identification and characterization of …...identiﬁcation and characterization of daurichromenic...

Identification and Characterization of DaurichromenicAcid Synthase Active in Anti-HIV Biosynthesis1

Miu Iijima,a Ryosuke Munakata,b,2 Hironobu Takahashi,c Hiromichi Kenmoku,c Ryuichi Nakagawa,a

Takeshi Kodama,d Yoshinori Asakawa,c Ikuro Abe,e Kazufumi Yazaki,b Fumiya Kurosaki,a andFutoshi Tauraa,3

aLaboratory of Medicinal Bioresources, Graduate School of Medicine and Pharmaceutical Sciences forResearch, University of Toyama, Sugitani, Toyama 930-0194, JapanbLaboratory of Plant Gene Expression, Research Institute for Sustainable Humanosphere, Kyoto University,Gokasho, Uji 611-0011, JapancInstitute of Pharmacognosy, Faculty of Pharmaceutical Sciences, Tokushima Bunri University, Yamashiro-cho,Tokushima 770-8514, JapandDepartment of Chemical and Biological Engineering, Akita National College of Technology, Iijimabunkyo-cho, Akita 011-8511, JapaneLaboratory of Natural Products Chemistry, Graduate School of Pharmaceutical Sciences, University of Tokyo,Tokyo 113-0033, Japan

ORCID IDs: 0000-0003-0045-7716 (H.T.); 0000-0001-8961-7362 (H.K.); 0000-0002-1847-412X (Y.A.); 0000-0002-3640-888X (I.A.);0000-0001-6551-123X (F.T.).

Daurichromenic acid (DCA) synthase catalyzes the oxidative cyclization of grifolic acid to produce DCA, an anti-HIVmeroterpenoid isolated from Rhododendron dauricum. We identified a novel cDNA encoding DCA synthase by transcriptome-based screening from young leaves of R. dauricum. The gene coded for a 533-amino acid polypeptide with moderate homologiesto flavin adenine dinucleotide oxidases from other plants. The primary structure contained an amino-terminal signal peptide andconserved amino acid residues to form bicovalent linkage to the flavin adenine dinucleotide isoalloxazine ring at histidine-112and cysteine-175. In addition, the recombinant DCA synthase, purified from the culture supernatant of transgenic Pichia pastoris,exhibited structural and functional properties as a flavoprotein. The reaction mechanism of DCA synthase characterized hereinpartly shares a similarity with those of cannabinoid synthases from Cannabis sativa, whereas DCA synthase catalyzes a novelcyclization reaction of the farnesyl moiety of a meroterpenoid natural product of plant origin. Moreover, in this study, wepresent evidence that DCA is biosynthesized and accumulated specifically in the glandular scales, on the surface of R. dauricumplants, based on various analytical studies at the chemical, biochemical, and molecular levels. The extracellular localization ofDCA also was confirmed by a confocal microscopic analysis of its autofluorescence. These data highlight the unique feature ofDCA: the final step of biosynthesis is completed in apoplastic space, and it is highly accumulated outside the scale cells.

Rhododendron dauricum (Ericaceae), distributed innortheastern Asia, produces unique secondary metab-olites, including daurichromenic acid (DCA; Fig. 1A), anovel meroterpenoid composed of orsellinic acid andsesquiterpene moieties (Kashiwada et al., 2001). DCAhas attracted considerable attention as a medicinalresource because this compound shows variouspharmacological activities (Iwata et al., 2004;Hashimoto et al., 2005). Especially, DCA has beenone of the most effective natural products with anti-HIV properties, as shown in experiments with acutelyinfected H9 cells, in which the EC50 value of DCA(15 nM) was smaller than that of the positive controldrug azidothymidine (44 nM; Lee, 2010). Thus, chem-ical synthesis of DCA has been studied extensivelyover the past few years (Liu andWoggon, 2010; Bukhariet al., 2015).

1 This work was supported in part by JSPS/MEXT KAKENHI(grant nos. 15K07994 and 17H05436 to F.T) and JSPS Core-to-Core Program, B, Asia-Africa Science Platforms (F.T. is one ofthe recipients).

2 Current address: Laboratoire Agronomie et Environnement, Uni-versité de Lorraine UMR 1121, 2 avenue de la forêt de Haye TSA40602, 54518 Vandœuvre-lès-Nancy, France.

3 Address correspondence to [email protected] author responsible for distribution of materials integral to the

findings presented in this article in accordance with the policy de-scribed in the Instructions for Authors (www.plantphysiol.org) is:Futoshi Taura ([email protected]).

F.T. conceived the research plans; M.I. performed most of the ex-periments; R.M. and K.Y. provided technical assistance to M.I.; H.T.and H.K. analyzed bioinformatics data; R.N. and T.K. performedchemical syntheses and phytochemical analyses; Y.A., I.A., K.Y.,and F.K. supervised this research; M.I., R.M., I.A., K.Y., F.K., andF.T. wrote the article.

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http://orcid.org/0000-0002-3640-888X

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http://orcid.org/0000-0002-3640-888X

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With respect to the biosynthesis of DCA, we previ-ously reported partial characterization of an oxidocy-clase, named DCA synthase, using a crude proteinextract from young leaves of R. dauricum (Taura et al.,2014). DCA synthase is an enzyme that catalyzes thestereoselective oxidative cyclization of the farnesylmoiety of grifolic acid to form DCA (Fig. 1A). UnlikeP450-type cyclases involved in glyceollin and fur-anocoumarin biosynthesis (Welle and Grisebach, 1988;Larbat et al., 2007), DCA synthase is a soluble proteinand does not need exogenously added cofactors for thereaction. Remarkably, these features are similar to thosereported for cannabinoid synthases from Cannabis sat-iva (Fig. 1B). Hitherto, three cannabinoid synthases,tetrahydrocannabinolic acid (THCA) synthase, canna-bidiolic acid (CBDA) synthase, and cannabichromenicacid (CBCA) synthase, have been identified and char-acterized (Taura et al., 1995, 1996; Morimoto et al.,1997). All cannabinoid synthases catalyze the oxido-cyclization of the geranyl group of a common substrate,cannabigerolic acid, to form novel ring systems. Previ-ous structural and biochemical studies have demon-strated that THCA synthase and CBDA synthase areflavoprotein oxidases belonging to the vanillyl alcoholoxidase (VAO) flavoprotein family (Sirikantaramaset al., 2004; Taura et al., 2007). These cannabinoid syn-thases catalyze reactions using covalently linked FADas the coenzyme and molecular oxygen as the finalelectron acceptor. CBCA synthase, which synthesizes achromene ring similar to that in DCA biosynthesis, hasnot been characterized at the molecular level, whereasCBDA synthase shares similar biochemical properties

with THCA synthase (Morimoto et al., 1997). In con-trast to these cannabinoid synthases, DCA synthase hasneither been cloned nor purified. Therefore, the struc-tural and functional characteristics of this enzyme re-main unclear.

VAO flavoprotein family members have divergentfunctions and are widely distributed among plants, an-imals, and microorganisms (Leferink et al., 2008). Apartfrom cannabinoid synthases, several VAO family en-zymes, involved in a variety of plant specialized path-ways, have been identified to date, such as berberinebridge enzyme (BBE), (S)-tetrahydroprotoberberineoxidase, and dihydrobenzophenanthridine oxidase inbenzylisoquinoline alkaloid biosynthesis (Kutchan andDittrich, 1995; Gesell et al., 2011; Hagel et al., 2012);AtBBE-like13 and AtBBE-like15 (monolignol oxidases)in lignin biosynthesis (Daniel et al., 2015); and carbohy-drate oxidases involved in the defense against microbialpathogens to supply hydrogen peroxide as a by-product(Custers et al., 2004). Recently, crystal structures ofEschscholzia californica BBE (EcBBE; Winkler et al., 2008),AtBBE-like15 (Daniel et al., 2015), and THCA synthase(Shoyama et al., 2012) have been determined, and thestructural basis of the enzymatic reactions was char-acterized in detail. As a common feature, these plantenzymes were proven to bicovalently bind to FAD co-enzyme via a novel 6-S-cysteinyl,8a-N1-histidyl linkage,which was first identified in a bacterial FAD oxidase,glucooligosaccharide oxidase from Acremonium strictum(Huang et al., 2005).

Among the identified members of the VAO family,THCA synthase and CBDA synthase, producing major

Figure 1. Biosynthesis of plant meroterpenoidsvia oxidative cyclization of isoprenoid moieties.A, DCA biosynthesis in R. dauricum catalyzedby DCA synthase. B, Cannabinoid biosynthesisin C. sativa catalyzed by THCA synthase, CBDAsynthase, and CBCA synthase.

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cannabinoids, have long been the only examples tocatalyze the oxidocyclization of the prenyl moiety in ameroterpenoid natural product (Baunach et al., 2015).Thus, DCA synthase, mediating a reaction similar tothose of cannabinoid synthases, is an interesting en-zyme in which to study the structural and functionalproperties. As the initial step to the detailed studies ofDCA synthase, we attempted to isolate the cDNA of thegene encoding DCA synthase, based on a homologysearch against the translated R. dauricum young leaftranscriptome, using cannabinoid synthases as queries.Heterologous expression of the recombinant proteins ina Pichia pastoris system provided evidence that one ofthe candidate cDNAs is of a gene that encodes an activeDCA synthase. In this study, we describe the molecularcloning and biochemical characterization of DCA syn-thase, a novel member of the meroterpenoid cyclase-type flavoprotein oxidases.In our previous study, we confirmed that the mer-

oterpenoid metabolites (DCA and grifolic acid) as wellas DCA synthase activity are localized predominantlyto young leaves of R. dauricum (Taura et al., 2014). Thistissue distribution also is notable because young leavesof R. dauricum, a kind of lepidote Rhododendron species,are covered with numerous trichomes called glandularscales (Desch, 1983). Glandular scale is a multicellularepidermal organ that contains secondary metabolitessuch as sesquiterpenoids, including germacrone, andthese metabolites are thought to participate in repellinginsects hazardous to this plant (Doss, 1984; Doss et al.,1986). However, little information is available on therelationship between glandular scales and DCA bio-synthesis. In this study, we provide detailed evidencethat DCA is biosynthesized and accumulated primarilyin the glandular scales of young leaves. We also discusspossible reasons why this specialized metabolite isproduced in the specialized epidermal organ, theglandular scales of R. dauricum.

