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First published online February 19, 2004; 10.1104/pp.103.030221 Plant Physiology 134:979-994 (2004) © 2004 American Society of Plant Biologists Molecular and Biochemical Analysis of Two cDNA Clones Encoding Dihydroflavonol-4-Reductase from Medicago truncatula1Plant Biology Division, The Samuel Roberts Noble Foundation, Inc., 2510 Sam Noble Parkway, Ardmore, Oklahoma 73402 (D.-Y.X., L.A.J., J.D.C., N.L.P.); and National Center for Natural Products Research, School of Pharmacy, University of Mississippi, University, Mississippi 38677 (D.F.)
Dihydroflavonol-4-reductase (DFR; EC1.1.1.219) catalyzes a key step late in the biosynthesis of anthocyanins, condensed tannins (proanthocyanidins), and other flavonoids important to plant survival and human nutrition. Two DFR cDNA clones (MtDFR1 and MtDFR2) were isolated from the model legume Medicago truncatula cv Jemalong. Both clones were functionally expressed in Escherichia coli, confirming that both encode active DFR proteins that readily reduce taxifolin (dihydroquercetin) to leucocyanidin. M. truncatula leaf anthocyanins were shown to be cyanidin-glucoside derivatives, and the seed coat proanthocyanidins are known catechin and epicatechin derivatives, all biosynthesized from leucocyanidin. Despite high amino acid similarity (79% identical), the recombinant DFR proteins exhibited differing pH and temperature profiles and differing relative substrate preferences. Although no pelargonidin derivatives were identified in M. truncatula, MtDFR1 readily reduced dihydrokaempferol, consistent with the presence of an asparagine residue at a location known to determine substrate specificity in other DFRs, whereas MtDFR2 contained an aspartate residue at the same site and was only marginally active on dihydrokaempferol. Both recombinant DFR proteins very efficiently reduced 5-deoxydihydroflavonol substrates fustin and dihydrorobinetin, substances not previously reported as constituents of M. truncatula. Transcript accumulation for both genes was highest in young seeds and flowers, consistent with accumulation of condensed tannins and leucoanthocyanidins in these tissues. MtDFR1 transcript levels in developing leaves closely paralleled leaf anthocyanin accumulation. Overexpression of MtDFR1 in transgenic tobacco (Nicotiana tabacum) resulted in visible increases in anthocyanin accumulation in flowers, whereas MtDFR2 did not. The data reveal unexpected properties and differences in two DFR proteins from a single species.
Flavonoids represent a large group of plant secondary metabolites with diverse biological activities, and the biochemical and genetic investigations of flavonoid biosynthesis have been well documented (Hahlbrock and Grisebach, 1975  ; Harborne, 1988; Stafford, 1990; Holton and Cornish, 1995; Dixon and Steele, 1999; Winkel-Shirley, 2001). Flavonoids are divided into several structural classes, including anthocyanins, which provide flower and leaf colors, catechins, and condensed tannins (CTs; proanthocyanidins), which contribute to resistance to microbes, and other derivatives with diverse roles in plant development and interactions with the environment (Harborne, 1988; Dixon and Paiva, 1995; Paiva, 2000). Several flavonoids are active ingredients in herbal medicines and appear to confer health benefits to humans when consumed regularly. Particular attention has been placed on the anthocyanins, catechins, and proanthocyanidins because of their antioxidant activities and their interactions with proteins (Tanner et al., 1994; Scalbert and Williamson, 2000; Ross and Kasum, 2002; Hou, 2003).
