HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

J. Biol. Chem., Vol. 281, Issue 52, 40076-40088, December 29, 2006

This Article Abstract Full Text (PDF) All Versions of this Article: 281/52/40076    most recent M605900200v1 Submit a Letter to Editor Alert me when this article is cited Alert me when eLetters are posted Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Youn, B. Articles by Kang, C. Search for Related Content PubMed PubMed Citation Articles by Youn, B. Articles by Kang, C. Social Bookmarking                      What's this?

Mechanistic and Structural Studies of Apoform, Binary, and Ternary Complexes of the Arabidopsis Alkenal Double Bond Reductase At5g16970^*

Buhyun Youn, Sung-Jin Kim, Syed G. A. Moinuddin, Choonseok Lee, Diana L. Bedgar, Athena R. Harper, Laurence B. Davin, Norman G. Lewis¹, and ChulHee Kang²

From the School of Molecular Biosciences, Washington State University, Pullman, Washington 99164-4660 and the Institute of Biological Chemistry, Washington State University, Pullman, Washington 99164-6340

Received for publication, June 20, 2006 , and in revised form, August 25, 2006.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION CONCLUDING REMARKS REFERENCES

 
In this study, we determined the crystal structures of the apoform, binary, and ternary complexes of the Arabidopsis alkenal double bond reductase encoded by At5g16970. This protein, one of 11 homologues in Arabidopsis thaliana, is most closely related to the Pinus taeda phenylpropenal double bond reductase, involved in, for example, heartwood formation. Both enzymes also have essential roles in plant defense, and can function by catalyzing the reduction of the 7-8-double bond of phenylpropanal substrates, such as p-coumaryl and coniferyl aldehydes in vitro. At5g16970 is also capable of reducing toxic substrates with the same alkenal functionality, such as 4-hydroxy-(2E)-nonenal. The overall fold of At5g16970 is similar to that of the zinc-independent medium chain dehydrogenase/reductase superfamily, the members of which have two domains and are dimeric in nature, i.e. in contrast to their original classification as being zinc-containing oxidoreductases. As provisionally anticipated from the kinetic data, the shape of the binding pocket can readily accommodate p-coumaryl aldehyde, coniferyl aldehyde, 4-hydroxy-(2E)-nonenal, and 2-alkenals. However, the enzyme kinetic data among these potential substrates differ, favoring p-coumaryl aldehyde. Tyr-260 is provisionally proposed to function as a general acid/base for hydride transfer. A catalytic mechanism for this reduction, and its applicability to related important detoxification mammalian proteins, is also proposed.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION CONCLUDING REMARKS REFERENCES

 
The phenylpropanoid pathway, which is essentially restricted to vascular plants, is of increasing significance and general importance because of the extensive medicinal/health protecting properties of many of its derivatives. For example, this pathway results in formation of the potent antiviral agent, podophyllotoxin (1), which also serves as a semi-synthetic source of the highly successful cancer chemotherapeutic treatments, teniposide, etoposide, and Etopophos® (2-4). Other examples include: matairesinol and secoisolariciresinol, which are some of the dietary sources of the "mammalian" lignans, enterolactone/enterodiol, these being protective against the onset of various malignancies (5); the nordihydroguaiaretic acid derivatives, which show considerable promise against refractory cancers of the neck and the head (6); the potent anti-oxidant chlorogenic acid, which has well documented anticancer properties (7, 8), as well as reducing the risk of cardiovascular disease (9); and the monomeric allyl/propenyl phenols such as chavicol and eugenol, which have well known antibacterial and analgesic properties (10). Other medicinally important phenylpropanoid (acetate) pathway metabolites include dihydroconiferyl alcohol (1; Fig. 1), a potentially useful anti-inflammatory agent (11), and phlorizin (5), which shows considerable promise for treatment of diabetes mellitus, obesity, and stress hyperglycemia (12).

Our recent studies have been directed toward establishing the various biochemical pathways associated with formation of such metabolites, including defining the catalytic mechanisms and high-resolution structures of the participating pathway enzymes. Examples of these include pinoresinol/lariciresinol reductases (13, 14), secoisolariciresinol dehydrogenase (15-17), phenylcoumaran benzylic ether reductase (14, 18), isoflavone reductase (14), cinnamyl alcohol dehydrogenases (19, 20), and chavicol/p-anol and eugenol/isoeugenol synthases (21, 22), as well as dirigent proteins (in the presence of auxiliary oxidative capacity) (23-25). These studies are part of broader goals aimed toward (i) systematically engineering selected enzyme substrate binding pockets in terms of potentially modifying them to be more specific for a particular metabolite/metabolic pathway and (ii) better understanding how these pathways in plants have evolved.

The objective of the study described herein was to determine the mechanism and structures of the enzyme involved in formation of medicinally promising dihydrophenylpropanoid derivatives, such as dihydroconiferyl alcohol (1). In the Pinaceae, e.g. loblolly pine (Pinus taeda), various dihydrophenylpropanoids accumulate as heartwood-forming constituents, which contribute to the color, quality, and durability of its woody tissues; these can have either propanol, propionic acid or propanaldehyde side chains (e.g. p-dihydrocoumaric (4)/dihydroferulic(3)acidsandp-dihydrocoumaryl(2)/dihydroconiferyl (1) alcohols (Fig. 1) in Picea glauca (26)). Interestingly, their amounts (e.g. 1 and 2) are known to increase in the galls of P. glauca upon aphid attack (e.g. by Adelges abietis) (26) in further support of roles in plant defense.

View larger version (14K): [in this window] [in a new window]   FIGURE 1. Dihydrophenylpropanoids (structures 1-4) and phlorizin (structure 5).

View larger version (18K): [in this window] [in a new window]   FIGURE 2. Reactions catalyzed by NADPH-dependent alkenal/alkenone/, double bond reductases in vitro. Reduction is shown of: A, p-coumaryl (6) and coniferyl (7) aldehydes by PtPPDBR and AtDBR1; B, pulegone (10) by PulR in Mentha piperita; C, various , unsaturated ketones in prostaglandin biosynthesis (e.g. 13) by guinea pig leukotriene B4 12-HD/PGR; D, 4-HNE (15) by AtDBR1.

View larger version (87K): [in this window] [in a new window]   FIGURE 3. Amino acid sequence alignment of AtDBR1 with other oxidoreductases. PtPPDBR from P. taeda, PulR from M. piperita, AOR from rat (Rattus norvegicus) liver, 12-HD/PGR from guinea pig (C. porcellus, PDB code 1V3V), and putative NADPH-dependent oxidoreductase from M. musculus (1VJ1). The nucleotide-binding domain is indicated by a thin dotted line; the conserved residues in the AXXGXXG motif and the conserved Tyr residues (Tyr-260 for AtDBR1) are highlighted in red and yellow, respectively. Secondary structural elements of AtDBR1 are highlighted in colored bars (orange for the -strands and green for the -helices).

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION CONCLUDING REMARKS REFERENCES

 
Materials—All solvents and chemicals used were reagent or HPLC grade unless otherwise stated. Chemical reactions were carried out under anhydrous conditions in a N₂ atmosphere using dry, freshly distilled solvents. Silica gel thin-layer chromatography (TLC) and silica gel column chromatography employed Partisil® PK5F (Silica gel 150 Å, 1 mm thickness), AL SIL G/UV₂₅₄ (Whatman, 20 x 20 cm, 0.25 mm), and silica gel 60 (EM Science), respectively, with UV absorption and heating with phosphomolybdic acid reagent (2.5% in H₂O) used for TLC plate visualization. NMR spectra were recorded on a Varian Mercury-Vx 300 MHz spectrometer operating at 300.1 (¹H) and at 75.5 (¹³C), with chemical shifts given in ppm relative to tetramethylsilane and J values in Hz, respectively.

