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J. Biol. Chem., Vol. 281, Issue 52, 40076-40088, December 29, 2006
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12
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.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONCONCLUDING REMARKSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONCONCLUDING REMARKSREFERENCES |
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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.
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double bond) reductase (PtPPDBR)3 (see Fig. 2A), for which the encoding gene (see Fig. 3) was cloned with the functionally recombinant protein obtained and characterized preliminarily (27). This enzyme, which is a member of the zinc-independent, medium chain dehydrogenase/reductase (MDR) superfamily, catalyzes the NADPH-dependent conversion of various monomeric and dimeric phenylpropenalaldehydes (e.g. 6 and 7; see Fig. 2A) into the corresponding phenylpropanaldehydes (e.g. 8 and 9). In terms of its amino acid similarity/identity, PtPPDBR has the closest homology to Arabidopsis thaliana At5g16970 (AtDBR1), as well as to a gene encoding (+)-pulegone reductase (PulR) from Mentha piperita (28), i.e. with similarities and identities of 63 and 43% and 62 and 44%, respectively (Fig. 3 and Table 1). In this study, we have described the characterization of the PtPPDBR homologue, At5g16970 (AtDBR1), which catalyzes the same conversions. By contrast, PulR from peppermint (M. piperita) (28) was not investigated; it is apparently specifically involved in a similar conversion of pulegone (10) to (+)-isomenthone (11) and (-)-menthone (12), with the latter predominating (Fig. 2B).
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, double bond reductions of various keto/aldehydic moieties in both prostaglandin metabolism of guinea pig (Cavia porcellus) kidney tissue, e.g. where 13 is converted to 14 by 12-hydroxydehydrogenase/15-oxo-prostaglandin 13-reductase (12-HD/PGR) (Fig. 2C and Fig. 3) (29), and in rat liver detoxification of 4-hydroxy-(2E)-nonenal (4-HNE (15)) (Figs. 2D and 3) (30). With the latter, this occurs through the action of a NAD(P)H-dependent alkenal/one oxidoreductase (AOR) to form 4-hydroxynonanal (4-HNA (16)) (30), in a manner analogous to the action of PtPPDBR. Both proteins, 12-HD/PGR and AOR, however, have only 51/34 and 51/33% similarity/identity to PtPPDBR and 89/79% similarity/identity to each other (Table 1). Interestingly, in humans, at physiological concentrations, 4-HNE (15), a product of lipid peroxidation, can induce apoptosis, affect cell-signaling pathways, and also form 4-HNE-protein adducts such as those found in Alzheimer disease and atherosclerotic plaques. Thus, there is a significant correlation in generation of these reactive intermediates to incidences of cancer, heart, and Alzheimer diseases in humans (30, 31).
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-crystallin protein family based on 
25-41% identity to mammalian -crystallins of unknown biochemical function (32, 33). Although this Arabidopsis protein was later shown capable of reducing diamide/quinone linkages (34), Mano et al. (35, 36) have since demonstrated that it can also reduce 4-HNE (15) and related potential substrates. This has led to a consideration that the Arabidopsis protein may, therefore, be involved in lipid-peroxidation derived alkenal reductions in response to oxidative stress.
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONCONCLUDING REMARKSREFERENCES |
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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 IntegrityTM 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 PackTM 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 MonoQTM 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).
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level for the apoform was 30,000 (95% completeness) between 15.0 and 2.5 Å resolution. The crystals of the NADP+ binary complexes did not diffract as well as the apoform and gave reflection numbers of 21,151 (above 2, 95% completeness) between 15.0 and 2.8 Å resolution. In addition, the ternary complex data of both p-coumaryl aldehyde (6) and 4-HNE (15) were collected between 15.0 and 2.8 Å resolution, 20,191 and 20,187 (above 2), respectively. The root mean square deviations (r.m.s.d.) (from ideal geometry) of the final coordinates corresponding to the apoform and the binary, ternary I, and ternary II complexes were 0.014, 0.015, 0.024, and 0.025 Å for bonds and 3.2, 3.5, 4.5, and 4.6° for angles, respectively. All AtDBR1 coordinates have been deposited in the Protein Data Bank (apoform, 2J3H; binary complex, 2J3I; ternary complex with p-coumaryl aldehyde (6), 2J3J; ternary complex with 4-HNE (15), 2J3K). |
RESULTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONCONCLUDING REMARKSREFERENCES |
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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).
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, double bond) reductions, including cinnamyl (17), 5-hydroxyconiferyl (18), and sinapyl (19) aldehydes. However, none of these closely related substrate analogues was converted into the corresponding dihydro derivatives (30-32), indicating that the substrate versatility of the AtDBR1 was fairly limited. This observation contrasts with reports that the AOR from rat liver is capable of reducing the , double bond of cinnamyl aldehyde (17) along with a very broad range of potential substrates; however, in that study the presumed conversion of 17 was evaluated indirectly by measuring the changes in absorbance at 340 nm (30) rather than by HPLC-MS of the resulting dihydro product (30). In an analogous manner, none of the hydroxycinnamic acids (20-24) were converted into the dihydrocinnamates (3, 4, 33-35) by AtDBR1, nor were any of the corresponding alcohols (25-29) directly reduced to afford 1, 2, 36-38.
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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).
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-strands from each subunit (F) in an anti-parallel manner, thereby forming an extended 12-stranded -sheet across the dimer interface (Fig. 5). Another smaller dimerforming interaction is at the opposite side of this F strand interaction and involves two hydrogen bonds between the two pseudo 2-fold related Val-268 residues (not shown). There are additional salt bridges and hydrogen bonds between the side chains and between the side chain and the main chains of individual subunits.
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 Fo - Fc 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).
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A and B (163AASGAVG169), which is known to participate in binding of the pyrophosphate group of NAD(P)+ or NAD(P)H through a micro-dipole of the helix (B) (54). Interestingly, PtPPDBR and PulR have a slightly different sequential motif, AAAGSVG, the significance of which (if any) is not yet known.
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).
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-carbon of the ,-unsaturated double bond of 4-HNE (15) in the two subunits are 3.8 and 4.0 Å, respectively, which are approximately the same distances as for the p-coumaryl aldehyde (6) complex. Moreover, as in the ternary complex with p-coumaryl aldehyde (6), the conserved Tyr-53 is now in a hydrophobic interaction with the corresponding aliphatic chain of the bound 4-HNE (15), thereby again facilitating orientation of the substrate within the specificity pocket (Fig. 7C). |
DISCUSSION |
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-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 pKa 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.
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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).
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CONCLUDING REMARKS |
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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.
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FOOTNOTES |
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* 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.
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ACKNOWLEDGMENTS |
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