w“ .3 ‘ LHiR‘AR y MIChlgan Sta” OVERDUE F INES: 25¢ per day per item RETURNING LIBRARY MATERIALS : Place in book return to remove charge from circulation records © 1981 CHARLES LEE HAUSWALD All Rights Reserved MUTATIONAL ACQUISITION OF O-FUCONATE CATABOLISM IN KLEBSIELLA PNEUMONIAE: ELUCIDATION OF THE PATHWAY AND CHARACTERIZATION OF D-GALACTONATE (D-FUCONATE) DEHYDRATASE By Charles Lee Hauswald A DISSERTATION Submitted to . Michigan State University in partial fulfillment of the requirements for the degree of MASTER OF SCIENCE Department of Biochemistry 1980 ABSTRACT MUTATIONAL ACQUISITION OF D-FUCDNATE CATABDLISM IN KLEBSIELLA PNEUMDNIAE: ELUCIDATION OF THE PATHWAY AND CHARACTERIZATION OF D-GALACTDNATE (D-FUCONATE) DEHYDRATASE By Charles Lee Hauswald The genetic and enzymatic basis for the acquisition of the ability of Klebsiella pneumoniae to use the xenobiotic D-fuconate as a sole carbon source was determined. A single mutation resulted in the con- stitutive production of D-galactonate (D-fuconate) dehydratase, a previouSly undescribed enzyme which is induced in the parental strain by D-galactonate but not by D-fuconate. The dehydratase product, 2-keto-3-. deoxy-D-fuconate (KDF), then induces a new class II aldolase, which cleaves KDF to pyruvate and D-lactaldehyde. This aldolase has no other known inducer. A mutant missing KDF aldolase failed to grow only on Defuconate (out‘of 24 compounds tested that supported parental-strain growth), whereas a mutant missing the dehydratase failed to grow on both D-fuconate and D-galactonate. D-Galactonate (D-fuconate) dehydratase (four percent of the cellular protein) was purified to electrophoretic homogeneity and characterized with respect to substrate specificity, kinetic constants, pH optimum, metal requirement, molecular weight, isoelectric point, and amino acid composition. ACKNOWLEDGEMENTS I acknowledge the invaluable assistance of Dr. Richard L. Anderson, my graduate advisor, for the helpful thought that went into the preparation of this thesis. In addition, I would like to thank the members of my guidance committee, Dr. Willis A. Wood, Dr. James L. Fairley, and the late Dr. John A. Boezi, who helped in the preparation of this manuscript. Finally, I would like to to give special notice to Dr. Arnold Revzin and Doris Bauer for their enthusiastic technical aid in the planning of experiments involving the use of the Beckman Model E Ultracentrifuge and the Beckman, model 121-3, automated amino acid analyzer, respectively. This thesis is dedicated to my dear parents, Charles A. and Bernice D. Hauswald, who gave both understanding and encouragement over the years and who now embark on a new journey of life, as do I. TABLE OF CONTENTS Page LIST OF TABLES ......................... . vii LIST OF FIGURES . . . ...................... ix LIST OF ABBREVIATIONS . ....... . ............. xii GENERAL INTRODUCTION. . . . . . . . . . . . . . . . . . . . . . . 1 LITERATURE REVIEW REGARDING GAIN MUTATIDNS, SUGAR ACID CATABDLIC PATHWAYS IN MICROORGANISMS, AND STRATEGIES FOR SUGAR CATABDLISM IN THE ENTERDBACTERIACEAE . . . . . . . . . . . . . . . . . . . . SECTION I ELUCIDATIDN OF THE METABOLIC FATE OF D-FUCDNATE INTRODUCTION. 0 I O O O O O O O O O O O O ...... O ..... 16 MATERIALS Am METHODS O O O O O O O O O O O O O O O O O O O O O O 17 Bacterial Strain, Cell Growth, and Preparation of Cell-Free ExtraCts O O O O O O C C O O O C O O C O C O C O O O 0 ~ 0 O 17 Bacterial Strain . . . . . . . . . . . . . . . . . . . . 17 "Edi a. O C O I O C O O O O O O O C O O O I O 17 C811 Growth. . . . . . . . . . . . . . ....... . . 17 Monitoring Cell Growth . ...... . . . . . . . . . . 13 Harvesting of Cells. . . . . . . . . . ...... . . 13 Centrifugation . . . . . . . . . . . . . . . . . . . 13 Preparation of Cell -Free Extracts. . . . . . . . . . . . 18 Protein Determination. . . . . . . . . . . . . . . . . 