathway in flavonoid synthesis (
Kašparová and Siatka, 2014). The accumulation of secondary metabolites is considered the finalconsequence of the biochemical changes induced by jasmonates (
Ishihara et al., 2002). The combinationof the jasmonic acid and ethylene signaling pathways is essentialfor plant defense responses to stresses. Ethylene may not be a commonsignal for plant induction, and the effect of ethylene on secondarymetabolite production is dependent on ethylene concentration (
Zhao et al., 2005).
Post-translational modification by transferases
Some of the identified proteins arecategorized as transferases. Transferases are involved in post-translationalmodification. For example, glycosyltransferases catalyze the transferof sugar moieties from activated donor molecules to specific acceptormolecules through the formation of O-, N-, S-, and C-glycosidic bondswith acceptors of small molecules such as sugars, lipids, proteins,or small molecules including phenylpropanoids. The glycosylation isregulated by the combination of regioselective glycosyltransferases(GTs) and glycoside hydrolases (GHs) (Le Roy et al., 2016). In thepresent study, the proteins detected in this category included putativebeta-1,3-galactosyltransferase 2, putative glycosyl hydrolase family10, putative beta-galactosidase, beta-galactosidase 5, and O-fucosyltransferasefamily protein. GTs that catalyze sugar conjugation of secondary metabolitesbelonging to the GT1 family are known as uridine diphosphate-glycosyltransferases(UGTs). UGTs are used by plants to synthesize flavonoids. UGTs facilitateglycosylation from a donor called uridine diphosphate glucose (UDPglucose).Sucrose synthase was also identified in this study. Sucrose synthaseis an enzyme (SuSy) used to catalyze a reversible sucrose conversionto fructose and UDPglucose in the presence of UDP (Bungaruang et al.,2013). In plants, the sugar donor is usually glucose, but it can alsobe galactose, xylose, rhamnose, arabinose, or glucuronic acid (Yonekura-Sakakibaraet al., 2008). Flavonoid glycosylation typically occurs at ring positionsbearing hydroxyl groups. When an acceptor has multiple binding sitesfor a sugar, UGTs exhibit regioselectivity by transferring the sugarto a specific position. Glycosylation is also regulated by glycosidehydrolases (GHs) in the hydrolysis and/or rearrangement of glycosidicbonds (Le Roy et al., 2016). Therefore, transferases are very importantfor vitexin synthesis in F. deltoidea. They are involved in the laterstage of translation after flavanone synthesis.
Many phenylpropanoid pathway-derivedproducts are toxic and unstable molecules, and therefore they seldomaccumulate as their aglycones in plants (
Alejandro et al., 2012;
Väisänen et al., 2015). Glycosylation cantherefore reduce phenylpropanoid toxicity and increase stability andsolubility and can also influence compartmentalization and biologicalactivity (
Le Roy et al., 2016). Among the diverse phenylpropanoids, flavonoids are probably thebest-characterized molecules in terms of glycosylation owing to theirbroad medical and commercial benefits. Natural flavonoids are presentin the form of O-glycosides or C-glycosides in plants (
Xiao et al., 2015).
C-glycosylated flavonoids have beenfound to possess better therapeutic properties than O-glycosylatedflavonoids. C-glycosides can act as siderophores, antibiotics, antioxidants,attractants, and feeding deterrents (
Brazier-Hicks et al., 2009). C-glycosylated flavonoidsare also more stable against the activity of glycosidases under hydrolyticconditions (
Rawat et al., 2009). Among the reported C-glycosides, flavone C-glycosides, especiallyvitexin, isoorientin, orientin, isovitexin, and their multi-glycosideshave been frequently mentioned in the literature. O-glycosides arelikely to have lower in vivo lifetimes (
Bungaruang et al., 2013).
Vitexin (apigenin-8-C-glucoside)and isovitexin (apigenin-6-C-glucoside) are the main C-glycosylatedflavonoid constituents of
F. deltoidea. Isovitexin is an isomer of vitexin, and its C-glycosylation happensat C-6. They are considered the chemical markers for this herb (
Azemin et al., 2014). Often, C-glycosylflavonoids are produced in greater amounts than O-glycosyl flavonoidsby weight (
Courts and Williamson,2015). The level of vitexin in the leaf extracts of
F. deltoidea was found to be higher thanthat of isovitexin. For instance, 12.31 mg/g vitexin and 4.81 mg/gisovitexin were reported in methanol extract, and 6.20 mg/g vitexinand 0.81 mg/g isovitexin were found in water extract (
Shafaei et al., 2012). Another studythat assessed the content of vitexin and isovitexin in
F. deltoidea varieties using the high-performanceliquid chromatographic method found that vitexin and isovitexin comprised1.53% and 0.9% of a 10 mg leaf sample (
Mohd et al., 2016).