HPLC analyses employed an Alliance 2695 HPLC system equipped with a diode array detector (Waters, Milford, MA), and GC-MS analyses utilized an HP 5973 MS detector (electron impact mode, 70 eV), an HP 6890 GC system, and a 7673 series injector equipped with a RESTEK-5Sil-MS (30 m x 0.25 mm x 0.25 µm) column. The carrier gas was helium with an initial flow of 1.4 ml min^-1 at a pressure of 11.65 p.s.i., with samples analyzed using the split injection mode and an injector temperature of 250 °C (split ratio, 10.1:1; split flow, 14.0 ml min^-1). The GC temperature program was initiated at 70 °C for 1 min, increasing to 170 °C at a rate of 8 °C/min, and held at 170 °C for 10 min. The mass range was scanned from m/z 50 to 800. HPLC electrospray ionization mass spectrometric analyses (LC-ESI-MS) were recorded on a Waters 2690 Alliance/Finnigan MAT LCQ, whereas electron impact mass spectra (EI-MS) were acquired on a Waters Integrity^TM HPLC-MS system at an ionization voltage of 70 eV.

Chemical Syntheses—(E)-p-Coumaryl aldehyde (6) was synthesized exactly as described in Kim et al. (19), and coniferyl aldehyde (7) was from Aldrich.

For dihydroconiferyl aldehyde (9), to a solution of 4-O-tert-butyldimethylsilyl-(E)-coniferyl aldehyde (19) (500 mg, 1.71 mmol) in dry MeOH (10 ml) was added 10% palladium on activated charcoal (70 mg) with the resulting suspension stirred at room temperature under H₂ for 6-8 h. The reaction mixture was then filtered, with the filtrate dried in vacuo. To the resulting 4-O-tert-butyldimethylsilyl dihydroconiferyl aldehyde derivative (450 mg, 1.53 mmol) dissolved in dry tetrahydrofuran (10 ml) under N₂ at 0 °C was next added a solution of tetrabutylammonium fluoride (TBAF) (1.0 M in tetrahydrofuran, 1.8 ml, 1.8 mmol) with the whole then allowed to stir for 45 min. The reaction mixture was quenched with a saturated NH₄Cl solution (30 ml) and extracted with dry diethyl ether (50 ml x 2), with the resulting combined organic solubles washed successively with water (30 ml x 2). The organic solubles were then dried (Na₂SO₄) and evaporated to dryness in vacuo. The residue so obtained was subjected to silica gel column chromatography (eluent: CHCl₃/MeOH, 9:1), and then preparative silica gel TLC eluted with CHCl₃/MeOH (9:1) to yield dihydroconiferyl aldehyde (9; 190 mg, 1.05 mmol, 70% yield). ¹H NMR (300 MHz, CDCl₃): 2.75 (2H, m, H-8), 2.89 (2H, m, H-7), 3.87 (3H, s, OMe), 6.66 (1H, dd, J = 8.7, 2.3 Hz, H-6), 6.70 (1H, d, J = 2.3 Hz, H-2), 6.84 (1H, d, J = 8.7 Hz, H-5), 9.81 (1H, t, J = 1.67 Hz, CHO). EI-MS (70 eV) m/z 180 [M⁺] (50), 137 (100).

p-Dihydrocoumaryl aldehyde (8) was synthesized exactly as described above for dihydroconiferyl aldehyde (9) by palladium on activated charcoal hydrogenation of 4-O-tert-butyldimethylsilyl-(E)-p-coumaryl aldehyde (19) (100 mg, 0.38 mmol) followed by deprotection with TBAF to afford p-dihydrocoumaryl aldehyde (8, 23.6 mg, 0.15 mmol, 52% yield). ESI-MS m/z 149.2 [M-H]^-. The NMR spectra of p-dihydrocoumaryl aldehyde (8) were in close agreement with reported data (38, 39).

4-HNE (15) was synthesized exactly as described by Gardner et al. (40) except for the purification/isolation steps. 4-HNE (15) was purified by silica gel preparative TLC using hexanes/diethyl ether (6:4) as eluants with detection by UV absorption and visualization of a bluish spot after staining and heating with phosphomolybdic acid reagent (2.5% (w/v) in H₂O). The band corresponding to 4-HNE (15) was excised, eluted with acetone/diethyl ether (1:2, 30 ml x 2) with the combined organic solubles filtered and evaporated to dryness under N₂ atmosphere by adding MeOH (1 ml) to avoid loss of 4-HNE (15) because of its volatility, and stored at -80 °C. 4-HNE (15) was obtained in 60-65% yield (46.7 mg, 0.3 mmol). GC-MS was employed next to determine the purity as well as the fragmentation pattern of 4-HNE (15) by preparing its trimethylsilyloxy derivative (41) using bis(trimethylsilyl)trifluoroacetamidechlorotrimethylsilane (99:1 Supelco, 50 µl) and pyridine (20 µl). The following fragments were observed at a retention time of 13.10 min (90% pure) from the GC-MS analyses: m/z 228 [M^·+], 213 [M⁺ - CH₃], 199 [M⁺ - CHO], 157 , 129 , 73 . The NMR spectra for HNE (15) were in close agreement with reported data (40, 42).

4-HNA (16) was synthesized via the diisobutylaluminium hydride reduction of -nonanoic lactone (43). The latter was synthesized by the Knoevenagel reaction of malonic acid with heptanal and lactonization with 85% sulfuric acid at 80 °C for 1 h (44, 45). The lactone (500 mg, 3.2 mmol) so formed was then reduced with a 1 M solution of diisobutylaluminium hydride in toluene (4.8 ml, 4.8 mmol) at -78 °C for 2 h exactly as described by Bloch and Gilbert (43). 4-HNA (16) was purified by silica gel preparative TLC using hexanes/diethyl ether (1:1) as eluants, with the band corresponding to 4-HNA (16) excised and eluted with acetone/diethyl ether (1:1, 30 ml x 2). The organic solubles were then combined, filtered, and dried under N₂ atmosphere to afford 4-HNA (16, 420 mg, 2.65 mmol, 80-85% yield). GC-MS analyses of 4-HNA (16) showed the following fragments at retention time = 11.57 and 11.64 min corresponding to both S-(16a) and R-(16b) isomers (95% pure): m/z 230 [M^·+], 215 [M⁺ - CH₃], 159 , 73 . The NMR spectrum for 4-HNA (16) was in close agreement with reported data (46, 47).