RESULTS

Molecular Cloning and Sequence Analysis of DCASynthase Candidates

Three cDNA contigs encoding FAD oxidase (FADOX)were identified as candidate cDNAs for DCA synthasefrom a young leaf transcriptome database ofR. dauricum,as described in “Materials andMethods,” and they weretentatively designated as RdFADOX1 toRdFADOX3. Toisolate their full-length cDNA sequences, we performed5ʹ- and 3ʹ-RACE to obtain the terminal regions of therespective genes. Then, the coding regions of RdFA-DOX1 to RdFADOX3 cDNAs were amplified usinggene-specific primers, which were subcloned into theexpression vector pPICZA, and the sequences werereconfirmed.The RdFADOX1 to RdFADOX3 genes contained

1,602-, 1,590-, and 1,587-bp open reading frames encoding533-, 529-, and 528-amino acid-long polypeptides,

respectively. The deduced amino acid sequences exhibi-ted ;50% identities among each other and showed from40% to 50% identities with functionally characterizedplant FAD oxidases, including THCA synthase, EcBBE,and AtBBE-like15 (Supplemental Fig. S1A). The RdFA-DOX amino acid sequences contained an FAD-bindingdomain 4 (pfam01565; Gesell et al., 2011) in theirN-terminal halves (Supplemental Fig. S1B). In addition,multiple sequence alignment confirmed that the His andCys residues that form the bicovalent linkage with theFAD coenzyme in structurally characterized FAD oxi-dases also are conserved in RdFADOXs (SupplementalFig. S1A).

The extracellular localization of RdFADOX1 toRdFADOX3 and the presence of N-terminal signalpeptides with 24, 24, and 21 amino acids, respectively,were predicted using online software such as PSORTand TargetP (Supplemental Fig. S1B). Thus, the maturepeptide sequences of RdFADOX1 to RdFADOX3 arepresumed to be of 509, 505, and 507 amino acids withmolecular masses of 57.2, 56.4, and 56.8 kD, respectively.In addition, several Asn glycosylation motifs werefound in respective RdFADOX primary structures.Based on these sequence characteristics, RdFADOX1to RdFADOX3 are predicted to be secreted FAD ox-idases, similar to cannabinoid synthases (Sirikantaramaset al., 2004; Taura et al., 2007).

Among these candidate proteins, RdFADOX1 wasrelatively remarkable because, in the phylogeneticanalysis based on the protein alignment with function-ally characterized plant FAD oxidases, the RdFADOX1sequence was the nearest neighbor to cannabinoid syn-thases to form a subclade, whereas RdFADOX3 wasclosest to carbohydrate oxidases and RdFADOX2 stoodindependently (Fig. 2). In addition, a relatively hightranscript level of RdFADOX1 in the young leaf tran-scriptome was estimated by calculating FPKM (frag-ments per kilobase of transcript per million fragmentsmapped) values (Trapnell et al., 2010) for the respectiveRdFADOX open reading frames. The FPKM value forRdFADOX1 was 74.7, which was much higher thanthose calculated for RdFADOX2 and RdFADOX3 (11.2and 29.6, respectively). Notably, the gene for orcinolsynthase (Taura et al., 2016; GenBank accession no.LC133082), the polyketide synthase involved in orsel-linic acid biosynthesis in the DCA pathway, also is ahighly expressed gene with an FPKM value of 81.9.

Heterologous Protein Expression in P. pastoris Culture

To examine which RdFADOX gene encodes DCAsynthase, the pPICZA plasmids containing full-lengthcDNAs were introduced to P. pastoris KM71H by elec-troporation. The transgenic P. pastoris colonies werecultured individually in minimal liquid medium, andprotein expression was induced by adding methanol.Enzyme assays using the cell extract and culture me-dium of the respective transgenic cultures clearlydemonstrated that the RdFADOX1 gene encodes an

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active DCA synthase, because the culture supernatantexhibited a clear DCA-producing activity from grifolicacid (Fig. 3A; 30–40 pkat mg21 protein). In contrast,the cell extract prepared from the same culture exhibitedno enzyme activity (Fig. 3A), suggesting that the re-combinant DCA synthase was effectively secreted fromP. pastoris, likely because the N-terminal signal peptidealso was functional in P. pastoris. The enzymaticallysynthesized DCA showed retention time and molecularmass identical to those of the authentic DCA sample onliquid chromatography (LC)-electrospray ionization(ESI)-mass spectrometry (MS) analysis (Fig. 3B). In ad-dition, a chiral HPLC analysis revealed that the recom-binant enzyme produced (+)-DCA as the predominantreaction product with an enantiomeric excess value of;92% (Supplemental Fig. S2). This stereoselectivity ofthe recombinant enzyme was similar to that of nativeDCA synthase (Taura et al., 2014). Based on these results,the RdFADOX1 gene cDNAwas confirmed to be the onecoding for DCA synthase. In contrast, neither the cellextract nor the culture medium prepared from thetransgenic P. pastoris harboring the RdFADOX2 orRdFADOX3 gene showed DCA synthase activity,suggesting that these proteins might contribute tooxidative reactions other than DCA biosynthesis inR. dauricum plants.

The recombinant DCA synthase in P. pastoris culturemedium was immediately purified by column chroma-tography on hydroxylapatite, followed by TOYOPEARLCM650M. On SDS-PAGE, the purified enzyme was ob-served as a broad protein band with an average molec-ular mass of;74 kD (Fig. 4). The native molecular massof the purified enzyme was estimated to be ;68 kD,based on the elution volume on gel filtration chroma-tography, demonstrating that DCA synthase is a mon-omeric protein, as reported for cannabinoid synthases(Taura et al., 1995, 1996). The N-terminal amino acidsequence of the purified protein was Ala-His-Thr, whichmatched to the tripeptide at the 25th position of DCAsynthase. Thus, the predicted 24-amino acid signalpeptide was correctly cleaved during the proteinsynthesis in P. pastoris. Because the molecular mass ofthe purified enzyme was apparently larger than thetheoretical value for the mature polypeptide (57.2 kD),carbohydrate attachment to the protein was suspected.As shown in Supplemental Figure S3, the purifiedenzyme was stained with periodic acid-Schiff sugarstaining reagent, and in addition, endoglycosidase Htreatment afforded an ;58-kD convergent proteinband that is insensitive to sugar staining, confirmingthat the recombinant DCA synthase was highly Asnglycosylated. This result was reasonable because the

Figure 2. Phylogenetic tree representing plant FAD oxidases. The bacterial enzyme Acremonium strictum glucooligosaccharideoxidase (AsGOOX) was used as an outgroup. Bootstrap values are presented at each node. The scale represents 0.5 aminoacid substitutions per site. Abbreviations for species are as follows: Am, Argemone mexicana; At, Arabidopsis thaliana;Bs, Berberis stolonifera; Bw, Berberis wilsoniae; Cj, Coptis japonica; Cs, Cannabis sativa; Ec, Eschscholzia californica;Ha, Helianthus annuus; Ls, Lactuca sativa; Ps, Papaver somniferum; Rd, Rhododendron dauricum. Abbreviations forenzymes are as follows: BBE, berberine bridge enzyme; CBDAS, CBDA synthase; CHO, carbohydrate oxidase; DBOX,dihydrobenzophenanthridine oxidase; DCAS, DCA synthase; FADOX, FAD oxidase; STOX, (S)-tetrahydroprotoberberineoxidase; THCAS, THCA synthase. The NCBI protein registration numbers are as follows. BsBBE, AAD17487.1; CjBBE,BAM44344.1; EcBBE, AAC39358.1; PsBBE, AAC61839.1; CsCBDAS, BAF65033.1; HaCHO, AAL77103.1; LsCHO,AAL77102.1; PsDBOX, AGL44334.1; AsGOOX, AAS79317.1; AmSTOX, ADY15027.1; BwSTOX, ADY15026.1; CjSTOX,BAJ40864.1; CsTHCAS, BAC41356.1.


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mature region of DCA synthase contains six Asn gly-cosylation motifs (Supplemental Fig. S1B).The concentrated solution of the recombinant

DCA synthase was yellowish, and transilluminationof the enzyme at 310 nm showed a clear in-gel auto-fluorescence as in the case of known flavoproteins(Kutchan and Dittrich, 1995; Sirikantaramas et al., 2004;Taura et al., 2007; Fig. 4). Because boiling of the enzymein a denaturing buffer for SDS-PAGE failed to releasethe compound showing fluorescence, it was suggestedthat the fluorescent molecule might be covalentlylinked to the enzyme. The purified DCA synthaseshowed absorption maxima at 385 and 450 nm, whichquite resembled that of authentic FAD (Fig. 5A). Inaddition, the fluorescence emission of DCA synthaseexhibited a maximum at 624 nm, and supplementation

with sodium dithionite, the flavin-reducing agent (Fox,1974), resulted in a complete quenching of fluorescence(Fig. 5B). Taken together, these properties were con-sistent with those of flavoproteins, indicating that DCAsynthase covalently binds to flavin coenzyme via con-served flavinylation residues (Supplemental Fig. S1).To obtain more structural information on the flavinmolecule attached to DCA synthase, we attempted tohydrolyze the flavin by boiling the enzyme in 10 mM

HCl for 10 min as described by Kutchan and Dittrich(1995). Consequently, AMP was clearly detected byHPLC and MS analyses in the acid-hydrolyzed sample(Supplemental Fig. S4), suggesting that the enzyme-bound FAD was hydrolyzed to release AMP as repor-ted for EcBBE (Kutchan and Dittrich, 1995). Therefore,the flavin molecule in DCA synthase is most likelyFAD.

