Identification of a Serine Hydrolase as a Key Determinant in the Microbial Degradation of Polychlorinated Biphenyls
2000; Elsevier BV; Volume: 275; Issue: 21 Linguagem: Inglês
10.1074/jbc.275.21.15701
ISSN1083-351X
AutoresStephen Y. K. Seah, Geneviève Labbé, Sven Nerdinger, Matthew R. Johnson, Victor Snieckus, Lindsay D. Eltis,
Tópico(s)Toxic Organic Pollutants Impact
ResumoThe ability of 2-hydroxy-6-oxo-6-phenylhexa-2,4-dienoate (HOPDA) hydrolase (BphD) ofBurkholderia cepacia LB400 to hydrolyze polychlorinated biphenyl (PCB) metabolites was assessed by determining its specificity for monochlorinated HOPDAs. The relative specificities of BphD for HOPDAs bearing chlorine substituents on the phenyl moiety were 0.28, 0.38, and 1.1 for 8-Cl, 9-Cl, and 10-Cl HOPDA, respectively,versus HOPDA (100 mm phosphate, pH 7.5, 25 °C). In contrast, HOPDAs bearing chlorine substituents on the dienoate moiety were poor substrates for BphD, which hydrolyzed 3-Cl, 4-Cl, and 5-Cl HOPDA at relative maximal rates of 2.1 × 10−3, 1.4 × 10−4, and 0.36, respectively, versus HOPDA. The enzymatic transformation of 3-, 5-, 8-, 9-, and 10-Cl HOPDAs yielded stoichiometric quantities of the corresponding benzoate, indicating that BphD catalyzes the hydrolysis of these HOPDAs in the same manner as unchlorinated HOPDA. HOPDAs also underwent a nonenzymatic transformation to products that included acetophenone. In the case of 4-Cl HOPDA, this transformation proceeded via the formation of 4-OH HOPDA (t 12 = 2.8 h; 100 mm phosphate, pH 7.5, 25 °C). 3-Cl HOPDA (t 12 = 504 h) was almost 3 times more stable than 4-OH HOPDA. Finally, 3-Cl, 4-Cl and 4-OH HOPDAs competitively inhibited the BphD-catalyzed hydrolysis of HOPDA (K ic values of 0.57 ± 0.04, 3.6 ± 0.2, and 0.95 ± 0.04 μm, respectively). These results explain the accumulation of HOPDAs and chloroacetophenones in the microbial degradation of certain PCB congeners. More significantly, they indicate that in the degradation of PCB mixtures, BphD would be inhibited, thereby slowing the mineralization of all congeners. BphD is thus a key determinant in the aerobic microbial degradation of PCBs. The ability of 2-hydroxy-6-oxo-6-phenylhexa-2,4-dienoate (HOPDA) hydrolase (BphD) ofBurkholderia cepacia LB400 to hydrolyze polychlorinated biphenyl (PCB) metabolites was assessed by determining its specificity for monochlorinated HOPDAs. The relative specificities of BphD for HOPDAs bearing chlorine substituents on the phenyl moiety were 0.28, 0.38, and 1.1 for 8-Cl, 9-Cl, and 10-Cl HOPDA, respectively,versus HOPDA (100 mm phosphate, pH 7.5, 25 °C). In contrast, HOPDAs bearing chlorine substituents on the dienoate moiety were poor substrates for BphD, which hydrolyzed 3-Cl, 4-Cl, and 5-Cl HOPDA at relative maximal rates of 2.1 × 10−3, 1.4 × 10−4, and 0.36, respectively, versus HOPDA. The enzymatic transformation of 3-, 5-, 8-, 9-, and 10-Cl HOPDAs yielded stoichiometric quantities of the corresponding benzoate, indicating that BphD catalyzes the hydrolysis of these HOPDAs in the same manner as unchlorinated HOPDA. HOPDAs also underwent a nonenzymatic transformation to products that included acetophenone. In the case of 4-Cl HOPDA, this transformation proceeded via the formation of 4-OH HOPDA (t 12 = 2.8 h; 100 mm phosphate, pH 7.5, 25 °C). 