Expression and Purification of AtDBR1—AtDBR1 (At5g16970), cloned into an Invitrogen pTrcHis2-TOPO® TA vector, was transformed into TOP10 Escherichia coli cells. Expression of AtDBR1 was induced by addition of isopropyl -D-thiogalactopyranoside to a 0.5 mM final concentration at mid-log phase (A₆₀₀ = 0.5). The induced cell suspension cultures were grown for 12 h at 25 °C with shaking at 250 rpm, with the cells subsequently harvested by centrifugation (3,000 x g for 20 min). The AtDBR1-derived pellet was suspended in lysis buffer (50 mM NaH₂PO₄, 300 mM NaCl, 10 mM imidazole, pH 8.0, 10% glycerol), sonicated (5 x 10 s, model 450 Sonifier®, Branson Ultrasonics), and centrifuged (20,000 x g for 40 min). After centrifugation, the supernatant was incubated with nickel-nitrilotriacetic acid agarose (Qiagen, Hilden, Germany) for 1 h in an overhead shaker at 4 °C. After washing with 10 column volumes of wash buffer (50 mM NaH₂PO₄, 300 mM NaCl, 20 mM imidazole, pH 8.0), the fusion protein was eluted stepwise with 50 mM NaH₂PO₄, 300 mM NaCl, 100-300 mM imidazole, pH 8.0. Thereafter, the AtDBR1-enriched fraction was subjected to anion exchange column chromatography (Self Pack^TM POROS® 10HQ, Applied Biosystems) pre-equilibrated in Buffer A (50 mM Tris-HCl, pH 8.0, containing EDTA (1 mM) and dithiothreitol (1 mM)) at a flow rate of 3 ml min^-1. Recombinant AtDBR1 was eluted using a NaCl step gradient (to 0.1, 0.2, 0.5, and 2.0 M, 50 ml each) with the corresponding fractions of interest (eluting at 0.1 M NaCl) desalted and concentrated into Buffer B (20 mM Tris-HCl, pH 7.5) by ultrafiltration in an Amicon 8050 cell with a 10-kDa cutoff membrane (Millipore). This fraction was applied to a MonoQ^TM GL10/100 anion exchange column (Amersham Biosciences) equilibrated in Buffer B at a flow rate of 2 ml min^-1 and eluted with a NaCl step gradient (0.05, 0.1, 0.2, 0.4, and 2.0 M; 20 ml each); the catalytically active AtDBR1 fraction eluted at 0.05 M NaCl. The AtDBR1 so obtained was concentrated with a final purity of >99% as estimated by SDS-PAGE (Coomassie Blue staining).

Kinetic Parameter Determinations—When p-coumaryl (6) and coniferyl (7) aldehydes were used as substrates, initial velocity kinetics were determined as follows. Assays consisted of MES buffer (100 mM, pH 6.25, 100 µl), 130 µl (3-8 µg) of AtDBR1 purified as described in Kim et al. (19) in 20 mM Tris-HCl, pH 7.5, aldehydes 6 or 7 (10-0.1 mM,10 µl), and NADPH (25 mM, 10 µl) in a total volume of 250 µl. Enzymatic reactions were initiated by the addition of AtDBR1, and after a 10-min incubation at 30 °C, they were stopped by the addition of glacial acetic acid (10 µl). An aliquot (80 µl) of each assay mixture was next subjected to reversed-phase HPLC analysis on a Symmetry Shield RP₈ column (Waters; 150 x 3.9 mm inner diameter, 5 µm particle size) with the following elution conditions at a flow rate of 1 ml min^-1 and detection at 280 nm: the column was pre-equilibrated in a 5:95 ratio of CH₃CN (solvent A) and 3% AcOH in H₂O (solvent B). After introduction of the sample, this composition was held for 1 min, after which a linear gradient to A:B (40:60) over 39 min was carried out followed by a linear gradient to A:B (5:95) in 5 min, this being held for 1 min. The amounts of products 8 and 9 formed were determined using pre-established calibration curves.

Assays with 4-HNE (15) as substrate were carried out as described above but with 0.5 µg of AtDBR1 and an incubation time of 2 min. After addition of glacial acetic acid (10 µl), 4-hydroxybenzaldehyde (10 mM, 5 µl) was added as an internal standard, with the mixture extracted with diethyl ether (1 ml x 3). After vortexing, the diethyl ether layers were removed and combined, the ether solubles were dried (over Na₂SO₄), and the volume was reduced to 100 µl under N₂, at which time 1,4-dioxane was added (100 µl). The resulting solution was transferred to a GC vial with the volume reduced to 100 µl under N₂, and the trimethylsilyloxy derivative was prepared. The samples were next subjected to GC-MS analyses as described under "Chemical Syntheses."

Molecular Mass Determination—Size exclusion chromatography/multiangle light scattering and dynamic light scattering were performed as described previously by Youn et al. (16, 20), with light scattering data acquired through accumulation (3 times) of 10 scans.

Crystallization of AtDBR1—For crystallization, a solution of purified AtDBR1 (48 mg ml^-1) in 20 mM Tris-HCl, pH 8.0, containing 1 mM EDTA and 1 mM dithiothreitol was prepared. Crystallization trials were performed using the hanging drop vapor diffusion method at two temperatures (277 and 293 K). ApoAtDBR1 crystals were obtained by mixing the above protein solution (1.5 µl) with an equal volume of reservoir solution containing 20% (w/v) polyethylene glycol 3350 and 0.2 M potassium chloride. Crystals usually appeared after 5 days, and larger crystals with dimensions of 0.3 x 0.5 x 0.8 mm were obtained after 2 weeks. Although these crystals were fairly large, they were hollow and had a diffraction limit of 3.5 Å. Crystals were stabilized, and the diffraction limit was increased (up to 2.5 Å) by slowly adding concentrated buffer solution to the drops in which crystals were grown. For most crystals, the final buffer composition was 30% (w/v) polyethylene glycol 3350, 0.3 M potassium chloride. The crystals of AtDBR1 belong to the orthorhombic space group, P2₁2₁2₁ (a = 49.46, b = 122.98, c = 148.00 Å), with two molecules in an asymmetric unit (Table 2). The binary complex (AtDBR1-NADP⁺) and the ternary complex (AtDBR1-NADP⁺-p-coumaryl aldehyde (6): ternary I) crystals were also produced under the same conditions except for the addition of 5 mM NADP⁺ and 5 mM p-coumaryl aldehyde (6) into the reservoir solution, respectively. Crystals of the other ternary complex (AtDBR1-NADP⁺-4-HNE (15): ternary II) were produced by soaking the binary complex crystal in a solution of 1 mM 4-HNE (15) (Alexis Biochemicals Inc.). All binary and ternary complexes crystallized in an orthorhombic space group, P2₁2₁2₁, with corresponding unit cells of a = 49.15, b = 122.66, c = 147.94 (binary), a = 49.04, b = 122.54, c = 147.65 Å (ternary I), and a = 49.08, b = 122.45, c = 147.84 (ternary II), respectively. Data for the apoform (2.5 Å resolution), the binary complex (2.8 Å), and both of the ternary complexes (2.8 Å) were collected from the Berkeley Advanced Light Source (ALS, beam line 8.2.1/apoform and ternary II), Chicago Advanced Photon Source (APS, beam line NE-CAT/8-BMD/binary complex) and Rigaku Saturn 92/MicroMax-007 (Washington State University/ternary I) at a temperature of 100 K. Before freezing, the corresponding crystals were soaked for 5 min in cryoprotectant (25% glycerol in each reservoir solution).

