Phytochemistry 71 (2010) 1040–1049

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Phytochemistry

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Review

Multifunctional flavonoid dioxygenases: Flavonol and anthocyanin biosynthesisin Arabidopsis thaliana L.

Stefan Martens a,*, Anja Preuß b, Ulrich Matern a

a Institut für Pharmazeutische Biologie, Philipps Universität Marburg, Deutschhausstr. 17A, D-35037 Marburg/Lahn, Germanyb Fachgebiet Biomolekulare Lebensmitteltechnologie, TU München, Hochfeldweg 1, 85354 Freising, Germany

a r t i c l e i n f o a b s t r a c t

Article history:Received 29 January 2010Received in revised form 14 April 2010Available online 8 May 2010

Keywords:Flavonoid pathwayDioxygenasesFlavonolsAnthocyaninsArabidopsis thaliana

0031-9422/$ - see front matter � 2010 Elsevier Ltd. Adoi:10.1016/j.phytochem.2010.04.016

* Corresponding author. Present address: FondazAgrario San Michele all’Adige, IASMA, Centro Ricercaazione, Via E. Mach 1, 38010 San Michele all’Adig615541; fax: +39 0461 615200.

E-mail address: stefan.martens@iasma.it (S. Marte

Flavonols and conditionally also anthocyanins, aside from flavonols, are the predominant polyphenolsaccumulated in various tissues of the model plant Arabidopsis thaliana L. In vitro experiments suggestedthat the dioxygenases involved in their biosynthesis, flavonol synthase and anthocyanidin synthase, are‘‘multifunctional” enzymes showing distinct side activities. The in vivo relevance of the additional activ-ities attributed to these enzymes, however, has remained obscure. In this review we summarize the mostrecent results and present final proof of the complementing activities of these synthases for flavonol andanthocyanidin formation in the model plant A. thaliana. The impact of their modification on the biosyn-thetic pathway and the pattern of flavonoids in different plant tissues are discussed.

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Contents

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10402. Physiological function in plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10413. Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1042

3.1. Grid of flavonoid biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10423.2. General features of 2-oxoglutarate dependent dioxygenases

(2-ODDs) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10433.3. Pivotal role of flavonol synthases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10433.4. Formation of anthocyanidins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1044

4. Flavonol and anthocyanidin biosynthesis in A. thaliana . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10455. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1047

Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1047References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1047

1. Introduction

Flavonoids represent a highly diverse class of polyphenolic sec-ondary metabolites which are abundant in spermatophytic plants,but have also been reported from primitive taxa, such as liverworts(Conocephalum conicum L., Conocephalaceae; Feld et al., 2003) andhorsetails (Equisetum arvense L., Equisetaceae; Oh et al., 2004).

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ione Edmund Mach, Istitutoe Innovazione, Area Aliment-e (TN), Italy. Tel.: +39 0461

ns).

These polyphenols, although usually not essential for growth, havebeen accredited with particular functions during plant develop-ment and propagation ranging from pigmentation over ultravioletscreening to signaling interactions with insects and microbes(Harborne and Williams, 2000; Ververidis et al., 2007a). Most nota-bly, they appear to be fundamental for environmental interactions,i.e. in the adaption to ecological niches or for coping with abioticstresses. Flavonoids accumulate constitutively or may be inducedupon challenge in flowers, fruits, green tissues and roots. Basedon the oxidation state and substitution pattern nine subgroups(Fig. 1) were distinguished: flavanones (synonym dihydroflav-ones), dihydroflavonols (DHFs), flavan-3,4-diols, anthocyanidins,flavones, flavonols, flavan-3-ols, proanthocyanidins (PAs) and

O1

2

3

45

6

7

8

2'

3'

4'

5'

6'

flavonoid skeletal structure

A C

B

A

flavanones

23

O

OC

dihydroflavonols

23

O

O

OHC

flavonols

23

O

O

OHC

flavones

23

O

OC

23

O

OHC+

anthocyanidinsisoflavones

23

CO

O

flavan-3,4-diols

23

CO

OHOH

flavan-3-ols

23

CO

OH

B

Fig. 1. Flavonoid structures. A. Flavan (C6–C3–C6) skeleton of flavonoids. B. The oxidation status and saturation of the C-heterocycle define the subgroups of flavonoids.

S. Martens et al. / Phytochemistry 71 (2010) 1040–1049 1041

isoflavonoids. Overall, about 10,000 flavonoids have been recordedwhich represent the third largest group of natural productsfollowing the alkaloids (�12,000) and terpenoids (�30,000)(Tahara, 2007; Ziegler and Facchini, 2008; Degenhardt et al.,2009). However, the number of flavonoid structures to be expectedin the plant kingdom might reach astronomical values assumingthat twelve of the flavan carbons (Fig. 1A) can be modified by arange of different substituents (Williams and Grayer, 2004). Withthe exception of flavan-3-ols (catechins) these compounds accu-mulate mainly in form of their glycosides and are often concen-trated in the upper epidermal cells of leaves and fruit skins. It iscommon that several types of flavonoids are present in a singleplant species.

Significant quantities of flavonoids are consumed as part of thedaily human diet, and their prospective health-promoting effectsprovide the rationale for further respective food supplements. Nev-ertheless, some concern was also raised about potentially deleteri-ous effects of a high flavonoid diet, but this has been revoked aslargely unfounded (Okamoto, 2005). Cardioprotective, anti-oxida-tive, anti-angiogenic, anti-inflammatory or neuroprotective prop-erties and even putative anticancer potential have been ascribedparticularly to flavonols, e.g. quercetin (Harborne and Williams,2000; Ross and Kasum, 2002; Havsteen, 2002). Rutin (quercetin3-O-rutinoside), the major flavonol in Citrus species (Rutaceae) orbuckwheat (Fagopyrum esculentum Moench., Polygonaceae) andmany other plants, has been shown to be an efficient nitric oxidescavenger, a property that averts nitric oxide-induced tissue dam-age (Haenen et al., 1997). These effects are in the focus of currentinvestigations. Moreover, other beneficial activities have also beendocumented, i.e. the inhibition of cyclooxygenase and/or 5-lipoox-ygenase activities involved in arachidonic acid metabolism as wellas the inhibition of the acetyltransferase forming the plateletaggregating factor, and thus flavonoids may act in a number ofways on blood components such as platelets, monocytes, or low-density lipoprotein (Harborne and Williams, 2000; Martens and

Mithöfer, 2005; Ververidis et al., 2007a). Much less is known aboutthe free-radical scavenging and pharmacological potentials ofanthocyanins (Williams and Grayer, 2004) which impress mostlyby their range of great colours. Accordingly, there is considerableinterest to enhance the level of bioactive flavonols in plants whichare commonly grown for human consumption (Luo et al., 2008).

