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α-KDOase Activity in Oyster and Synthesis of α- and β-4-Methylumbelliferyl Ketosides of 3-Deoxy-d-manno-octulosonic Acid (KDO)

1997; Elsevier BV; Volume: 272; Issue: 42 Linguagem: Inglês

10.1074/jbc.272.42.26419

1083-351X

Yu‐Teh Li, Lai‐Xi Wang, Nadejda V. Pavlova, Su‐Chen Li, Yuan C. Lee,

Polyamine Metabolism and Applications

Although α- and β-linked 3-deoxy-d-manno-octulosonic acid (KDO) is found in lipopolysaccharides (LPSs) of Gram-negative bacteria, capsular polysaccharides of microorganisms, and plants, very little is known about its degradation. Using both thin-layer chromatography and the periodate-thiobarbituric acid reaction, we found that the hepatopancreas of oyster (Crassostrea virginica) contained an enzyme (α-KDOase) capable of releasing α-linked KDO from LPSs. To facilitate the studies of α-KDOase, we have carried out the synthesis of 4-methylumbelliferyl-α-KDO (α-KDO-MU) by conjugating the glycosyl chloride of the per-O-acetylated methylester of KDO with methylumbelliferone by the SN2 type reaction and the catalyzed phase-transfer. In both cases, the β-anomer was obtained as the major product with a yield of about 80%, whereas the yield of α-anomer was only about 7%. Attempts to increase the yield of α-anomer were not successful. α-KDO-MU was used as substrate to follow the purification of α-KDOase from oyster hepatopancreas. The pH optimum for oyster α-KDOase was determined to be 4.5 using Re-LPS as substrate and 3.0 using α-KDO-MU as substrate. The enzyme was found to be stable in the pH range of 3–8. This enzyme released KDO from different LPSs, including Re-LPS from Escherichia coliand Salmonella minnesota, Rd-LPS from S. minnesota, and de-O-acyl-Re-LPS (Kiang, J., Szu, S. C., Wang, L.X., Tang, M., and Lee, Y. C. (1997)Anal. Biochem. 245, 97–101). Although α- and β-linked 3-deoxy-d-manno-octulosonic acid (KDO) is found in lipopolysaccharides (LPSs) of Gram-negative bacteria, capsular polysaccharides of microorganisms, and plants, very little is known about its degradation. Using both thin-layer chromatography and the periodate-thiobarbituric acid reaction, we found that the hepatopancreas of oyster (Crassostrea virginica) contained an enzyme (α-KDOase) capable of releasing α-linked KDO from LPSs. To facilitate the studies of α-KDOase, we have carried out the synthesis of 4-methylumbelliferyl-α-KDO (α-KDO-MU) by conjugating the glycosyl chloride of the per-O-acetylated methylester of KDO with methylumbelliferone by the SN2 type reaction and the catalyzed phase-transfer. In both cases, the β-anomer was obtained as the major product with a yield of about 80%, whereas the yield of α-anomer was only about 7%. Attempts to increase the yield of α-anomer were not successful. α-KDO-MU was used as substrate to follow the purification of α-KDOase from oyster hepatopancreas. The pH optimum for oyster α-KDOase was determined to be 4.5 using Re-LPS as substrate and 3.0 using α-KDO-MU as substrate. The enzyme was found to be stable in the pH range of 3–8. This enzyme released KDO from different LPSs, including Re-LPS from Escherichia coliand Salmonella minnesota, Rd-LPS from S. minnesota, and de-O-acyl-Re-LPS (Kiang, J., Szu, S. C., Wang, L.X., Tang, M., and Lee, Y. C. (1997)Anal. Biochem. 245, 97–101). 3-Deoxy-d-manno-octulosonic acid (KDO) 1The abbreviations used are: KDO, 3-deoxy-d-manno-octulosonic acid; LPS, lipopolysaccharide; DeOA-LPS, de-O-acylated Re-LPS; MU, 4-methylumbelliferone; TLC, thin-layer chromatography; DMF,N,N-dimethylformamide; α-KDO-MU, MU α-ketoside of KDO; β-KDO-MU, MU β-ketoside of KDO; MS, mass spectrometry. is a major component of LPSs in Gram-negative bacteria (1Ghalambor M.A. Levine E.M. Heath E.C. J. Biol. Chem. 1966; 241: 3207-3215Abstract Full Text PDF PubMed Google Scholar, 2Droge W. Lehmann V. Luderitz O. Westphal O. Eur. J. Chem. 1970; 14: 175-184Crossref Scopus (135) Google Scholar, 3Le Dur A. Caroff M. Chaby R. Szabo L. Eur. J. Biochem. 