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION CONCLUDING REMARKS REFERENCES

 
It was first instructive to obtain the needed kinetic data for the recombinant (His-tagged) AtDBR1 using as substrates p-coumaryl aldehyde (6), coniferyl aldehyde (7), and 4-HNE (15), respectively. In this regard, all of the required substrates were synthesized as follows. (E)-p-Coumaryl aldehyde (6) was obtained in a 79% overall yield by the lithium aluminum hydride reduction of the 4-O-tert-butyldimethylsilyl derivative of methyl-4-hydroxycinnamate followed by oxidation with 2,3-dichloro-5,6-dicyano-1,4-benzoquinone and finally by deprotection with TBAF (19). (E)-Coniferyl aldehyde (7) was purchased from Sigma-Aldrich, and the unstable 4-HNE (15) was synthesized by epoxidation of 3-(Z)-nonenal with 3-chloroperoxybenzoic acid followed by periodinane oxidation and treatment with 1.3 M NaOH (40). The reduced products, p-dihydrocoumaryl (8) and dihydroconiferyl (9) aldehydes, were also individually synthesized via palladium on activated charcoal hydrogenation of the corresponding 4-O-tert-butyldimethylsilyl aldehyde derivatives with subsequent deprotection using TBAF in 60-70% yield (see "Experimental Procedures"). 4-HNA (16) was obtained via diisobutylaluminium hydride reduction of -nonanoic lactone to afford the desired product (R and S isomers) in an 1:1 ratio as evidenced by GC-MS analyses. Additionally, a variety of monomeric phenylpropanoid substrate analogues and their dihydro products were prepared, e.g. cinnamyl (17)/5-hydroxyconiferyl (18)/sinapyl (19) aldehydes, the corresponding dihydrocinnamyl (30)/5-hydroxydihydroconiferyl (31)/dihydrosinapyl (32) aldehydes, as well as p-coumaric (20), caffeic (21), ferulic (22), 5-hydroxyferulic (23), and sinapic (24) acids and the potential dihydroproducts (3, 4, 33-35). In an analogous manner, p-coumaryl (25), caffeyl (26), coniferyl (27), 5-hydroxyconiferyl (28), and sinapyl (29) alcohols and the dihydro derivatives 1, 2, 36-38 were also synthesized (see Figs. 1 and 4 for structures).⁴ Each of the purified potential substrates (6, 7, 17-29) and products (1-4, 30-38) was then used to establish standard curves using HPLC, and GC-MS was used for 4-HNE (15)/4-HNA (16) analyses, i.e. to directly quantify substrate turnover and product formation. By contrast, all previous studies of 4-HNE (15) enzymatic reductions were carried out indirectly by measuring changes in UV absorption at 340 nm.

Kinetic Parameters/Substrate Versatility—With all of the corresponding potential substrates needed for the study in hand, the AtDBR1 cDNA was cloned into a pTrcHis-TOPO vector containing a N-terminal polyhistidine (His₆) tag, with the plasmid used to transform E. coli TOP10 cells for gene expression. AtDBR1 was purified to apparent homogeneity (evaluated by SDS-PAGE with silver staining) following metal chelate affinity column chromatography as described by Kim et al. (19).

View larger version (28K): [in this window] [in a new window]   FIGURE 4. Phenylpropanoids (structures 17-29) and dihydrophenylpropanoids (structures 30-38).

Overall Structure of AtDBR1—To understand in greater detail the mechanistic/structural basis for the AtDBR1 kinetic properties, the recombinant AtDBR1 was next crystallized in its apoform and NADP⁺ binary and ternary complexes using the two different substrates p-coumaryl aldehyde (6) and 4-HNE (15), respectively. Crystals of both ternary complexes were obtained by mixing AtDBR1 with NADP⁺/p-coumaryl aldehyde (6) and by soaking the AtDBR1/NADP⁺ binary complex crystals in a solution containing 4-HNE (15), respectively. The apoform of AtDBR1 was determined at 2.5 Å resolution by the molecular replacement method using coordinates of the afore-mentioned 12-HD/PGR (PDB code 1V3V) from guinea pig (C. porcellus) (29), which shows 56/41% similarity/identity to AtDBR1 (Table 1). Additionally, the binary and ternary complex structures of AtDBR1 were determined at 2.8 Å resolution using the coordinates of the deduced structure of the AtDBR1 apoform.

As shown in Fig. 5, AtDBR1 is a homodimer with two subunits arranged through a noncrystallographic 2-fold axis. The two subunits are virtually superimposable, with an r.m.s.d. of 0.67 Å between the corresponding C atoms of the two subunits, without including the residues from 60 to 70 in the case of the apoform. The loop region of residues 60-70 connecting the 1 and 4 regions is disordered in one subunit, whereas the corresponding area of the other subunit is ordered because of crystal packing interactions. Both static and dynamic multiangle laser light-scattering data of AtDBR1 confirmed its dimeric nature (Fig. 6, A and B).

View larger version (41K): [in this window] [in a new window]   FIGURE 5. Ternary complex of AtDBR1 homodimer with bound NADP+/p-coumaryl aldehyde (structure 6) (A) and NADP+/4-HNE (structure 15) (B). The substrate- and nucleotide-binding domains are depicted in green and violet in the lower subunit, and in brown and light blue in the upper subunit, respectively. NADP+ is shown in dark blue, with p-coumaryl aldehyde (6) or 4-HNE (15) in dark brown. The active site in the lower subunit is outlined by a dotted box, and the secondary structure elements of AtDBR1 are numbered sequentially as 1-3/A-G and 1-10/A-F.

The overall fold of AtDBR1 belongs to that of the zinc-independent MDR superfamily. Like other MDRs, each monomer is composed of two domains: a substrate-binding domain (residues 1-137 and 306-345) and a nucleotide-binding domain (Rossman fold, residues 138-305) (Figs. 3 and 5). The substrate-binding domain has three -helices and ten -strands forming two -sheets, one of which is a highly twisted, eight-stranded, -barrel-like structure as observed previously for the putative NADP-dependent oxidoreductase from Mus musculus (Fig. 3; PDB code 1VJ1) (50). The nucleotide-binding domain of AtDBR1 also has seven -helices and six -strands forming a typical six-stranded parallel -sheet flanked by three helices on each side. In the binary and ternary complexes, the corresponding NADP⁺ and substrates were located at the active site clefts between the substrate- and nucleotide-binding domains, as described in detail below. Upon NADP⁺ and/or substrate binding, no significant changes were detected in either overall conformation or of the amino acids in the binding pockets as reflected in small r.m.s.d. values of 0.42-0.66 Å (not shown) among the C atoms of the apoform and the binary and ternary complexes. A minor positional change in the backbone was observed in the area of residues 36-39, which connects 2 and 3.

Structural Alignment—In general, the MDR superfamily can be divided into two subgroups (51). One subgroup contains no Zn²⁺, as seen for AtDBR1, PtPPDBR, 12-HD/PGR, and AOR, whereas the other has catalytic and/or structural Zn²⁺, such as in the well studied horse liver alcohol dehydrogenase (52) and cinnamyl alcohol dehydrogenase (19, 20). As noted earlier (Fig. 3), amino acid sequence alignments had also revealed that the plant enzymes AtDBR1 and PulR are the most closely related, although no three-dimensional structures had been reported until now.

A Dali search (53) of the PDB data base indicated that the highest match was to the 12-HD/PGR from guinea pig C. porcellus (PDB code 1V3V) with a Z-score of 43.0; this was followed by several quinone reductases, including the human -crystalline (PDB code 1YB5) with a Z-score of 32.3 (not shown), and the putative NADP-dependent oxidoreductase from M. musculus (1VJ1) of 31.4. The latter two show only 30 and 19% identity, however, to AtDBR1, based on amino acid sequence comparisons. Nevertheless, all of these high Z-scored proteins belong to the zinc-independent MDR superfamily.

The sequence alignment among the above-mentioned enzymes (Fig. 3) also revealed that AtDBR1 has several areas of deletion and insertion when compared with the other enzymes, which include several loops and even some secondary structural elements. Especially, one loop region that is composed of residues 30-40 connecting 2 and 3 is quite different, particularly when compared with the mammalian enzymes (Fig. 3). In addition, a highly disordered loop region (amino acids 60-89) in the structure of AtDBR1 is not aligned well among the other enzymes.

In general, however, all of the enzymes show a high degree of sequence similarity for most of the nucleotide-binding domain up to the conserved ²⁵⁵GXXS²⁵⁸ motif; this is known to stabilize both the adenine and nicotinamide moieties of the cofactor in the NADPH-bound form of quinone oxidoreductase (50). On the other hand, the remaining part of the nucleotide-binding domain (residues 262-305) has a low level of sequence similarity and participates mainly in subunit interactions.