2. Physiological function in plants

Colourless flavonols are among the most abundant flavonoidsfound in plants, usually conjugated in form of mono-, di- or trigly-cosides (Böhm et al., 1998). Their physiological relevance presum-ably differs across different plant species, but the absorptionmaximum at 280–320 nm predicts a major function in tissue-pro-tection from UV radiation (Stracke et al., 2007 and referencestherein). This can be inferred, for example, from soybean cultivars(Glycine max L., Fabaceae) with enhanced tolerance to UV–B irradi-ation which accumulated significantly more flavonols than sensi-tive cultivars, and the expression of flavonol biosynthesis wasinduced upon UV treatment (Reed et al., 1992; Kim et al., 2008).Sunlight also enhanced the flavonol accumulation in grape vine(Vitis vinifera L., Vitaceae) (Fujita et al., 2006). Alternatively, flavo-nols may act as ultraviolet flower pigments of attractive and defen-sive functions for insects (Gronquist et al., 2001). Furthermore,flavonols possess a planar ring-architecture and may act as copig-ments sandwiched between anthocyanin molecules to shift thecolour of flowers and fruits (Yoshitama et al., 1992; Nielsen et al.,2002). Flavonols might be essential for male fertility, i.e. pollengermination and tube growth of Zea mays L. (maize, Poaceae) andPetunia hybrida Hort. ex Vilm. (petunia, Solanaceae), but not ofArabidopsis thaliana (L.) Heynh. (mouseear cress, Brassicaceae) orEustoma grandiflorum Grise. (lisianthus, Gentianaceae) (Mo et al.,1992; van der Meer et al., 1992; Burbulis et al., 1996; Nielsenet al., 2002). They were also proposed to be involved in auxin

1042 S. Martens et al. / Phytochemistry 71 (2010) 1040–1049

metabolism and transport as well as in the defense against mi-crobes (Stafford, 1991; Stracke et al., 2007 and references therein).Other functions attributed to flavonols concern the influence onlight and gravity responses of plants or the repression of bud out-growth affecting lateral branching. These effects have been ana-lyzed in mutants only (Stracke et al., 2007 and referencestherein; Owens et al., 2008a).

Anthocyanin pigments of pink, red, orange, scarlet, purple, blueor blue-black and also yellow appearance have been reported(Davies et al., 2003), and the intensity and hue of tissue colourdepends largely on the hydroxylation and glycosylation patternsas well as on the co-pigmentation with other flavonoids, e.g. flavo-nols. The bright colour of these pigments is often a critical factorfor insect-mediated pollination of plants, and the pigments areimplicated in further processes such as modulation of hormone re-sponses, UV–B protection, and photo-perception of autumn foliage(Reddy et al., 2007 and references therein). Anthocyanins havebeen reported not only from flower petals, but also from leaves,stems, roots, tubers, fruits and seeds (Williams and Grayer,2004). In many instances colourless plant tissues are capable ofaccumulating anthocyanins under stress conditions such as lowavailability of nitrogen and phosphate, wounding, pathogen infec-tion, treatment with methyl jasmonate, water stress or irradiationby ultraviolet, visible and far-red light (Stewart et al., 2001). Lowtemperature also induces anthocyanin synthesis in numerousplant species including A. thaliana and maize (Rowan et al., 2009).

3. Biosynthesis

3.1. Grid of flavonoid biosynthesis

The principles of flavonoid biosynthesis, i.e. the biochemistryand molecular genetics, have been thoroughly studied over the lastdecades, and flavonoids belong to the most intensively investi-gated plant secondary metabolites (Davies and Schwinn, 2007;Ververidis et al., 2007b). Advanced technologies including the re-combinant expression enabled the in vitro classification of individ-ual enzymes and, in particular, the determination of substrate andproduct specificities. The basal structure of a flavanone (Fig. 1),consisting of a substituted 2,3-dihydro-2-(4-hydroxyphenyl)-4H-1-benzopyran-4-one as in naringenin, is composed from p-couma-royl-CoA contributing nine carbons forming the B- and C-rings,while the six carbons of the A-ring are added by three cycles of apolyketide chain elongation reaction requiring malonyl-CoA as

3 x malonyl-CoA

p-coumaroylCoA

Chalcones Flavanones Dihydrflavon

CHI FHT

Isoflavones

Flavones Flavon

IFS/HID

FNSI&IIF2H/ HFD

CHS

Fig. 2. General outline of the flavonoid pathway. CHS, chalcone synthase; CHI, chalcone i(ANS), anthocyanidin synthase; FGT, flavonoid glycosyltransferase; FNS, flavone synthareductase; IFS, isoflavone synthase; HID, 2-hydroxyisoflavanone dehydratase, F2H, flava

the co-substrate. p-Coumaroyl-CoA, supplied by the consecutiveaction of the shikimate and general phenylpropanoid pathways,serves as a starter substrate for condensation which is catalyzedby a type III polyketide synthase known as chalcone synthase(CHS). The cyclization of the A-ring yielding the chalcone concom-itantly with the polycondensation is a peculiar feature of CHSwhich is the first enzyme committed to flavonoid biosynthesis.Subsequently, chalcone isomerase (CHI) catalyzes the stereospe-cific intramolecular cyclization (Michael type addition) of thechalcone to the respective (2S)-flavanone, although the non-stereospecific cyclization occurs readily at room temperature.