3-Cl HOPDA (t 12 = 504 h) was almost 3 times more stable than 4-OH HOPDA. Finally, 3-Cl, 4-Cl and 4-OH HOPDAs competitively inhibited the BphD-catalyzed hydrolysis of HOPDA (K ic values of 0.57 ± 0.04, 3.6 ± 0.2, and 0.95 ± 0.04 μm, respectively). These results explain the accumulation of HOPDAs and chloroacetophenones in the microbial degradation of certain PCB congeners. More significantly, they indicate that in the degradation of PCB mixtures, BphD would be inhibited, thereby slowing the mineralization of all congeners. BphD is thus a key determinant in the aerobic microbial degradation of PCBs. polychlorinated biphenyl HOPDA hydrolase 2,3-dihydroxybiphenyl DHB dioxygenase dichlorobiphenyl heteronuclear multiple bond correlation 2-hydroxy-6-oxo-6-phenylhexa-2,4-dienoate high pressure liquid chromatography nuclear Overhauser effect spectroscopy trichlorobiphenyl PCBs1 are one of the most widely distributed classes of chlorinated pollutants in the environment (1.Tiedje J.M. Quensen III, J.F. Chee-Sanford J. Schimel J.P. Boyd S.A. Biodegradation. 1993; 4: 231-240Crossref PubMed Scopus (112) Google Scholar). Commercial mixtures of PCBs typically consist of over 60 of the 209 possible congeners, differing in the number and position of the chloro substituents. Although their production and widespread usage has been banned in the industrial world for over 20 years, the destruction of PCBs is of continued relevance due to their pervasiveness and persistence in soils and sediments, their impact on fragile ecosystems, such as those found in the Arctic, and human health concerns (2.McFarland V.A. Clarke J.U. Environ. Health Perspect. 1989; 81: 225-239Crossref PubMed Scopus (618) Google Scholar). The persistence of PCBs is due in part to their thermodynamic stability, structural diversity, and strong adsorption to organic matter. The discovery that a variety of microorganisms transform some PCB congeners has fueled interest in harnessing microbial catabolic capabilities to remediate sites contaminated by these pollutants. Effective bioremediation strategies require increasing the bioavailability of these compounds and optimizing the microbial catabolic activities involved in their mineralization (3.Head I.M. Microbiology. 1998; 144: 599-608Crossref Scopus (122) Google Scholar,4.Focht D.D. Curr. Opin. Biotechnol. 1995; 6: 341-346Crossref Scopus (67) Google Scholar). Many PCBs are transformed aerobically by the bph pathway, which is found in a wide variety of soil isolates (5.Furukawa K. Tomizuka N. Kamibayashi A. Appl. Environ. Microbiol. 1979; 38: 301-310Crossref PubMed Google Scholar, 6.Bedard D.L. Unterman R. Bopp L.H. Brennan M.J. Haberl M.L. Johnson C. Appl. Environ. Microbiol. 1986; 51: 761-768Crossref PubMed Google Scholar). The upperbph pathway consists of four enzymatic activities that act sequentially to transform biphenyl to benzoate and 2-hydroxypenta-2,4-dienoate via dihydroxylation of one of the aryl rings and the production of a catecholic intermediate (4.Focht D.D. Curr. Opin. Biotechnol. 1995; 6: 341-346Crossref Scopus (67) Google Scholar). The ability of the bph pathway to transform PCB congeners is strain-dependent (5.Furukawa K. Tomizuka N. Kamibayashi A. Appl. Environ. Microbiol. 1979; 38: 301-310Crossref PubMed Google Scholar, 6.Bedard D.L. Unterman R. Bopp L.H. Brennan M.J. Haberl M.L. Johnson C. Appl. Environ. Microbiol. 1986; 51: 761-768Crossref PubMed Google Scholar, 7.Bedard D.L. Haberl M.L. Microbiol. Ecol. 1990; 20: 87-102Crossref Scopus (138) Google Scholar). However, even the so called “best” PCB-degrading strains, such as Burkholderia cepacia LB400, transform few highly chlorinated congeners and do not effectively mineralize all lightly chlorinated congeners (5.Furukawa K. Tomizuka N. Kamibayashi A. Appl. Environ. Microbiol. 1979; 38: 301-310Crossref PubMed Google Scholar, 6.Bedard D.L. Unterman R. Bopp L.H. Brennan M.J. Haberl M.L. Johnson C. Appl. Environ. Microbiol. 1986; 51: 761-768Crossref PubMed Google Scholar, 7.Bedard D.L. Haberl M.L. Microbiol. Ecol. 1990; 20: 87-102Crossref Scopus (138) Google Scholar, 8.Seeger M. Timmis K.N. Hofer B. Appl. Environ. Microbiol. 