NADP⁺-binding Site—As seen for other members of the MDR superfamily, the electron density corresponding to NADP⁺ in AtDBR1 was located in the cleft between the two domains formed by the carboxyl ends of A, B, D, E, F, and the loop connecting F and G. There was also a clear and continuous electron density in the initial F_o - F_c map (Fig. 7A, inset); all of the side chains around this cofactor were thus in a well defined electron density that is highly conserved among the MDR family. The corresponding binding pocket for NADP⁺, however, is filled with water molecules in the apoform. The coordinates for the two NADP⁺ molecules were thus refined to the same relative position in each subunit, with the bound NADP⁺ molecules adopting the anti configuration for both the ribose-nicotinamide and ribose-adenine glycosidic bonds (Fig. 7A).

View larger version (12K): [in this window] [in a new window]   FIGURE 6. Molecular mass of AtDBR1 in solution. A, elution profile of AtDBR1 monitored by multiangle laser light scattering (2 mg ml-1). The elution profile is shown as molecular mass versus elution time. The thin solid lines represent changes in refractive index on an arbitrary scale that is proportional to protein concentration. The thick solid line indicates the calculated molecular masses. B, dynamic light scattering data of AtDBR1 (2 mg ml-1). The calculated molecular radius and molecular mass are 3.94 nm and 84 kDa, respectively, indicating its dimeric nature.

The pyrophosphate group of NADP⁺ in AtDBR is within hydrogen bonding distance of both the backbone amide nitrogens of residues Ala-167 and Val-168, which reside inside the AXXGXXG motif, as well as the side chains of two residues, Asn-334 and Lys-337 (Fig. 7A). In addition, the backbones of Cys-254 in E and Phe-284/Val-286 adjacent to F are within hydrogen bonding distance of the amide moiety of the nicotinamide ring (Fig. 7A). Interestingly, the above-mentioned ²⁵⁵GXXS²⁵⁸ motif, which is also located near the nicotinamide ring, can be extended to "CGXXSXY" among the compared zinc-independent MDR enzymes. The net effect is that the freedom of the nicotinamide ring is very restricted, presumably fixing the position of the C-4 atom during catalysis.

Notably, both the hydroxyl group of the Tyr-260 and the backbone nitrogen of Tyr-53, which are completely conserved among all of the oxidoreductases compared (see Fig. 3), are within hydrogen bonding distance of the O-2' of the nicotinamide ribose ring. In contrast to the well anchored nicotinamide ring, however, the adenine ring does not have much interaction with the surrounding amino acids and thus should be relatively free to move, as indicated by its high B-value.

The AtDBR1, NADP⁺, and p-Coumaryl Aldehyde (6) Ternary Complex—In the ternary complex, the observed distances between the C-4 atom of the nicotinamide ring and the -carbon of the , (7,8)-unsaturated double bond of p-coumaryl aldehyde (6) in the two subunits are 3.8 and 4.0 Å, respectively. Thus, p-coumaryl aldehyde (6), located at the active site cleft between the substrate- and nucleotide-binding domains, is in the proper orientation to the nicotinamide ring for the well established A-face-specific hydride transfer from C-4 to the corresponding substrate reaction center (Fig. 7B).

Unlike the NADP⁺ molecule deeply buried inside the binding pocket with many interactions involving various amino acid residues, p-coumaryl aldehyde (6) is relatively exposed to the solvent; thus a weaker binding constant can be expected when compared with that of the cofactor. The inner wall of this exposed substrate-binding site is also surrounded by the nicotinamide ring, as well as Tyr-53, Tyr-81, Met-138, Tyr-260, Ser-287, and Tyr-290 from one subunit and Ile-275 and Tyr-276 from another, indicative of its quite polar nature (Fig. 7B). In either the apoform or the NADP⁺ binary complex, however, this substrate-binding site is filled with water molecules, reflecting its somewhat hydrophilic nature.

Of particular note is the fact that the phenolic hydroxy group of Tyr-260 is within hydrogen bonding distance to the aldehydic oxygen of the substrate (6), in addition to the 2'-hydroxyl group of the nicotinamide ribose described previously. Therefore, we considered that this conserved Tyr-260 residue is hydrogen bonded to both. In addition, the Tyr-81 hydroxyl group is also potentially within hydrogen bonding distance of the phenolic hydroxyl group (Fig. 7B), whereas the Ser-287 hydroxyl group is 3.9 Å away, thus probably too far away for hydrogen bond formation. However, because of their location in the flexible (high temperature factor) loops, there provisionally remained a possibility that hydroxyl groups in Tyr-81 and Ser-287 were potentially involved in facilitating substrate binding.

Lastly, the phenol ring of the highly conserved Tyr-53, which is located between a relatively conserved short loop and one-turn -helix, is in a potential stacking interaction with the corresponding phenol ring of the bound p-coumaryl aldehyde (6), thereby probably further facilitating orientation of the substrate within the specificity pocket (Fig. 7B).

View larger version (56K): [in this window] [in a new window]   FIGURE 7. Stereo view of the substrate-binding pocket of AtDBR1 (viewed from the bulk solvent). A, binary complex. B, ternary complex with NADP+ and p-coumaryl aldehyde (6). C, ternary complex with NADP+ and 4-HNE (15). The glycosidic bond angles of the bound NADP+ for all cases adopt an anti conformation. In the insets, experimental difference Fourier maps (|Fo| - |Fc|) corresponding to NADP+, p-coumaryl aldehyde (6), and 4-HNE (15) in binary and ternary complexes of AtDBR1 are shown at a contour level of 3.0 . The participating residues in the binding substrate and cofactor are shown with their residue position numbers. The residues are depicted in light blue and orange to represent their belonging to two different subunits.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION CONCLUDING REMARKS REFERENCES

 
In contrast to two other oxidoreductases involved in related aspects of phenylpropanoid metabolism, i.e. secoisolariciresinol dehydrogenase (15-17) and cinnamyl alcohol dehydrogenase 5 (19, 20), the crystal structure of AtDBR1 shows that it belongs to the Zn²⁺-independent MDR family. This class of enzymes, as indicated above, includes PtPPDBR, PulR, 12-HD/PGR, AOR, the putative NADP-dependent oxidoreductase from M. musculus (PDB code 1VJ1), and the -crystallins. The enzymatic reaction mechanism of this class of enzymes that lack Zn²⁺ is, however, still unclear, e.g. when compared with the very well established Zn²⁺-containing MDRs such as alcohol dehydrogenase. Indeed, a majority of the enzymes provisionally belonging to this group are still depicted as putative gene products (55).

In terms of the overall distribution of secondary structural elements and the local active site, AtDBR1 displays a striking similarity to 12-HD/PGR. As discussed earlier, the latter catalyzes the reduction of a very broad array of ,-unsaturated ketones and aldehydes, including toxic products of lipid peroxidation in addition to that of the 13-14-double bond of 15-oxoprostaglandins (29, 56). Accordingly, most of the residues constituting the NADP(H)-binding sites are very similar between AtDBR1 and 12-HD/PGR. Indeed, the overall shapes of the binding pockets for the cofactor are also very similar, and the bound cofactor in both reductases adopts the same sugar conformation.