(2S)-Flavanone is the common substrate for the various flavo-noid branch pathways which rely on three major reactions: arylmigration, oxidation or hydroxylation. Isoflavones emerge from(2S)-flavanones by C-2 to C-3 aryl migration concomitant with C-2 hydroxylation. This reaction is catalyzed by isoflavone synthase(IFS; synonym 2-hydroxyisoflavanone synthase, 2HIS) which hasbeen characterized in detail from several legume plants as amembrane-bound cytochrome P450 (Cyt P450) monooxygenase(Hagmann and Grisebach, 1984; Davies and Schwinn, 2007).Dehydration of the IFS product delivers the isoflavone (Fig. 1).The oxidation of (2S)-flavanones introducing a double bond at C-2/C-3 to yield flavones is accomplished by flavone synthase I or II(FNS I and II) (Figs. 1B and 2). FNS I was characterized as a 2-oxo-glutarate-dependent dioxygenase (2-ODD) and has been reportedfrom species of the Apiaceae family only (Martens et al., 2001;Gebhardt et al., 2005, 2007) except for one recent report from rice(Lee et al., 2008). In contrast, the Cyt P450 monooxygenase FNS IIwas expected to be more widely distributed than FNS I (Martensand Mithöfer, 2005) and has been described from various plantfamilies. Dihydroflavonols (Fig. 1B and 2) arise from (2S)-flavanon-es by the action of flavanone 3ß-hydroxylase (FHT, synonym F3H),another member of the 2-ODD class of enzymes. FHT occupies akey position in flavonoid metabolism, competing with FNS I or IIand controlling the flux of flavanones into branch pathways forend products of distinct physiological functions (Owens et al.,2008b) (Fig. 2). Colourless or yellowish flavonols (Fig. 2) may beformed from dihydroflavonols by the action of flavonol synthase(FLS) which again belongs to the 2-ODD class of enzymes. In com-petition with FLS, jointly acting dihydroflavonol 4-reductase (DFR)and leucoanthocyanidin dioxygenase (LDOX; synonym anthocy-anidin synthase, ANS) may utilize dihydroflavonols for the forma-tion of anthocyanidins (Fig. 2) or proanthocyanidins (Davies et al.,2003; Davies and Schwinn, 2007). Flavonoids and particularlyanthocyanidins may be glucosylated by UDP-glucose:flavonoid

o-ols

Leuco-anthocyanins

Antho-cyanins

Antho-cyanidins

DFR LDOX FGT

ols trans-Flavan-3-ols

cis-Flavan-3-ols

Proanthocyanidins

FLS LAR ANR

somerase; FHT, flavanone 3-b-hydroxylase; DFR, dihydroflavonol 4-reductase; LDOXse; FLS, flavonol synthase; LAR, leucoanthocyanidin reductase; ANR, anthocyanidinnone 2-hydoxylase; HFD, 2-hydroxyflavanone dehydratase.

S. Martens et al. / Phytochemistry 71 (2010) 1040–1049 1043

O-glucosyltransferases (UFGT) for stable storage of pigments(Davies and Schwinn, 2007).

Modifications of the flavonoid skeleton enable the formation ofnumerous metabolites. These modifications require precisely regu-lated and coordinated hydroxylation, oxidoreduction, methylation,acetylation or glycosylation reactions. For example, flavonoid 30

and 30, 50-hydroxylases (F30H; F3050H) which both belong to theCyt P450 family catalyze the single or double hydroxylation ofthe B-ring (Fig. 1A) of flavanones or other flavonoids, which signif-icantly affect the physiological quality of flavonols or anthocyaninsas the end products of the branch pathways (Davies and Schwinn,2007). Multiple activities are necessary to synthesize the full spec-trum of natural flavonoids, but only a few of these enzymes have sofar been thoroughly examined at the biochemical and molecularlevel. It has become obvious, however, that genes selected bymutational studies or homology cloning require the functionalexpression and biochemical characterization of the polypeptidesfor unequivocal annotation. The marginal difference of only 10%in the DNA sequences of parsley FHT and FNS I is a striking exam-ple, emphasizing that base alignments are insufficient to provefunctionality (Gebhardt et al., 2005, 2007). Unfortunately, a thor-ough biochemical characterization of enzymes attributed to flavo-noid metabolism has often been neglected.

3.2. General features of 2-oxoglutarate dependent dioxygenases(2-ODDs)

A plethora of 2-ODDs is known from bacteria, fungi, plants orvertebrates, which catalyze diverse reactions including aliphatichydroxylation, epoxidation, desaturation and desaturating cycliza-tion, respectively. Most of these enzymes are involved inbiosynthetic processes leading to collagen or other modifiedpolypeptides and amino acids, alkaloids, ethylene, gibberellinsand ß-lactam antibiotics, i.e. penicillins and cephalosporins(Prescott, 1993; DeCarolis and DeLuca, 1994; Prescott and John,1996; Schofield and Zhang, 1999; Prescott and Lloyd, 2000).Accordingly, studies on these dioxygenases were inspired by themedicinal and industrial relevance and focused initially on thearchitecture and mode of action of microbial and human 2-ODDs,but in recent years considerable insight has also been achieved intothe functional organisation of plant 2-ODDs. A single plant may ex-press numerous putative 2-ODDs, as was revealed by sequenceanalysis of the A. thaliana genome coming up with about 100isogenes (Wilmouth et al., 2002).

The activities of 2-ODDs depend on ferrous iron which is re-quired for the reduction of molecular oxygen. Two electrons areprovided in this process by the decarboxylation of a cosubstratewhich is most commonly 2-oxoglutarate. The stabilization of iro-n(II) in the active site of these labile soluble enzymes is broughtabout by the unique conformation of the enzyme polypeptide.Eight amino acid residues are strictly conserved in all of these diox-ygenases which include two histidines (His221, His277; numbers re-fer to AtFLS1 sequence) and one acidic amino acid (Asp223) as wellas one arginine (Arg287) and one serine (Ser289) proposed to bind 2-oxoglutarate. Crystallography of 2-ODDs revealed that theconserved motifs Hx(D/E)xnH and RxS are part of a characteristicdouble-stranded beta helix (Clifton et al., 2006) and two sets offour anti-parallel beta sheets forming a sandwich-like structure(jelly roll topology) which shield the ferrous iron in the active cen-ter for oxygen activation and catalysis (Koehntop et al., 2005). Theactive-site iron is coordinated to the histidine-rich motif whichmore often shows the Hx(D)xnH composition (Clifton et al.,2006). Four amino acids (Gly68, His75, Gly261, Pro207) are conservedwith no obvious functionality but most likely required for properfolding of the polypeptide (Wellmann et al., 2002). Computationalanalysis (Chua et al., 2008) suggested further five residues

(His132, Phe134, Lys202, Phe293 and Glu295), albeit less conserved,for substrate binding.