1995; 61: 2654-2655Crossref PubMed Google Scholar). These different catabolic capabilities have been studied largely as a function of biphenyl dioxygenase, which catalyzes the initial dihydroxylation of the PCB congener. Accordingly, this enzyme has been studied with a view to augmenting its utility for the degradation of PCBs (9.Mondello F.J. Turcich M.P. Lobos J.H. Erickson B.D. Appl. Environ. Microbiol. 1997; 63: 3096-3103Crossref PubMed Google Scholar, 10.Kimura N. Nishi A. Goto M. Furukawa K. J. Bacteriol. 1997; 179: 3936-3943Crossref PubMed Google Scholar, 11.Hurtubise Y. Barriault D. Sylvestre M. J. Bacteriol. 1998; 180: 5828-5835Crossref PubMed Google Scholar, 12.Kumamaru T. Suenaga H. Mitsuoka M. Watanabe T. Furukawa K. Nat. Biotechnol. 1998; 16: 663-666Crossref PubMed Scopus (183) Google Scholar, 13.Brühlmann F. Chen W. Biotechnol. Bioeng. 1999; 63: 544-551Crossref PubMed Scopus (89) Google Scholar). In contrast to these efforts, there have been very few reports on the ability of other bph pathway enzymes to transform chlorinated metabolites. Such studies are essential to understanding the basis of microbial catabolic activities and improving these activities for PCB degradation. Studies in which intact cells were incubated with individual PCB congeners indicate that the fourth enzyme of the bph pathway limits the degradation of PCBs by this pathway. This hydrolase, BphD (EC 3.7.1.8), catalyzes carbon-carbon bond cleavage of HOPDA (14.Catelani D. Colombi A. Biochem. J. 1974; 143: 431-434Crossref PubMed Scopus (37) Google Scholar), yielding 2-hydroxypenta-2,4-dienoate and benzoate (SchemeFS1) (15.Omori T. Sugimura K. Ishigooka H. Minoda Y. Agric. Biol. Chem. 1986; 50: 931-937Google Scholar). Incubation of two different biphenyl-degrading strains with 2,4′-diClB (2,4,4′-triClB and 2,4′,5-triClB, respectively) resulted in the accumulation of chlorinated HOPDAs (5.Furukawa K. Tomizuka N. Kamibayashi A. Appl. Environ. Microbiol. 1979; 38: 301-310Crossref PubMed Google Scholar). Similar results have been reported in several other strains, including B. cepacia LB400 (7.Bedard D.L. Haberl M.L. Microbiol. Ecol. 1990; 20: 87-102Crossref Scopus (138) Google Scholar, 8.Seeger M. Timmis K.N. Hofer B. Appl. Environ. Microbiol. 1995; 61: 2654-2655Crossref PubMed Google Scholar). However, in none of these studies were the accumulated chlorinated HOPDAs well characterized. Other congeners, such as 2,3,3′-triClB, have been reported to give rise to chloroacetophenones (7.Bedard D.L. Haberl M.L. Microbiol. Ecol. 1990; 20: 87-102Crossref Scopus (138) Google Scholar, 16.Bedard D.L. Haberl M.L. May R.J. Brennan M.J. Appl. Environ. Microbiol. 1987; 53: 1103-1112Crossref PubMed Google Scholar). The transformations leading to chloroacetophenones have yet to be described. BphD belongs to a family of enzymes involved in the aerobic degradation of aromatic compounds, hydrolyzing vinylogous 1,5-diketones formed by the dioxygenative meta-ring cleavage of arenes (17.Hofer B. Eltis L.D. Dowling D.N. Timmis K.N. Gene. 1992; 130: 47-55Crossref Scopus (154) Google Scholar). These enzymes have an α/β hydrolase fold as well as conserved serine, histidine, and aspartate residues, reminiscent of the catalytic triad present in a number of other hydrolytic enzymes (18.Ollis D.L. Cheah E. Cygler M. Dijkstra B. Frolow F. Franken S.M. Harel M. Remington S.J. Silman I. Schrag J. Sussman J.L. Verschueren K.H.G. Goldman A. Protein Eng. 1992; 5: 197-211Crossref PubMed Scopus (1821) Google Scholar). Mutagenesis studies on XylF, the hydrolase of the toluene and xylene degradation (TOL) pathway, and BphD of Comamonas testosteroni B-356 have confirmed that these residues are critical for catalysis (19.Diaz E. Timmis K.N. J. Biol. Chem. 1995; 270: 6403-6411Abstract Full Text Full Text PDF PubMed Scopus (70) Google Scholar, 20.Ahmad D. Fraser J. Sylvestre M. Larose A. Khan A. Bergeron J. Juteau J.M. Sondossi M. Gene. 1995; 156: 69-74Crossref PubMed Scopus (30) Google Scholar). Interestingly, the conserved serine residue appears to act as a base (21.Fleming S.M. Robertson T.A. Langley G.J. Bugg T.D.H. Biochemistry. 