For guinea pig 12-HD/PGR and rat liver AOR, a ketoreductase reaction mechanism has been proposed with a conjugated enolate as the reaction intermediate (29, 55). Apparently, the latter enzyme is also able to reduce one of the phenylpropanoids, cinnamyl aldehyde (17) (30), whereas AtDBR1 does not reduce it. It is tempting to speculate, therefore, that the hydrophobic nature of the phenyl moiety of the cinnamyl aldehyde (17) can be accommodated in the hydrophobic substrate-binding pocket of 12-HD/PGR but not in the AtDBR binding pocket, which is more hydrophilic in nature. In addition, the intrinsic difference in terms of rotational freedom around the C-1-C-7 bond between cinnamyl aldehyde (17) and p-coumaryl aldehyde (6) (resulting from an ability of the latter to more readily form resonance hybrids because of the phenol functionality at C-4) may also provisionally explain the substrate preferences in binding and catalysis between 12-HD/PGR and AtDBR1.

Yet, in contrast to the similarities between the 12-HD/PGR and AtDBR1, the previously proposed candidate catalytic residue (Tyr-262) for the former could not be further substantiated. In part, this is because this residue (Tyr-276 in AtDBR1) is not conserved in the other sequences (see Fig. 3), and from the analysis of the crystal structure, it is too far away (5.5-5.9 Å) in AtDBR1 to be able to hydrogen bond with the aldehydic group of the substrate.

On the other hand, the residue Tyr-260 is conserved in all enzymes shown in Fig. 3. Hence, from the observed hydrogen bonding pattern and the strong conservation of Tyr-260 among related enzymes, we propose that this residue serves as a general acid/base by stabilizing the enol form of the transition state (Fig. 8, A and B). Tyr-260 is also apparently hydrogen bonded to the 2'-hydroxyl group of the nicotinamide ribose, thereby potentially enabling the pK_a of its hydroxyl group to be modulated further. A similar catalytic mechanism can also be envisaged for 12-HD/PGR, PtPPDBR, PulR, AOR, and the putative NADPH-dependent oxidoreductase from M. musculus (1VJ1).

From our studies using [4R-³H]- and [4S-³H]NADPH, AtDBR1⁵ and PtPPDBR (37) catalyze the transfer of the pro-R hydride from NADPH to that of the double bond between C-7 and C-8, with this bond being polarized due to its neighboring carbonyl group. In agreement with this finding, the observed interactions between the carbonyl oxygen of both substrates, p-coumaryl aldehyde (6) and 4-HNE (15), and the Tyr-260 residue can also potentially stabilize the transient oxyanion enolate intermediate, thereby facilitating pro-R hydride transfer from the C-4 atom of the NADPH nicotinamide to the substrate (Fig. 8, A and B). As anticipated, however, AtDBR1 was not able to reduce the corresponding unpolarized double bond of the various phenylpropenols 25-29, as shown from our kinetic data.

View larger version (18K): [in this window] [in a new window]   FIGURE 8. Observed potential interactions in the ternary complex of AtDBR1 with NADP+/p-coumaryl aldehyde (structure 6) (A), NADP+/4-HNE (structure 15) (B), and their corresponding schematic reaction mechanisms (C and D). Arrows indicate the stacking interaction between phenol rings. The phenolic group of Tyr-81 is within a hydrogen bond distance of the p-coumaryl aldehyde (6) phenolic group, and the phenolic group of Tyr-260 is within hydrogen bonding distance of the carbonyl group of both bound p-coumaryl aldehyde (6) and 4-HNE (15).

In addition, the C-7=C-8 unsaturated double bond of p-coumaryl aldehyde (6) is in conjugation with both the allylic aldehyde moiety and the aromatic ring. Accordingly, the highly conserved Tyr-53 of AtDBR1 is thus possibly in a stacking position with the phenolic ring of the substrate. Therefore, - interactions between the phenolic substrate (6) and the nearby Tyr-53 residue could potentially stabilize further the propenal transition state, i.e. by withdrawing electron density away from C-8 (Fig. 8, A and C). This potential interaction may also explain why AtDBR1 can catalyze p-coumaryl aldehyde (6) reduction more efficiently than 4-HNE (15), as indicated from the kinetic data (Table 3).

	CONCLUDING REMARKS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION CONCLUDING REMARKS REFERENCES

 
With the discovery of the physiological function of PtPPDBR, it was possible to study the overall substrate versatility and catalytic mechanism of Arabidopsis At5g16970 (AtDBR1). This established first that p-coumaryl (6)/coniferyl (7) aldehydes were the preferred substrates over that of 4-HNE (15). Additionally, it identified Tyr-260 (i.e. in AtDBR1) as responsible for general acid/base catalysis and thus enabled us to define the overall catalytic mechanism of the alkenal double bond reductases in vascular plants. These findings also provide a much fuller understanding of the catalytic mechanism of the more distantly related mammalian double bond reductase homologues such as 12-HD/PGR and AOR.

Careful inspection of the AtDBR1 substrate-binding site in both the apoform and the ternary complex also revealed that it has several factors favoring p-coumaryl aldehyde (6) as a substrate, even though it can accommodate (and reduce) other phenylpropanoids and 2-alkenals of longer/shorter chains. For example, because the substrate binding pocket is polar in nature, the presence of hydroxyl groups in p-coumaryl (6)/coniferyl (7) aldehydes, as well as in 4-HNE (15), enhance their binding affinities. By contrast, similar size molecules, such as cinnamyl aldehyde (17), which lacks a phenolic/hydroxyl group, have a much lower affinity for the pocket as suggested by the lack of activity. Additionally, although not depicted in the proposed catalytic mechanism, the phenolic group of p-coumaryl aldehyde (6) is fully conjugated to the allylic aldehydic moiety, and this (because of a potentially reduced level of rotational freedom in the C-1-C-7 bond of cinnamyl aldehyde (17) versus p-coumaryl aldehyde (6)) may provide a further rationale as to why 17 does not serve as a substrate. Finally, the relatively tight fit of the overall binding pocket might also serve to explain why more highly substituted potential substrates, such as 5-hydroxyconiferyl (18) and sinapyl (19) aldehydes are unable to undergo reductive conversions (i.e. when the phenolic group at C-4 is flanked by o,o' substituents). Future work will be directed toward more comprehensively establishing the range of physiological functions of AtDBR1 and PtPPDBR, respectively.

	FOOTNOTES

 
The atomic coordinates and structure factors (codes 2J3H, 2J3I, 2J3J, and 2J3K) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).

^* This research was supported by grants from the National Institutes of Health, NIGMS (to C.-H. K. and N. G. L.); Grants MCB-9976684 and MCB-0417291 from the National Science Foundation; Agricultural Plant Biochemistry Grant 2006-03339 from the United States Department of Agriculture; and by grants from McIntire-Stennis, the Murdock Charitable Trust, and the G. Thomas and Anita Hargrove Center for Plant Genomic Research. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

¹ To whom correspondence may be addressed: Inst. of Biological Chemistry, Washington State University, Pullman, WA 99164-6340. Tel.: 509-335-2605; Fax: 509-335-8206; E-mail: lewisn{at}wsu.edu. ² To whom correspondence may be addressed: School of Molecular Biosciences, Washington State University, Pullman, WA 99164-4660. Tel.: 509-335-1409; Fax: 509-335-9688; E-mail: chkang{at}wsunix.wsu.edu.

³ The abbreviations used are: PtPPDBR, P. taeda phenylpropenal (, double bond) reductase; AOR, alkenal/one oxidoreductase; AtDBR1, A. thaliana double bond reductase 1; EI-MS, electron impact mass spectra; ESI, electrospray ionization; GC-MS, gas chromatography-mass spectrometry; 12-HD/PGR, 12-hydroxydehydrogenase/15-oxo-prostaglandin 13-reductase; 4-HNA, 4-hydroxynonanal; 4-HNE, 4-hydroxy-(2E)-nonenal; HPLC, high pressure liquid chromatography; MDR, medium chain dehydrogenases/reductases; MES, 4-morpholineethanesulfonic acid; PDB, Protein Data Bank; r.m.s.d., root mean square deviation; PulR, (+)-pulegone reductase; TBAF, tetrabutylammonium fluoride.