19 Gas-Liquid Chromatography. . . . . . . . . . . . . ..... 19 Preparation of Ion Exchange Resins . . . . . . . . . . . 20 Colorimetric Assays. . . . . . . . . . . . . . . . . . . 20 Reducing Sugar Assay . . . . . . . . . . . . . . . . 21 Aldonic Acid Assay . . . . . . . . . . . . . . . . . . 21 Aldehyde Colorimetric Assays. . . . . . . . . . . . . . 21 Alpha-Keto Acid Determination. . . . . . . . . . . 21 2- -Keto-3-Deoxy Sugar Acid Determination. . . . . . . . . 22 Preparation of Substrates. . . . . . . . . . . . . . . . . . 22 Aldonic Acid Synthesis . . . . . . . .......... 22 Preparation of Oxaloacetic Acid. ..... . ..... . 24 Synthesis of D-Lactaldehyde ............... Chemical Synthesis of 2-Keto-3-Deoxy-D-Fuconate ..... (i) Portsmouth Synthesis ................ (i i) Dahms Synthesis . . . ..... . . . ...... (iii) Summary of the Synthetic Methods . . ....... (iv) Alkaline Lability of the Portsmouth Synthetic Products .......... . . . . Preparation of Other 2- Keto- 3- -Deoxy Sugar Acids ..... Enzymatic Assays . . . . . . . . . . . . . . ..... Dehydrogenase Assays . . . . . . . . . . . . . . D-Galactonate (D-Fuconate) Dehydratase Assay . . . . . 2-Keto-3-Deoxy-D-Galactonate Kinase Assay. . . . . . Z-Keto-B-Deoxy-6-Phospho-D-Galactonate Aldolase Assay. 2-Keto-3-Deoxy-D-Fuconate Aldolase Assay . . . . . . End-Point Assays for Pyruvate, D-Lactaldehyde, and Oxaloacetic acid . . . . . . . . . . . . . . . . Selection of D-Fuconate and D-Galactonate-Negative Mutants Revertants for-the Dehydratase-Negative Mutant . . . . . IMViC Tests for the Bacteriological Classification of all strai ns. C O O O O O I O I O O C O O O O O O O O O 0 sources of Material 5 O O O O O O O O O O O O O 0 O O O 0 RESULTS 0 C O O O O O O O O O O O O O O O O O O O O O O O O 0 Selection of Mutant Strain CH-lOl (D-FuconateaPositive) from Klebsiella Pneumoniae PRL-R3, U' and Gross Characterization . . . . . . . . . . . . . . . . . Elucidation of the Enzymatic Reactions Involved in the Catabolism of D- Fuconate . . . . . . . . . . . . . . 'Enzymatic Reactivity of D-Fuconate . . . . . . . . . Enzymatic Reactivity of 2- Keto-B-Deoxy-D- Fuconate. . Inducibility of the D-Fuconate and D-Galactonate Pathway Enzymes on Various Carbohydrates . . . . . . . . . . Mutant Analysis and Verification of Enzyme Deficiency. . Procurement of Revertants for Growth on D-Galactonate from Strai n CH‘103 O O O O O O O O O O O O O O 0 Classification of the Bacterial Strains. . . . . . . DISCUSSION ' ........................... SECTION II PURIFICATION AM) CHARACTERIZATION OF D—GALACTONATE (D-FUCONATE) DEHYDRATASE INTRODUCTION. . . . . . . . . . . . . . . . . . . . . . . . . . MATERIALS AND METHODS . . . . . . . . . . . ........... Bacterial Strain and Culture Conditions. . . . . . . . . . Bacterial Strain . . . . . . . . . . . . . . . . . . Medi um O O I O O 0 O O O O O O O ..... O 0 O O O O 0 iv 49 49 57 57 59 67 73 80 81 86 92 94 94 94 94 Page Starter Culture Preparation ............... 94 Cell Growth ....................... 95 Harvesting Cell Culture ............. . . . . 95 Protein Determination. . . . . . . . . ........... 95 D-Galactonate (D-Fuconate) Dehydratase Assay ........ 96 Enzyme Purification Procedure. . . . . . . . ....... . 96 DEAE-Cellulose Chromatography. . ............ 96 Sepharose A- SH Chromatography. . . . . . . . . . . . . . 97 Hydroxyapatite Chromatography. . . . . . . 97 Sephadex G- 200 and Bio-Gel P- 300 Chromatography. . . . 98 Pressure Dialysis Concentration of Protein . . . . . 98 Conductivity Measurements. . . . . . . . . . . . . . . . 99 Polyacrylamide Gel Electrophoresis . . . . . . . . . . . . 99 General Procedures . . . . . . . . . . . . . . . . . 99 Preparation of Native Gels . . . . . . . . . . . . . . . 100 Preparation of SDS Gels. . . . . . . . . . . . . . . . . 102 Isoelectric Focusing . . . . . . . . . . . . . . . . . . . . 103 Enzymatic Assays for Molecular Height Standards. . . . . . . 