2000; 39: 1522-1531Crossref PubMed Scopus (54) Google Scholar). More specifically, mechanistic studies on MhpC, a hydrolase involved in the degradation of phenylpropionate, indicate that the substrate is first bound in the enol form and undergoes tautomerization to the keto form before being subject to base-catalyzed attack by water (21.Fleming S.M. Robertson T.A. Langley G.J. Bugg T.D.H. Biochemistry. 2000; 39: 1522-1531Crossref PubMed Scopus (54) Google Scholar, 22.Lam W.W.Y. Bugg T.D.H. Biochemistry. 1997; 36: 12242-12251Crossref PubMed Scopus (48) Google Scholar, 23.Henderson I.M.J. Bugg T.D.H. Biochemistry. 1997; 36: 12252-12258Crossref PubMed Scopus (38) Google Scholar). It has been proposed that the keto form of the substrate binds weakly and can dissociate from the active site prior to attack by water. Steady-state kinetic studies on BphD of B. cepacia LB400 (BphDLB400) indicate that this enzyme is also “leaky” (24.Seah Y.K.S. Terracina G. Bolin J.T. Riebel P. Snieckus V. Eltis L.D. J. Biol. Chem. 1998; 273: 22943-22949Abstract Full Text Full Text PDF PubMed Scopus (67) Google Scholar). To investigate the significance of BphD in the microbial degradation of PCBs, the specificity of BphDLB400 for monochlorinated HOPDAs was determined by steady-state kinetics. The HOPDAs used in these studies were generated enzymatically from the extradiol cleavage of the corresponding chlorinated dihydroxybiphenyls and were characterized using UV-visible and NMR spectroscopies. The stability and acid dissociation constants of the HOPDAs in aqueous buffer were determined. The transformation products of the HOPDAs in the absence and presence of BphD were investigated by HPLC. The results of these studies are discussed in terms of the degradation of PCBs, as well as the mechanism of BphD-related hydrolases. DHB, 4-Cl, 5-Cl, 6-Cl, 2′-Cl, 3′-Cl, and 4′-Cl DHBs were synthesized by a combined directed orthometalation-cross coupling strategy, purified to greater than 99% purity, and fully characterized as described elsewhere (25.Nerdinger S. Kendall C. Marchhart R. Riebel P. Johnson M.R. Yin C-F. Eltis L.D. Snieckus V. Chem. Commun. 1999; 22: 2259-2260Crossref Scopus (28) Google Scholar). All other chemicals were of analytical grade. BphDLB400 was purified to apparent homogeneity from Escherichia coli DH5α containing the plasmid pSS314 as described previously (24.Seah Y.K.S. Terracina G. Bolin J.T. Riebel P. Snieckus V. Eltis L.D. J. Biol. Chem. 1998; 273: 22943-22949Abstract Full Text Full Text PDF PubMed Scopus (67) Google Scholar). DHBD was purified anaerobically from B. cepacia LB400 containing the plasmid pLEBD4 as described previously (26.Vaillancourt F.H. Han S. Fortin P.D. Bolin J.T. Eltis L.D. J. Biol. Chem. 1998; 273: 34887-34895Abstract Full Text Full Text PDF PubMed Scopus (79) Google Scholar). Aliquots of concentrated enzymes were flash frozen in liquid nitrogen and stored at −80 °C. All buffers were made with water purified to a resistivity of 17 to 18 MΩ-cm using a NANOpure UV water purifier (Barnstead, Dubuque, IA). The pH was determined using a model PHM93 reference pH meter (Radiometer, Copenhagen, Denmark). Stock solutions of HOPDAs in 100 mm ionic strength of potassium phosphate, pH 7.5, were prepared immediately prior to use. HOPDAs was prepared by dissolving the corresponding DHB in a small volume of 95% ethanol and diluting to the desired volume with buffer (ethanol constituted less than 0.1% of the enzymatic assay solution). To this solution was added a sufficient amount