⁴ S.-J. Kim, S. G. A. Moinuddin, D. L. Bedgar, C. Lee, L. B. Davin, and N. G. Lewis, manuscript in preparation.

⁵ S.-J. Kim, S. G. A. Moinuddin, D. L. Bedgar, L. B. Davin, and N. G. Lewis, unpublished observations.

	ACKNOWLEDGMENTS

 
We thank C. Ogata (Advanced Photon Source, Northeastern Collaborative Access Team beam line) and C. Ralston (Advanced Light Source, beam line 8.2.1) for assistance.

	REFERENCES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION CONCLUDING REMARKS REFERENCES

 

1. Canel, C., Moraes, R. M., Dayan, F. E., and Ferreira, D. (2000) Phytochemistry 54, 115-120[CrossRef][Medline] [Order article via Infotrieve]
2. Hande, K. R. (1998) Eur. J. Cancer 34, 1514-1521[CrossRef][Medline] [Order article via Infotrieve]
3. Imbert, T. F. (1998) Biochimie (Paris) 80, 207-222
4. Meresse, P., Dechaux, E., Monneret, C., and Bertounesque, E. (2004) Curr. Med. Chem. 11, 2443-2466[Medline] [Order article via Infotrieve]
5. Thompson, L. U., Seidl, M. M., Rickard, S. E., Orcheson, L. J., and Fong, H. H. S. (1996) Nutr. Cancer 26, 159-165[Medline] [Order article via Infotrieve]
6. Chang, C. C., Heller, J. D., Kuo, J., and Huang, R. C. C. (2004) Proc. Natl. Acad. Sci. U. S. A. 101, 13239-13244[Abstract/Free Full Text]
7. Kasai, H., Fukada, S., Yamaizumi, Z., Sugie, S., and Mori, H. (2000) Food Chem. Toxicol. 38, 467-471[CrossRef][Medline] [Order article via Infotrieve]
8. Feng, R., Lu, Y., Bowman, L. L., Qian, Y., Castranova, V., and Ding, M. (2005) J. Biol. Chem. 280, 27888-27895[Abstract/Free Full Text]
9. Bazzano, L. A., He, J., Ogden, L. G., Loria, C. M., Vupputuri, S., Myers, L., and Whelton, P. K. (2002) Am. J. Clin. Nutr. 76, 93-99[Abstract/Free Full Text]
10. Burt, S. (2004) Int. J. Food Microbiol. 94, 223-253[CrossRef][Medline] [Order article via Infotrieve]
11. Karonen, M., Hämäläinen, M., Nieminen, R., Klika, K. D., Loponen, J., Ovcharenko, V. V., Moilanen, E., and Pihlaja, K. (2004) J. Agric. Food Chem. 52, 7532-7540[CrossRef][Medline] [Order article via Infotrieve]
12. Ehrenkranz, J. R. L., Lewis, N. G., Kahn, C. R., and Roth, J. (2005) Diabetes-Metab. Res. Rev. 21, 31-38[CrossRef][Medline] [Order article via Infotrieve]
13. Dinkova-Kostova, A. T., Gang, D. R., Davin, L. B., Bedgar, D. L., Chu, A., and Lewis, N. G. (1996) J. Biol. Chem. 271, 29473-29482[Abstract/Free Full Text]
14. Min, T., Kasahara, H., Bedgar, D. L., Youn, B., Lawrence, P. K., Gang, D. R., Halls, S. C., Park, H., Hilsenbeck, J. L., Davin, L. B., Lewis, N. G., and Kang, C. (2003) J. Biol. Chem. 278, 50714-50723[Abstract/Free Full Text]
15. Xia, Z.-Q., Costa, M. A., Pélissier, H. C., Davin, L. B., and Lewis, N. G. (2001) J. Biol. Chem. 276, 12614-12623[Abstract/Free Full Text]
16. Youn, B., Moinuddin, S. G. A., Davin, L. B., Lewis, N. G., and Kang, C. (2005) J. Biol. Chem. 280, 12917-12926[Abstract/Free Full Text]
17. Moinuddin, S. G. A., Youn, B., Bedgar, D. L., Costa, M. A., Helms, G. L., Kang, C., Davin, L. B., and Lewis, N. G. (2006) Org. Biomol. Chem. 4, 808-816[CrossRef][Medline] [Order article via Infotrieve]
18. Gang, D. R., Kasahara, H., Xia, Z. Q., Vander Mijnsbrugge, K., Bauw, G., Boerjan, W., Van Montagu, M., Davin, L. B., and Lewis, N. G. (1999) J. Biol. Chem. 274, 7516-7527[Abstract/Free Full Text]
19. Kim, S. J., Kim, M. R., Bedgar, D. L., Moinuddin, S. G. A., Cardenas, C. L., Davin, L. B., Kang, C., and Lewis, N. G. (2004) Proc. Natl. Acad. Sci. U. S. A. 101, 1455-1460[Abstract/Free Full Text]
20. Youn, B., Camacho, R., Moinuddin, S. G. A., Lee, C., Davin, L. B., Lewis, N. G., and Kang, C. (2006) Org. Biomol. Chem. 4, 1687-1697[CrossRef][Medline] [Order article via Infotrieve]
21. Koeduka, T., Fridman, E., Gang, D. R., Vassão, D. G., Jackson, B. L., Kish, C. M., Orlova, I., Spassova, S. M., Lewis, N. G., Noel, J. P., Baiga, T. J., Dudareva, N., and Pichersky, E. (2006) Proc. Natl. Acad. Sci. U. S. A. 103, 10128-10133[Abstract/Free Full Text]
22. Vassão, D. G., Gang, D. R., Koeduka, T., Jackson, B., Pichersky, E., Davin, L. B., and Lewis, N. G. (2006) Org. Biomol. Chem. 4, 2733-2744[CrossRef][Medline] [Order article via Infotrieve]
23. Davin, L. B., Wang, H.-B., Crowell, A. L., Bedgar, D. L., Martin, D. M., Sarkanen, S., and Lewis, N. G. (1997) Science 275, 362-366[Abstract/Free Full Text]
24. Halls, S. C., Davin, L. B., Kramer, D. M., and Lewis, N. G. (2004) Biochemistry 43, 2587-2595[CrossRef][Medline] [Order article via Infotrieve]
25. Davin, L. B., and Lewis, N. G. (2005) Curr. Opin. Biotechnol. 16, 398-406[CrossRef][Medline] [Order article via Infotrieve]
26. Kraus, C., and Spiteller, G. (1997) Phytochemistry 44, 59-67
27. Kasahara, H., Davin, L. B., and Lewis, N. G. (March 9, 2004) U. S. Patent 6,703,229
28. Ringer, K. L., McConkey, M. E., Davis, E. M., Rushing, G. W., and Croteau, R. (2003) Arch. Biochem. Biophys. 418, 80-92[CrossRef][Medline] [Order article via Infotrieve]
29. Hori, T., Yokomizo, T., Ago, H., Sugahara, M., Ueno, G., Yamamoto, M., Kumasaka, T., Shimizu, T., and Miyano, M. (2004) J. Biol. Chem. 279, 22615-22623[Abstract/Free Full Text]
30. Dick, R. A., Kwak, M. K., Sutter, T. R., and Kensler, T. W. (2001) J. Biol. Chem. 276, 40803-40810[Abstract/Free Full Text]