Biochemical Properties of the DCA Synthase Reaction

Using the purified recombinant enzyme, the bio-chemical properties of the DCA-producing reactionwere analyzed. First, as described previously for thenative enzyme (Taura et al., 2014), the DCA synthasereaction did not need exogenously added redox coen-zymes such as FAD (Supplemental Table S1). Divalentmetal ions, which are often used as cofactors for class Iand class II terpene cyclases (Baunach et al., 2015), alsoexhibited little effect on enzyme activity (SupplementalTable S1). In contrast, the reaction absolutely requiredmolecular oxygen, because an oxygen-removing treat-ment using Glc oxidase and catalase in the presence ofGlc (Fabian, 1965) completely abolished DCA synthaseactivity (Table I). Molecular oxygen would participateas the final electron acceptor in the reaction, because

Figure 4. SDS-PAGE of the purified recombinant DCA synthase. LaneM, Molecular mass standards with indicated molecular masses; lane 1,purified DCA synthase (2 mg); lane 2, the same sample as in lane 1 vi-sualized by transillumination at 310 nm.

Figure 3. Product analyses of the reaction catalyzed by enzyme solu-tion prepared from the transgenic P. pastoris expressing DCA synthase.A, HPLC elution profiles of DCA standard (top), the reaction mixturewith culture medium (middle), and the reaction mixture with cellularextract (bottom). mAU, Milliabsorbance unit. B, ESI-MS (negativemode) of DCA standard (left) and DCA synthesized by culture mediumas an enzyme (right).











hydrogen peroxide, almost in equal quantities to thoseof DCA, was detected in the reaction mixture with amolar ratio of 1.07 (hydrogen peroxide) versus 1 (DCA).Thus, the reaction stoichiometry of DCA synthase issummarized as follows: grifolic acid + O2 → DCA +H2O2. Furthermore, the oxidative reaction is likely toproceed through an ionic intermediate rather thanradical species, because two typical radical trap agents,2-methyl-2-nitrosopropane (Makino et al., 1981) andN-acetyl-Cys (Ates et al., 2008), did not inhibit the enzymeactivity (Supplemental Table S1). These biochemicalproperties of the recombinant DCA synthase werequite similar to those reported previously for FADoxidases, including cannabinoid synthases (Kutchanand Dittrich, 1995; Sirikantaramas et al., 2004; Tauraet al., 2007).

The substrate specificity of DCA synthase was ana-lyzed using grifolic acid and its analogs, and the kineticconstants for each substrate were calculated. First, therecombinant DCA synthase oxidized grifolic acid withlowest Km and highest kcat values among the substratestested herein (Table II). The resulting catalytic efficiency(kcat/Km) was 5.61, which is similar to that reported forEcBBE (Gaweska et al., 2012) and more than 1,000-foldhigher than those of THCA synthase and CBDA syn-thase (Supplemental Table S2; Taura et al., 1995, 1996).Therefore, the oxidative cyclization of grifolic acid byDCA synthase seems to be efficient enough to producea large amount of DCA that accumulates in young

leaves of R. dauricum (Taura et al., 2014). In contrast, therecombinant enzyme did not accept grifolin, thedecarboxylated form of grifolic acid, as a substrate(Table II), suggesting that the carboxyl group is essen-tial for substrate recognition of this enzyme. Likewise,DCA synthase did not catalyze the oxidation of can-nabigerolic acid (Table II), the substrate for cannabinoidsynthases. Cannabigerolic acid is a kind of prenylatedalkylresorcylic acid like grifolic acid, but it has a largeralkyl chain and a shorter prenyl group compared withgrifolic acid.

Next, to evaluate the effects of prenyl side chains, weprepared two prenyl chain analogs of grifolic acid,namely cannabigerorcinic acid (3-geranyl orsellinicacid; Shoyama et al., 1978) and 3-geranylgeranylorsellinic acid, by acid-catalyzed prenylation of orsel-linic acid (Crombie et al., 1988) and used them as sub-strates. Remarkably, DCA synthase could catalyze theoxidocyclization of both synthetic substrates to produceDCA analogs, although the calculated catalytic effi-ciencies were apparently lower than that for grifolicacid (Table II). The product obtained form cannabi-gerorcinic acid was cannabichromeorcinic acid, whichwas reported previously as a synthetic cannabinoid(Shoyama et al., 1984). The product obtained from3-geranylgeranyl orsellinic acid was a novel DCA an-alog harboring a diterpenoid portion; therefore, wenamed it diterpenodaurichromenic acid.

With respect to the kinetic constants for each analog,the kcat value for cannabigerorcinic acid was compara-ble with that of grifolic acid; however, the Km value wasmore than 50-fold higher than that for grifolic acid(Table II). Thus, the farnesyl-to-geranyl conversion ofthe prenyl moiety in the substrate greatly lowered theaffinity between enzyme and substrate. In contrast,when 3-geranylgeranyl orsellinic acid was used as asubstrate, the kcat value was apparently reduced,whereas the Km value was of the same order as thatfor grifolic acid (Table II), suggesting that, although3-geranylgeranyl orsellinic acid is an acceptable sub-strate for DCA synthase, it is rather hard to be cyclizedin the active site of this enzyme. These results demon-strated that DCA synthase was relatively tolerant of theprenyl chain length of the substrate but mostly pre-ferred grifolic acid, presumably because the farnesylgroup is best suited for the active site. In contrast, DCAsynthase strictly recognized the alkyl chain length be-cause cannabigerolic acid with the pentyl group wasnot a suitable substrate for this enzyme.

Table I. Oxygen requirement for the DCA synthase reaction

Substrate and enzyme solutions were preincubated individually with the indicated additives at 30°C for1 h prior to initiation of the reactions. Data are means6 SD of triplicate measurements. Relative activity of100% was 57.4 nkat mg21 protein. U, Units.

Treatment Relative Activity (%)

None 100.0 6 4.540 mM Glc + 5 U of Glc oxidase + 10 U of catalase Not detected40 mM Glc + boiled Glc oxidase + 10 U of catalase 97.5 6 4.2

Figure 5. Spectral measurements of the purified recombinant DCAsynthase. A, Absorption spectrum of the enzyme (0.23 mg mL21) dis-solved in 10 mM sodium phosphate buffer (pH 7). B, Fluorescenceemission spectrum of the same sample as in A. The sample was irradi-ated at 310 nm. The fluorescence emission at 624 nmwas quenched bythe addition of sodium dithionite.


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Homology Modeling of DCA Synthase

The homology model for DCA synthase was pre-pared using the crystal structure of THCA synthase asthe template. As presented in Figure 6A, DCA synthaseadopts almost the same overall topology as THCAsynthase (Shoyama et al., 2012). In addition, the DCAsynthase model places conserved His and Cys residues

near the 8a and 6 positions, respectively, of the FADisoalloxazine ring (Fig. 6B). This result strongly sug-gested that DCA synthase also features a bicovalentlinkage with FAD via His-112 and Cys-175, as in thecase of THCA synthase (Shoyama et al., 2012). Al-though the overall structure as well as the covalentlinkage with the cofactor appear to be conserved, thereare significant differences between the predicted active

Figure 6. Homology model of DCA synthaseconstructed with the crystal structure of C. sativaTHCA synthase (PDB ID: 3vte) as a template. A,Ribbon model for the overall structure of DCAsynthase (green) superimposed on that of THCAsynthase (red). B, Closeup views of the enzymeactive sites. DCA synthase (green) was super-imposed on the corresponding position of THCAsynthase (red). The FAD molecule bicovalentlylinked to THCA synthase is depicted as a yellowstick model. The amino acid residues lining theactive sites are indicated as one-letter codes ingreen (DCA synthase) or red (THCA synthase).

Table II. Steady-state kinetic parameters of the recombinant DCA synthase

Data are means 6 SD of triplicate measurements.

Km kcat kcat/Km

Substrate Structures Product Structures mM s21 s21 mM21

1.19 6 0.20 6.53 6 0.12 5.61 6 0.82

Grifolic acid DCA

Product not detected - - -

Grifolin

Product not detected - - -

Cannabigerolic acid

66.1 6 12.7 2.14 6 0.39 0.0326 0.001

Cannabigerorcinic acid Cannabichromeorcinic acida

7.46 6 1.85 0.13160.029 0.01860.002

3-Geranylgeranyl orsellinic acid Diterpenodaurichromenic acida

aThe absolute configuration of these products was not determined.





site residues of DCA synthase and THCA synthase (Fig.6B). For example, Tyr-484 (top center), the general base inthe THCA synthase reaction (Shoyama et al., 2012), isreplaced by Cys-477 at the corresponding position ofDCA synthase. Likewise, several amino acid residuessurrounding FAD in THCA synthase also are substitutedsimultaneously in DCA synthase. These amino acidchanges would be responsible for the differences in sub-strate coordination and/or catalytic processes betweenDCA synthase and THCA synthase. We perceive thatx-ray crystallographic analysis of the enzyme-ligandcomplex would be required for detailed understandingof the unique catalytic features of DCA synthase.