Five 2-ODDs have been identified in the context of flavonoidbiosynthesis. FHT (hydroxylating), FLS (desaturating) and LDOX(hydroxylating/dehydrating) activities (Fig. 2) are widely distrib-uted (Davies and Schwinn, 2007). FNS I (desaturating) appears tobe confined to species of the Apiaceae (Gebhardt et al., 2005,2007) with a sole finding in rice (Lee et al., 2008), while Cyt P450take over the FNS I-equivalent function (Fig. 2) in other plants(Davies and Schwinn, 2007). It remains to be established, whethercatalysis by these Cyt P450s follows the same mode, because ahydroxylated substrate intermediate was ruled out for the FNS Ireaction (Britsch, 1990; Martens and Mithöfer, 2005). It is note-worthy that an analogous scenario was reported for flavonoid 6-hydroxylation which is catalyzed by a 2-ODD in Chrysospleniumamericanum Schwein. ex Hook. (American golden saxifrage,Saxifragaceae; Anzelotti and Ibrahim, 2004) but a Cyt P450 fromsoybean (Latunde-Dada et al., 2001). Among the flavonoid-com-mitted 2-ODDs most mechanistic and molecular investigationshave been conducted on LDOX or FLS, and X-ray diffraction ofLDOX-naringenin co-crystals already revealed the putative sitesof substrate binding (Welford et al., 2001; Turnbull et al., 2004;Gebhardt et al., 2007).

3.3. Pivotal role of flavonol synthases

The full perception of factors and enzyme activities regulatingthe tissue-level of flavonols is a prerequisie for breeding of flavo-nol-enriched fruits and vegetables, but determines also the per-spectives of metabolic engineering of plants or microorganisms(Ververidis et al., 2007b). FLS catalyzing the oxidation of dihydrofl-avonols to flavonols competes at a crucial branch point withdihydroflavonol reductase (DFR) in the anthocyanin/PA pathway(Fig. 2). FLS was reported first from irradiated parsley cells andcharacterized as a soluble 2-ODD requiring 2-oxoglutarate, FeII

and ascorbate for full activity (Britsch et al., 1981). This enzymewas considered to be specific for natural (+)-(2R, 3R)-dihydroflavo-nol substrates. Much later, a putative FLS cDNA was cloned from P.hybrida and functionally expressed in yeast and plants (Holtonet al., 1993), which was followed by FLS sequences from variousother plants, such as A. thaliana (Pelletier et al., 1997; Wismanet al., 1998), Citrus unshiu Marc. (satsuma mandarin, Rutaceae;Moriguchi et al., 2002), E. grandiflorum (Nielsen et al., 2002), Sola-num tuberosum L. (potato, Solanaceae; van Eldik et al., 1997), Malus� domestica Borkh. (apple, Rosaceae; Lee et al. direct submission toGenbank), Matthiola incana (L.) R. Br. (stock, Brassicaceae; Henkeland Forkmann, direct submission to Genbank), G. max (Takahashiet al., 2007), Petroselinum crispum L. (parsley, Apiaceae; Martenset al., 2003a) or Fragaria � ananassa Duch. (strawberry, Rosaceae;Almeida et al., 2007). The translated FLS sequences show a remark-able degree of conservation (about 85% similarity, �50% identity)and share partial similarity with LDOX (50–60%), but much lesswith the related 2-ODD enzymes FNS I or FHT. It is noteworthy thatFNS I is capable of oxidizing flavanones, i.e. (2S)-naringenin (Fig. 3),to flavones analogous to the 2,3-desaturation of dihydroflavonols(Fig. 1B) by FLS. However, FNS I does not accept dihydroflavonolsas substrates, and provisional assays indicated that FLS does notform flavones from flavanones (Fig. 1B).

The first detailed FLS-study was carried out on the FLS from C.unshiu expressed in Escherichia coli (Wellmann et al., 2002). The re-combinant enzyme surprisingly accepted (2S)-naringenin as sub-strate which was almost completely converted to kaempferol.Minute amounts of dihydrokaempferol recovered from these as-says suggested that (2R, 3R)-dihydrokaempferol is an intermediateen route to kaempferol, and thus FHT/FLS bifunctionality (Fig. 3)must be assigned to FLS oxidizing both (+)-trans-dihydroflavonols

FHT

DFR

LDOX

FLS1 & 3

R1 = H; naringeninR1 = OH; eriodictyol

FLS1LDOX

LDOX

FLS1/LDOX

R1 = H; dihydrokaempferolR1 = OH; dihydroquercetin

R1 = H; leucopelargonidinR1 = OH; leucocyanidin

R1 = H; pelargonidinR1 = OH; cyanidin

R1 = H; kaempferolR1 = OH; quercetin

OH O

OH

R1OH

OHO

23

3'

4'

OH

O

O

OH

OH

R1

OH3

3'

4'

2

OH O

OH

OH

R1

OHOH

3

3'

4'

2OH O

OH

OH

R1

OH3

3'

4'

2

OH

O

O

OH

OH

R1

23

3'

4'

Fig. 3. Postulated scheme of flavonoid biosynthesis in A. thaliana. The side activities of LDOX replace partially the function of FLS1 and/or 3 in the formation of flavonols andanthocyanidins/proanthocyanidins. Full arrows – common pathway; dotted arrows – side activities.

1044 S. Martens et al. / Phytochemistry 71 (2010) 1040–1049

and (2S)-flavanones to the respective flavonols (Lukacin et al.,2003). Moreover, unnatural (2R)-naringenin was converted to(�)-trans-dihydrokaempferol by the FLS from C. unshiu yieldingonly traces of kaempferol, and equivalent specificities at lowerconversion rates were observed with FLSs from P. crispum,F. � ananassa or A. thaliana (Prescott et al., 2002; Turnbull et al.,2004; Almeida et al., 2007; Preuß et al., 2009). Nevertheless, allFLSs examined so far completely lacked FNS I activity. Thus, FLS be-longs to a group of 2-ODDs with broad specificities, whereas FNS Iand FHT show narrow substrate specificity and appear to form aseparate group of dioxygenases in accordance with their high de-gree of sequence homology (Martens et al., 2003a; Gebhardtet al., 2005, 2007).

3.4. Formation of anthocyanidins

Some information on the molecular regulation of anthocyanidinproduction is available from A. thaliana, where the PRODUCTION OFANTHOCYANIN PIGMENTS1 (PAP1) gene, a conserved MYB regulatorof phenylpropanoid biosynthesis, plays a key role in mediating theenvironmental regulation of anthocyanins but not flavonols(Rowan et al., 2009). The reduction of dihydroflavonols by DFR tocolourless cis-(2R, 3S, 4S) leucoanthocyanidins (flavan-3,4-diols)marks the beginning of the anthocyanidin-specific branch pathway(Fig. 2), and LDOX was shown to catalyze in planta the subsequentconversion to coloured, labile anthocyanidins (Fig. 2) (Saito et al.,1999; Welford et al., 2001). The latter step formally involves thedehydrogenation at C-2 and C-3/C-4 dehydration. Putative LDOXgenes and cDNAs have been isolated from Antirrhinum majus L.