The common precursors in the biosynthesis of all classes of flavonoids are malonyl-CoA and p-coumaroyl-CoA, condensed into chalcone intermediates by the action of chalcone synthase (CHS; Fig. 1). Single genes and multigene families encoding CHS, chalcone isomerase, and flavanone 3-hydroxylase have been studied extensively from many plant species (Holton and Cornish, 1995
Dihydroflavonol 4-reductase (DFR; EC1.1.1.219) is a later key enzyme controlling flux into biosynthetic pathway branches leading to anthocyanins and CTs. DFR preparations from several plants catalyze the reduction of the three dihydroflavonols dihydrokaempferol (DHK), dihydroquercetin (DHQ), and dihydromyricetin (DHM) into leucoanthocyanidins, which are common precursors for anthocyanins and CT synthesis (Fig. 1). DFR proteins in certain species such as petunia (Petunia hybrida) and Cymbidium hybrida do not accept the monohydroxylated DHK and, therefore, cannot produce the corresponding monohydroxylated pelargonidin anthocyanins (Meyer et al., 1987
Anthocyanidin synthase catalyzes the conversion of leucoanthocyanidins to anthocyanidins as the first step of anthocyanin (anthocyanidin-3-O-glucoside) biosynthesis (Nakajima et al., 2001
The genetics and regulation of DFR have been studied extensively in several plant species, where it has usually been found as a single gene or a small gene family. DFR was reported as a single gene in the genomes of barley (Hordeum vulgare), Arabidopsis, tomato (Lycopersicon esculentum), grape (Vitis vinifera), snapdragon (Antirrhinum majus), and rice (Oryza sativa; Kristiansen and Rohde, 1991
Knowledge of the details of DFR biochemistry is very important to understanding aspects of flavonoid biosynthesis, especially how plants regulate CT and anthocyanin biosynthesis and composition and different stereochemical features of flavan-3-ols and related compounds. Early in vitro enzyme assays using cell-free extracts of Gingko biloba and Douglas fir (Pseudotsuga menziesii) showed that DFR converted DHQ to leucocyanidin and DHM to leucodelphinidin (Stafford and Lester, 1982
M. truncatula is a popular model legume species for which many cDNA and genomic sequence databases are being developed (Cook, 1999
Characterization of M. truncatula DFR cDNA Clones
By searching the BLASTX results (Bell et al., 2001
Despite the high DNA sequence similarities (79.6% identical at the nucleotide level in the coding regions when aligned using ClustalW), the two cDNA clones differ in their restriction maps, including a HindIII restriction site present only in MtDFR1 [near the poly(A+) tail] and an EcoRI restriction site present only in MtDFR2 (position 451 in the coding region). Although the nucleotide sequences are highly conserved in the 5' end of the coding regions, the sequence similarity is sufficiently low in 3' regions (data not shown) to allow the design of gene-specific hybridization probes, and these regions were selectively amplified using PCR. Southern hybridization showed that each gene specific probe only hybridized with the corresponding cDNA clone, and no cross hybridization occurred (Fig. 3, A and B, first and second lanes). Each probe hybridized to only a single band in EcoRI- or HindIII-digested M. truncatula genomic DNA, indicating that MtDFR1 and MtDFR2 are each present as a single copy in the M. truncatula cv Jemalong line A-17 genome (Fig. 3, A and B). Although the sizes of the bands detected in the EcoRI-digested DNA are similar with both MtDFR1 and MtDFR2 probes, simultaneous hybridization with both probes confirmed that these were two distinct bands (Fig. 3C). When either the MtDFR1 or MtDFR2 complete coding regions were used as probes, no additional hybridizing bands were observed, indicating that no additional DFR genes are present in this species. The "DFR-like" ANR (Xie et al., 2003
Although several DFR genes have been identified by genetic studies, and DFR cDNAs have been characterized and expressed in plants, little has been published regarding the biochemical characterization of the proteins encoded by these clones, especially multiple DFR enzymes from the same species. To functionally characterize the two M. truncatula DFR enzymes, we subcloned the coding regions of MtDFR1 and MtDFR2 into pSE380 (Brosius, 1989
Enzyme extracted from cultures expressing either MtDFR1 or MtDFR2 protein converted (±)-taxifolin to a product eluting earlier in the HPLC system, whereas this product did not accumulate in reactions carried out with protein extracted from cultures harboring the empty pSE380 expression vector (Fig. 4). The relative retention time (Stafford and Lester, 1984
The activities of MtDFR1 and MtDFR2 were assayed at (±)-taxifolin and (+)-taxifolin concentrations ranging from 0 to 200 µg reaction-1 (0–1.32 mM final assay concentration) in an attempt to estimate the binding constants for the substrates. Plots of reaction velocity versus substrate concentration and corresponding double-reciprocal plots from three independent determinations (data not shown) revealed patterns inconsistent with classical Michaelis-Menton kinetics, in some cases indicating inhibition of DFR activity by high substrate and/or product concentrations. Similar inhibition was observed for VR with structurally similar isoflavanone substrates (Guo et al., 1994
The activity of MtDFR1 and MtDFR2 was also assayed at NADPH concentrations from 0 to 4 mM. MtDFR1 and MtDFR2 exhibited Km values of approximately 0.8 and 1 mM NADPH, respectively. No reduction products were observed when NADH was substituted for NADPH, as was previously reported for some DFR preparations (Stafford, 1990 Dramatic differences were observed in the temperature and pH dependence of MtDFR1 and MtDFR2. MtDFR1 activity exhibited a sharp temperature optimum at 45°C, whereas the activity of MtDFR2 was maximal over the broad range of 30°C to 45°C (Fig. 5A). The optimum pH values for MtDFR1 were from 6.6 to 7.0 (Fig. 5B), whereas the optimum pH values for MtDFR2 were in the range from 5.4 to 6.2 (Fig. 5C).