1978; 84: 579-589Crossref PubMed Scopus (37) Google Scholar). In addition, KDO was also found in bacterial capsular polysaccharides (4Taylor P.W. Biochem. Biophys. Res. Commun. 1974; 61: 148-154Crossref PubMed Scopus (21) Google Scholar, 5Bhattacharjee A.K. Jennings H.J. Kenny C.P. Biochemistry. 1978; 17: 645-651Crossref PubMed Scopus (100) Google Scholar) and plant cell walls (6York W.S. Darvill A.G. McNeil M. Albersheim P. Carbohydr. Res. 1985; 138: 109-126Crossref Scopus (218) Google Scholar). Despite the wide occurrence of KDO, nothing is known about its degradation. Using both TLC and the periodate-thiobarbituric acid reaction (7Unger F.M. Adv. Carbohydr. Chem. Biochem. 1981; 38: 323-388Crossref Scopus (367) Google Scholar), we found that the hepatopancreas of oyster,Crassostrea virginica, contained an enzyme (α-KDOase) capable of releasing α-linked KDO from LPSs. To facilitate the studies of α-KDOase, we have carried out the synthesis of α-KDO-MU by conjugating the glycosyl chloride of per-O-acetylated methylester of KDO with methylumbelliferone using the SN2 coupling reaction and the catalyzed phase-transfer reaction. In both cases, the β-anomer was obtained as the major product. This paper describes the preparation of α- and β-KDO-MU (Fig. 1) and the use of α-KDO-MU as a substrate to follow the isolation of α-KDOase from oyster hepatopancreas. KDO (ammonium salt) was prepared by alkaline conjugation of oxalacetic acid and d-arabinose (7Unger F.M. Adv. Carbohydr. Chem. Biochem. 1981; 38: 323-388Crossref Scopus (367) Google Scholar). The following were purchased from commercial sources: oysters (C. virginica) were from P&J Oyster Company (New Orleans, LA); KDO, octyl-Sepharose, and phenylmethylsulfonyl fluoride were from Sigma; precoated silica gel-60 and 60 F254 TLC plates, silica gel, and Fractogel EMD SP-650(S) were from E. Merck (Darmstadt, Germany); and Sephacryl S 200-SF and Con A-Sepharose were from Pharmacia LKB Biotechnology. Re-LPS from Escherichia coli K12, D31 m4, lot 5 (KDO content 11.7%); Re-LPS from Salmonella minnesota R595, lot 30437B (KDO content 11.3%); and Rd-LPS from S. minnesota R7, lot 3 (KDO content 16.5%) were from List Biological Laboratories (Campbell, CA). DeOA-LPS prepared from Re-LPS R595 by mild hydrazinolysis as described previously (8Kiang J. Szu S.C. Wang L.X. Tang M. Lee Y.C. Anal. Biochem. 1997; 245: 97-101Crossref PubMed Scopus (17) Google Scholar) was a gift from Dr. S. C. Szu (National Institutes of Health, Bethesda, MD). Melting points were determined with a Fisher-Johns apparatus and were not corrected. NMR spectra were recorded at 25 °C with a Bruker AMX-300 spectrometer at 300 and 100 MHz for 1H NMR and13C NMR, respectively. Mass spectra were recorded with a VG 70-S mass spectrometer in a chemical ionization mode (reagent gas, NH3) or a fast atom bombardment-positive mode (matrix: 3-nitrobenzyl alcohol). TLC was carried out on precoated silica gel 60 F254 plates, and the carbohydrate components were detected by charring at 140 °C after spraying the plates with 15% H2SO4 in 50% ethanol or by UV absorption. Column chromatography was performed on silica gel. Ratios of solvents for TLC and column chromatography were expressed by volume. All evaporation and concentration was carried out below 40 °C under reduced pressure using a water aspirator unless specified otherwise. Compound 1, methyl 2,4,5,7,8-penta-O-acetyl-3-deoxy-α-d-manno-2-octulopyranosonate, was prepared by the acetylation of 20 g of KDO (ammonium salt) with acetic anhydride:pyridine (1:1; 160 ml). After 20 h at 25 °C, the reaction mixture was evaporated and co-evaporated with toluene, and the residue was partitioned between water (50 ml) and CHCl3 (150 ml). The organic layer was separated and washed with 5% HCl and water, dried (Na2SO4), and filtered. The filtrate was evaporated to give a pale yellow foam. TLC (ethyl acetate:AcOH:H2O, 3:2:1) revealed that it contained the 2,4,5,7,8-penta-O-acetyl-3-deoxy-α-d-manno-2-octulopyranosonic acid (R F 0.5) as the major component, together with several unknown, fast-moving components. Treatment of the