Nancy Barnabe

NAD-linked formate dehydrogenase from methanol-grown Pichia pastoris NRRL-Y-7556

Abstract An NAD-linked formate dehydrogenase (EC 1.2.1.2.) from methanol-grown Pichia pastoris NRRL Y-7556 has been purified. The purification procedure involved ammonium sulfate fractionation, hollow-fiber H1P10 filtration, ion-exchange chromatography, and gel filtration. Both dithiothreitol (10 m m ) and glycerol (10%) were required for stability of the enzyme during purification. The final enzyme preparation was homogeneous as judged by polyacrylamide gel electrophoresis and by sedimentation pattern in an ultracentrifuge. The enzyme has a molecular weight of 94,000 and consists of two subunits of identical molecular weight. Formate dehydrogenase catalyzes specifically the oxidation of formate. No other compounds tested can replace NAD as the electron acceptor. The Michaelis constants were 0.14 m m for NAD and 16 m m for formate (pH 7.0, 25 °C). Optimum pH and temperature for formate dehydrogenase activity were around 6.5–7.5 and 20–25 °C, respectively. Amino acid composition of the enzyme was also studied. Antisera prepared against the purified enzyme from P. pastoris NRRL Y-7556 form precipitin bands with isofunctional enzymes from different strains of methanol-grown yeasts, but not bacteria, on immunodiffusion plates. Immunoglobulin fraction prepared against the enzyme from yeast strain Y-7556 inhibits the catalytic activity of the isofunctional enzymes from different strains of methanol-grown yeasts.

Identification and purification of a nicotinamide adenine dinucleotide-dependent secondary alcohol dehydrogenase from C1-utilizing microbes

Phenazine methosulfate (PMS)-dependent methanol dehydrogenase has been reported from many methylotrophic bacteria [l-4]. This enzyme oxidizes primary alcohols from Cr-C,, but does not oxidize secondary alcohols. Nicotinamide adenine dinucleotide (NAD)dependent alcohol dehydrogenases have been reported from liver and yeast [5]. These alcohol dehydrogenases oxidize primary alcohols and acetaldehyde but have no activity on methanol. In addition, the alcohol dehydrogenases from yeast and liver also oxidize some secondary alcohols at a very low rate (<l% of their ethanol activity). NAD(P)-dependent alcohol dehydrogenases were also reported inPseudomonas [6,7], Escherichiu coli [8] and Leuconostoc [9]. However, these enzymes were active only on long-chain primary alcohols or hydroxy fatty acids [7]. To our knowledge, no secondary alcohol-specific alcohol dehydrogenase (SADH) has been reported. We have recently identified an NAD-linked, secondary alcohol-specific, alcohol dehydrogenase in cell-free extracts of various gaseous hydrocarbon-utilizing microbes. This enzyme is also found in cells grown on methanol. It specifically and stoichiometrically oxidizes secondary alcohols to their corresponding methyl ketones. This enzyme has been purified 2600-fold and shows a single protein band on acrylamide gel electrophoresis.

Biodegradation of xanthan by salt-tolerant aerobic microorganisms

SummaryThree salt-tolerant bacteria which degraded xanthan were isolated from various water and soil samples collected from New Jersey, Illinois, and Louisiana. The mixed culture, HD1, contained aBacillus sp. which produced an inducible enzyme(s) having the highest extracellular xanthan-degrading activity found. Xanthan alone induced the observed xanthan-degrading activity. The optimum pH and temperature for cell growth were 5–7 and 30–35°C, respectively. The optimum temperature for activity of the xanthan-degrading enzyme(s) was 35–45°C, slightly higher than the optimum growth temperature. With a cell-free enzyme preparation, the optimum pH for the reduction of solution viscosity and for the release of reducing sugar groups were different (5 and 6, respectively), suggesting the involvement of more than one enzyme for these two reactions. Products of enzymatic xanthan degradation were identified as glucose, glucuronic acid, mannose, pyruvated mannose, acetylated mannose and unidentified oligo- and polysaccharides. The weight average molecular weight of xanthan samples shifted from 6.5·106 down to 6.0·104 during 18 h of incubation with the cell-free crude enzymes. The activity of the xanthan-degrading enzyme(s) was not influenced by the presence or absence of air or by the presence of Na2S2O4 and low levels of biocides such as formaldehyde (25 ppm) and 2,2-dibromo-3-nitrilopropionamide (10 ppm). Formaldehyde at 50 ppm effectively inhibited growth of the xanthan degraders.