of DHBD to completely transform the DHB to the corresponding HOPDA. Extinction coefficients of the yellow-colored HOPDAs were determined by adding excess quantities of the ring cleavage enzyme to solutions containing weighed amounts of DHB and determining the resultant absorbance spectrophotometrically. In the preparation of stock solutions of HOPDAs for the hydrolase assays, complete conversion was verified by ensuring that (i) there was no further increase in absorbance at the wavelength of maximum absorbance upon addition of fresh ring cleavage enzyme to a diluted sample, and (ii) the absorbance expected from the amount of DHB present and the determined extinction coefficient was obtained. HOPDAs used for steady-state kinetic studies were prepared enzymatically, divided into aliquots, frozen at −80 °C, and used within 8 h of being frozen. For all HOPDAs except 4-Cl HOPDA, individual aliquots were thawed as needed and used within 1 h of being thawed. In the case of 4-Cl HOPDA, aliquots were used immediately after being thawed. To determine the configuration of chlorinated HOPDAs in solution, HOPDAs were prepared from the respective DHBs as described above, acidified to pH 3 with 12 n HCl, and then extracted three times with ethyl acetate. The pooled organic phase was dried over anhydrous magnesium sulfate, which was then removed by filtration. To prepare the free acid of the respective HOPDAs, the ethyl acetate fraction was evaporated to dryness under reduced pressure. To prepare the sodium salt of the respective HOPDAs, a stoichiometric amount of sodium hydroxide dissolved in ethanol was added to the ethyl acetate fraction. The precipitated salt was washed with ethyl acetate and then dried under reduced pressure. The free acid and sodium salt forms of the HOPDAs were stored in a dessicator at 4 °C. For NMR analysis, the free acid and sodium salt of the HOPDAs were dissolved in deuterated acetone-d6 or deuterated phosphate buffer, pH 7.5, respectively, to a final concentration of about 40 mm. To determine pK a values, each HOPDA was generated by the cleavage of the corresponding DHB, as described above, in 20 ml of unbuffered water. DHBD was removed by ultrafiltration using a stirred cell equipped with a YM10 membrane (Amicon). The enzyme-free ultrafiltrate, which contained approximately 0.5 mm HOPDA, was acidified with 12 n HCl to pH 3. This solution was titrated with aliquots of 10 μl 50 mm NaOH, and the pH was determined after the addition of each aliquot. The pK a values were determined from plots of pHversus amount of base added. The stability of each HOPDA was determined by spectrophotometrically monitoring the decrease in absorbance of a solution of the HOPDA (100 mm phosphate, pH 7.5, 25 °C) at the wavelength of maximum absorbance of the enolate tautomer of the HOPDA. The experiments were repeated using three different concentrations of each HOPDA (0.03, 0.3, and 0.7 mm). Half-lives were determined by fitting the data to a first-order decay using Excel (Microsoft, Redmond, WA). To prepare transformation products, 0.1 to 0.2 mm samples of freshly prepared HOPDA (100 mm phosphate, pH 7.5) were incubated in the absence or presence of purified BphD. When present, BphD was added to a final concentration of 0.1 μg/ml, and the samples were monitored for up to 1 h. When no BphD was present, the samples were monitored for up to 4 days. The transformation of the HOPDA was monitored spectrophotometrically at the appropriate wavelengths. Transformation products were analyzed by HPLC as described below. Experiments were done in duplicate. 