In addition to taxifolin, DFR proteins present in different plant species also catalyze the reduction of DHK or DHM, although in some cases multiple DFR proteins or isoforms may be present in individual cells or enzyme extracts (Stafford, 1990
Reduction products were observed with all five of the dihydroflavonols tested, irrespective of the number of hydroxyl groups on the B ring, or the presence or absence of the 5-hydroxyl group (Fig. 6B). No reaction was observed with substrates lacking the hydroxyl group at the 3 position or with a double bond present between carbons 2 and 3 (flavone/flavonol derivatives), including quercetin, eriodictyol, kaempferol, and apigenin. LC-MS confirmed that the products of DFR acting on the dihydroflavonols had all gained exactly 2 mass units relative to the substrates (data not shown), consistent with a simple reduction using NADPH. Relative to (±)-taxifolin, (+)-taxifolin was converted more extensively (150%–160%) by both MtDFR1 and MtDFR2, confirming the previously observed stereospecificity of these enzymes. (±)-DHM (trihydroxy B ring) was converted less efficiently than taxifolin by both enzymes [10%–30% of (±)-taxifolin value]. Although (±)-DHK (monohydroxy B ring) was converted much more by MtDFR1 [250% of (±)-taxifolin value], MtDFR2 utilized this substrate much less efficiently than taxifolin [60% of (±)-taxifolin value]. (±)-Fustin, the 5-deoxy analog of (±)-taxifolin, has not been reported previously to be a substrate of DFR but was reduced 5 times faster than (±)-taxifolin by MtDFR2 and approximately 3 times faster by MtDFR1. MtDFR2 showed twice the relative activity on (-)-fustin as did MtDFR1. The reduction of (±)-fustin produced two large product peaks (retention times of 7.5 and 9.0 min), having identical UV profiles and Mrs. When (±)-fustin was used as the substrate, the two product peaks were similar in area, but when (-)-fustin [possessing the same absolute stereochemistry as (+)-taxifolin] was used as the substrate, the later eluting peak contained 90% of the product area. This suggests that the DFRs can accept both isomers of fustin and that the resulting diastereomeric products are resolved in the HPLC system. (±)-DHR, the 5-deoxy analog of (±)-DHM, was reduced as efficiently as (±)-taxifolin by both enzymes. Two product peaks were also observed for (±)-DHR. Reduction of the carbonyl group in fustin or DHR would yield the compounds commonly known as leucofisetinidin and leucorobinetinidin, known constituents of certain leguminous plants (Harborne, 1988
Patterns of anthocyanin accumulation in leaf tissues vary among M. truncatula ecotypes and nearisogenic lines. In M. truncatula cv Jemalong line A-17, a central portion of the upper surface of each leaflet of the trifoliate leaves, is intensely colored by red anthocyanins when the plants are grown under certain conditions (Fig. 7A). The anthocyanin accumulation is generally higher when plants are grown under high light (greater than 200 µE) and low or moderate nitrogen levels, such as those provided by nitrogen fixation after nodulation by Rhizobium meliloti (data not shown). Anthocyanin levels are greatly reduced in leaves grown under high inorganic nitrogen (such as greater than 20 mM nitrate) and low light and approach zero in leaves of plants grown from seedlings that were vernalized at 4°C for 12 to 15 d, a treatment that is commonly used to accelerate seed set. Anthocyanin accumulation is also inducible in the epidermis of hypocotyls of vernalized seedlings by using higher light treatments, as has been demonstrated for many plant species.
The flowers of M. truncatula contain mainly yellow pigments, most likely xanthophylls or carotenoid derivatives (Fig. 7B), although fine red veins of anthocyanins run the length of the standard petal, similar to patterns observed for alfalfa flower coloration (Hanson et al., 1988 A preliminary characterization of the red leaf pigments was carried out, primarily to confirm the anthocyanin nature and to determine the structure of the anthocyanidin core. Red centers of leaflets were dissected away from the surrounding green regions, and each was extracted and analyzed for anthocyanin content by HPLC. Using detection at 515 nm to selectively detect red chromophores, three major anthocyanin peaks were detected at approximately 21 (predominant peak), 15, and 13 min (Fig. 7E). Diode array scans revealed that the three peaks had almost identical UV/visible absorption patterns, with maxima at 280 and 520 to 525 nm (Fig. 7F), consistent with common anthocyanins in acidified solvents.