104 Calibration Standards. . . . . . . . . . . . . . . . . . 104 Lactate Dehydrogenase. . . . . . . . . . . . . . . . . . 104 Pyruvate Kinase. . . . . . . . . . . . . . . . . . . . . 104 Alkaline Phosphatase . . . . . . . . . . . . . . . . . . 104 Catalase . . . . . . . . . . . . . . . . . . .“. . . . . 105 Fumarase . . . . . . . . . . . . . . . . . . . 105 Methods for the Determination of Molecular Height. . . . . . 105 Sucrose Density Gradient Centrifugation. . . . . . . . 105 Sedimentation Velocity and Sedimentation Equilibrium Analysis . . . . . . . . . . . . . . . . . . . . . . 106 (i) Sedimentation Velocity . . . . . . . . . . . . . . . 106 (ii) Sedimentation Equilibrium . . . . . . . . . . . . . 105 Amino Acid Composition Determination . . . . . . . . . . . . 107 Sources of Materials . . . . . . . . . . . . . . . . . . 107 RESULTS Purification of D-Galactonate (D-Fuconate) Dehydratase . . . 109 Cell Extract . . . . . . . . . . . . . . . . . . . . . . 109 Protamine Sulfate Fractionation. . . . . . . . . . . . . 111 Ammonium Sulfate Precipitation . . . . . . . . . . . . . 111 DEAE-Cellulose Chromatography I. . . . . . . . . . . . . 111 Sepharose A-5M Chromatography. . . . . . . . . . . . . . 114 Hydroxyapatite Chromatography. . . . . . . . . . . . . . 114 DEAE-Cellulose Chromatography II . . . . . . . . . . . . 114 Sephadex G-ZDO Chromatography. . . . . . . . . . . . 119 Determination of Purity of the Dehydratase Preparation . . . 113 $05 Polyacrylamide Gel ElectrOphoresis . . . . . . Native Gel Electrophoresis to Show Co-Migration of Protein with Dehydratase Activity. . . . . . . . . 128 Characterization of D-Galactonate (D-Fuconate) Dehydratase . 133 Stability. . . . . . . . . . . . . . . . . . . . . . . . 133 pH Optimum . . . . . . . . . . . . . . . . . . . . . . . 133 Substrate Specificity and Kinetic Constants. . . . 133 Divalent Cation Activation in the Presence of EDTA. : . 140 Page Isoelectric Point ................... 145 Molecular weight Determination. . . .......... 145 (i) Analytical Gel Filtration ............. 145 (ii) Sucrose Density Gradient Sedimentation ...... 152 (iii) Sedimentation Velocity. . . . . . . ....... 152 (iv) Sedimentation Equilibrium. . . . . . . . . . . . . 157 Subuni.t Molecular Weight. . . . . . . . 150 Summary of Both Native and Subunit Molecular Weight Determinations. . . . . . . . . . . . . 150 Amino Acid Composition of the Dehydratase . . . . . . . 150 Preparation and Identification of the Products of the Dehydratase-Catalyzed Reaction. . . . . . . . . . . . . 172 Enzymatic Preparation of the Dehydratase Products . . . 172 Identification of the Products of the Dehydratase- Catalyzed Reaction. . . . . . . . . . . . . . . . . 180 (1) Formation of Characteristic Chromogens. . . . . . . 180 (ii) Co-Chromatography of Isomeric Mixtures . . . . . . 133 (iii) Periodate Consumption and the Rates of Periodate Oxidation . . . . . . . . . . . -,- . . . . . . . . 189 (iv) Confirmation of the Biological Activity of the Purified Dehydratase Product. . . . . . . . . . 192 Summary of the Product Identification Studies . . . . . 194 DISCUSSION . . . . . . . . . . . . . . . . . . . . . . . . . . . 195 REFERENCES . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 vi 11. 12. 13. 14. 15. 16. LIST OF TABLES Retention times for carbohydrates by gas-liquid ChromatographyI I I I I I I I I I I I I I I I I I I I I I Growth yield of strain CH-IDI on D-glucose, D-galactonate, Page and D‘fuconateI I I I I I I I I I I I I I I I I I I I I I I KDF aldolase reaction products: measurement of the reaction products with appropriate coupling enzymes. . . . . . . . . Effect of MgClz on aldolase activities in EDTA-treated extracts. I I I I I I I I I I I I I I I I I I I I I I I I I Additive enzymatic activity studies on keto-deoxy sugar mi xtures With KDF aldOI aSEI I I I I I I I I I I I I I I I I Inducibilities for enzymes of the D-fuconate and D-ga13ctonate pathways. 