mixture with excess diazomethane in CH2Cl2 (200 ml), followed by silica gel column chromatography using toluene:ethyl acetate (2:1) as the eluent, gave compound 1 (18.8 g, 52% from KDO) as a white solid: R F 0.30 (toluene:ether, 1:1); m.p. 154–156 °C; m.p. (literature) 155–158 °C (9Unger F.M. Stix D. Schulz G. Carbohydr. Res. 1980; 80: 191-195Crossref Scopus (102) Google Scholar); MS (m/z, chemical ionization-positive mode) 480 ((M + NH4)+, 100%), 420 ((M − AcOH + NH4)+, 8%), and 403 ((M − AcOH + H)+, 26%); 1H NMR (CDCl3) δ 3.818 (s, 3 H, OCH3), 2.151, 2.120, 2.057, 2.010, and 2.006 (each s, each 3 H, 5 CH3CO). The data for sugar protons are listed in TablesI and II. To prepare compound 2, dry hydrogen chloride gas was bubbled into a solution of 1 (3.85 g, 7.96 mmol) in acetyl chloride (40 ml) in a 100-ml flask at 0 °C for 30 min, then the flask was sealed and kept at 4 °C. After 24 h the solution was evaporated and the residual solvent was co-evaporated with toluene to give compound 2 (3.6 g, quantitative) as a colorless oil,R F 0. 55 (toluene:ether, 1:1). This glycosyl chloride of 1 was used as the glycosyl donor (2) without further purification.Table I1H NMR data for the sugar protons of KDO derivativesHydrogenChemical shift (δ)1-aChemical shift (δ) was expressed in ppm. and multiplicity1-bMultiplicity: s, singlet; d, doublet; t, triplet; m, multiplet.13456H-3a2.209 t2.459 t2.237 t2.115 t1.938 tH-3e2.261 dd2.488 dd2.410 dd2.483 dd2.224 ddH-45.335 ddd5.014 ddd5.548 ddd3.865 m4.255 dddH-55.407 m5.394 m5.452 m4.002 m4.032 mH-64.184 dd4.588 d4.193 dd4.044 d3.466 dH-75.232 ddd5.231 ddd5.289 ddd3.936 m3.894 dddH-8a4.488 dd4.428 dd4.356 dd3.819 m3.706 ddH-8b4.123 dd4.372 dd4.076 dd3.819 m3.123 ddThe NMR spectra were measured at 300 MHz in CDCl3 (for 1, 3, and 4) or D2O (for 5 and6).1-a Chemical shift (δ) was expressed in ppm.1-b Multiplicity: s, singlet; d, doublet; t, triplet; m, multiplet. Open table in a new tab Table IIVicinal proton coupling constants (J values) for KDO derivativesCoupled HJ value13456Hz3a, 3e12.012.012.512.512.83a, 411.211.512.212.512.03e, 45.75.85.64.65.44, 53.02.83.0—2-aNot analyzed because of the overlapping of signals.2.55, 61.3<1.01.2<1.0<1.06, 79.89.59.59.310.07, 8a2.32.62.2—2-aNot analyzed because of the overlapping of signals.2.27, 8b3.83.43.2—2-aNot analyzed because of the overlapping of signals.6.88a, 8b12.412.512.5—2-aNot analyzed because of the overlapping of signals.11.52-a Not analyzed because of the overlapping of signals. Open table in a new tab The NMR spectra were measured at 300 MHz in CDCl3 (for 1, 3, and 4) or D2O (for 5 and6). The conjugation of 2 with 4-methyl-umbelliferone was accomplished by SN2 coupling in DMF and catalyzed phase-transfer reaction as described below. To a solution of2 (2.5 g, 5.7 mmol) in DMF (20 ml), 4-methylumbelliferone (sodium salt) (1.70 g, 8.5 mmol) was added. The mixture was stirred at 25 °C for 4 h, and TLC (toluene:ethyl acetate, 1:1) indicated the disappearance of 2. DMF was evaporated in vacuo, and the residue was partitioned between water (20 ml) and CHCl3 (80 ml). The organic layer was separated and washed with aqueous NaHCO3 and water, dried (Na2SO4), and filtered. The filtrate was evaporated, and the residue was subjected to silica gel column chromatography using toluene:ethyl acetate (3:1) as the eluent to give a mixture of two UV-absorbing and carbohydrate-containing compounds that migrated closely on TLC (toluene:ethyl acetate, 2:1 or toluene:ether, 1:1). The mixture was further fractionated by silica gel column chromatography using toluene:ether (1:1) as the eluent to obtain the β-anomer 3 (2.5 g, 76%) and the α-anomer4 (224 mg, 6.8%). Alternatively, the two anomers could be simply separated by agitation of the mixture in toluene. The β-anomer3 is quite soluble in toluene but the α-anomer4 is not. Filtration of the suspension gave pure3 in toluene and pure 4 as a solid. A mixture of 2(440 mg, 1.0 mmol), 4-methylumbelliferone (sodium salt) (324 mg, 1.5 mmol), and tetrabutylammonium bisulfate (399 mg, 1.0 mmol) in