Phytochemical Analysis of Meroterpenoid Constituentsin R. dauricum

DCA synthase could accept prenyl chain analogs ofthe physiological substrate grifolic acid. Based on this

in vitro result, we suspected that the prenyl chain analogsof DCA might actually be components in R. dauricumplants. To confirm this presumption, the methanol ex-tract of young leaves was examined by HPLC and LC-PDA-ESI-MS analyses. As shown in Figure 7, the elutionprofile of the extract clearly showed peakswith retentiontimes identical to those for cannabichromeorcinic acidand diterpenodaurichromenic acid, respectively, eventhough the peak intensities were apparently lower ascompared with that of DCA. In addition, their MS andUV spectra, summarized in Table III, matched those ofauthentic samples. Thus, the presence of these DCAanalogs was clearly identified in R. dauricum. Regardingtheir precursors, cannabigerorcinic acid as well as3-geranylgeranyl orsellinic also were detected as minorconstituents along with grifolic acid in the same leafextract (Fig. 7; Table III). Therefore, R. dauricum containsmeroterpenoids with C10, C15, and C20 isoprenoid moi-eties. The typical contents of the respectivemeroterpenoids

Figure 7. HPLCanalyses of theR. dauricumyoung leaf extract to detect meroterpenoidconstituents. A, Standard samples (top)and leaf extract (bottom) eluted with 90%acetonitrile. The instrument conditions aredescribed in “Materials and Methods.” B,The same samples eluted using 60% ace-tonitrile so that the peak (a) in the extractcould be well resolved. The peaks are asfollows: (a), cannabigerorcinic acid; (b)grifolic acid; (c) cannabichromeorcinicacid; (d) 3-geranylgeranyl orsellinic acid;(e) DCA; (f) diterpenodaurichromenicacid. mAU, Milliabsorbance unit.


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in the young leaves, quantified by HPLC, are presented inSupplemental Figure S5. Presumably, DCA synthase cat-alyzes the biosynthesis of not only DCA but also prenylchain analogs of DCA based on its in vitro substratespecificity, as described above.

Secretion Property of DCA Synthase in Planta

To estimate the subcellular localization of DCA syn-thase in plants, we stably transformed cultured tobacco(Nicotiana tabacum) ‘Bright Yellow 2’ (BY-2) cells to ex-press a fusion protein comprising the full-length DCAsynthase and GFP and analyzed its distribution byconfocal laser microscopy, western blotting, and en-zyme assay. As shown in Figure 8A, the control cellsexpressing only GFP showed clear fluorescence in thecytoplasm and nucleus, whereas the fluorescence of theDCA synthase fusion protein was apparently weak butdetectable in the endomembrane system of the trans-genic BY-2 cells. However, this clearly suggested thatmost of the fusion protein was not retained in theendomembrane but secreted into the medium, as thewestern blotting using anti-GFP antibody detected an;89-kD band, which is the predicted size for the fusionprotein in the sample prepared from culture superna-tant (Fig. 8B, lane 1). However, the cell extract affordedno detectable signal in the western analysis, probablybecause the fusion protein in the endomembrane sys-tem was under the detection limit (Fig. 8B, lane 2).Furthermore, DCA synthase activity also was observedalmost exclusively in the culture supernatant, as sum-marized in Table IV: the specific activity in the culturesupernatant was 1,678 pkat mg21 protein, whereasapparently weak enzyme activity (0.862 pkat mg21

protein) was obtained for the cell extract, indicating thatBY-2-derived DCA synthase was catalytically activeand secreted efficiently even when it was fused to GFP.

Consequently, DCA synthase was confirmed to behaveas a secretory protein not only in P. pastoris cells but alsoin BY-2 cells, implying an extracellular localization ofDCA synthase in R. dauricum plants.

DCA Is Biosynthesized and Accumulated in the GlandularScales of R. dauricum

R. dauricum is a typical lepidote Rhododendron speciesbearing numerous glandular scales on both the abaxialand adaxial surfaces of leaves (Desch, 1983). The scale isa multicellular structure of epidermal origin consistingof a stalk, attached to a leaf epidermis, and a circular-shaped expanded cap (Desch, 1983). Figure 9A showsthe top view of the abaxial surface of a young leaf of R.dauricum covered with glandular scales. The cap of thescale is composed of relatively small central cells con-nected to stalk cells and surrounding inner and outerrim cells. During themicroscopic observation of the leafsurface, we found that the scales showed light bluishautofluorescence under UV irradiation (Fig. 9A), sug-gesting that a fluorescent metabolite is accumulatedspecifically in the scales. Most of the scales showed aregularly compartmented fluorescence pattern, butsome of them exhibited intense and diffused signalsimplying fluorescence leakage into the cytosol, proba-bly due to physical damage in the scale structure.

In order to analyze the subscale localization ofautofluorescence more precisely, we detached thescales using an adhesive tape and observed the patternusing confocal fluorescence microscopy. It was quite aninteresting observation that the autofluorescence wasdetected only in the periphery of scale cells showing amesh-like pattern (Fig. 9B), suggesting that the fluo-rescent constituent is most likely localized in the apo-plast of the scale. The observed scale was partiallydisrupted to its right (Fig. 9B), where cytoplasmic

Table III. LC-PDA-ESI-MS analyses of meroterpenoid constituents in young leaves of R. dauricum

Peak

Identifier in

Figure 7 Analyte

Retention

Time/Mobile Phasea HR-ESI-MS MS/MSb lmax

min m/z nm(a) Cannabigerorcinic acid 8.5/A (55.7/B) 303.15985 [M-H]2 (calculated

for C18H23O42, 303.15964)

259.2 [M-H-CO2]2 223, 267, 303

285.2 [M-H-H2O]2

(b) Grifolic acid 13.0/A 371.22217 [M-H]2 (calculatedfor C23H31O4

2, 371.22224)327.4 [M-H-CO2]

2 225, 267, 305353.2 [M-H-H2O]2

(c) Cannabichromeorcinic acid 14.3/A 301.14413 [M-H]2 (calculatedfor C18H21O4

2, 301.14399)257.2 [M-H-CO2]

2 256283.2 [M-H-H2O]2

(d) 3-Geranylgeranylorsellinic acid

25.9/A 439.28470 [M-H]2 (calculatedfor C28H39O4

2, 439.28484)395.4 [M-H-CO2]

2 224, 261, 302421.4 [M-H-H2O]2

(e) DCA 28.9/A 369.20673 [M-H]2 (calculatedfor C23H29O4

2, 369.20659)325.3 [M-H-CO2]

2 256351.2 [M-H-H2O]2

(f) Diterpenodaurichromenicacid

70.4/A 437.26834 [M-H]2 (calculatedfor C28H37O4

2, 437.26919)393.4 [M-H-CO2]

2 256419.3 [M-H-H2O]2

aLC separations were performed isocratically using 90% acetonitrile (A) or 60% acetonitrile (B), both containing 0.1% formic acid, at a flow rate of0.25 mL min21. The instrument conditions are described in “Materials and Methods.” bPrecursor ions are the [M-H]2 ions in high-resolution(HR)-ESI-MS.






shrinkage alsowas observed for some cells, as indicatedwith the arrows (Fig. 9B, enlarged images in bottomrow). The fluorescence at the corresponding positionpartly leaked into the cytosol; however, it largelyremained in the mesh-like pattern even after cytoplas-mic shrinkage (Fig. 9B). Therefore, we assumed that thefluorescent material in the apoplast of the scale mightbe partly associated with the cell wall components.

To identify the fluorescent compound, the methanolextract of scales was analyzed by thin-layer chroma-tography (TLC). The major fluorescent component inglandular scales was identified to be DCA, as the spotcorresponding to DCA gave clear autofluorescence onthe TLC plate under irradiation at 365 nm (Fig. 9C). Incontrast, grifolic acid did not show fluorescence (Fig.9C), indicating that the chromenic acid substructure inDCA contributes to the fluorescent property. In addition,HPLC analysis of the scale extract and scale-removed leaf

extract clearly showed that DCA and grifolic acid areaccumulated mainly in scales rather than in the leaf body(Fig. 9D). Likewise, protein extract prepared from glan-dular scales exhibited considerably higher DCA synthaseactivity than that of leaf body extract (Fig. 9E), stronglysuggesting that DCA is biosynthesized and accumulatedin glandular scales.

We also studied the tissue-specific expression of theDCA synthase gene by means of semiquantitativeRT-PCR using gene-specific primers. Agarose gelanalysis demonstrated that the DCA synthase gene isclearly expressed at a higher level in young leaves andin lesser amounts in mature leaves and twigs (Fig. 10).This tissue distribution pattern agreed well with thepreviously reported DCA content in each tissue (Tauraet al., 2014). In addition, a much larger amount of PCRproduct was detected for the sample from scales incomparison with the sample from scale-removedyoung leaves (Fig. 10), demonstrating that the DCAsynthase gene is expressed predominantly in the glan-dular scales. Based on all results of the assays at thechemical, enzymatic, and genetic levels, we concludedthat the glandular scales are the site of DCA biosyn-thesis in R. dauricum plants. The extracellular localiza-tion of DCA autofluorescence seemed to be reasonablebecause DCA synthase is a secretory biosynthetic en-zyme (Fig. 8).