1H NMR spectroscopy was performed on a Bruker Avance DMX 500 MHz spectrometer, a Bruker Avance DPX 300 MHz spectrometer, and a Bruker AC300 spectrometer. The Avance models were interfaced with Silicon Graphics workstations. HOPDA samples were dissolved in acetone-d6, and solvent peaks were used as internal standards (1H = 2.04 ppm). NOESY and HMBC spectra were recorded using standard 2D sequences. In the NOESY sequence (recycle – π/2 – Δ – π/2 – τm – π/2 – acquire), the recycle delay and mixing time (τm) were 1 s. The variable delay increment was optimized to cover the useful NMR frequency range. The F2 dimension FID was defined with 2K points, and the F1 dimension was defined with 128 increments. Eight scans were summed for each increment. In the HMBC sequence (recycle – π/2 (P1) – Δ (½J) – π/2 (P3) – Δ (D6) – π/2 (P3) – (Dφ) – π – (Dφ) – π/2 (P3) – acquire), the recycle delay was 1 s, the evolution delay (D6) was 100 μs, and the increment delay (Dφ) was 3 μs. Twelve scans were summed for each increment. All final two-dimensional maps were symmetrized about the diagonal. Spectra were simulated using the ACD Labs software package (Advanced Chemistry Development Inc., Toronto, Ontario, Canada). Elemental analyses were performed by Canadian Microanalytical Service Ltd. (Delta, British Columbia, Canada). HOPDA transformation products were analyzed using a Hewlett Packard model HP1050 HPLC (Mississauga, Ontario, Canada) equipped with a Hewlett Packard ODS Hypersil C18 column (125 × 4 mm) and operated at a flow rate of 1 ml/min. Samples of 10 μl were injected into the HPLC. Samples analyzed for benzoates were eluted with 55% methanol:0.22% phosphoric acid:44.78% water, and the eluate was monitored at 230 nm. Samples analyzed for acetophenones were eluted with 40% methanol:0.3% phosphoric acid:59.7% water or 35% methanol:0.325% phosphoric acid:64.675% water, and the eluate was monitored at 250 nm. The retention times, peak areas, and absorption spectra of the eluted compounds were compared with standards containing known concentrations (100, 200, and 400 μm) of benzoate, acetophenone, or their chlorinated derivatives. Enzymatic activity was measured by following the consumption of the yellow substrate using a Varian Cary 3 spectrophotometer equipped with a thermojacketed cuvette holder. The spectrophotometer was interfaced to a microcomputer and controlled by Cary OS/2 multitasking software. The amount of enzyme used in each assay was adjusted so that the progress curve was linear for at least 2 min. Initial velocities were determined from a least squares analysis of the progress curves using the kinetics module of the Cary software. Specificity and inhibition experiments were carried out in a total volume of 1.0 ml of 100 mm ionic strength potassium phosphate, pH 7.5, 25.0 ± 0.1 °C. The reaction was initiated by adding 5–10 μl of an appropriately diluted enzyme preparation to the reaction cuvette. A reaction mixture prepared without the hydrolytic enzyme served as a reference. Reactions were monitored at the wavelengths indicated in Table I. For specificity experiments, initial velocities were determined at substrate concentrations that ranged from 0.3 to 10 times the K m for that substrate. For inhibition experiments, HOPDA was used as a substrate and the concentration of inhibitor was varied from at least 0.7 to 4 times the K i for that compound. At each concentration of inhibitor, the concentration of HOPDA was varied from 0.3 to 10 times its apparent K m. Initial velocities determined at different substrate and inhibitor concentrations were fitted to the appropriate equations using the least squares and dynamic weighting options of LEONORA (27.Cornish-Bowden A. Analysis of Enzyme Kinetic Data. Oxford University Press, New York1995Google Scholar). The validity of the kinetic models was evaluated using residual plots and analyses of variance (F statistic).Table IMolar extinction coefficients of chlorinated