(snapdragon, Scrophulariaceae), Perilla hybrids, Torenia hybrids L.(torenia, Scrophulariaceae), Perilla frutescens L. (beefsteak plant,Lamiaceae), A. thaliana, Gerbera hybrida L. (gerbera, Asteraceae)and Z. mays (Saito et al., 1999; Nakajima et al., 2001; Turnbullet al., 2001; Wellmann et al., 2006). The essence of LDOX for leuco-anthocyanidin-anthocyanidin conversion was demonstrated byrestoration/complementation experiments in LDOX-minus linesof maize (A2 mutant) and Zantedeschia aethiopica L. (calla lily, Ara-ceae; natural LDOX block in white spathe), where overexpressionof LDOX cDNAs from maize, rice and Gerbera or transient expres-sion by particle bombardment restored the anthocyanin formation(Menssen et al., 1990; Martens et al., 2003b and unpublished;Reddy et al., 2007). The alternative suppression of LDOX by anti-sense, sense or RNAi techniques clearly reduced the anthocyaninaccumulation in flowers of torenia (Nakamura et al., 2006).Nevertheless, the exact biochemical function of LDOX remainedunknown till the recombinant enzyme from P. frustescens was as-sayed with leucoanthocyanidin as substrate (Saito et al., 1999).Translated LDOX sequences from different plants share 48–78%similarity and less, albeit significant, similarity with FLS (50–60%)and FHT or FNS I (about 30%), respectively.

The physiological role of LDOX has been put into question, be-cause the recombinant enzyme from A. thaliana was classified asa non-specific 2-ODD catalyzing primarily the desaturation oftrans-dihydroquercetin to quercetin, analogous to FLS, and theformation of 3,4-cis-dihydrokaempferol as a major product from(+/�)-naringenin (Turnbull et al., 2001; Welford et al., 2001;Lukacin et al., 2003; Martens et al., 2003a; Wellmann et al., 2006).Furthermore, the proposed natural substrate cis-(2R,3S,4S)leucocy-

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anidin and trans-(2R,3S,4R)leucocyanidin were oxidized to querce-tin as the predominant product (>85%) and cis-dihydroquercetinwith only trace amounts of cyanidin (Turnbull et al., 2003). Thein vivo significance of the findings has remained obscure, but theobserved side activities of LDOX and also FLS raised the possibilitythat these enzymes provide an alternative route to flavonols in situin mutant lines lacking FHT or FLS activity. However, the proof ofconcept is still missing.

4. Flavonol and anthocyanidin biosynthesis in A. thaliana

The flavonoid pattern in A. thaliana is simple and consists solelyof flavonol glucosides derived from kaempferol, quercetin or isorh-amnetin, beside anthocyanins and PAs (Pelletier et al., 1997, 1999;Veit and Pauli, 1999; Peer et al., 2001; Broun, 2005; Routaboulet al., 2006; Stracke et al., 2009). The flavonols accumulate mainlyin the vacuoles of epidermal cells (Feinbaum and Ausubel, 1988;Hartmann et al., 1998; Landry et al., 1995), whereas PAs are foundin specialized tissues, i.e. the seed testa (Lepiniec et al., 2006), andthe accumulation of anthocyanins might be induced by low tem-perature in correlation with the abundance of flavonoid-commit-ted transcripts (Leyva et al., 1995). The common substrates forall flavonoids of A. thaliana are dihydroflavonols which fuel thebranch pathways to flavonols, depending on FLS, and to anthocy-anidins/PAs, catalyzed by DFR and LDOX (Fig. 2).

While most of the biosynthetic enzymes appear to be encodedby single copy genes, e.g. LDOX, the A. thaliana genome containsa small family of up to six FLS isogenes (Pelletier et al., 1997;Stracke et al., 2007) as is also the case in S. tuberosum (van Eldiket al., 1997), E. grandiflorum (Nielsen et al., 2002) and V. vinifera(Fujita et al., 2006). The FLS isogenes were tentatively identifiedby their homology to 2-ODDs and clustering with FLSs. All sixgenes are located on chromosome 5 with AtFLS2, -3, -4, and -5 ar-ranged in a 7.5-kb tandem array. The four clustered genes are notmore closely related to each other than to the other two genes (62–73% identity at the nucleotide level) with AtFLS2 being most distant(48–51% identity), which suggests ancient duplications events(Owens et al., 2008a; Stracke et al., 2009). The putative A. thalianaFLS gene family and the LDOX gene were examined in detail onlyrecently using genetic and biochemical approaches (Owens et al.,2008a; Stracke et al., 2009; Preuß et al., 2009). Recombinantexpression and enzyme assays revealed activities of only AtFLS1and AtLDOX (Wisman et al., 1998; Saito et al., 1999; Winkel-Shir-ley, 2001; Prescott et al., 2002), whereas AtFLS2-6 were assignedinactive (Owens et al., 2008a; Stracke et al., 2009). Nevertheless,the contribution of AtFLS isoenzymes of variant substrate specific-ities or differential regulation had been suggested as a means ofcontrol of flavonoid synthesis in A. thaliana, which might explainthe variable composition of flavonol glucosides in different tissuesor upon a change in environmental conditions (Pelletier et al.,1997; Mehrtens et al., 2005; Owens et al., 2008a and referencestherein).

The differential biosynthesis of flavonols and anthocyanidins inA. thaliana, just like in maize and grape, is also controlled by spe-cific transcription factors (TFs) (Mehrtens et al., 2005; Fujitaet al., 2006). Regulation of anthocyanins and PA accumulation viacombinatorial action of MYB and basic helix-loop-helix (bHLH)-type TFs is a conserved feature in plants, and the R2R3-MYB factorMYB12 was identified recently as a flavonol-specific regulator offlavonoid biosynthesis in A. thaliana. MYB12 activates coordinatelyAtCHS, AtCHI, AtFHT and AtFLS1, the four genes required for the for-mation of a flavonol. However, a preference for either one AtFLSisogene was not observed and the question whether MYB 12 is in-volved in any of the many stress signaling pathways that result inflavonol accumulation is still unanswered (Mehrtens et al., 2005).