A small amount of the most abundant anthocyanin (21-min retention time, Fig. 7E) was partially purified and hydrolyzed by heating in acid. Two major product peaks were detected after hydrolysis. One product peak was cyanidin, based on co-elution with a commercial standard (at 24.5 min) and identical UV/visible scans. A second product peak matched the retention time and absorption spectrum of the 15-min peak. Partially purified samples of the 21- and 15-min peaks were subjected to LC/MS and LC/tandem MS analysis in positive ion mode. (The quantities of the purified 13-min peak were insufficient to allow analysis.) The parent ion observed in the 21-min peak was m/z = 667 atomic mass units (amu), with major fragments at m/z = 625, 449, and 287, whereas the 15-min peak had a parent ion of m/z = 625 amu, with major fragments at m/z = 287 and 449. The fragment masses of 287 and 449 are equal to the mass of cyanidin and possible cyanidin-3-O-glucoside ions, respectively. The 42-amu mass difference between the two parent ions is consistent with the loss of an acetyl group as ketene. Several anthocyanins and other flavonoids have been reported to be acetylated, and others that are malonylated in vivo spontaneously decarboxylate during isolation to form the acetate derivatives, especially in the presence of HCl (Harborne, 1988
The tissue specificity of the expression of the two M. truncatula DFR genes was initially examined by RNA gel-blot analysis (Fig. 8A). RNA was extracted from a variety of M. truncatula tissues, especially those accumulating CTs or anthocyanins, known products of DFR activity. A soybean 18S ribosomal RNA probe was used to confirm equal loading and transfer of the total RNA samples (Fig. 8A). The northern analysis results indicated that both MtDFR1 and MtDFR2 transcripts were highly abundant in YS tissues, the source tissue for the original cDNA library, and a known site of CT biosynthesis in alfalfa (Koupai-Abyazani et al., 1993
Because of concerns regarding possible crosshybridization of the two highly similar DFR transcripts during the above analysis, the same sets of tissue RNAs were analyzed by RT-PCR with primer pairs specific to each DFR transcript. The first two lanes of Figure 8B demonstrate the high specificity of the primer pairs for the amplification of their respective cDNA clones. RT-PCR using the MtDFR1 gene-specific primer pair produces one band 413 bp in size (Fig. 8B), whereas the MtDFR2 gene specific primer pair produces one band 682 bp in size (Fig. 8B). RT-PCR using a set of primers common to both DFR transcripts provides an estimate of the combined DFR gene expression in each tissue (Fig. 8B). Because the larger band size would automatically cause the intensity of the MtDFR2 bands to appear brighter than the MtDFR1 bands for the same amount of template, thus possibly biasing the direct visual interpretation of the results, the signals from the ethidium bromide-stained gels were also quantitated and normalized against the strongest signal observed for each primer pair (Fig. 8C) to facilitate a comparison of the RT-PCR results from the two primer pairs. The results of these semiquantitative RT-PCR experiments paralleled those obtained by northern analysis but also revealed subtle differences in the expression patterns of the two DFR genes. For both genes, transcripts were at or near relative maximal levels in the YS mRNA pool, and no transcripts were detected with either primer pair in the two root samples or dark-grown hypocotyls. Although both MtDFR1 and MtDFR2 transcripts were detected in open flowers and flower buds, MtDFR2 had a 60% to 100% higher relative expression level than MtDFR1 in these floral samples, and MtDFR2 was slightly more highly expressed in open flowers than in the YS mRNA pool. In younger, folded leaves, the relative levels of MtDFR2 transcripts were approximately one-half of the maximal levels observed, whereas MtDFR1 transcripts were much lower. MtDFR2 transcripts were twice as abundant as MtDFR1 transcripts in young folded leaves actively accumulating anthocyanins (red spot folded leaves), and MtDFR2 transcripts were 20 times higher than the levels of MtDFR1 transcripts in young folded leaves not accumulating anthocyanins (non-red spot folded leaves). In contrast, MtDFR1 transcripts were at about one-half of the maximal levels, approximately 4 times more abundant than MtDFR2 transcripts, in fully expanded, mature (unfolded) leaves that had accumulated anthocyanins (red spot unfolded leaves). Neither DFR gene was highly expressed in mature leaves containing no anthocyanins (non-red spot unfolded leaves). Both MtDFR1 and MtDFR2 transcript levels were higher in light-grown hypocotyls, particularly after 30 h of light exposure, compared with dark-grown hypocotyls, where no transcripts were detected.