CH2Cl2 (10 ml) and aqueous NaHCO3(0.5 m, 10 ml) was shaken vigorously at 25 °C for 12 h. Then, the organic layer was separated and washed with brine and water, dried (Na2SO4), and filtered. The filtrate was evaporated and the products were purified by the procedure described above to give the β-anomer 3 (463 mg, 80%) and α-anomer 4 (44 mg 7.6%). R F 0.24 (toluene:ether, 1:1); m.p. 77–80 °C; MS (m/z, chemical ionization mode) 596 ((M +NH4)+, 100%) and 579 ((M + H)+, 11%); 13C NMR (CDCl3, set CDCl3 at δ 77.23): δ 170.22, 169.92, 169.37, 169.27, and 167.46 (5 carboxyl), 160.30, 156.35, 154.12, and 152.01 (C-2′, 4′, 7′, 9′), 125.43, 115.87, 115.56, 113.16, 107.16, and 100.32 (C-2, 3′, 5′, 6′, 8′, 10′), 71.64, 67.71, 66.40, 63.61, and 61.92 (C-4, 5, 6, 7, 8), 52.94 (OCH3), 31.79, 30.42, 20.33, 20.26, 18.24, and 18.04 (C-3 and 5 CH3); 1H NMR (CDCl3): δ 7.515 (d, 1 H, J = 9.4 Hz, H-5′), 7.078 (dd, 1 H, J = 2.4 and 9.4 Hz, H-6′), 7.070 (d, 1 H, J = 2.4 Hz, H-8′), 6.210 (s, 1 H, H-3′), 3.731 (s, 3 H, OCH3), 2.417 (s, 3 H, CH3 at C-3′), 2.160, 2.142, 2.038, and 2.015 (each s, each 3 H, 4 COCH3). The data for sugar protons are listed in Tables I and II. R F 0.22 (toluene:ether, 1:1); m.p. 226–228 °C; MS (m/z, chemical ionization mode) 596 ((M + NH4)+, 100%) and 579 ((M + H)+, 16%); 13C NMR (CDCl3, set CDCl3 at δ 77.23) δ 170.44, 170.14, 170.00, 169.67, and 167.11 (5 carboxyl), 160.74, 156.90, 154.81, and 152.02 (C-2′, 4′, 7′, 9′), 125.86, 115.52, 113.54, 113.34, 105.48, and 99.99 (C-2, 3′, 5′, 6′, 8′, 10′), 69.56, 67.51, 65.99, 64.31, and 62.10 (C-4, 5, 6, 7, 8), 53.58 (OCH3), 33.16, 20.86, 20.78, 20.71, 20.18, and 18.70 (C-3 and 5 CH3);1H NMR (CDCl3) δ 7.494 (d, 1 H,J = 8.7 Hz, H-5′), 6.994 (d, 1 H, J = 2.4 Hz, H-8′), 6.956 (dd, 1 H, J = 2.4 and 8.7 Hz, H-6′), 6.210 (q, 1 H, J = 1.2 Hz, H-3′), 3.788 (s, 3 H, OCH3), 2.407 (d, 3 H, J = 1.2 Hz, CH3 at C-3′), 2.143, 2.035, 1.973, and 1.545 (each s, each 3 H, 4 COCH3). The data for sugar protons are listed in Tables I and II. MeONa/MeOH (0.5 m, 2 ml) was added to a solution of3 (1.0 g, 1.73 mmol) in MeOH (40 ml), and the mixture was stirred at 25 °C. After 1.5 h, the reaction mixture was concentrated to 10 ml and diluted with water (50 ml). The solution was adjusted to pH 11 and maintained at this pH by adding 2 mNaOH. TLC (CHCl3:MeOH:H2O:AcOH, 65:25:4:1) indicated the complete saponification of the methyl ester after 2 h. The solution was neutralized by Dowex 50W-X8 (H+ form) to pH 4 and filtered. The filtrate was then adjusted to pH 8 by adding 0.1 m NH4OH and evaporated. The residue was applied onto a Sephadex G-10 column (2.5 × 95 cm), which was equilibrated and eluted with 50 mm NH4OH, to obtain β-KDO-MU (5) as its ammonium salt (601 mg, 84%):R F 0.26 (CHCl3:MeOH:H2O:AcOH, 65:25:4:1); m.p. 117–120 °C; MS (m/z, fast atom bombardment-positive mode) 419 ((M + Na)+, 2%), 397 ((M + H)+, 3%), 154 (100%); 13C NMR (D2O set the external dioxane at δ 66.67): δ 173.73 (C-1), 164.85, 158.52, 156.68, and 153.95 (C-2′, 4′, 7′, 9′), 126.77, 117.83, 116.00, 111.99, 107.81, and 103.56 (C-2, 3′, 5′, 6′, 8′, 10′), 75.51, 70.21, 68.00, 66.43, and 64.86 (C-4, 5, 6, 7, 8), 36.09 (C-3), 18.70 (CH3); 1H NMR (D2O, set the internal HDO at δ 4.778): δ 7.573 (d, 1 H, J = 8.5 Hz, H-5′), 7.060 (dd, 1 H, J = 2.2 and 8.5 Hz, H-6′), 7.046 (d, 1 H, J = 2.2 Hz, H-8′), 6.114 (s, 1 H, H-3′), 2.320 (s, 3 H, CH3). The data for sugar protons are listed in Tables I and II. The de-O-acetylation and alkaline saponification of4 (200 mg, 0.346 mmol) followed by Sephadex G-10 chromatography were performed according to the procedures described for the preparation of β-KDO-MU to give α-KDO-MU (6) as its ammonium salt (130 mg, 91%): R F 0.21 (CHCl3:MeOH:H2O:AcOH, 65:25:4:1); m.p. 129–131 °C; MS (m/z, fast atom bombardment-positive mode) 419 ((M + Na)+, 2%), 397 ((M + H)+, 4%), 154 (100%); 13C NMR (D2O, set external dioxane at 66.67): δ 174.18 (C-1), 164.63, 157.73, 156.37, and 153.73 (C-2′, 4′, 7′, 9′), 126.28, 114.57, 114.51, 110.88, 104.39, and 101.18 (C-2, 3′, 5′, 6′, 8′, 10′), 72.86, 69.10, 66.23, 65.76, and 63.34 (C-4, 5, 6, 7, 8), 34.83 (C-3), 17.94 (CH3). 