HOPDAsCompoundWavelengthεt 1/2pK anmmm −1 cm −1hHOPDA43425.7587.33-Cl HOPDA43240.65046.14-Cl HOPDA409.526.8179a4-Cl HOPDA was transformed to 4-OH HOPDA (t 1/2 = 2.8 h), which was further transformed (t 1/2 = 179 h) as described under “Results.”5-Cl HOPDA40240.13606.18-Cl HOPDA39340.31976.59-Cl HOPDA43628.2506.810-Cl HOPDA43826.3486.9Molar extinction coefficients and half-lives were determined in 100 mm potassium phosphate buffer, pH 7.5, 25 °C using freshly generated HOPDAs. The pK a values of the respective enolates were determined in unbuffered water.a 4-Cl HOPDA was transformed to 4-OH HOPDA (t 1/2 = 2.8 h), which was further transformed (t 1/2 = 179 h) as described under “Results.” Open table in a new tab Molar extinction coefficients and half-lives were determined in 100 mm potassium phosphate buffer, pH 7.5, 25 °C using freshly generated HOPDAs. The pK a values of the respective enolates were determined in unbuffered water. Solutions of the freshly prepared HOPDAs, produced from the DHBD-catalyzed cleavage of the corresponding DHBs in 100 mm phosphate, pH 7.5 (25 °C), all had the characteristic yellow color of the enolate anion. Accordingly, the absorption spectra of these solutions had wavelengths of maximal absorbance between 390 and 440 nm (TableI). The extinction coefficient at this wavelength for each HOPDA are given in Table I. This table also summarizes the pK a values for the enolic hydroxyl group for HOPDA and each chlorinated HOPDA as determined by titration. The pK a value was relatively unaffected by chloro substituents at C-9 or C-10. In contrast, chloro substituents at C-3, C-5, or C-8 lowered the pK a value by 0.7 to 1 unit. As described below, the pK a of 4-Cl HOPDA could not be reliably determined due to its nonenzymatic transformation, which is accelerated at high pH (>9). All HOPDAs were observed to undergo a spontaneous transformation in aqueous buffer. The half-lives of the HOPDAs, determined spectrophotometrically by monitoring the decrease in absorbance of the enolate anion, indicated that their stability varied according to the position of chloro substituent (Table I). Interestingly, this stability appears to correlate with the pK a of the enolic group. Thus, HOPDA, 9-Cl HOPDA, and 10-Cl HOPDA, the enolic pK avalues of which are between 6.8 and 7.3, had half-lives of about 50 h. In contrast, 3-Cl, 5-Cl, and 8-Cl HOPDA, with pK a between 6.1 and 6.5, had half-lives of over 200 h. It appears that the compounds with lower pK a values are more stable due to the presence of higher proportion of enolate species at pH 7.5. 4-Cl HOPDA differed from the other HOPDAs in that it underwent two successive transformation in aqueous solution. When prepared freshly from 5-Cl DHB, 4-Cl HOPDA had a maximal absorption at 409.5 nm at pH 7.5, typical of an enolate anion (compound A). Compound A underwent an initial transformation to compound B, the absorption spectrum of which was characterized by a maximum at 300 nm and isosbestic points at 262, 281, and 344 nm with respect to the initial enolate (Fig.1). The half-life of the transformation of compound A to B was 2.8 h at pH 7.5 and decreased at basic pH. Compound B underwent a second transformation with a half-life of 179 h. Addition of 0.6 m sodium borate to a mixture of compounds A and B at pH 7.5 yielded a spectrum with features at 395 and 409.5 nm (Fig. 1), the relative intensities of which depended on the proportion of compounds A and B in the mixture. Addition of borate to a solution of freshly prepared 4-Cl HOPDA yielded a spectrum that was similar to that of the enolate anion. In contrast, addition of borate to a solution containing 95% compound B yielded a spectrum with an absorbance maximum at 395 nm and a shoulder around 409 nm. Borate promotes the