The chemical name of vitexin is 8-D-glucosyl-4′,5,7-trihydroxy-flavone,and is known as ‘Mujingsu’ in Chinese. It is an activecomponent in many traditional Chinese medicines. To our knowledge,no different pharmacological properties of vitexin and isovitexinhave been reported to date.
Praveenaet al. (2013) reported that C-8 glycosylation decreasedthe negative charge on the oxygen atom at the C-3 position, leadingto better antioxidant potency of vitexin as compared to apigenin.However,
Brazier-Hicks and Edwards(2013) revealed that vitexin and isovitexin exhibitedsimilar pharmacological effects, partly owing to their similarityin chemical structures. Both C-glycosides were found to possess antioxidant(
Farsi et al., 2011)and anti-inflammatory properties (
Zunoliza et al., 2009), as well as a–amylase and a-glucosidase inhibition (
Farsi etal., 2011;
Choo et al.,2012), which may reduce postprandial hyperglycemia anddiabetic complications.
Secondary mechanism of vitexin synthesis
A large number of plant secondarymetabolites derive from phenylalanine and tyrosine as precursors viathe phenylpropanoid pathway (
Le Royet al., 2016). Hence, flavonoids are considered to bephenylpropanoid-derived compounds (
Stafford, 1991). Flavonoids are synthesized from thecondensation of p-coumaroyl-CoA with three malonyl-CoA molecules bychalcone synthase (CHS), which in turn produces a flavanone containinga 2-phenylchroman backbone (
Le Royet al., 2016). The 2-phenylchroman backbone is the basicstructure of flavanols, isoflavonoids, flavonols, flavones, and anthocyanidins.In the present study, flavonol synthase (FLS) was detected; FLS isused to synthesize flavonols through dihydroflavonols as intermediates.FLS expression can be induced by several abiotic stresses includingUV-B, abscisic acid, cold, sucrose, salicylic acid, and ethephon.Previous studies indicate that detection of FLS also explains thepresence of flavonols such as myricetin and kaempferol (
Dzolin et al., 2015).
Xu et al. (2012) revealed that FLScould also catalyze the formation of dihydrokaempferol to kaempferol(flavonol) and the conversion of kaempferol from naringenin (flavanone).
Biosynthesis of vitexin in F. deltoidea follows four major steps ina mechanistic pathway: condensation by plant polyketide chalcone synthase(CHS), isomerization either spontaneously or catalyzed by chalconeisomerase (CHI), oxidation by cytochrome P450 (CYP P450) to convertflavanone to flavone, and finally the transfer of sugar moiety byC-glycosyl transferase (CGT), followed by a dehydration step to produceflavone-6-C-glucosides, as illustrated in Fig. 2.
CHS is one of the plant type IIIpolyketide synthases (PKS). CHS is the first committed enzyme in thebiosynthesis of flavonoids and directs carbon flux from the generalphenylpropanoid metabolism to the flavonoid pathway (
Saito et al., 2013;
Zhang et al., 2017). It initiatesthe loading of p-coumaroyl-CoA to its active site, followed by twomalonyl-CoA units, until a linear tetraketide chain is created, whichis then circularized to form the chalcone product through an aromatase-likemechanism (
Austin and Noel, 2003). The spontaneous cyclization of the triketide intermediate resultsin the formation of naringenin chalcone. Lussier et al., (2013) suggestedthat naringenin chalcone was produced via a Claisen cyclization reaction.CHS initiates the elongation of p-coumaroyl-CoA to a C15 skeletonresulting branch of the phenylpropanoid pathway, and produces a varietyof stress-induced compounds and pigments. Light is one of the mostimportant factors triggering flavonoid biosynthesis and inductionof light-responsive gene expression. This may explain why flavonoidsare scarcely produced in plants grown in the dark, since there isa lack of genes encoding expression for CHS (
Kaltenbach et al., 1999).
The second step of flavone synthesisis the isomerization of chalcone to flavanone by chalcone isomerase.CHI catalyzes the stereospecific cyclization of naringenin chalconeto (2S)-naringenin (
Saito et al.,2013). Although this step can occur spontaneously, CHI-catalyzedisomerization is approximately 107-fold more efficient than spontaneousisomerization (Bednar and Hadcock, 1988). The non-enzymatic conversionof chalcones yields racemic (2R/S)-flavanones. Since only (2S)-flavanonesare intermediates of the subsequent flavonoid pathway, CHI specificityguarantees efficient formation of biologically active (2S)-flavanone(Bednar and Hadcock, 1988;
Cheng etal., 2011). The expression of genes encoding for CHIis also upregulated by UV-B irradiation (
Cheng et al., 2011).