Interestingly, different flavonols predominate in the various tissueswith quercetin being the primary seed flavonol and kaempferol themain flavonol in whole flowers, whereas similar levels of these twoflavonols are found in stamens (Pelletier et al., 1997). Only traces ofisorhamnetin are found in seedlings and leaves (Stewart et al.,2001; Routaboul et al., 2006). Moreover, limitations in the supplyof nitrogen or phosphorus induce higher concentrations of flavo-nols in seedling and vegetative tissue of A. thaliana (Stewartet al., 2001). The final composition of the flavonol contents in dif-ferent tissues of A. thaliana depends on the responsiveness of genesthat encode flavonol glucosyltransferases and other enzymesdownstream (i.e. modifying kaempferol) in the biosynthetic path(Stracke et al., 2007).

Experiments with fht mutant lines (transparent testa 6, tt6) orfls1 knock-out lines of A. thaliana revealed some unforeseen accu-mulation of flavonol, anthocyanin and PA, whereas chs mutants(tt4) literally lacked flavonoids (Burbulis et al., 1996; Wismanet al., 1998; Pelletier et al., 1999; Routaboul et al., 2006; Strackeet al., 2007; Owens et al., 2008a,b; Ric de Vos, personal communi-cation). The fht and fls genotypes are generally characterized bypale-brown rather than yellow seeds, indicating the accumulationof anthocyanins and/or PAs. The Atfls1 plants also exhibited a muchmore intense red colouration of the hyptocotyl and cotyledons dur-ing germination and at the base of the stalk of mature plants com-pared with the wild-type (Owens et al., 2008a; Stracke et al., 2009).A similar shift in metabolites of branch pathways has been re-ported for the flavonoid mutant banyuls which is deficient in anth-ocyanidin reductase (ANR; Fig. 2) (Devic et al., 1999). Furthermore,the synthesis of flavonols was not eliminated entirely in Atfls1plants lacking a functional flavonol-specific TF MYB12 factor(Mehrtens et al., 2005).

The unexpected flavonol accumulation in mutants togetherwith the observed multifunctionality of FLS and ANS in vitro andthe presence of FLS isogenes led to the following speculations:

(a) AtFLS isozymes with different substrate specificities controlthe amount and type of flavonols present in specific tissues,

(b) flavonols present in a given tissue or different AtFLS genesmay be regulated independently of one another in a devel-opmental- or tissue-specific manner,

(c) an additional AtFLS isogene and/or AtLDOX side activity isresponsible for flavonol formation under in situ conditions,

(d) AtFLSs and/or AtLDOX can complement the FHT or FLS step.

First support for the hypothesis of contributing FLS/LDOX sideactivities in flavonoid biosynthesis in planta was recently providedby overexpressing LDOX in rice plants (Reddy et al., 2007), by thebiochemical characterization of AtFHT from A. thaliana mutant linett6 (Owens et al., 2008b), and from metabolomic and genetic anal-yses in A. thaliana (Stracke et al., 2009). The accumulation of a mix-ture of flavonols and anthocyanins increased concomitantly with adecrease of PAs in transgenic rice plants overexpressing homolo-gous LDOX cDNA (Reddy et al., 2007). The authors suggested thatLDOX may execute more than one dioxygenase activities on differ-ent flavonoid substrates. The ‘‘leaky” phenotype of Atfht mutant al-leles (tt6), producing pale-brown rather than yellow seeds, wasbiochemically and genetically characterized and provided evidencethat FLS and LDOX compensate in vivo, at least partially, for the dis-rupted FHT enzyme. Similar ‘‘leaky” phenotypes apparentlyblocked in FHT were also reported from G. max and Dianthus cary-ophyllus L. (carnation, Asteraceae) suggesting an analogous com-pensation by related 2-ODDs in these plants as well (Owenset al., 2008b). The findings of Fujita et al. (2006) may support thesisb for grape-vine, because each of the five FLS genes was suggestedto encode functional enzyme due to highly conserved amino acidresidues at the relevant binding sites. Furthermore the transcrip-

1046 S. Martens et al. / Phytochemistry 71 (2010) 1040–1049

tion patterns of the five FLSs varied with the organ and develop-mental stage.

In a set of experiments employing A. thaliana tissue from seed-lings and adult plants of wild-type (Col-0) and mutant lines (fls1-2mutant, ldox/fls1-2 double mutant) the role of LDOX and/or FLSs forflavonoid accumulation was investigated (Preuß et al., 2009). A.thaliana ldox/fls1-2 double mutant revealed a significant reductionin flavonol level as compared to the wild-type and the fls1-2 in cor-relation with the genetic data (Stracke et al., 2009). Additionally, atthe expense of flavonols a significant accumulation of dihydrofl-avonol glycosides was observed (Stracke et al., 2009). Similar find-ings were recorded from petunia and lisianthus flowers expressinghomologous antisense-FLS constructs (Holton et al., 1993; Nielsenet al., 2002). It must be emphasized again that extracts of all threegenotypes contained flavonols, albeit at greatly different levels,which assigns FLS-like enzyme activities to all genotypes (Strackeet al., 2009). Crude protein preparations from these plant tissueswere prepared (Martens et al., 2003a), and standard enzyme assayswith [14C]naringenin or [14C]dihydrokaempferol as substrateclearly revealed FHT and FLS activities in wild-type extracts, con-verting both substrates to kaempferol, whereas extracts of thefls1-2 single mutant and the ldox/fls1-2 double mutant lacked theseactivities (Preuß et al., 2009). LDOX activity, however, has neverbeen demonstrated in crude plant extracts of Arabidopsis or anyother species, which can be explained by the instability of the en-zyme during isolation and in vitro assay (Saito et al., 1999). Plant 2-ODDs are commonly rather labile enzymes which might suffer ra-pid partial digestion after extraction prohibiting their functionalidentification (Lukacin et al., 2000a). Furthermore, low FLS activityof either one of the other putative FLS polypeptides or LDOX incrude extracts might have escaped the detection by in vitro assayswhich were commonly run for 30 min only (Martens et al., 2003a),while other groups used extended periods of incubation, i.e. up to3 h with recombinant A. thaliana FLS1 (Prescott et al., 2002), for thedetection of products on subsequent HPLC analysis.