tautomerization of α-ketoacids to α-enol acids through the formation of borate-enol complexes (28.Knox W.E. Pitt B.M. J. Biol. Chem. 1957; 225: 675-688Abstract Full Text PDF PubMed Google Scholar, 29.Koester R. Rotermund G.W. Liebigs Ann. Chem. 1965; 689: 40-64Crossref Scopus (28) Google Scholar). It was thus concluded that 4-Cl HOPDA was initially transformed to a structurally similar α-ketoacid. Compound B was identified as 4-OH HOPDA by NMR and elemental analysis as described below. The 1H NMR spectra of HOPDAs in acetone indicated that the monochlorinated HOPDAs exist in different tautomeric forms dependent on the position of the chloro substituent on the dienoate (TableII). The spectrum of HOPDA was in good agreement with the reported spectrum and indicated that this compound exists as an enol in a trans transoid configuration (14.Catelani D. Colombi A. Biochem. J. 1974; 143: 431-434Crossref PubMed Scopus (37) Google Scholar) (Scheme FS1). The spectrum of 5-Cl HOPDA revealed two doublets at 7.59 and 6.68 ppm. These correspond to H-3 and H-4, respectively, and indicate that 5-Cl HOPDA also exists in a single enolic form in acetone. The coupling constants of the H-3 and H-4 protons are consistent with a transoid configuration. In contrast, the1H NMR spectrum of 3-Cl HOPDA demonstrated that this compound exists in both enol and keto tautomeric forms in a ratio of 2:1 respectively (Table II). Identification of the keto form was based on the occurrence of a doublet at 4.41 ppm, corresponding to the saturated CH2 of the keto tautomer (H-5K) and a triplet at 7.52 ppm, corresponding to the β proton of the α/β unsaturated carbonyl (H-4K). Significantly, the1H NMR spectrum of 3-Cl HOPDA in deuterated phosphate buffer indicated that this compound exists as an enol in aqueous solution, consistent with the electronic absorption spectrum.Table IIAssignment of 1 H NMR of HOPDAsCompoundProton345orthometaparaHOPDAδ (ppm)6.54 (d)7.90 (dd)7.40 (d)8.07 (d)7.53–7.68J (Hz)11.811.8, 15.115.17.33-Cl HOPDAδ (ppm)8.58 (bd)7.12 (bd)7.997.70–7.48 EnolJ(Hz)—aThe coupling constant was not determined due to the broadness of the peak.—aThe coupling constant was not determined due to the broadness of the peak.3-Cl HOPDAδ (ppm)7.52 (t)4.41 (d)8.087.70–7.48 KetoJ (Hz)7.57.54-Cl HOPDAδ (ppm)7.29 (s)6.98 (s)8.05 (d)7.67–7.50J(Hz)7.14-OH HOPDAδ (ppm)6.42 (s)6.61 (s)7.79 (d)7.67–7.50J (Hz)7.35-Cl HOPDAδ (ppm)7.59 (d)6.68 (d)7.73 (d)7.54 (dd)7.63 (d)J(Hz)11.211.277.7, 7.47.4Spectra were recorded in acetone-d6, and chemical shifts are reported relative to solvent peaks as internal standards.a The coupling constant was not determined due to the broadness of the peak. Open table in a new tab Spectra were recorded in acetone-d6, and chemical shifts are reported relative to solvent peaks as internal standards. The 1H NMR and 13C NMR spectra of 4-Cl HOPDA preparations in acetone revealed the presence of two structurally similar compounds, A and B, consistent with the absorption spectra of borate complexes of these preparations (Figs.2 and 3). Elemental analysis of compound B demonstrated that it contained no chlorine, consisting of 60.93% C, 4.45% H, and 34.36% O. Based on simulated NMR spectra, the elemental analysis, and chemical considerations, it was proposed that compounds A and B correspond to 4-Cl and 4-OH HOPDAs, respectively. In the 1H NMR spectrum of a mixture of compounds A and B, resonances corresponding to 4-Cl HOPDA were observed at 7.29 ppm (H-3A) and 6.98 ppm (H-5A). A NOESY spectrum (Fig. 2) revealed a cross-peak between H-5A (6.98 ppm) and the two orthoprotons on the aromatic ring (8.05 ppm), confirming the assignment of the singlet at 7.29 ppm to H-3A. Similarly, resonances corresponding
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