Although it was evident that the two DFR proteins exhibited different kinetic properties in vitro and had slightly different expression patterns in M. truncatula, overexpression in transgenic tobacco plants was used to assess whether or not the two DFR proteins would perform differently in plant cells. Although selected M. truncatula varieties have been reported to be transformed and regenerated, cv Jemalong A-17 is fairly recalcitrant, and the process requires several months. Binary vectors containing the coding regions of the MtDFR1 and MtDFR2 cDNA clones under the control of the CaMV35S promoter were derived from pBI121 (Jefferson et al., 1987
Tobacco (Nicotiana tobacum cv Xanthi), used for transformation, produces pale pink flowers under standard greenhouse conditions. When the transgenic lines were allowed to bloom, several plants harboring the MtDFR1 overexpression construct produced much darker pink flowers than were observed on untransformed plants or on plants harboring either the MtDFR2 vector or the pBI121 control vector (Fig. 9A). The anthocyanins were extracted from the corollas of flowers from each of the three types of transformants and were roughly quantitated spectrophotometrically. Although the anthocyanin content varied from flower to flower on the same plant, at least four plants harboring the MtDFR1 overexpression construct produced flowers containing significantly higher amounts of anthocyanins on a fresh weight basis (Fig. 9B). None of the MtDFR2-transformed lines showed significantly increased levels of anthocyanins relative to pBI121-transformed lines based on visual observations or spectrophotometric analysis. Northern analysis confirmed that several MtDFR2-transformed lines did accumulate MtDFR2 transcripts to the same levels as MtDFR1-transformed lines accumulated MtDFR1 transcripts (data not shown), indicating that the differences in anthocyanin accumulation were because of posttranscriptional events. Together, this indicates that the enzyme encoded by MtDFR1 successfully interacts with the endogenous tobacco anthocyanin biosynthetic pathway enzymes to increase anthocyanin accumulation in vivo, whereas the enzyme encoded by MtDFR2 is unable to increase the flux toward anthocyanin products. These data suggests that the endogenous DFR levels are rate limiting for anthocyanin biosynthesis at some stage in tobacco flower development, and MtDFR1 can increase the pathway flux to anthocyanins at that stage. A similar enhancement of anthocyanin accumulation was observed in vegetative tissues of Forsythia Intermedia cv "Spring Glory" transformed with Antirrhinum majus DFR (Rosati et al., 1997
Two DFR cDNA clones were isolated from a YS EST library indicating the presence of a small DFR gene family in M. truncatula. Both DFR clones were functionally expressed in E. coli, confirming that they both encode active proteins and providing a means to explore substrate specificity of the proteins in vitro. Recently, three other reports of microbial expression of plant DFR clones were published (Martens et al., 2002 ; Peters and Constabel, 2002; Fischer et al., 2003). One reported no success with two E. coli expression systems followed by excellent expression of a G. hybrida DFR cDNA clone in yeast (Martens et al., 2002). Apple (Malus domestica) and pear (Pyrus communis) DFR cDNA clones also were expressed successfully using the same yeast system (Fischer et al., 2003). A third group (Peters and Constabel, 2002) successfully expressed DFR activity from a trembling aspen (Populus tremuloides) cDNA clone in E. coli using a system very similar to the one successfully utilized herein. The variable success in E. coli expression may be a function of subtle variations in protein sequences among the DFR clones. Although the recombinant proteins used in this study were prepared under identical conditions and levels of protein accumulation and initial activity were similar, a dramatic loss of DFR activity was observed after repeated freezing and thawing of bacterial extracts prepared using the MtDFR1 clone, whereas no loss of activity was observed in MtDFR2 extracts (data not shown).
Preliminary chemical characterization of the major foliar red pigments in M. truncatula indicated that these are derivatives of cyanidin diglucosides (3',4'-dihydroxylated anthocyanins). Cyanidin derivatives are widespread through the plant kingdom (Harborne, 1988
MtDFR1 and MtDFR2 provide a natural pair of dihydroflavonol reductases from the same species with slightly differing amino acid sequences to examine the active site features that determine the relative substrate preferences of these isozymes (Fig. 10). Following alignments of several DFR clones isolated from species varying in their anthocyanin hydroxylation patterns, earlier workers had postulated that a 26-amino acid region in DFR controlled the substrate specificity of the reductases, which in turn controls anthocyanin composition in some species (Beld et al., 1989
Our in vitro assays showed that each of the recombinant M. truncatula DFR proteins can catalyze the reduction of all three of the most common dihydroflavonol substrates in nature (DHK, DHQ, and DHM) containing one hydroxyl, two hydroxyls, and three hydroxyls on the B ring, respectively. However, both MtDFR1 and MtDFR2 enzymes most efficiently reduced fustin, a 5-deoxy-dihydroflavonol analog of taxifolin, to leucofisetinidin, and reduced DHR, a 5-deoxy analog of DHM, to leucorobinetinidin. To our knowledge, this is the first report of DFR converting 5-deoxy-dihydroflavonols more efficiently than the 5-hydroxy-dihydroflavonols. DFR proteins from pear and apple were also able to reduce fustin and 5-deoxydihydrokaempferol (garbanzol), but these substrates were less efficiently processed than DHQ, and 5-deoxyflavonols do not naturally occur in these Rosaceae species (Fischer et al., 2003
The two MtDFRs have relative substrate preferences different from those of individual DFRs from other plant species. For example, DFR-A in petunia reduces DHM more readily than taxifolin, unlike either of the M. truncatula DFRs (Forkmann and Ruhnau, 1987
Any differences in the respective biosynthetic roles of the two M. truncatula DFRs are unclear at this time. In many plant species, the expression of a single DFR gene accounts for the observed complex patterns of metabolite accumulation (Gerats et al., 1990
Both MtDFR1 and MtDFR2 appear to be equally expressed in seed and flower leucoanthocyanidin synthesis. Further dissection of the tissues, in parallel with a more extensive phytochemical analysis of the tissues, may reveal the association of the two enzymes with different metabolic pathways or expression in adjacent regions. The M. truncatula flowers used to isolate RNA were not dissected because of their small size and the fused architecture of the floral components, and although the majority of the mass is from petals, a number of other tissues (calyx, anthers, stigmas, and developing ovules) were present, possibly obscuring differences in DFR gene expression. Similarly, RNA was isolated from whole developing seeds, although different parts of the seed or layers of the seed coat may accumulate different metabolites. In Arabidopsis, CTs and anthocyanins accumulate in a single endothelium cell layer in the developing seed coat (Devic et al., 1999
Plant Materials
To produce immature seeds for cDNA library construction, Medicago truncatula cv Jemalong line A17 (Cook, 1999
Total RNA was extracted from immature seeds using a commercial RNA isolation kit (Plant RNAeasy Midi kit, Qiagen, Valencia, CA) with the alternative guanidine hydrochloride buffer recommended for high starch tissues. mRNA was isolated from 1 mg of total RNA using immobilized oligo(dT) [Oligotex poly(A+) mRNA purification kit, Qiagen]. A primary cDNA library (YS library) containing over 6 x 106 phage was constructed following the manufacturer's instructions using the UniZapXR cDNA library kit (Stratagene, La Jolla, CA).