DCA and Grifolic Acid Are Cell Death-InducingPhytotoxic Metabolites

As described above, DCA is accumulated in the ex-tracellular compartment, the apoplast of glandularscales. With respect to the reason for this specific lo-calization, we assumed that DCA might be a toxicmetabolite and, thus, has to be excluded from plantcells. To confirm this possibility, we investigated thephytotoxic properties of DCA and its precursors,grifolic acid and orsellinic acid, using tobacco BY-2cells as our experimental system. First, the cell via-bility of BY-2 cells after the treatment with eachcompound was measured by Trypan Blue staining.As shown in Figure 11A, 50 mMDCA and grifolic acidinduced almost 100% cell death in BY-2 culturewithin 24 h of treatment, whereas orsellinic acid, thephenolic precursor, had no significant effects on theviability of BY-2 cells, suggesting that the prenyla-tion of orsellinic acid is crucial for the phytotoxiceffects. In addition, it was of interest that DNA laddering

Figure 8. Analyses of the DCA synthase-GFP fusion protein expressedin suspension cultured cells of tobacco BY-2. A, Full-length DCA syn-thase was fused at its C-terminus to GFP and stably expressed in BY-2cells. Fluorescence was observed with confocal microscopy. The toprow represents cells expressing freeGFP, whereas the bottom row showsfluorescence based on the fusion protein. Bars = 10mm. B,Western-blotanalysis with an anti-GFP antibody of the protein preparations fromtransgenic BY-2 expressing the fusion protein. Lane M, Prestainedmarkers with the indicated molecular masses; lane 1, 20 mL of theculture medium; lane 2, 20 mL of the cellular protein extract.

Table IV. Distribution of DCA synthase activity between culture supernatant and cell extract preparedfrom the transgenic tobacco BY-2 cell culture expressing a DCA synthase-GFP fusion protein

Data are means 6 SD of triplicate measurements.

Enzyme Preparation Total Activity Total Protein Specific Activity

pkat in mL of culture mg in mL of culture pkat mg21 proteinCulture supernatant 161.6 6 7.5 0.096 6 0.013 1,678 6 35.8Cell extract 0.556 6 0.051 0.643 6 0.034 0.862 6 0.041


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as well as smearingwere detectable for genomic DNAsamples from DCA- and grifolic acid-treated cells,wherein the laddering was more obviously detected forgrifolic acid-treated cells (Fig. 11B). Therefore, DCA- andgrifolic acid-induced cell death may be associated withthe apoptotic pathway, at least in part. Although a de-tailed study should be conducted to clarify the efficacyand precise mechanism of their cytotoxicity, this reportprovides obvious evidence that DCA and grifolic acidare cell death-inducing phytotoxic meroterpenoids, simi-lar to the cannabinoids such as THCA and cannabigerolicacid sequestered in the trichome head of C. sativa(Sirikantaramas et al., 2005).

DISCUSSION

Molecular and Biochemical Characterization ofDCA Synthase

Meroterpenoids, produced by plants, bacteria, andfungi, are natural products that partially originate fromthe isoprenoid pathway (Matsuda andAbe, 2016). Plant

meroterpenoids include many prenylated aromatics,such as flavonoids, isoflavonoids, stilbenoids, couma-rin, quinone, phloroglucinol, and resorcinol harboringisoprenoid portions, and are a remarkable class ofnatural products because of their unique pharmaco-logical and biological activities (Singh and Bharate,2006; Rivière et al., 2012; Veitch, 2013; Chen et al., 2014;Mechoulam et al., 2014; Dugrand-Judek et al., 2015;Widhalm and Rhodes, 2016). Thus, biosynthetic studiesof plant meroterpenoids have been conducted exten-sively in recent years, especially on aromatic prenyl-transferases (Yazaki et al., 2009) and oxidocyclasesbelonging to FAD oxidases (Sirikantaramas et al., 2004;Taura et al., 2007) and P450 enzymes (Welle andGrisebach,1988; Larbat et al., 2007). Physiologically, many plantmeroterpenoids serve as chemical defense compo-nents in plants. For example, sophoraflavanone G, alavandulyl flavanone from the genus Sophora, showsa potent antibacterial activity (Tsuchiya and Iinuma,2000), whereas furanocoumarins, derived from preny-lated phenylpropanoids, are defensive against variouskinds of herbivores (Zangerl and Berenbaum, 1990). It is

Figure 9. Analyses of glandular scales on the youngleaves of R. dauricum. A, Closeup view of the ab-axial side of a young leaf with numerous glandularscales under bright-field (left) and UV (right) irradi-ation. Bar = 100 mm. B, Confocal images of a de-tached scale under UV and bright-field irradiation.The bottom row shows enlarged images of the boxedpart of the images in the top row. Arrows indicatecytoplasmic shrinkage in a rim cell. Bars = 20mm.C,TLC analysis of scale extract and standard mer-oterpenoids visualized under irradiation at 254 nm(left) and 365 nm (right). Lane 1, DCA standard; lane2, glandular scale extract; lane 3, grifolic acid stan-dard. D, Distribution of meroterpenoid constituentsbetween scales and scale-removed leaf tissue ana-lyzed by HPLC. Black bars, DCA; white bar, grifolicacid. E, DCA synthase activity in the crude proteinextract prepared from glandular scales (left) andscale-removed leaf tissue (right).





also of interest that these hydrophobic meroterpenoids,in addition to cannabinoids, are secreted into apoplasticcompartments after their biosynthesis (Yamamoto et al.,1996; Sirikantaramas et al., 2005; Voo et al., 2012), as isthe case with various hydrophobic terpenoids accumu-lated in the secretory cavity of glandular trichomes (Lange,2015). In contrast to these well-documented examples,the biosynthetic mechanism, tissue localization, andphysiological function of DCA had remained unclearlargely because the gene responsible for DCA biosynthe-sis was not characterized.

In this study, we cloned a cDNA coding for DCAsynthase from young leaves of R. dauricum. Phyloge-netic analysis and amino acid alignment demonstratedthat the primary structure of DCA synthase has ap-preciable homologies to plant FAD oxidases and con-tains conserved residues essential for novel bicovalentflavinylation (Leferink et al., 2008). VAO family FADoxidases arewidely distributed across plant species andare divergently evolved to catalyze oxidative reactionsin various specialized metabolic pathways (Leferinket al., 2008). Among them, DCA synthase is the thirdexample of meroterpenoid oxidocyclases, the first twobeing THCA synthase and CBDA synthase. However,DCA synthase has a novel characteristic as the enzymethat specifically cyclizes the farnesyl group in a naturalmeroterpenoid product of plant origin. It is noteworthythat flavoprotein oxidases with similar structure andfunction have evolved in genetically distinct plantspecies, C. sativa and R. dauricum, to produce respec-tive pharmacologically valuable constituents. Hitherto,

CBCA synthase in C. sativa and glyceollin synthase inGlycine max also have been identified to catalyze prenylto chromene cyclization like DCA synthase (Welle andGrisebach, 1988; Morimoto et al., 1997); however, theseenzymes have not been characterized at the molecularlevel.

Although the reactions catalyzed by VAO flavopro-teins are variable, they all share common structuralfeatures and FAD-dependent reaction mechanisms(Leferink et al., 2008). Similarly, DCA synthase alsocontains covalently linked flavin as a coenzyme. Inaddition, this enzyme has an absolute requirement formolecular oxygen and releases hydrogen peroxideequal to the produced DCA, as reported for FAD oxi-dases. Based on these observations, we propose a re-action mechanism for DCA synthase in a flavin- andoxygen-dependent manner, as illustrated in Figure 12.With thismechanism, first, the flavinmolecule accepts ahydride from the benzylic position of grifolic acid; si-multaneously, a basic residue of the enzyme abstracts aproton from a phenolic hydroxyl group. These stepsproduce a reduced flavin and an ionic intermediate inthe active site. The subsequent step is the stereospecificelectrocyclization to form DCA and the hydride iontransfer from the reduced flavin to molecular oxygen,resulting in hydrogen peroxide formation and reox-idation of the flavin for the next reaction cycle.

In the case of THCA synthase, for which a crystalstructure was elucidated, Tyr-484 contributes as a cata-lytic base to accept proton from the substrate (Shoyamaet al., 2012). This residue is not conserved in DCAsynthase, as depicted in the molecular model structure(Fig. 6B). It is notable, however, that the side chain ofTyr-114 in DCA synthase alternatively protrudes to

Figure 10. Analysis of the tissue distribution of DCA synthase tran-scripts in R. dauricum by semiquantitative RT-PCR. A, Agarose gelelectrophoresis of a DCA synthase gene fragment (557 bp) amplifiedusing gene-specific primers. The 18S rRNA gene fragment (663 bp) wasamplified as a housekeeping control gene. Numbers indicate youngleaves (1), mature leaves (2), twigs (3), flowers (4), roots (5), glandularscales (6), and scale-removed leaves (7). B, The band densities of DCAsynthase fragments were normalized with that of 18S rRNA and arepresented as means 6 SD of triplicate assays, in which the expressionlevel in young leaves was set to 1. Numbers indicate the same tissues asshown in A.

Figure 11. Phytotoxic effects of DCA and grifolic acid on suspensioncultured tobacco BY-2 cells. A, Trypan Blue staining of DCA-, grifolicacid-, or orsellinic acid-treated cells. Five-day-old cell cultures wereincubated with 50 mM of each indicated compound for 24 h and thenstained with Trypan Blue. Control cells were treated with the solvent(DMSO) only. Bars = 100 mm. B, Agarose gel electrophoresis of ge-nomicDNA fromBY-2 cells treated as in A. DNAwas separated on a 2%agarose gel and stained with ethidium bromide. Lane M, Marker DNAwith the indicated sizes; lane 1, control cells; lane 2, DCA-treated cells;lane 3, grifolic acid-treated cells; lane 4, orsellinic acid-treated cells.