0 o o o o o o o o o .0 o o o o o o 0 Screening of mutagenized cultures for growth on D-fuconate andD‘galaCtonateooooooooooooooooooooo Mutant screening on various carbohydrates . . . . . . . Enzyme activities in mutant and revertant studies . . . Genealogy and phenotype of the Klebsiella pneumoniae Strai "S I I I I I I I I I I I I I I I I I I I I I I I I Growth studies of the Klebsiella pneumoniae PRL-R3,U‘ and derived mutant strains. . . . . . . . . . . . . . . Standard IMViC tests for the Klebsiella pneumoniae PRL-R3,U‘- Strain am derivm strainSI I I I I I I I I I I I I I I Purification of D-galactonate (D-fuconate) dehydratase. Effect of dialysis of the dehydratase against EDTA. . . Metal ion activation of the dehydratase in the presence 0f EDTA I I I I I I I I I I I I I I I I I I I I I I I I Summary data for the molecular weight of D-galactonate (D-fuconate) dehydratase. . . . . . . . . . . . . . . . vii 25 61 79 83 B4 110 145 147 . 165 Table 17. 18. I9. 20. 21. Amino acid composition of the dehydratase following acid hYdr01y5i50 o o o o o o o o o o o o o o o o o ooooooo 168 Total amino acid composition of the D-galactonate (D-fuconate) dehydratase. . . . . . . . . . . . . ..... 171 Recoveries of chromatographic mixtures used to identify the D-fuconate dehydratase product. . . . . . . . . . . . . . . 188 Periodate uptake of the pentonic acids produced by hydrogen peroxide treatment of the D-fuconate product and chemically synthesized isomers . . . . . . . . . . . . . . . . . . . . 191 Relative rates of periodate oxidation of the dehydratase product and synthetic isomers . . . . . . . . . . . . . . . 193 viii Figure 10. 11. 12. 13. 14. 15. LIST OF FIGURES Page Elution profile for 3,6-dideoxy-D-hexulosonic acids from the Portsmouth synthesis ............... 29 Elution profile for 3,6-dideoxy-D-hexulosonic acids from the Dahms synthesis ................. 33 Alkaline lability of the chemically synthesized isomers of the Portsmouth synthesis ............... 33 Growth of the Klebsiella pneumoniae PRL-R3,U' (parental) strain on D-glucose, Dlgalactonate, and D-fuconate . . . . 51 Growth of the D-fuconate-positive mutant, strain CH-IDI, on D-glucose, D—galactonate, and D-fuconate ....... 53 Linearity of the D-fuconate dehydratase assay with respect to time. . . . .................. 58 Proportionality of the D-fuconate dehydratase activity with protein concentration . . . ........ . . . 53 pH optimum of the KDF aldolase .............. 52 Metal ion requirement of the KDF aldolase ........ 64 Linearity of the KDF aldolase assay with respect to time . 55 Proportionality of the KDF aldolase activity with protein concentration ......... . ......... 56 The D-fuconate pathway as elucidated in this study . . . . 87 DEAE-cellulose chromatography I. . . . . ...... . . . 112 'Sepharose A-SM chromatography of the DEAE-cellulose I fractions. 0 o o o o o o o _o o o o oooooooooooo 115 Hydroxyapatite chromatography of the combined Sepharose A-SM pooled fractions. . ................. 117 ix Figure 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 22. 28. 29. DEAE-cellulose II chromatography of the hydroxyapatite pooled fractions .................... Page 120 Sephadex G-ZOO chromatography of the pooled DEAE-cellulose II step fractions ................... SDS polyacr lamide gel electrophoresis of D-galactonate (D-fuconate dehydratase in the presence of molecular weight standards .................... SDS polyacr lamide gel electrophoresis of D~galactonate (D-fuconate dehydratase for the determination of Purity ......................... Native polyacrylamide gel electrophoresis of D-galac- tonate (D-fuconate) dehydratase for the determination of pur'i ty ....................... Plot of dyed protein absorbence and dehydratase activity versus gel length and slice number on a native polyacrylamide gel ................... Effect of pH and buffer composition on D-galactonate (D-fuconate) dehydratase activity . ; . . ;‘ ...... Lineweaver-Burk plot using D-fuconate as substrate in the dehydratase reaction mixture ------------ Lineweaver-Burk plot using D-galactonate as substrate in the dehydratase reaction mixture .......... Effect of EDTA on the dehydratase in the absence of metal in the enzymatic reaction mixture ........ Effect of EDTA on the dehydratase rate parameters in the presence of MgCl2 Elution profile of pH and dehydratase activity from a 110.0-ml volume isoelectric focusing column ----- Elution profile of molecular weight standards and D-galactonate (D-fuconate) dehydratase as chromatographed on P-300 ---------------- Bio-Gel P-300 chromatography of D-galactonate (D-fuconate) dehydratase with molecular weight stan- dards: plot of molecular weight versus corresponding peak fraction number .................. 122 124 126 129 131 134 136 138 141 143 148 150 153 Figure 30. 31. 32. 33. 34. 3'5. 36. 37. 38. 39. Sucrose density sedimentation of the D-galactonate (D-fuconate) dehydratase in the presence of marker proteins ........................ Sedimentation velocity determination of the sedimentation constant for the D-galactonate (D-fuconate) dehydratase ................ Sedimentation equilibrium Log C versus r2 Determination of the subunit molecular weight of D-galactonate (D-fuconate) dehydratase by SDS polyacrylamide gel electrophoresis ........... Time course study for the enzymatic preparation of the dehydratase product from D-fuconate and D-galactonate ..................... Purification of the D-fuconate dehydratase product. . . . Purification of the D-galactonate dehydratase product . . Absorption spectra of semicarbazone derivatives of the purified dehydratase products ........... Absorption spectra of the thiobarbituric acid assay (TBA) chromogen of the purified dehydratase products. . . Chromatographic identification of the D-fuconate dehydratase product .................. xi plot ..... Page 155 158 161 163 174 176 178 181 184 186 MN ATP ADP PEP NAD+ NADH NADP+ NADPH mg u9 ml 111 M 9 11111 LIST OF ABBREVIATIONS molecular weight adenosine 5'-triphosphate adenosine 5'-diphoSphate Phosphoenol pyruvate nicotinamide adenine dinucleotide reduced nicotinamide adenine dinucleotide nicotinamide adenine dinucleotide phosphate reduced nicotinamide adenine dinucleotide phosphate acceleration of gravity - gram milligram microgram liter milliliter microliter molar millimolar micromolar centimeter millimeter micrometer nanometer xii EDTA DEAE- TEMED Tris HEPES PIPES SDS TBA asKeto B-formyl pyruvate KDF KDQ KDGal D,L-KDA KDPGal ethylenediamine tetraacetic acid diethylaminoethyl- N,N,N‘,N'-tetramethylethylenediamine tris (hydroxymethyl) aminomethane N-Z-hydroxyethylpiperazine-N'-2-ethane-sulfonic acid 1,4-piperazinediethane sulfonic acid absorbance sedimentation coefficient (Svedberg; 1 x 10'13 second) sodium dodecyl sulfate thiobarbituric acid alpha-keto (2-keto) beta-formyl pyruvate (3-formyl pyruvate) 2-keto-3-deoxy-D-fuconic acid 2-keto-3-deoxy-D-quinovonic acid Z-keto-3-deoxy-D-galactonic acid Z-keto-B-deoxy-D,L-arabonic acid 2-keto-3-deoxy-6-phospho—D-galactonic acid xiii GENERAL INTRODUCTION The purpose of this work was to determine the basis for the mutational acquisition of the ability of Klebsiella pneumoniae to use D-fuconate (6-deoxy-D-galactonate) as the sole carbon and energy source for growth. The fact that D-fuconic acid apparently does not exist in Nature outside of serving as an intermediate of D-fucose metabolism in a pseudomonad, coupled with the inability of Klebsiella species to use the aldose D-fucose as a growth substrate, would indicate that D-fuconate catabolism in Klebsiella could serve as a model for a study of the degradation of xenobiotics (compounds that are rarely, if ever, found in Nature). As will be pointed out in the Literature Review, the