Purification and properties of a NAD-linked 1,2-propanediol dehydrogenase from propane-grown Pseudomonas fluorescens NRRL B-1244

NAD-dependent 1,2-propanediol dehydrogenase (EC 1.1.1.4) activity was detected in cell-free crude extracts of various propane-grown bacteria. The enzyme activity was much lower in 1-propanol-grown cells than in propane-grown cells of Pseudomonas fluorescens NRRL B-1244, indicating that the enzyme may be inducible by metabolites of propane subterminal oxidation. 1,2-Propanediol dehydrogenase was purified from propane-grown Ps. fluorescens NRRL B-1244. The purified enzyme fraction shows a single-protein band upon acrylamide gel electrophoresis and has a molecular weight of 760,000. It consists of 10 subunits of identical molecular weight (77,600). It oxidizes diols that possess either two adjacent hydroxy groups, or a hydroxy group with an adjacent carbonyl group. Primary and secondary alcohols are not oxidized. The pH and temperature optima for 1,2-propanediol dehydrogenase are 8.5 and 20-25 degrees C, respectively. The activation energy calculated is 5.76 kcal/mol. 1,2-Propanediol dehydrogenase does not catalyze the reduction of acetol or acetoin in the presence of NADH (reverse reaction). The Km values at 25 degrees C, pH 7.0, buffer solution for 1,2-propan1,2-propanediol dehydrogenase are 8.5 and 20-25 degrees C, respectively. The activation energy calculated is 5.76 kcal/mol. 1,2-Propanediol dehydrogenase does not catalyze the reduction of acetol or acetoin in the presence of NADH (reverse reaction). The Km values at 25 degrees C, pH 7.0, buffer solution for 1,2-propan1,2-propanediol dehydrogenase are 8.5 and 20-25 degrees C, respectively. The activation energy calculated is 5.76 kcal/mol. 1,2-Propanediol dehydrogenase does not catalyze the reduction of acetol or acetoin in the presence of NADH (reverse reaction). The Km values at 25 degrees C, pH 7.0, buffer solution for 1,2-propanediol and NAD are 2 X 10(-2) and 9 X 10(-5) M, respectively. The 1,2-propanediol dehydrogenase activity was inhibited by strong thiol reagents, but not by metal-chelating agents. The amino acid composition of the purified enzyme was determined. Antisera prepared against purified 1,2-propanediol dehydrogenase from propane-grown Ps. fluorescens NRRL B-1244 formed homologous precipitin bands with isofunctional enzymes derived from propane-grown Arthrobacter sp. NRRL B-11315, Nocardia paraffinica ATCC 21198, and Mycobacterium sp. P2y, but not from propane-grown Pseudomonas multivorans ATCC 17616 and Brevibacterium sp. ATCC 14649, or 1-propanol-grown Ps. fluorescens NRRL B-1244. Isofunctional enzymes derived from methane-grown methylotrophs also showed different immunological and catalytic properties.

Xanthan‐Degrading Enzymes

Xanthan is an anionic extracellular polysaccharide produced by Xanthomonas campestris NRRL B-1459.’ The high viscosity of this polymer solution is relatively insensitive to temperature, ionic strength, shear, and pH. For this reason, xanthan finds commercial use as a viscosity-enhancing agent for aqueous solutions. The primary structure of xanthan was established by Jansson et al.‘ It consists of a main chain of /3-1,4-linked D-glucose units, as in cellulose, but with a three-sugar side chain attached to alternate glucose residues. Xanthan is inert to attack by microbes or by presently available enzymes. Rinaldo and Milas6 were the first to show partial hydrolysis of xanthan by cellulase only in the absence of salt, where xanthan is in an unordered conformation. More recently, Cadmus et al.‘ reported biodegradation of xanthan by a Bacillus sp. in the presence of salt. The “xanthanase” they obtained was a mixture of enzymes that attacked all of the side chain linkages in the xanthan molecule, including the one involving ( I 3)-linkage of acetylated mannose to the glucosidic backbone. They found no endocellulase type of activity in their cultures. Sutherlandg described an enzyme system hydrolyzing the polysaccharides of xanthomonas species. The /3-glucanohydrolase hydrolyzed both p1.3and @-1,4-linked polymers with side-chains or other substituents. In this paper, we describe the isolation of a salt-tolerant xanthan-utilizing culture HD1 and the purification and properties of a novel xanthan depolymerase.