1H NMR (D2O, set the internal HDO at δ 4.778): δ 7.640 (d, 1 H, J = 8.8 Hz, H-5′), 7.001 (dd, 1 H, J= 2.2 and 8.8 Hz, H-6′), 6.958 (d, 1 H, J = 2.2 Hz, H-8′), 6.147 (s, 1 H, H-3′), 2.356 (s, 3 H, CH3). The data for sugar protons are listed in Tables I and II. When the KDO-cleaving activity of α-KDOase was assayed using Re-LPS or DeOA-LPS as substrate, 40 μg of LPS were incubated with an appropriate amount of enzyme in 70 μl of 50 mm sodium acetate buffer pH 4.5 at 37 °C for a predetermined time. The amount of KDO released was determined by high performance anion exchange chromatography (8Kiang J. Szu S.C. Wang L.X. Tang M. Lee Y.C. Anal. Biochem. 1997; 245: 97-101Crossref PubMed Scopus (17) Google Scholar) and/or the periodate-thiobarbituric acid method (7Unger F.M. Adv. Carbohydr. Chem. Biochem. 1981; 38: 323-388Crossref Scopus (367) Google Scholar). For detection of the free KDO released from LPS or α-KDO-MU by TLC, the reaction mixture contained 5 nmol of LPS or 20 nmol of α-KDO-MU in 40 μl of 50 mm sodium formate buffer pH 4.5 or 3.0 and an appropriate amount of the enzyme. After incubation at 37 °C for a preset time, the reaction was stopped by adding 40 μl of ethanol followed by brief centrifugation. The supernatant was evaporated to dryness using a SpeedVac, dissolved in 12 μl of methanol:water (1:2) and analyzed by TLC using chloroform:methanol: 12 mmMgCl2 (45/40/12) as the developing solvent. The plate was sprayed with diphenylamine reagent (10Harris G. MacWilliams I.C. Chem. Ind. 1954; : 249Google Scholar) and heated at 120 °C for 20 min to reveal glycoconjugates. The fluorometric assay of α-KDOase activity using α-KDO-MU as substrate was carried out according to the procedure described by Potier et al. (11Potier M. Mameli L. Bélisle M. Dallaire L. Melancon S.B. Anal. Biochem. 1979; 94: 287-296Crossref PubMed Scopus (745) Google Scholar). The enzyme was incubated with 0.5 mm α-KDO-MU in 50 mmsodium formate buffer, pH 3.0, in a total volume of 100 μl at 37 °C. After a preset time, 1.5 ml of 0.2 m sodium borate buffer, pH 9.8, was added to the reaction mixture to stop the reaction. The released MU was determined using a Sequoia-Turner Model 450 fluorometer. One unit of α-KDOase is defined as the amount of the enzyme that liberates 1 nmol of KDO per min at 37 °C. All operations were carried out at a temperature between 0 and 5 °C. Centrifugations were routinely carried out at 20,000 ×g for 30–40 min using a Sorvall RC5C refrigerated centrifuge. Ultrafiltration was carried out with an Amicon stirred cell using a PM10 membrane. The hepatopancreas (450 g) dissected from 1 gallon of fresh oysters was homogenized with 3.2 liters of cold acetone using a Polytron homogenizer, quickly filtered through a Buchner funnel, and immediately dried under vacuum to obtain 90 g of the acetone powder of oyster hepatopancreas. The acetone powder was extracted with 2.25 liters of water containing 1 mm EDTA and 1 mmphenylmethylsulfonyl fluoride as protease inhibitors using a Polytron homogenizer, followed by centrifugation. The pH of the water extract was adjusted to 4.3 with a saturated solution of citric acid, and the precipitate was removed by centrifugation. The supernatant was brought to 45% saturation with solid ammonium sulfate (277 g/liter). After standing for 2 h, the precipitate was removed by centrifugation, and the supernatant was brought to 85% saturation with ammonium sulfate (295 g/liter). After standing overnight, the precipitate was collected by centrifugation, dissolved in 40 ml of 50 mmsodium acetate buffer, pH 4.2, and applied onto a Sephacryl S-200 column (5 × 100 cm) equilibrated with 50 mm sodium acetate buffer, pH 4.2, containing 0.15 m NaCl. The column was eluted with the same buffer at 1 ml/min, and 20-ml fractions were collected. The fractions containing α-KDO-MU cleaving activity, as shown in Fig. 2 A, were pooled, concentrated to 5.0 ml, and applied onto a SP-Fractogel column (1.5 × 10 cm) equilibrated with 25 mm