ability of a microorganism to utilize a given xenobiotic is often dependent upon ' mutation and selection. In most cases, the mutations are found not to result in the formation of new enzymes, but rather in alterations in the inducibility of an enzyme. In the present case, the mutation resulted in the constitutive production of a dehydratase normally induced by D-galactonate but not by D-fuconate. The product of the dehydratase reaction, Zéketo-B-deoxy-D-fuconate, induces the formation of an aldolase which catalyzes the cleavage of the intermedate to pyruvate and D-lactaldehyde. Curiously, the aldolase has no known inducer other than this intermediate of the xenobiotic catabolism. This thesis is divided into two sections. Section I deals with the isolation of a D-fuconate-positive mutant of 5, pneumoniae, elucidation of the pathway of D-fuconate catabolism, and determination of the genetic and enzymatic basis for the acquired function. Section II deals with the purification and characterization of a previously undescribed enzyme, D-galactonate (D-fuconate) dehydratase, which is the passkey in the gained ability to use D-fuconate. LITERATURE REVIEW REGARDING GAIN MUTATIONS, SUGAR ACID CATA- BOLIC PATHWAYS IN MICROORGANISMS, AND STRATEGIES FOR SUGAR CATABOLISM IN THE ENTEROBACTERIACEAE' Gain mutations The constitutive production of enzymes for which novel compounds serve as fortuitous substrates is one of several biochemical bases by which gain mutations augment the growth of a bacterium on the novel. compounds. Examples of this are the utilization of L-arabitol via ribitiol dehydrogenase (1), D-arabinose and L-xylose via L-fucose isomerase (2), mannitol via D-arabitol dehydrogenase (3), and B-glycerol phosphate via alkaline phosphatase (4). A more complex situation involv- ing this type of mutation is the xylitol pathway in Aerobacter aerogenes (1.5.6). A single mutation leading to the derepression of ribitol dehydr- ogenase enables this bacterium to use xylitol as a fortuitous substrate but not its inducer (1,5). Through further mutation, a derivative strain with an improved rate of growth on xylitol was obtained from the first mutant made possible by the production of an altered ribitol dehydrogen- aSe (6). This type of structural-gene mutation has been shown to oper- ate in other instances which augmented growth of a bacterium on a novel compound (9). In addition, a third mutant with an even faster growth rate on xylitol was obtained. This strain constitutively produced an active transport system that accepted xylitol as well as D-arabitol (6). and produced the same altered ribitol dehydrogenase as seen in the second mutant. Rigby et al. (7) have shown that continuous culturing of the 3 constitutive strain, using xylitol as the sole carbon source, evolves faster growing strains which show elevated levels of ribitol dehydrog- enase activity. Ribitol dehydrogenase has been observed to comprise up to 25% of the cell protein. Through examination of these strains with respect to growth on xylitol, kinetic parameters of the purified ribitol dehydrogenase, and mutation frequency for ribitol dehydrogenase, these authors concluded that the increased activity was due to duplica- tion or amplification of the §gh.gene. Such amplification occurs at frequencies much higher than mutational alteration of substrate affin- ity, and this allows the bacterial population to adapt to short-term environmental stress without altering the genetic constitution (7,8). The acquisition of a catabolic pathway for L-1.2-propanediol by mutant strains of Escherichia coli K-12 providesryet another instance in which constitutive production of an enzyme plays a role in the gain mutation. Mutants selected through repeated transfer to medium contain- ing L-1,2-propanediol have been found to lose the fucose permease, fucose isomerase, fuculose kinase, and fuculose-I-phosphate aldolase, normally functional in the catabolism of L-fucose in the wild-type or- ganism (10). However, production of the propanediol oxidoreductase and the L-lactaldehyde dehydrogenase (normally acting in the breakdown of L-lactaldehyde to pyruvate, aerobically, and of L-lactaldehyde to propanediol, anerobically, respectively) were found in this bacterial mutant. 