Arabidopsis AtFLS6 and AtFLS4 had been identified before aspseudogenes or non-functional copies (Owens et al., 2008a;Stracke et al., 2009). Therefore, only AtFLS1, AtFLS2, AtFLS3 andAtFLS5 as well as AtLDOX were expressed for further functionalcharacterization in microbial systems. Standard assays of the re-combinant polypeptides were conducted employing crude extractsor the purified enzyme fraction. As expected, AtFLS1 efficientlyconverted dihydrokaempferol or dihydroquercetin to kaempferoland quercetin, respectively, with a clear bias towards dihydroka-empferol (kaempferol/quercetin ratio 1:0.7). These results are fullycompatible with previous findings (Prescott et al., 2002), although‘‘unnatural” (2S,3S)-dihydrokaempferol does not appear to be anFLS substrate (Lukacin et al., 2003). AtLDOX also oxidized the sub-strates to the corresponding flavonols, albeit to a lower extent(kaempferol/quercetin ratio 0.7:1). However, neither one of theother putative AtFLSs, tested in crude extracts or after purification,provided any detectable product under standard assay conditions.These results suggested that the FLS-like activity of AtLDOX signif-icantly contributed to the formation of flavonols in the A. thalianafls1-2 mutant. Additional support for this assumption was gainedby the tightened reduction of flavonol contents in the ldox/fls1-2double mutant (Stracke et al., 2009). Nevertheless, the small, butsignificant, residual amount of flavonols in the ldox/fls1-2 mutant(Stracke et al., 2009) required further explanation. The minorFLS-like activity of this double mutant might thus be due to eitherone of the FLS isogenes which had been proposed from in vitro as-says to encode catalytically inactive proteins or to other membersof the 2-ODD family of enzymes with as yet unknown function(Owens et al., 2008a,b).

Sequence alignments of AtFLSs and AtLDOX indicated thatAtFLS2, analogous to truncated pseudogenes AtFLS4 and AtFLS6,

likely encodes a non-functional polypeptide because of a truncatedC-terminus and lack of appropriate iron and 2-oxoglutarate bind-ing sites (Owens et al., 2008a; Stracke et al., 2009; Preuß et al.,2009). A limited loss of C-terminal amino acids may be tolerated,because, at least in Petunia FHT (Lukacin et al., 2000b), this reducedthe activity only. However, marginal changes in the binding sites ofsubstrate, ferrous iron and oxoglutarate cosubstrate strongly affectthe substrate and/or product specificity of 2-ODD enzymes (Wil-mouth et al., 2002). The crucial role of these residues for enzymeactivity had been documented through knock down of AtFLS1activity in studies mutating some of the residues proposed for fer-rous iron or 2-oxoglutarate binding, and these mutations appar-ently affected the positioning of 2-oxoglutarate in the active sitecausing inefficient binding or total loss of function (Chua et al.,2008). Alignment of the AtFLS5 polypeptide with AtFLS1 also re-vealed substantial exchanges which likely affect the fixation ofsubstrate through hydrogen bonds and thus the catalytic activity(Owens et al., 2008a). In AtFLS3, however, the proposed substrateand cofactor binding sites appear fully conserved. Molecular de-tails of substrate binding sites are not yet available for the FLSs,but LDOX may be used as a model enzyme. The high sequence sim-ilarity of LDOX and FLS implies a relationship much closer than tothe other flavonoid 2-ODDS, FHT and FNS I (Gebhardt et al., 2005,2007). Furthermore, the mode of action of FLS must be very similarto that of LDOX, because the ‘‘unnatural” oxidation of dihydroqu-ercetin to quercetin catalyzed in vitro by LDOX is identical to thereaction catalyzed by FLS. Since the crystal structure of LDOXwas reported (Turnbull et al., 2001; Wilmouth et al., 2002), homol-ogy modeling might identify the amino acid residues essential forFLS activity. This approach had been used for FHT and FNS I fromparsley as well as for AtFLS1 (Gebhardt et al., 2007; Chua et al.,2008) and should be amenable to describe the substrate andproduct selectivities of FLS2, 3 and 5. Model structures of AtLDOX-naringenin (2brt) (Welford et al., 2005) and AtLDOX-dihydroquerce-tin complexes (Wilmouth et al., 2002) were employed for homologymodeling of AtFLS1, AtFLS3 and AtFLS5 (Prescott et al., 2002;Martens et al., 2003a). The conserved binding sites for 2-oxoglutar-ate and iron are distributed at an equivalent spacing in AtFLSspolypeptides. Furthermore, the models endorse the role of Glu327

in substrate binding and the lack of activity of AtFLS5 (Preuß,2009). Notably, AtFLS3 differs from AtFLS1 by replacement ofLeu136 through Tyr136, and the AtFLS3-substrate model complexpredicted that this phenolic residue changes the orientation ofnearby His149 which is involved in substrate binding (Preußet al., 2009). Thus, a shift in location of substrate toward the activeferryl species is expected and the kinetic parameters of enzymecatalysis are likely affected. Point mutations in putative substratebinding sites commonly decrease considerably the activity of en-zymes, which in turn confirms their relevance for catalytic turn-over. However, replacing His132 by phenylalanine in thedihydroquercetin binding site of AtFLS generated a mutant ofapproximately 20% higher activity as compared to the wild-type(Chua et al., 2008). In this instance, kinetic analyses indicated animproved substrate binding affinity.

Taking advantage of the in vivo bioconversion protocol yeasttransformants expressing recombinant AtFLSs or AtLDOX werefed with various flavanones and dihydroflavonols (Preuß et al.,2009). This method allows testing of a large number of potentialsubstrates and may define the in vivo substrate specificities withrespect to the types of flavonols accumulating in different tissues.Moreover, improved stability of enzymes is expected in situ and as-says can be extended significantly in time. Recent studies with re-combinant E. coli overexpressing LDOX accordingly demonstratedthe formation of plant-specific anthocyanins and flavonols uponfeeding relevant precursors, which confirms the reliability of theprocedure (Yan et al., 2005). Bioconversions were performed by

S. Martens et al. / Phytochemistry 71 (2010) 1040–1049 1047

incubating cultures of AtFLS or AtLDOX yeast transformants in thepresence of 5 mg of either one of the potential substrates (Witteet al., 2009) for 17 h, and similar experiments were run with E. colitransformants for 4 h. All dihydroflavonols and flavanones appliedwere converted to a variable degree and formation of the corre-sponding flavonols was detected without any intermediates. AtL-DOX transformants showed a considerably lower conversion ratethan cells transformed with AtFLS1, whereas none of the transfor-mants harbouring AtFLS2, 3 or 5 constructs was found to convertany of the dihydroflavonol substrates under these conditions.Therefore, bioconversions were extended to two days, which final-ly revealed a significant conversion of dihydrokaempferol ordihydroquercetin to kaempferol and quercetin, respectively, onlyin the incubations of AtFLS3-yeast transformants. AtFLS3 was there-fore be annotated as a second functional gene in A. thaliana. Never-theless, further site-directed mutagenesis is necessary to identifythose amino acids which determine the turnover rate of FLSs inArabidopsis. Furthermore, AtFLS3 is likely responsible for the for-mation of residual flavonols in seedlings of the ldox/fls1-2 mutantline, but final proof requires the analysis of the respective triplemutant (ldox/fls1-2/fls3) (Preuß et al., 2009).