After isomerization of chalcone toflavanone by chalcone isomerase (CHI), the subsequent pathway branchesto several different flavonoid classes, including aurones, dihydrochalcones,flavanonols (dihydroflavonols), isoflavones, flavones, flavonols,leucoanthocyanidins, anthocyanins, and proanthocyanidins (
Mierziak et al., 2014). Flavonesare synthesized from flavanones by the introduction of a double bondbetween the C-2 and C-3 positions by flavone synthase (FNS) (Martensand Mithofer, 2005). FNS converts naringenin either directly to apigenin(flavones) or 2-hydroxyflavanones by FNS II (Fig. 2). The conversionof naringenin to 2-hydroxyflavanones is catalyzed by flavanone-2-hydroxylase(F2H) (
Yonekura-Sakakibara and Hanada,2011). It is noteworthy that FNS II can also have F2Hor FNS activity. FNS II functions as oxygen- and NADPH-dependent CYPP450 membrane-bound monooxygenases, which are widespread among higherplants. FNS II belongs to the CYP 93B subfamily for dicots and theCYP 93G subfamily for monocots (Martens and Mithofer, 2005). CYP P450-linkedenzymes are implicated in the biosynthesis of various structural,growth regulatory, and protective substances in plant cells via numerousmetabolic pathways. They contribute to the stable equilibrium of phytohormonesand signaling molecules by controlling their biosynthesis and catabolism.They are involved in the biosynthesis of pigments, volatiles, antioxidants,allelochemicals, and defense compounds, including phenolics and theirconjugates, flavonoids, coumarins, lignans, glucosinolates, cyanogenicglucosides, benzoxazinones, isoprenoids, and alkaloids (
Morant et al., 2003;
Mizutani and Sato, 2011).
In this study, five CYP P450s-basedproteins were detected (Table 1). CYP 706 is involved in terpenoidmetabolism. CYP 71A28 was previously known as CYP 713A2, belongingto the subfamily of CYP 71 A, which has been shown to have monoterpenehydroxylase activity. CYP 87 is involved in plant hormone metabolism,and LACERATA (CYP 86A8) is involved in fatty acid metabolism (
Bak et al., 2011). Plant CYP P450sare bound to membranes, usually anchored on the cytoplasmic surfaceof the endoplasmic reticulum. They need to be coupled to electron-donatingproteins such as CYP P450 reductases or CYP b5 for activation (
Bak et al., 2011). Even though allCYP P450s found in this study are related to terpenoid, hormone, andfatty acid biosynthesis, proteins of CYP P450s are also known to havemulti-catalyzing functions. Therefore, the identified CYP P450s mightalso be involved in the introduction of double bond to the C-2 andC-3 positions in order to convert naringenin to 2-hydroxyflavanonesin third step of vitexin biosynthesis.
However, little is known about flavone-C-glycosidebiosynthesis. Flavanone, which is the core intermediate of the flavonepathway, is most likely to be a precursor. This reaction is mediatedby C-glycosyltransferase (CGT), which catalyzes the formation of flavone-C-glycosidesfrom flavanone precursors (
Brazier-Hickset al., 2009). Figure 2 shows that flavanones are hydroxylatedby F2H/FNS II to form 2-hyroxylflavanones, and then glycosylated to2-hydroxylflavanone C-glycosides by CGT. This C-glycosylated 2-hydroxyflavanoneis consequently dehydrated to produce flavone C-glycosides.