sodium phosphate buffer, pH 6.2. After washing with the same buffer, the column was eluted with a linear NaCl gradient from 0 to 0.5m (total volume, 170 ml). The fractions containing α-KDO-MU-cleaving activity as shown in Fig. 2 B were pooled, concentrated, and dialyzed against 25 mm sodium phosphate buffer, pH 7.0. This preparation (1.4 ml) was applied to a Con A-Sepharose column (1.0 × 5 cm) that had been equilibrated with 25 mm sodium phosphate buffer, pH 7.0. The column was washed with the same buffer; the α-KDOase activity was then eluted with 0.5 m methyl-α-mannoside. Fractions with α-KDO-MU-cleaving activity were concentrated, dialyzed against 1.5m ammonium sulfate, and applied to an octyl-Sepharose column (1.5 × 9 cm) that had been equilibrated with 1.5m ammonium sulfate. After washing with the same buffer, the column was successively eluted with 1.0 m ammonium sulfate, 0.5 m ammonium sulfate, and water. The majority of α-KDO-MU-cleaving activity was eluted with 1.0 m ammonium sulfate. This α-KDOase preparation was concentrated, dialyzed against 25 mm sodium acetate buffer, pH 5.0, and used for subsequent studies. Table IIIsummarizes the purification of α-KDOase.Table IIIPurification of α-KDOase from 450 g of the hepatopancreas of C. virginicaStepProteinTotal activitySpecific activityPurificationRecoverymgunitsunits/mg-fold%Crude extract22,000134,2006.1110040–85% (NH4)2SO41,955100,080518.475Sephacryl S20043569,4031602652SP-Fractogel5937,99064410628Con A-Sepharose16.313,30081613410Octyl-Sepharose5.88,31214332356.2 Open table in a new tab The glycosyl chloride of the per-O-acetylated methylester of KDO (2) was chosen as the glycosyl donor for the synthesis of α- and β-KDO-MU. Two procedures, which have been used for the preparation of 4-methylumbelliferyl ketoside of NeuAc, were applied for the coupling of 2 with the sodium salt of 4-methylumbelliferone, namely, the direct coupling in a dipolar aprotic solvent such as DMF (12Myers R.W. Lee R.T. Lee Y.C. Thomas G.H. Reynolds L.W. Uchida Y. Anal. Biochem. 1980; 101: 166-174Crossref PubMed Scopus (148) Google Scholar) and the catalyzed phase transfer reaction (13Rotherrnel J. Failland H. Carbohydr. Res. 1990; 196: 29-40Crossref Scopus (37) Google Scholar). In both cases, the methyl (4-methylumbelliferyl 4,5,7,8-tetra-O-acetyl-3-deoxy-β-d-manno-2-octulopyranoside)onate (3) was obtained as the major product (∼80%), together with the α-anomer (4) as the minor product (∼7%). Compounds 3 and 4 could be separated either by repeated column chromatography or by taking advantage of their differential solubility in toluene. The β-anomer 3 was more soluble in toluene than the α-anomer 4. The structures of 3 and 4 were determined by mass spectrometry and by 1H and 13C NMR spectroscopy. Although it is difficult to deduce the anomeric configuration by 1H NMR analysis, we assigned the major product as the β-anomer 3 based on the following considerations. First, it is reasonable to assume that due to the anomeric effect, compound 2 should exist predominantly in the α-anomeric configuration. Second, the coupling reaction under a given condition should occur by the usual SN2 mechanism to give the product an anomeric inversion. This assignment was confirmed by the proton-coupled 13C NMR signals for C-1 in the final products. Several attempts to improve the yield of the α-anomer 4were unsuccessful. For example, silver triflate-catalyzed reaction of2 with anhydrous 4-methylumbelliferone in CH2Cl2 in the presence of molecular sieves (MS4A) led to the formation of methyl 4,5,7,8-tetra-O-acetyl-2,6-anhydro-3-deoxy-d-manno-oct-2-enonate (14Claesson A. Luthrnan K. Acta Chem. Scand. Ser. B. 1982; 36: 719-720Crossref Google Scholar) (the glycal derivative of KDO) in 82% yield instead of the coupling product. It is noteworthy that the reaction of 2with the sodium salt of 4-methylumbelliferone in DMF in the presence of tetrabutylammonium chloride (2 molecular equivalents) did not improve the yield of 4. Therefore, the formation of the α-anomer4 did not result from in situ anomerization of the glycosyl chloride by chloride ion that was generated during the coupling reaction, but instead, the α-anomer must have come from the contaminated β-glycosyl chloride. It should be pointed out that the formation of isomeric products was not reported in the similar preparation of MU-ketoside of sialic acid (12Myers R.W. Lee R.T. Lee Y.C. Thomas G.H. Reynolds L.W. Uchida Y. Anal. Biochem. 