31. Liu, Q., Smith, M. A., Avila, J., DeBernardis, J., Kansal, M., Takeda, A., Zhu, X., Nunomura, A., Honda, K., Moreira, P. I., Oliveira, C. R., Santos, M. S., Shimohama, S., Aliev, G., de la Torre, J., Ghanbari, H. A., Siedlak, S. L., Harris, P. L. R., Sayre, L. M., and Perry, G. (2005) Free Rad. Biol. Med. 38, 746-754[CrossRef][Medline] [Order article via Infotrieve]
32. Babiychuk, E., Kushnir, S., Belles-Boix, E., Van Montagu, M., and Inzé, D. (1995) J. Biol. Chem. 270, 26224-26231[Abstract/Free Full Text]
33. Mano, J., Yoon, H., Asada, K., Babiychuk, E., Inzé, D., and Mikami, B. (2000) Biochim. Biophys. Acta 1480, 374-376[CrossRef][Medline] [Order article via Infotrieve]
34. Mano, J., Babiychuk, E., Belles-Boix, E., Hiratake, J., Kimura, A., Inzé, D., Kushnir, S., and Asada, K. (2000) Eur. J. Biochem. 267, 3661-3671[Medline] [Order article via Infotrieve]
35. Mano, J., Torii, Y., Hayashi, S.-i., Takimoto, K., Matsui, K., Nakamura, K., Inzé, D., Babiychuk, E., Kushnir, S., and Asada, K. (2002) Plant Cell Physiol. 43, 1445-1455[Abstract/Free Full Text]
36. Mano, J., Belles-Boix, E., Babiychuk, E., Inzé, D., Torii, Y., Hiraoka, E., Takimoto, K., Slooten, L., Asada, K., and Kushnir, S. (2005) Plant Physiol. 139, 1773-1783[Abstract/Free Full Text]
37. Kasahara, H., Jiao, Y., Bedgar, D. L., Kim, S.-J., Patten, A. M., Xia, X.-Q., Davin, L. B., and Lewis, N. G. (2006) Phytochemistry 67, 1765-1780[CrossRef][Medline] [Order article via Infotrieve]
38. Herbert, R. B., Kattah, A. E., and Knagg, E. (1990) Tetrahedron 46, 7119-7138[CrossRef]
39. Herbert, R. B., and Kattah, A. E. (1990) Tetrahedron 46, 7105-7118[CrossRef]
40. Gardner, H. W., Bartelt, R. J., and Weisleder, D. (1992) Lipids 27, 686-689[CrossRef]
41. Schneider, C., Tallman, K. A., Porter, N. A., and Brash, A. R. (2001) J. Biol. Chem. 276, 20831-20838[Abstract/Free Full Text]
42. Sugamoto, K., Matsushita, Y.-i., and Matsui, T. (1997) Lipids 32, 903-905[CrossRef][Medline] [Order article via Infotrieve]
43. Bloch, R., and Gilbert, L. (1987) J. Org. Chem. 52, 4603-4605[CrossRef]
44. Bates, H. A., and Farina, J. (1985) J. Org. Chem. 50, 3843-3845[CrossRef]
45. Pyysalo, H., Enqvist, J., Honkanen, E., and Pippuri, A. (1975) Finn. Chem. Lett. 129-135
46. Bates, R. W., Díez-Martín, D., Kerr, W. J., Knight, J. G., Ley, S. V., and Sakellaridis, A. (1990) Tetrahedron 46, 4063-4082[CrossRef]
47. Trzeciak, A. M., Wolszczak, E., and Ziólkowski, J. J. (1996) New J. Chem. 20, 365-370
48. Navaza, J. (1994) Acta Crystallogr. Sect. A 50, 157-163[CrossRef]
49. Brünger, A. T., Adams, P. D., Clore, G. M., DeLano, W. L., Gros, P., Grosse-Kunstleve, R. W., Jiang, J.-S., Kuszewski, J., Nilges, M., Pannu, N. S., Read, R. J., Rice, L. M., Simonson, T., and Warren, G. L. (1998) Acta Crystallogr. Sect. D 54, 905-921[CrossRef][Medline] [Order article via Infotrieve]
50. Levin, I., Schwarzenbacher, R., McMullan, D., Abdubek, P., Ambing, E., Biorac, T., Cambell, J., Canaves, J. M., Chiu, H. J., Dai, X., Deacon, A. M., DiDonato, M., Elsliger, M.-A., Godzik, A., Grittini, C., Grzechnik, S. K., Hampton, E., Jaroszewski, L., Karlak, C., Klock, H. E., Koesema, E., Kreusch, A., Kuhn, P., Lesley, S. A., McPhillips, T. M., Miller, M. D., Morse, A., Moy, K., Ouyang, J., Page, R., Quijano, K., Reyes, R., Robb, A., Sims, E., Spraggon, G., Stevens, R. C., van den Bedem, H., Velasquez, J., Vincent, J., von Delft, F., Wang, X., West, B., Wolf, G., Xu, Q., Hodgson, K. O., Wooley, J., and Wilson, I. A. (2004) Proteins 56, 629-633[CrossRef][Medline] [Order article via Infotrieve]
51. Edwards, K. J., Barton, J. D., Rossjohn, J., Thorn, J. M., Taylor, G. L., and Ollis, D. L. (1996) Arch. Biochem. Biophys. 328, 173-183[CrossRef][Medline] [Order article via Infotrieve]
52. Eklund, H., and Brändén, C. I. (1987) in Biological Macromolecules and Assemblies (Jurnak, F. A., and McPherson, A., eds) Vol. 3, pp. 75-142, John Wiley & Sons, London
53. Holm, L., and Sander, C. (1993) J. Mol. Biol. 233, 123-138[CrossRef][Medline] [Order article via Infotrieve]
54. Barbosa Pereira, P. J., Macedo-Ribeiro, S., Párraga, A., Pérez-Luque, R., Cunningham, O., Darcy, K., Mantle, T. J., and Coll, M. (2001) Nat. Struct. Biol. 8, 215-220[CrossRef][Medline] [Order article via Infotrieve]
55. Dick, R. A., and Kensler, T. W. (2004) J. Biol. Chem. 279, 17269-17277[Abstract/Free Full Text]
56. Ensor, C. M., Zhang, H., and Tai, H.-H. (1998) Biochem. J. 330, 103-108
57. Fromm, H. J. (1983) in Contemporary Enzyme Kinetics and Mechanism (Purich, D. L., ed) pp. 233-251, Academic Press, New York

CiteULike

Complore

Connotea

Del.icio.us

Digg

Reddit

Technorati

This article has been cited by other articles:

				S. Porte, E. Valencia, E. A. Yakovtseva, E. Borras, N. Shafqat, J. E. Debreczeny, A. C. W. Pike, U. Oppermann, J. Farres, I. Fita, et al. Three-dimensional Structure and Enzymatic Function of Proapoptotic Human p53-inducible Quinone Oxidoreductase PIG3 J. Biol. Chem., June 19, 2009; 284(25): 17194 - 17205. [Abstract] [Full Text] [PDF]

This Article Abstract Full Text (PDF) All Versions of this Article: 281/52/40076    most recent M605900200v1 Submit a Letter to Editor Alert me when this article is cited Alert me when eLetters are posted Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Youn, B. Articles by Kang, C. Search for Related Content PubMed PubMed Citation Articles by Youn, B. Articles by Kang, C. Social Bookmarking                      What's this?