Kerscherand Franz (1987) reported that CGT from
Fagopyrum esculentum seedlings catalyzedthe transfer of glucose to 2-hydroxyflavanones. Naringenin, naringenin-chalcone,and flavones such as apigenin and chrysin cannot act as glucosyl acceptorsin C-glycosyl-flavonoid biosynthesis. C-glycosylation takes placeafter dehydration of 2-hydroxynaringenin. This finding was also supportedby another study on cereal crops where C-glycosylated flavones weresynthesized through the action of CGT and dehydratase on 2-hydroxyflavanones(
Brazier-Hicks et al., 2009). The dehydration step from unstable 2-hydroxyflavanones can occurspontaneously or can be catalyzed by an enzyme (
Akashi et al., 2005). However, flavone-6-C-glucosidesare preferentially formed in plants. 2-hydroxyflavanone conjugatesundergo spontaneous dehydration to yield a mixture of flavone-6-C-and flavone-8-C-glucosides (
Brazier-Hickset al., 2009). Isoflavone is formed by 1,2-eliminationof water from 2-hydroxyisoflavanone catalyzed by 2-hydroxyisoflavanonedehydratase (HID). HID displays clear substrate specificity and isdistinguishable from differently substituted 2-hydroxyisoflavanone.Histidine and aspartic acid are critical residues for HID catalysis(
Hakamatsuka et al., 1998;
Akashi et al., 2005). The spontaneous dehydration to form isoflavones is slow comparedto the enzyme-catalyzed reaction. This suggests that production offlavones in plants primarily depends on enzymes. The non-enzymaticslow production of isoflavone in plants becomes an alternative process(
Akashi et al., 2005).
CGT from
Zea mays displays O-glycosylation activity toward naringenin,but C-glycosylation activity toward 2-hydroxynaringenin (
Falcone Ferreyra et al., 2013). Thisindicates that the activity of CGT is highly specific to 2-hydroxynaringeninand its derivatives, such as 2,4′,5,7-tetrahydroxyflavanoneand 2,5,7-trihydroxyflavanone. CGT does not accept flavanones, flavones,or flavonols as glucose acceptors (
Kerscher and Franz, 1987;
Nagatomo et al., 2014). Vitexin and isovitexin havebeen found to be present only in certain plants, especially in plantswith medicinal values (
He et al.,2016). Medicinal plants such as pearl millet (
El Amrani et al., 2004), hawthorn(
Akashi et al., 1999),pigeon pea (
Brazier-Hicks et al.,2008;
Crosby et al.,2011), mung bean (
Crozieret al., 2009), mosses (
Day et al., 2003;
Dixonet al., 2009), Passiflora (
Du et al., 2010a, 2010b), bamboo (
Dürr et al., 2004;
François et al., 2004), mimosa (
Ha et al., 2010), wheat leaves (
Halpin et al., 1999), and chaste tree or chaste berry (
Hasegawa et al., 2007) have been previously reportedto contain flavones.
PKS has three processing domains;namely, the ketoreductase (KR), dehydratase (DH), and enoylreductase(ER) domains (
Li et al., 2015). The dehydratase (DH) domain of PKS was identified in this study.Dehydratase domains of PKSs function to generate an a,b-unsaturatedbond through a dehydration reaction in a
cis or
trans configuration (Akeyet al., 2010). This catalyzes the formation of the unsaturated triketideintermediate using malonyl-CoA as the chain extension substrate (
Wu et al., 2005). The double bondsare formed by DH domains through abstraction of the a-proton and concomitant protonation of the b-hydroxyl group of the nascent b-hydroxyacyl-ACP polyketide intermediate,resulting in loss of one water molecule (
Li et al., 2015). The active site of DH contains ahistidine residue from the N-terminal and an aspartate residue fromthe C-terminal. Histidine and aspartic are conserved across DH-containingenzymes (
Wu et al., 2005; Akey et al., 2010;
Ishikawa etal., 2012;
Li et al.,2015). The products of all DH reactions contain a, b-doublebonds conjugated with the thioester carbonyl (acetyl-CoA) (Akey etal., 2010). Several similarities have been observed between DH domainof PKS and HID. Both proteins possess conserved histidine and asparticacid residues at their active sites for catalytic activity. They catalyzeformation of an a, b-double bond by the loss of a water molecule(
Li et al., 2015). Theyalso display high substrate specificity, where double bond confirmationdepends on the chirality of the b- OH substrate. Thus, the DH domainof PKS identified from this study might be responsible for the dehydrationstep to produce corresponding vitexin in
F.deltoidea leaves.
Conclusions
Vitexin biosynthesis in F. deltoidea was predicted to follow a four-stepmechanism involving plant polyketide chalcone synthase (CHS); isomerization,either spontaneous or catalyzed by chalcone isomerase (CHI); oxidationby CYP P450 to convert flavanone to flavone; and the transfer of sugarmoiety by C-glycosyltransferase (CGT), followed by dehydration toproduce flavone-8-C-glucosides. The detection of the proteins alsosupports previous findings that vitexin is present in medicinal plants. F. deltoidea is a traditional medicinal plantthat is widely used by indigenous peoples in South-east Asian countries.Further studies are required to elucidate the related biochemicalpathways in order to trigger vitexin production.
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