1980; 101: 166-174Crossref PubMed Scopus (148) Google Scholar, 13Rotherrnel J. Failland H. Carbohydr. Res. 1990; 196: 29-40Crossref Scopus (37) Google Scholar). De-O-acetylation of 3 and 4, followed by alkaline hydrolysis of the methyl ester and purification by Sephadex G-10 chromatography, gave the β-anomer 5 (84%) and the α-anomer 6 (91%), respectively, as their ammonium salts. The 1H NMR spectra (Fig. 3) of 5 and 6 revealed that the sugar protons of the two anomers had distinctly different chemical surroundings. In5, the signals for H-4, 5, 6, 7, and 8 were overlapping in a narrow area ranging from δ 4.04 to 3.82; whereas in 6, the signals for each proton of 4, 5, 6, 7, 8 were completely separated in a wide range (δ 4.26–3.12). Some empirical 1H NMR rules have been used for deducing the anomeric configurations of KDO derivatives. One such rule is that the difference in chemical shift between H-3a and H-3e in β-anomer is usually bigger than that in α-anomer (9Unger F.M. Stix D. Schulz G. Carbohydr. Res. 1980; 80: 191-195Crossref Scopus (102) Google Scholar). However, these empirical rules often lead to ambiguous assignments. For example, in the present case, the difference between δ (H-3e) and δ (H-3a) in β-anomer 5 is bigger than that in α-anomer6, whereas the difference between δ (H-3e) and δ (H-3a) in β-anomer 3 is actually smaller than that in α-anomer4. This is due to the fact that the substituents could greatly influence the chemical shifts of neighboring protons. A definitive determination of the anomeric configurations of KDO derivatives could be achieved by comparison of the proton-coupled13C NMR signals of the C-l in α- and β-anomers (9Unger F.M. Stix D. Schulz G. Carbohydr. Res. 1980; 80: 191-195Crossref Scopus (102) Google Scholar). In a typical 5C2 chair conformation of KDO derivatives, the dihedral angles of (C-1)–(C-2)–(C-3)–(H-3a) in α- and β-anomers are nearly 60° and 180°, respectively. Therefore, the α-anomer would give a small value for the coupling constant between C-l and H-3a (J C-1, H-3a < 1 Hz), and the β-anomer would give a relatively large coupling constant (J C-1, H-3a = 5–6 Hz), according to the Karplus relationship (15Schwarcz J.A. Perlin A.S. Can. J. Chem. 1972; 50: 3667-3670Crossref Google Scholar). In the proton-coupled 13C NMR spectra (Fig. 4), the C-1 signal of 5appeared at δ 173.73 as a doublet (J C-1, H-3a= 5.5 Hz), whereas the C-1 signal of 6 appeared at δ 174.18 as a broad singlet (J C-1, H-3a < 1 Hz). Accordingly, 5 should be in the β-d-configuration and 6 in the α-d-configuration. This method was previously used for the determination of the anomeric configurations of sialic acid derivatives (16Hori H. Nakajima T. Nishida Y. Ohrui H. Meguro H. Tetrahedron Lett. 1988; 29: 6317-6320Crossref Scopus (134) Google Scholar). The present study shows that this assignment is equally applicable for KDO derivatives. Under mildly acidic conditions (0.2 macetate buffer, pH 4.5, 70 °C), the β-anomer was found to be much more labile than the α-anomer (Fig. 5). A similar observation was reported for the anomeric pairs of the ketosides of sialic acid, in which the equatorial aglycon was hydrolyzed much faster than the axial aglycon (17Itoh M. Shitori Y. Chem. Pharm. Bull. 1986; 34: 1479-1484Crossref Scopus (23) Google Scholar, 18Furuhata K. Anazawa K. Itoh M. Shitori Y. Ogura H. Chem. Pharm. Bull. 1986; 34: 2725-2731Crossref Scopus (24) Google Scholar). KDO-containing glycoconjugates have been found to contain both α- and β-linked KDO (7Unger F.M. Adv. Carbohydr. Chem. Biochem. 