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the putative substrate-binding pocket (si-side of FAD;Fig. 6B), implying that Tyr-114 is a possible candidatefor the catalytic base in the DCA synthase reaction.Actually, the catalytic base residues in plant FAD ox-idases are not necessarily conserved at the specificposition (Supplemental Fig. S1), as discussed previ-ously by Daniel et al. (2015). Nevertheless, we regardthat crystal structure determination and site-directedmutational studies of DCA synthase are essential for aprecise understanding of the structure-function rela-tionship of the reaction. These ongoing projects woulddemonstrate the structural basis underlying the differ-ences in substrate recognition and cyclization processesbetween DCA synthase and cannabinoid synthases andmight provide valuable insights into the rational designof enzyme active sites to synthesize novel unnaturalmeroterpenoids.Kinetic studies using various substrates clearly

demonstrated that DCA synthase is highly specific togrifolic acid. The catalytic efficiency (kcat/Km) for DCAformation was 5.61, which was much higher than thoseof THCA synthase and CBDA synthase (Taura et al.,1995, 1996). This result may be reasonable because theoxidocyclization process catalyzed by DCA synthase israther simple: it proceeds without the rearrangement ofthe geometrical configuration of the prenyl group pro-posed for the reaction schemes of THCA synthase andCBDA synthase (Sirikantaramas et al., 2004; Tauraet al., 2007). Unlike grifolic acid, DCA synthase did notaccept grifolin and cannabigerolic acid as a substrate.These results are relevant to the fact that R. dauricumdoes not produce neutral meroterpenoids without car-boxyl group and cannabinoid-type metabolites with apentyl side chain (Taura et al., 2014).Of particular interest, DCA synthase showed enzyme

activity on substrate analogs containing differentprenyl groups to produce the corresponding chromenicacid derivatives, namely cannabichromeorcinic acidand diterpenodaurichromenic acid. Consequently,DCA synthase is a novel oxidocyclase that could cyclizemultiple prenyl moieties in plant meroterpenoid

biosynthesis. In addition, phytochemical analysis usingLC-PDA-ESI-MS identified cannabichromeorcinic acidand diterpenodaurichromenic acid, probably producedby DCA synthase, as minor constituents in leaf extractof R. dauricum, although the absolute configuration ofthese DCA analogs has yet to be characterized. Theseresults indicated the possibility that an unidentifiedaromatic prenyltransferase involved in the DCA path-way might promiscuously accept prenyl donor sub-strates to produce orsellinic acid derivatives withgeranyl, farnesyl, and geranylgeranyl portions as thesubstrates for DCA synthase. The corresponding pre-nyltransferase is exceptionally remarkable becausemost of the aromatic prenyltransferases involved inplant specialized metabolism are strictly selective toeither dimethylallyl pyrophosphate or geranyl pyro-phosphate (Munakata et al., 2014).

Localization and Possible Physiological Function of DCAin R. dauricum

Rhododendron species are classified into two types,lepidote or elepidote Rhododendron, based on whetherthey have glandular scales or not (Desch, 1983).Morphologically, the stalked architecture of glandu-lar scales is somewhat similar to the capitate stalkedtrichomes on plants belonging to Lamiaceae andCannabaceae (Lange, 2015); however, glandular scalesdevelop the expanded cap comprising the central andrim cells and do not have an obvious secretory cavity,although they contain hydrophobic sesquiterpenoids(Doss, 1984; Doss et al., 1986). In this study, we alsodemonstrated that DCA is produced and accumulatedspecifically in the glandular scales of R. dauricum basedon several lines of evidence: DCA synthase gene ex-pression, enzyme activity, as well as DCA content beingobserved predominantly in glandular scales rather thanscale-removed leaf tissues. In addition, the extracellulardistribution of DCA-based autofluorescence and thesecreted nature of the recombinant DCA synthaseexpressed in a plant cell culture suggested the possi-bility that the DCA synthase reaction takes place in theapoplast of the glandular scales, like the THCA syn-thase reaction that occurs in the secretory cavity ofglandular trichomes (Sirikantaramas et al., 2005).

Such an extracellular biosynthesis and accumulationof DCA presumably provides several advantages for R.dauricum plants. (1) DCA and grifolic acid could effec-tively serve as chemical barriers in the apoplast ofglandular scales, the outermost layer of the plant, be-cause these meroterpenoids exhibit antimicrobial ac-tivities (Hashimoto et al., 2005). (2) Hydrogen peroxide,the by-product of DCA synthase, also may take part inchemical defense because of its antimicrobial propertyas described in relation to FAD-dependent carbohy-drate oxidases (Custers et al., 2004). (3) As demon-strated in this study, grifolic acid and DCA arepotentially phytotoxic metabolites. Thus, they shouldbe stored in the apoplast to avoid cellular damage in

Figure 12. Proposed reaction mechanism of DCA synthase. Two elec-trons from the substrate are accepted by enzyme-bound flavin and thentransferred to molecular oxygen to reoxidize flavin. DCA is synthesizedfrom the ionic intermediate via stereoselective cyclization by the en-zyme. B indicates the proposed basic residue of the enzyme.






R. dauricum plants. Therefore, grifolic acid has to beeffectively secreted once it is biosynthesized by an ar-omatic prenyltransferase. Many hydrophobic special-ized metabolites are secreted, probably to serve in plantdefense and/or to avoid self-poisoning, although only afew studies have been reported with respect to themechanisms for transportation of these components(Shitan, 2016). For example, a diterpenoid sclareol inNicotiana plumbaginifolia and monolignols in Arabidopsis(Arabidopsis thaliana) are transported to the apoplast byATP-binding cassette transporters (Stukkens et al., 2005;Alejandro et al., 2012). In addition, a recent study usinghairy root culture of Lithospermum erythrorhizon hasdemonstrated that shikonin, a kind of naphthoquinonemeroterpenoid, is secreted utilizing pathways common tothe ADP ribosylation factor/guanine nucleotide ex-change factor system and actin filament polymerization(Tatsumi et al., 2016).We assume that a specificmolecularmechanism, which might share similarity with either ofthem, would transport grifolic acid into the apoplast ofglandular scales to circumvent self-poisoning and for thefollowing DCA biosynthesis.

In conclusion, we conducted molecular and bio-chemical characterization of DCA synthase, a noveloxidocyclase specific to grifolic acid, to produce DCAwith anti-HIV properties. The DCA synthase gene andthe recombinant enzyme would be applicable for thesemisynthesis and biomimetic heterologous productionof DCA, because grifolic acid can be obtained in a largequantity from the inedible mushroom Albatrellus dis-pansus (Hashimoto et al., 2005), and more recently, a denovo production system of grifolic acid has beenestablished using Aspergillus oryzae transformed with agrifolic acid-producing gene cluster from Stachybotrysbisbyi (Li et al., 2016). In addition, the ecological sig-nificance of DCA biosynthesis in plants also would bean interesting research subject. For example, DCAmight participate in chemical defense against insects, ascannabinoids, structurally related to DCA, have insect-repellent properties (Rothschild and Fairbairn, 1980).

MATERIALS AND METHODS

Plant Materials and Reagents

Rhododendron dauricum plants were cultivated at the Experimental Stationfor Medicinal Plant Research at the University of Toyama. Suspension-culturedtobacco (Nicotiana tabacum) BY-2 cells were obtained from the RIKEN Bio-resource Center and cultured in a modified Murashige and Skoog medium asdescribed previously (Sirikantaramas et al., 2005).

Chemical reagentswere purchased fromWako Pure Chemicals unless statedotherwise. DCA, grifolic acid, grifolin, and cannabigerolic acid were from ourlaboratory collection (Taura et al., 2014). Previously reported meroterpenoids,cannabigerorcinic acid and cannabichromeorcinic acid, were synthesizedas described by Shoyama et al. (1978, 1984). The standard sample of3-geranylgeranyl orsellinic acid was chemically synthesized, using p-toluene-sulfonic acid as a catalyst, from orsellinic acid and geranylgeraniol (Sigma-Aldrich), as described by Crombie et al. (1988). Diterpenodaurichromenicacid in racemic form was synthesized from 3-geranylgeranyl orsellinic acid bychemical oxidation using dichlorodicyanobenzoquinone (Hashimoto et al.,2005). The structures were verified by obtaining their physical data as sum-marized below using a JNM-ECA500 NMR spectrometer (JEOL) and an

LC-PDA-ESI-MS system (Thermo Fisher Scientific). The 1H-NMR spectra of3-geranylgeranyl orsellinic acid and diterpenodaurichromenic acid were al-most identical to those reported for grifolic acid and DCA (Kashiwada et al.,2001; Li et al., 2016), respectively, except for signals corresponding to prenylchain differences (Supplemental Figs. S6 and S7).

3-Geranylgeranyl orsellinic acid: 1H-NMR (500MHz, CDCl3): d 11.85 (s, 1H),6.26 (s, 1H), 5.28 (t, J = 7.0 Hz, 1H), 5.08 to 5.10 (m, 3H), 3.43 (d, J = 7.0 Hz, 2H),2.52 (s, 3H), 2.01 to 2.06 (m, 12H), 1.82 (s, 3H), 1.68 (s, 3H), 1.59 (overlapped, 9H).HRMS (ESI, negative): m/z calculated for C28H39O4 [M-H]2: 439.28483, found439.28439. lmax (PDA): 224, 261, 303 nm. Diterpenodaurichromenic acid:1H-NMR (500MHz, CDCl3): d 11.85 (s, 1H), 6.73 (d, J = 10.0 Hz, 1H), 6.22 (s, 1H),5.48 (d, J = 10.0 Hz, 1H), 5.08 to 5.12 (m, 3H), 2.52 (s, 3H), 2.04 to 2.09 (m, 6H),1.95 to 1.98 (m, 4H), 1.73 to 1.78 (m, 2H), 1.68 (s, 3H), 1.60 (s, 3H), 1.59 (s, 3H),1.57 (s, 3H), 1.40 (s, 3H). HRMS (ESI, negative):m/z calculated for C28H37O4 [M-H]2: 437.26919, found 437.26838. lmax (PDA): 255 nm.