A portion of the YS primary cDNA library (10,000 plaque forming units) was mass excised using Stratagene's protocol. The resulting ExAssist phagemids were used to transform SOLR cells, and transformants containing pBluescript SK- plasmids with cDNA inserts were selected on Luria-Bertani agar plates supplemented with 100 µg mL-1 ampicillin. Individual colonies were randomly selected and used to inoculate 1.5 mL of Terrific Broth medium with 100 µg mL-1 ampicillin in 96-well 2-mL capacity plates. After 24 h of growth with shaking at 37°C, plasmids were isolated in a 96-well format using a modified alkaline lysis protocol (Roe et al., 1996
MtDFR1
MtDFR2
The constructs containing MtDFR1 and MtDFR2 ORFs in the expression vector pSE380 were used to transform E. coli BL21-Gold competent cells (Stratagene). Single colonies harboring expression constructs pSE380-MtDFR1, pSE380-MtDFR2, or pSE380 (vector control) were cultured in Luria-Bertani medium with 100 µg mL-1 ampicillin, induced with 1 mM isopropyl- The initial standard enzyme assay conditions included incubation at 30°C for 30 min in 0.5-mL total volume containing 370 µL of 100 mM Tris-HCl buffer (pH 7.0), 70 µL of protein extract (1.5 µg protein µL-1), 50 µL of 10 mM NADPH, and 10 µL of (±)-taxifolin (10 µg µL-1 methanol) in 1.5-mL centrifuge tubes. Enzyme reactions were stopped by adding 1 mL of ethyl acetate and vortexing. Samples were centrifuged for 2 min at 10,000g, and 0.75 mL of ethyl acetate extract was removed to a new tube and evaporated with a stream of nitrogen gas. Residues were dissolved in 150 µL of methanol and used directly for HPLC analysis (20-µL injection). The area of the leucoanthocyanidin product peaks was integrated using 32-Karat software (Beckman Instruments, Fullerton, CA). (±)-Taxifolin (Sigma, St. Louis) was used as the initial substrate to test the effects of substrate concentration, assay pH, temperature, duration, and NADPH concentration on enzyme activity. Studies of pH dependence were conducted at 30°C for 30 min in a 0.5-mL volume consisting of 370 µL of 50 mM citrate/phosphate buffer (pH 4.6–7.0) or 100 mM Tris-HCl buffer (pH 7.0–8.8), 70 µL of protein extract, 50 µL of 10 mM NADPH, and 10 µL of 33 mM (±)-taxifolin.
Dihydroflavonols including (±)-taxifolin (DHQ; Sigma), (+)-taxifolin, (-)-fustin (Dr. Daneel Ferreira), racemic (±)-fustin (Indofine, Hillsborough, NJ), DHK, DHM, and DHR (Apin, Abingdon, Oxon, UK) were dissolved in methanol at 10 mg mL-1. The flavonols, flavanones, or flavones, i.e. kaempferol, quercetin, eriodictyol, and apigenin, were dissolved in methanol at 10, 10, 2.5, and 1 mg mL-1, respectively. Reactions for MtDFR1 were conducted at 30°C for 30 min using 370 µL of 100 mM Tris-HCl buffer (pH 7.0), 70 µL of MtDFR1 enzyme extract, 10 µL of substrate, and 50 µL of 20 mM NADPH, whereas reactions for MtDFR2 substituted 370 µL of 50 mM citrate/phosphate buffer (pH 6.2), so that enzymes were assayed at their respective optimum pH values.