1981; 38: 323-388Crossref Scopus (367) Google Scholar). Therefore, α- and β-KDO-MU should be useful for studying α- and β-KDOases, and the availability of these two enzymes will facilitate the studies of the structure and function of KDO-containing glycoconjugates. Despite the medical importance of LPSs, virtually nothing is known about their degradation. The enzyme (α-KDOase) that cleaves α-linked KDO from LPSs has never been reported. The hepatopancreas of the oyster, C. virginica, was found to be rich in various glycoconjugate-cleaving enzymes. Using the periodate-thiobarbituric acid reaction (7Unger F.M. Adv. Carbohydr. Chem. Biochem. 1981; 38: 323-388Crossref Scopus (367) Google Scholar) and TLC, the crude extract of oyster hepatopancreas was found to liberate KDO from Re-LPS prepared from E. coli and S. minnesota. Since KDO residues in LPSs have been shown to be α-ketosidically linked (19Strain S.M. Fesik S.W. Armitage I.M. J. Biol. Chem. 1983; 258: 13466-13477Abstract Full Text PDF PubMed Google Scholar, 20Strain S.M. Fesik S.W. Armitage I.M. J. Biol. Chem. 1983; 258: 2906-2910Abstract Full Text PDF PubMed Google Scholar), we used the synthesized α-KDO-MU as substrate to follow the enzyme activity to purify α-KDOase from the crude extract of oyster hepatopancreas. Before α-KDO-MU was synthesized, we used Re-LPS as substrate, and the liberated KDO was determined by the periodate-thiobarbituric acid reaction. Initially, this method was used to monitor the purification of α-KDOase during Sephacryl-S-200 gel filtration. We subsequently found that in the column fractions, the enzyme activity detected by the Re-LPS/periodate-thiobarbituric acid reaction coincided well with that detected using α-KDO-MU as substrate. The availability of α-KDO-MU greatly facilitated the purification of α-KDOase. The oyster α-KDOase was found to be stable in the pH range of 3–8. Interestingly, the pH optima for this enzyme using the synthetic and the natural substrates were found to be significantly different. The pH optimum of this enzyme was determined to be 4.5 for releasing KDO from Re-LPS, whereas that for the hydrolysis of α-KDO-MU was 3.0 (Fig.6). Thus, the optimal pH for the hydrolysis of KDO from KDO-containing glycoconjugates cannot be inferred from the hydrolysis of α-KDO-MU. The time courses of the liberation of KDO from DeOA-LPS by α-KDOase and by acid hydrolysis are compared in Fig.7. The oyster α-KDOase was able to completely detach the KDO from DeOA-LPS. The amount of KDO released by the enzyme was found to be comparable to that liberated by acid hydrolysis. Fig. 8 shows the TLC analysis of the release of KDO from DeOA-LPS and α-KDO-MU. α-KDOase isolated from oyster hepatopancreas was able to cleave KDO efficiently from all LPS substrates tested, including Re-LPS from E. coli, Re-LPS from S. minnesota R595, Rd-LPS from S. minnesotaR7, and DeOA-LPS. However, β-KDO-MU was found to be refractory to this enzyme. Thus, the specificity of α-KDOase also supports the assignment of the anomeric configurations of the two KDO-MU anomers. The oyster α-KDOase represents the first α-KDO-cleaving enzyme, and the presence of such an enzyme in the hepatopancreas of oyster may suggest the wide occurrence of KDO in marine organisms. α-KDOase capable of liberating KDO from LPSs should be important and useful for studying the structure and function of LPSs and other α-KDO-containing glycoconjugates.Figure 8TLC analysis showing the liberation of KDO from DeOA-LPS and α-KDO-MU by oyster α-KDOase. Lane 1,α-KDOase; lane 2, DeOA-LPS; lane 3, DeOA-LPS + α-KDOase; lane 4, KDO; lane 5, α-KDO-MU + α-KDOase; lane 6, α-KDO-MU. In all cases, 1 unit of the enzyme was used. For the hydrolysis of DeOA-LPS, the incubation was carried out at pH 4.5 for 1 h, whereas for cleaving α-KDO-MU, the incubation was carried out at pH 3.0 for 15 min under the assay conditions described under “Experimental Procedures.”View Large Image Figure ViewerDownload Hi-res image Download (PPT) The authors thank Dr. S. C. Szu for a sample of De-OAc-LPS from Re-LPS and Mei Tang for analysis of KDO by high performance anion exchange chromatography.