Transcriptome Sequencing and Screening of CandidateGenes for RdFADOX

Total RNA was extracted from young leaves of R. dauricum using theRNAqueous kit according to the manufacturer’s instructions (Thermo FisherScientific). Then, the cDNA librarywas prepared using the TruSeq RNA samplepreparation kit with a low-throughput protocol (Illumina). The cDNA clustersgenerated on a Single-Read flow cell were sequenced on the Illumina GenomeAnalyzer IIx as 100-bp single-end reads to generate a data set consisting of35,093,217 reads. The resulting sequence reads were assembled de novo usingRnnotator version 2.4.12 (Martin et al., 2010) with the default parameters toobtain 71,944 cDNA contigs with an average length of 535 bp. These tran-scriptome data were used as the DNA database for a local tBLASTn homologysearch using the BioEdit program version 7.2.5 (http://www.mbio.ncsu.edu/BioEdit/bioedit.html). Three putative nucleotide sequences encoding R. daur-icum FAD oxidases (RdFADOX1–RdFADOX3) were mined from the databasesusing amino acid sequences of cannabinoid synthases as queries. The FPKMvalue for each gene was calculated according to the reported scheme (Trapnellet al., 2010).

cDNA Cloning of RdFADOX1 to RdFADOX3

The gene-specific oligonucleotide primers and PCR conditions listed inSupplemental Table S3 were used for amplification of the RdFADOX1 toRdFADOX3 gene fragments. To obtain complete cDNA sequences, includinguntranslated regions, 3ʹ- and 5ʹ-RACE reactions were conducted as describedpreviously (Taura et al., 2016). The cDNA template was prepared by RT ofyoung leaf RNA using ReverTra Ace (Toyobo) and dT17AP primer. The 3ʹ-RACE products for RdFADOX1 to RdFADOX3 genes were obtained using theadapter primer AP and specific primers FADOX(1–3)-3R, respectively. Thetemplate for 5ʹ-RACE was prepared by polyadenylation of the cDNA usingterminal deoxynucleotidyl transferase (Takara Bio) in the presence of deoxy-adenine. First-round PCRs were conducted with dT17AP primer and specificprimers FADOX(1–3)-5R1. Then, nested PCR using primers AP and FADOX(1–3)-5R2 amplified the 5ʹ-RACE products for RdFADOX1 to RdFADOX3genes. All RACE products were subcloned into pMD-19 (Takara Bio) andsequenced on a 3130 DNA sequencer (Thermo Fisher Scientific). The full-length RdFADOX1 to RdFADOX3 coding sequences were amplified withprimers FADOX(1–3)-Fw and FADOX(1–3)-Rv, respectively, and subclonedinto the expression vector pPICZA (Thermo Fisher Scientific) predigestedwith EcoRI and SalI using the In-Fusion HD cloning reagent (Clontech). Thepartial Kozak sequence (AAAACA) was inserted prior to the translationalstart ATG codon for optimal yeast expression (Hamilton et al., 1987). Theexpression vectors were designed to express the recombinant proteins inPichia pastoris in the presence of methanol.

Computational Analyses of RdFADOX Sequences

The subcellular location and protein-sorting signal were predicted with theonline programs TargetP (http://www.cbs.dtu.dk/services/TargetP/) andPSORT (http://www.psort.org/). Protein motifs were scanned using PfamScan(http://www.ebi.ac.uk/Tools/pfa/pfamscan/) and ScanProsite (http://prosite.expasy.org/scanprosite/). Multiple alignment of FAD oxidases, in-cluding RdFADOX1 to RdFADOX3, was carried out using the Clustal Omegaprogram (http://www.ebi.ac.uk/Tools/msa/clustalo/). A neighbor-joining


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http://www.mbio.ncsu.edu/BioEdit/bioedit.html

http://www.mbio.ncsu.edu/BioEdit/bioedit.html


http://www.cbs.dtu.dk/services/TargetP/

http://www.psort.org/

http://www.ebi.ac.uk/Tools/pfa/pfamscan/

http://prosite.expasy.org/scanprosite/

http://prosite.expasy.org/scanprosite/

http://www.ebi.ac.uk/Tools/msa/clustalo/


phylogenetic tree was drawn by 1,000 bootstrap tests using MEGA6.06 soft-ware (http://www.megasoftware.net). The homology model of DCA syn-thase (RdFADOX1) was generated with SWISS-MODEL software (https://swissmodel.expasy.org) using the crystal structure of THCA synthase fromCannabis sativa (Protein Data Bank identifier 3vte) as the template. The modelquality was evaluated by a Ramachandran plot, using RAMPAGE (http://mordred.bioc.cam.ac.uk/;rapper/rampage.php), to confirm that most aminoacid residues were grouped in the energetically allowed regions. All proteinfigures were rendered with PyMOL (http://www.pymol.org).

Heterologous Expression of RdFADOX1 to RdFADOX3in P. pastoris

The pPICZA expression vectors, containing one of the full-length RdFADOXgenes, were introduced individually to P. pastoris strain KM71H (Thermo FisherScientific) by electroporation, and the transformants were selected on YPD agarplates containing 1 mg mL21 zeocin (InvivoGen). Protein expression in P. pas-toris was accomplished essentially as described by Weis et al. (2004). The threeminimal media used herein contained 200 mM potassium phosphate buffer (pH6), 1.34% yeast nitrogen base (Difco Laboratories), and 4 3 1025% D-biotin anddiffered with respect to the carbon source of 10 g L21 Glc or 1% or 5%methanolfor BMD, BMM2, or BMM10, respectively. A single colony was inoculated intoErlenmeyer flasks containing 10 mL of BMD and cultivated at 25°C for 60 h.Then, 10 mL of BMM2 was added to the culture to initiate protein expression,and 2 mL of BMM10 was fed every 24 h after the onset of the induction. After96 h of induction, the culture was centrifuged to obtain culture supernatant andcell pellet. The cells were washed twice with water and suspended in buffer A(10 mM potassium phosphate buffer [pH 7] containing 3 mM mercaptoethanol).Then, cellular extract was prepared by vortexing vigorously in the presence ofglass beads. DCA synthase activity in the culture medium and cell extract wasassayed as described below.

Purification and Biochemical Properties of theRecombinant DCA Synthase

The P. pastoris strain harboring the DCA synthase (RdFADOX1) gene wasprecultured in a large volume (400 mL) of BMD medium, and protein expres-sion was induced by adding BMM2 (400 mL) followed by BMM10 (80 mL)every 24 h. The culture was harvested after 96 h of induction and centrifuged at20,000g for 15 min. The supernatant (;1,000 mL) was diluted 3-fold with bufferA and applied to a hydroxylapatite column (Nacalai Tesque; 1 cm 3 10 cm)equilibrated with buffer A. The column was washed with three column vol-umes of the same buffer, and bound proteins were eluted with a 400-mL lineargradient of buffer A to 0.5 M potassium phosphate buffer (pH 7) containing3 mM mercaptoethanol. Fractions containing DCA synthase activity were con-centrated and dialyzed against buffer B (10 mM sodium citrate buffer [pH 5.5]containing 3 mM mercaptoethanol). The dialysate was applied to a 1-cm 3

10-cm column containing TOYOPEARL CM-650M (Tosoh) equilibrated withbuffer B. The columnwaswashedwith the same buffer, and DCA synthase waseluted with a 300-mL gradient of NaCl (0–0.5 M) in buffer B. The most activefractions eluted ;0.2 M NaCl and were collected and used for the followingbiochemical characterization. The typical yield of the purified DCA synthasewas 100 to 120 mg from 1,000 mL of culture supernatant.

The purity and subunit molecular mass of the recombinant DCA synthasewere verified by SDS-PAGE analysis using a 12.5% acrylamide gel. The nativemolecular mass of DCA synthase was determined by gel filtration chromatogra-phy on a 2.5-3 75-cmSephacryl S-200HR column (GEHealthcare) calibratedwithstandard proteins. The N-terminal amino acid sequence of the purified DCAsynthase was determined on a PPSQ-21 protein sequencer (Shimadzu). The car-bohydrate staining of the enzyme was performed using the GlycoproteinWesternDetection Kit (Clontech). The absorption and fluorescent emission spectra of theenzyme, dissolved in buffer A at a concentration of 0.23 mg mL21, were recordedon a NanoDrop 2000c (Thermo Scientific) and a F-4500 (Hitachi Hi-Technologies)spectrophotometer, respectively. AMP was released from the purified DCA syn-thase by hydrolysis at 99°C for 10 min in the presence of 10 mMHCl (Kutchan andDittrich, 1995) and then detected by a reverse-phase HPLC system, as describedbelow, using 1% acetonitrile containing 10 mM ammonium formate as a mobilephase at a flow rate of 1 mL min21. The mass spectrum of the AMP also was an-alyzed by an LC-ESI-MS system as described below.

The molecular oxygen requirement was assayed by DCA synthase reactionusing substrate and enzyme solutions preincubated in the presence of 40 mM

Glc, 5 units of Glc oxidase, and 10 units of catalase at 30°C for 1 h. The hydrogenperoxide produced by a by-product of DCA synthase was quantified using theAmplex Red Hydrogen Peroxide/Peroxidase Assay Kit as described by themanufacturer (Thermo Fisher Scientific).