Compounds were resolved on a C18-silica HPLC column (5 µm, narrow pore, 4.6 x 250 mm, Bakerbond, J.T. Baker, Phillipsburg, NJ) with detection at 280 nm, the maximum absorbance wavelength for most of the substrates and products. UV spectra were recorded with a UV diode array detector (model 168, Beckman Instruments). For routine HPLC quantitation, the solvents were 1% (v/v) H3PO4 in water (A) and methanol (B), and separation methods were developed by adapting the systems reported previously (Stafford and Lester, 1984
To prepare large quantities of anthocyanins for preliminary structural characterization, fresh leaves and stems grown under high-light conditions were extracted for 12 to 24 h with neutral acetone to remove the bulk of the chlorophyll, lipids, and other substances without extracting the majority of the anthocyanins (data not shown). The red anthocyanin sectors were still clearly visible against the decolorized portions of the leaves. The acetone was decanted and replaced with acidic methanol (1 mL HCl L MeOH-1), and anthocyanins were extracted for 1 to 2 d at room temperature with shaking in the dark. The MeOH was removed by rotary evaporation, and the red residue was redissolved in a small volume of MeOH. Anthocyanins were quantitated using HPLC (4.6- x 250-mm Bakerbond C18-silica column, as above; J.T. Baker) with detection at 515 nm and normalized against the fresh weight values. The solvents were 1% (v/v) H3PO4 in water (A) and CH3CN (B) with column equilibration at 5% (v/v) B and a flow rate of 0.8 mL min-1. Following injection (20 µL), a linear gradient from 5% to 40% (v/v) B in 35 min was initiated. The anthocyanins were partially purified by diluting the crude solution with water to less than 10% (v/v) MeOH and adsorbing to a disposable C-18 cartridge (Discovery DSC-18 SPE, Supelco, St. Louis). After rinsing with 10% (v/v) MeOH, 30% (v/v) CH3CN was used to elute the anthocyanins. The eluate was concentrated and subjected to semipreparative HPLC (Econosil C-18, 22- x 250-mm column, Alltech, Deerfield, IL) using a gradient and solvents identical to the analytical method but with a 20 mL min-1 flow rate. Fractions containing anthocyanins were pooled and CH3CN was removed using a stream of nitrogen gas. The remaining aqueous solution was extracted with EtOAc, then the EtOAc phase back-extracted with water to remove traces of H3PO4, and EtOAc was removed by evaporation. The residue was dissolved in MeOH and subjected to LC-MS analysis. A portion of each purified anthocyanin sample was hydrolyzed by heating in 10% (v/v) HCl at 95°C for 4 h to release the anthocyanidin core.
To compare the anthocyanin content of small tissue samples from M. truncatula (100–300 mg fresh weight) or tobacco flowers, tissues were extracted with acidic MeOH (5 mL) by shaking at room temperature in the dark for 48 h. For M. truncatula tissues, extracts were dried and redissolved in 200 µL of MeOH and subjected to HPLC analysis as above. For tobacco flowers, total anthocyanins were quantitated by spectrophotometry at 528 nm (Xie et al., 2003
For the (±)-taxifolin product, 200-fold scaled-up reactions (100-mL volume) were performed at 30°C for 3 h in 50 mM citrate/phosphate buffer (pH 6.2) with 14 mL of MtDFR2 enzyme extract, 20 mg of (±)-taxifolin, and 10 mL of 20 mM NADPH. The reaction mixture was extracted six times with 100-mL aliquots of ethyl acetate. The pooled extracts were concentrated, and the residue was dissolved in 3 mL of methanol for further purification or LC-MS analysis. For (-)-fustin, (±)-fustin, and other substrates, similar large-scale reactions (up to 6-mL reaction volume) and HPLC and LC-MS analyses were carried out. For LC-MS analysis of the DFR products, an HP1100 series HPLC system (Agilent, Palo Alto, CA) with UV/Vis diode array detection followed by a Esquire-LC00098 mass spectrometer system (Bruker, Billerica, MA) was used. H3PO4 was omitted from the HPLC solvents without substantially altering the retention times of the DFR products. The total ion chromatograms were obtained using MS detection with negative ionization and scans of peaks were stored from m/z of 50 to 1,000 amu. For LC-MS analysis of the extracted and hydrolyzed anthocyanins, the anthocyanin quantitation method was modified by substituting volatile 1% (v/v) acetic acid for 1% (v/v) H3PO4. Total ion chromatograms were obtained using MS detection with positive ionization as is recommended for anthocyanins, and scans of peaks were stored from m/z of 50 to 2200 amu.