5. Conclusion

A. thalíana has served as a model plant for various biochemicaland genetic studies of secondary metabolism including the flavo-noid pathway (Winkel-Shirley, 2001). Flavonols (quercetin,kaempferol, isorhamnetin glucosides) and the corresponding fla-van-3-ols or dihydroflavonols (Fig. 1) as well as PAs and anthocya-nins were reported from wild-type plants. At least thirty-five genescommitted to flavonoid biosynthesis have been identified: tenstructural genes (e.g. FLS, LDOX), ten genes coding for modifyingenzymes and 12 regulatory genes (transcription factors; e.g. TT2,TT8), besides three genes involved in flavonoid compartmentation(Yonekura-Sakakibara et al., 2008). A single LDOX gene and a fam-ily of six FLS genes were identified in the Arabidopsis genome, butonly AtFLS1 was shown to yield a functional FLS (Wisman et al.,1998). LDOX and FLS were attributed to different branch pathways(Fig. 2), and FLS should correlate with the accumulation of flavo-nols. Investigations of the expression levels of AtFLS1, 2, 3 and 5 re-vealed a wide tissue-distribution of AtFLS1 with highest transcriptamounts during the reproductive stages. In contrast, the inactiveAtFLS2 appears to be expressed at high levels in the shoot apexand lower stem, where AtFLS1 transcript was absent, and only toa low extent in flowers and siliques. AtFLS5 and 3 were expressedat much lower intensities or extremely low and even falling toundetectable levels. Moreover, all AtFLS isogenes have tissue-and cell-type specific promoter activities that overlap with thoseof AtFLS1 and interact with other flavonoid proteins in yeast twohybrid assays (Owens et al., 2008a).

Recent reports identified AtFLS3 as another active enzyme(Preuß et al., 2009) and assigned a novel FLS-activity in situ to AtL-DOX (Stracke et al., 2009). Both these enzymes and their expres-sion patterns provide an ample explanation for the significantquantities of flavonols produced in fls1-2 and fls1-2/ldox mutantlines of A. thaliana (Stracke et al., 2009; Preuß et al., 2009) andmust be accounted for in the scheme of flavonoid biosynthesis(Figs. 2 and 3). Nevertheless, the activities of FLS3 and LDOX areinsufficient to fully substitute for FLS1 in fls1-mutant Arabidopsisplants. This may reflect structural differences that preclude effi-cient interaction with other flavonoid-specific enzymes in the formof larger protein complexes. The data derived from Arabidopsis onFLS and on the FLS-like activity of LDOX are likely relevant alsofor other plant species and need to be considered, when metabolicengineering studies are conducted in plants or microorganisms.

The broad substrate specificity of multifunctional FLS and LDOXmay be the result of incomplete evolution (Turnbull et al., 2001).Based on previous reports, the Arabidopsis FLS gene family was pre-sumed to have originated from recent gene duplication events(Stracke et al., 2009; Owens et al., 2008a). Such duplications maylead to differentially regulated genes encoding enzymes of variablesubstrate specificities, which develop preferences for the synthesisof selected flavonols and meet the dynamic physiological needs ofthe plants (Owens et al., 2008a). However, isogenes encoding inac-tive polypeptides and expressed independently of flavonoid accu-mulation should no longer be assigned to the FLS gene family.The preservation of these non-functional pseudogenes (AtFLS4and 6) over substantial evolutionary time rather suggests theirformer functional relevance, whereas functional AtFLS3 or AtFLS2and 5, although coding for polypeptides lacking FLS activity, werestill expressed in patterns overlapping with that of AtFLS1 andinteract with other flavonoid genes. Based on the recent resultswe postulate that a pseudogenisation/mutation process currentlyeliminates ‘‘unnecessary” genes and their protein functions. Alter-natively, AtFLS2, 3 and 5 may have acquired another function whileAtFLS3 at least preserved some FLS activity.

Acknowledgments

Funding by EU Project ‘FLAVO’ (FOOD CT-2004-513960) and byDeutsche Forschungsgemeinschaft is grateful acknowledged.

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Stefan Martens has earned a Ph.D. in plant breeding atthe Technical University of Munich Freising-Weihenst-ephan. After two years Post Doc. in the Institute of Prof.Dr. Gert Forkamnn in 2002 he joined the Institute ofPharmaceutical Biology at the Philipps University ofMarburg (AG Prof. Ulrich Matern) where he establishedan independent research group. In 2009 he wasappointed Group Leader for Biochemistry and MolecularBiology of Soft Fruits at the Edmund Mach Foundationin San Michele all ‘Adige (TN, Italy). His scientificinterest is focused on the biosynthesis of secondarymetabolites not only in soft fruits but also in apple,

grapes, herbs and other plant species including breeding, metabolomics, enzy-mology and genomics.

Anja Preuß studied Biochemistry (1999–2004) at Ernst-Moritz-Arndt University of Greifswald. From 2005–2008 she was Ph.D. student at Institute for Pharma-ceutical Biology of Philipps-University Marburg. Since2008 she is Post Doc at Biomolecular Food TechnologyTU-Munich in Freising.

Ulrich Matern, Prof. (em.) holds a pharmacy license and

received his Ph.D. in biochemistry (1972) from FreiburgUniversity, Germany (supervisor Hans Grisebach). Afterpostdoctoral training at the Dept. of Plant Pathology,Montana State University (1976/77), and the Dept. ofChemistry, University of Minnesota (1979), he returnedto Freiburg University to continue studies in plant bio-chemistry and, after a second thesis (Habilitation,1983), was entitled Professor of biochemistry. In 1995,he joined the faculty at Philipps-University Marburg,Germany, as Professor and Director of the Institute ofPharmaceutical Biology till retirement in 2008. His

research focused on the biochemistry of secondary metabolites, ranging fromantibiotics over phytotoxins to flavonoids, coumarins and other phenylpropanoids,which has been published in numerous reports and was acknowledged by the

Phytochemistry Pioneer Award (2008).