Standard Assay Conditions for DCA Synthase

The standard reaction mixture consisted of 10 mM potassium phosphatebuffer (pH 6), 20 mM grifolic acid (or substrate analog), and the protein solutionin a total volume of 100 mL. The reaction was incubated at 30°C for 30 min andterminated by adding 100mL ofmethanol. The 10-mL aliquotswere analyzed byHPLC and LC-PDA-ESI-MS to quantitate and characterize the reaction product,respectively. The stereoselectivity of the DCA-producing reaction was exam-ined by a chiral HPLC device using a CHIRALPAK AD-H column (Daicel) asdescribed previously (Taura et al., 2014).

HPLC and LC-PDA-ESI-MS Analyses of theReaction Products

The enzyme reaction products were routinely analyzed and quantified by anHPLC system (Tosho) equipped with a Cosmosil 5C18-MS-II column (4.6 mm3150 mm; Nacalai Tesque), as described previously (Taura et al., 2014). Elutionwas performed isocratically with aqueous acetonitrile containing 0.1% formicacid at a flow rate of 1 mLmin21. The acetonitrile concentration was set for eachproduct: 85% for DCA, 75% for cannabichromeorcinic acid, and 95% forditerpenodaurichromenic acid. The products were detected by absorption at254 nm and quantified using calibration curves of the standard compounds.The samples also were analyzed by an LC-PDA-ESI-MS system (Thermo FisherScientific), composed of an Accela 600 HPLC pump, an Accela PDA detector,and an LTQ-Orbitrap-XL ETDHybrid Ion Trap-Orbitrap mass spectrometer, tocharacterize the products in detail. The column andmobile phasewere the sameas those used for the HPLC analysis. The high-resolution MS (negative), MS/MS, and UV spectra were collected as described previously (Taura et al., 2016).

Enzyme Kinetics

The enzyme reactions were conducted in a similar manner to the standardassay conditions, using six concentrations (5, 10, 15, 20, 25, and 30 mM) ofsubstrates, in the presence of 0.37 mg of the purifiedDCA synthase. The reactionproducts were quantified by HPLC, and the kinetic constants were calculatedby fitting the velocity data at each substrate concentration to Hanes-Woolfplots.

Phytochemical Analysis of Young Leaf Extract

Fresh young leaves (100mg)were frozen in liquid nitrogen andground into afine powder by pestle andmortar. The powder was then extracted with 1 mL ofmethanol, and the resulting sample was subjected to HPLC as described abovefor quantitative analyses. The chromatographic separations were performedwith 90%acetonitrile containing 0.1% formic acid at aflow rate of 0.25mLmin21.Alternatively, 60% acetonitrile containing 0.1% formic acid was used as a mo-bile phase for the better separation of cannabigerorcinic acid from other con-stituents. The LC-PDA-ESI-MS analysis alsowas performed as described above,to identify the respective metabolites.

GFP Fusion Gene Construction and Expression in TobaccoBY-2 Cells

TheGFPgeneused in this study is the synthetic variant S65T cloned inpUC18(Niwa et al., 1999). First, the GFP coding sequence was amplified with primersGFP-Fw and GFP-Rv and then introduced into the binary vector pBI121(Clontech) between BamHI and SacI restriction sites to create pBI121-GFP. Full-length DCA synthase cDNA, amplified with DCAS-Fw and DCAS-Rv, wassubcloned into pBI121-GFP by use of XbaI and BamHI restriction sites to con-struct pBI121-DCAS-GFP. The partial Kozak sequence (AACA; Lütcke et al.,1987) was inserted prior to the start codon of DCA synthase via the primerDCAS-Fw. pBI121-GFP and pBI121-DCAS-GFP were introduced into Agro-bacterium tumefaciens LBA4404 by electroporation. BY-2 cells were transformedby A. tumefaciens infection as described (An, 1987) and selected on solidifiedMurashige and Skoog medium containing kanamycin (100 mg mL21). The




http://www.megasoftware.net

https://swissmodel.expasy.org

https://swissmodel.expasy.org

http://mordred.bioc.cam.ac.uk/~rapper/rampage.php



http://www.pymol.org


suspension cultures of the transgenic cells were established in liquidMurashigeand Skoog medium containing kanamycin and subcultured every 7 d. The 5-d-old culture (1 mL) was centrifuged to obtain culture supernatant and cellpellet. Cells were suspended in 1 mL of buffer A, and the extract wasprepared using a microtube BioMasher homogenizer (Nippi), for DCAsynthase assay and western blotting, along with the culture supernatant.Western blotting was performed using an anti-GFP antibody (MBL) asdescribed previously (Kurosaki and Taura, 2015). The GFP fluorescence inthe transgenic cells was observed by confocal microscopy as describedbelow.

Collection and Analyses of the Glandular Scales fromYoung Leaves of R. dauricum

The glandular scales from both sides of young leaves were detached using asemitransparent adhesive tape (Scotch; 3M). The scales were suspended in PBSby gentle sonication, collected by centrifugation at 3,000g for 1 min, and thenused for the following analyses. The methanol extract was prepared using aBioMasher, and the constituents were analyzed by TLC using a silica gel F254

plate (Merck) developed with chloroform:methanol (2:1). The DCA andgrifolic acid contents in the extract also were analyzed by HPLC as describedabove. The crude protein extract of the collected scales was prepared withbuffer A, using a BioMasher, and used for DCA synthase assay. The auto-fluorescence image of an isolated glandular scale was taken as describedbelow.

Microscopy Analyses

Twodifferentfluorescencemicroscopeswereused in this study for respectivepurposes. A BX-50 microscope equipped with a BX-FLA fluorescent unit(Olympus) was used to observe epifluorescence images of the R. dauricum leafsurface with a band-pass excitation filter at 330 to 385 nm and a long-passfilter at 420 nm. Confocal images of isolated glandular scales were takenwith a laser-scanning fluorescence microscope (LSM 700; Carl Zeiss), usinga filter set of 405 nm excitation and 420 to 480 nm emission, whereas GFPsignals expressed in BY-2 cells were excited at 488 nm and collected at505 to 530 nm.

Expression Analysis of the DCA Synthase Gene bySemiquantitative RT-PCR

TotalRNAwas isolatedfromyoungleaves,mature leaves, twigs,flowers, roots,glandular scales, and scale-deficient young leaves of R. dauricum as describedabove. The first-strand cDNA was then synthesized from 0.5 mg of each RNAsample as described above, except that a random hexamer (Toyobo) was used asthe primer. The DCA synthase gene fragment (557 bp) was amplified with theprimers DCAS-RT-Fw and DCAS-RT-Rv, whereas the 18S rRNA gene fragment(GenBank accession no. AB973224.1; 663 bp) was amplified with 18S-Fw and18S-Rv as a housekeeping gene control. The PCR products were resolved on a 2%agarose gel and stainedwith ethidiumbromide. Each banddensity from triplicateexperiments was analyzed by ImageJ version 1.49 (http://imagej.nih.gov/ij/).

Analysis of the Phytotoxic Effects of DCA andGrifolic Acid

Five-day-old BY-2 cells were incubated with 50 mM DCA, grifolic acid, ororsellinic acid for 24 h. The viability of the cells was tested by adding TrypanBlue solution (Sigma-Aldrich). Genomic DNA was isolated from treated BY-2cells by the CTABmethod (Murray and Thompson, 1980). Then, electrophoresiswas performed on a 2% agarose gel.

Accession Numbers

Sequence data from this article can be found in the GenBank/EMBL/DDBJdatabases. The nucleotide sequences of RdFADOX1 to RdFADOX3,whichwererenamed DCA synthase, DCA synthase-like1, and DCA synthase-like2, wereregistered with accession numbers LC184180, LC184181, and LC184182, re-spectively. The transcriptome data set of R. dauricum young leaves was de-posited to the DDBJ Sequence Read Archive with BioProject accession numberPRJDB5618.

Supplemental Data

The following supplemental materials are available.

Supplemental Figure S1. Amino acid sequence analyses of RdFADOX1 toRdFADOX3.

Supplemental Figure S2. Stereoselectivity of the recombinant DCA syn-thase reaction evaluated by a chiral HPLC analysis.

Supplemental Figure S3. SDS-PAGE and carbohydrate staining of the pu-rified recombinant DCA synthase.

Supplemental Figure S4. Analyses of AMP released from DCA synthaseby acid hydrolysis.

Supplemental Figure S5. Quantitative analysis of meroterpenoid constit-uents in the young leaves of R. dauricum.

Supplemental Figure S6. 1H-NMR analysis of 3-geranylgeranyl orsellinicacid.

Supplemental Figure S7. 1H-NMR analysis of diterpenodaurichromenicacid.

Supplemental Table S1. DCA synthase activity under various conditions.

Supplemental Table S2. Steady-state kinetic parameters of cannabinoidsynthases.

Supplemental Table S3. Oligonucleotide primers and PCR conditionsused in this study.

ACKNOWLEDGMENTS

We thank Dr. Y. Niwa (Shizuoka University) for pUC18 plasmid containingthe S65T-type GFP gene; H. Fujino, Y. Tatsuo, Y. Takao, and Y. Murakami, atthe Experimental Station for Medicinal Plant Research of the University ofToyama, for breeding R. dauricum plants; and Dr. T. Obita (University ofToyama), T. Matsumoto (University of Toyama), and Dr. T. Matsui (TohokuUniversity) for helpful advice on instrumental analyses.

Received May 1, 2017; accepted June 28, 2017; published July 5, 2017.

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