Genomic DNA of M. truncatula was extracted from young leaves using the Plant DNAeasy Maxi kit (Qiagen). DNA (10 µg) was digested overnight with HindIII and EcoRI and resolved by electrophoresis in a 0.8% (w/v) agarose gel in Tris-acetic acid-EDTA buffer. Blotting to GeneScreenPlus (DuPont, Wilmington, DE) nylon membranes and hybridization at 65°C were performed as previously described (Church and Gilbert, 1984
Gene-specific probes for MtDFR2 or MtDFR1 were designed from the DNA sequences of the 3' portions of the ORFs. The 225-bp DNA fragment for the MtDFR1-specific probe was amplified using the PCR primer pairs of 5'GCACCAATAAGTAATGGTGTCAC3' and 5'CGTGTAGCTCTCAATAAGG3', whereas the 173-bp DNA fragment for the MtDFR2 specific probe was amplified using 5'CCTAAAGTTACAGAGACTCCGG3' and 5'CAGACAACGTGACCCATAAAC3'. The PCR products were labeled with
To assess MtDFR1 and MtDFR2 expression patterns in M. truncatula, total RNA was extracted from different organs including roots, hypocotyls, leaves, flower buds, open flowers, and YSs using Tri-Reagent (Molecular Research Center, Cincinnati, OH). Root samples were harvested from plants grown in perlite wetted with a nutrient solution (Hipskind and Paiva, 2000
Total RNA samples (15 µg) were electrophoresed in 1.5% (w/v) agarose MOPS-formaldehyde gels and transferred to GeneScreen Plus (NEN/DuPont, Wilmington, DE) membranes as described by Sambrook et al. (1989
One primer pair was designed from conserved regions in MtDFR1 and MtDFR2 to amplify a 329-bp PCR product from both cDNA clones. The primers were 5'CCTATGGATTTTGAGTCCAAGGACCC3' (corresponding to bases 315–340 of MtDFR1 and 259–284 of MtDFR2) and 5'GGACCAACAACAAGAGGTGG3' (corresponding to bases 643–624 of MtDFR1 and 587–568 of MtDFR2). MtDFR1 and MtDFR2 gene-specific primer pairs were designed based on alignments of the cDNA sequences. The MtDFR1 primers were 5'CCTCAAGGCCAAAACTGTCC3' (bases 398–417) and 5'GATACCTCCCTTCTACTTCC3' (bases 810–791), yielding a 413-bp PCR product. The MtDFR2 primers were 5'GACTTATGGAGCGCGGCTACACA3' (bases 72–93) and 5'GTATCTCCCATGTGCTTTAGGG3' (bases 753–732), yielding a 682-bp PCR product. Total RNA (1 µg) was used for first strand cDNA synthesis, and 5 µL of the first strand cDNA was used for each PCR reaction (50 µL) using the Advantage RT-for-PCR kit (CLONTECH Laboratories, Palo Alto, CA). For the conserved primers for both MtDFR1 and MtDFR2, PCR conditions were 94°C for 2 min, 30 cycles of 94°C for 30 s, 60°C for 30 s, and 72°C for 1 min, followed by 72°C for 10 min. Identical conditions were used for MtDFR1 gene specific primers, whereas the annealing temperature was increased to 65°C for MtDFR2 gene-specific primers. PCR products (20 µL) were resolved on 0.8% (w/v) agarose Tris-acetic acid-EDTA gels, visualized with ethidium bromide, and band intensities were quantitated using ImageQuant software (Molecular Dynamics). Band intensities were normalized for each primer pair by expressing the results relative to the highest band intensity observed in each experiment.
Using modified PCR primers, a BamHI site was introduced immediately upstream of the start codons in MtDFR1 and MtDFR2, and a SacI site was introduced approximately 40 bp downstream of the stop codons. These modified coding regions from MtDFR1 and MtDFR2 were subcloned into the BamHI and SacI sites of the binary vector pBI121 (Jefferson et al., 1987
We would like to thank Dr. Bettina Deavours (The Samuel Roberts Noble Foundation, Ardmore, OK) for critical reading of the manuscript. We would also like to thank David Huhman (The Samuel Roberts Noble Foundation, Ardmore, OK) for expert assistance in acquiring LC-MS data. Received July 15, 2003; returned for revision August 17, 2003; accepted October 28, 2003.
Article, publication date, and citation information can be found at http://www.plantphysiol.org/cgi/doi/10.1104/pp.103.030221.
1 This work was supported by The Samuel Roberts Noble Foundation (Ardmore, OK) and by Forage Genetics (West Salem, WI).
2 Present address: Department of Physical Sciences, Southeastern Oklahoma State University, 1405 N. 4th Avenue, Durant, OK 74701. * Corresponding author; e-mail nlpaiva{at}alum.mit.edu; fax 580–745–7494.
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-D-thio-galactoside-induced cells were prepared and assayed for enzyme activity using methods previously used for isoflavone reductase (Paiva et al., 1991
0.05; three lines passed at P 
10,000g at 4°C. 
-32P-dCTP using random primer labeling (Prime-a-Gene, Promega, Madison, WI) and used as probes, although either can cross-hybridize to some extent to both DFR genes. 