'These changes work in concert with an enhanced facilitated diffusion across the cell membrane in the trapping of propanediol by hastening its rate of entry and conversion to lactate. Other mutations, such as the formation of new enzymes (11) in a few instances, and the type of mutation leading to the use of L-mannose. a toxic and unnatural sugar, in A_aerogenes (12) are uncommon. Consider- ing the mutational acquisition of the ability of A;_aerogenes to use L-mannose, the wild-type organism can not use this sugar despite the fact that there is an inducible metabolic pathway for its degradation. Apparently the natural breakdown of L-mannose involves the formation of a metabolite that inhibits cell growth. However, through a series of mutations a strain can be obtained which overcomes this innate toxicity of the metabolite through some unknown process (13). Other examples of gain mutations in bacteria are cited in published reviews (9, 14). Sugar Acid Catabolic Pathways in Microorganisms Since the original report of MacGee and Doudoroff (15) on the ident- ification of 2-keto-3-deoxy-6-phospho-D-gluconic acid as an intermediate in the metabolism of D-glucose in Pseudomonas saccharophila, there is growing evidence that the production of 2-keto-3-deoxy sugar acids repre- sent a general trend in the bacterial and mammalian utilization of mono- saccharides-and their respective acid derivatives. Thus, the metabolism of such compounds as D-glucose (16), D-galactose (17-19), D-arabinose (20), D-mannose (21), L-arabinose-(22-23), D-gluconate (24-27), D-galac- tonate (28-29), L-galactonate (30), D-fucose and D-fuconate (31-34), L-fucose and L-fuconate (35-38), D-glucarate and D-galactarate (39-41). D-glucosaminic acid (42), and D-glucuronic and D-galacturonic acids (43), have been found to involve formation of analogous keto-deoxy intermediates possessing common structural features. Review of the literature shows that the formation of such Z-keto- 3-deoxy sugar acids and sugar-acid phosphates have generally been effect- ed through the action of specific dehydratases some of which require a phosphorylated and some a non-phosphorylated substrate. Such enzy- matic dehydrations are found to occur d-B to the carboxyl group of aldonic. aldaric. and uronic acids. Pbosphorylation reactions leading to the formation of sugar-acid phosphate intermediates, as effected by Specific kinases, can occur either before or after such dehydrations. As for the degredation of such keto-deoxy intermediates, reactions in- volving aldol-type cleavage reactions are most common; other cleavages such as decarboxylation or dehydrogenation reactions are less commonly found. ‘ With these observations, four basic routes for the degradation of sugar acids can be enumerated: (i) phosphorylation of the sugar acid followed by dehydration and then cleavage; (ii) dehydration of the sugar acid followed by phosphorylation and then cleavage; (iii) dehydration of the sugar acid, without phosphorylation, followed by cleavage; and (iv) successive dehydration followed by decarboxylation or dehydrogena- tion type cleavages to give the end products. Regarding the first case. most evidence has been gained through the study of D-glucose and D-gluconate metabolism in bacteria. Entner and Doudoroff (16) found that cell-free extracts of Pseudomonas saccharo- philg_ could convert D-glucose to pyruvate and triose phosphate through the following entirely new pathway involving a dehydratase and aldolase: D-gl ucose—-- D-gl ucose-G-phosphate -—-6-phospho-D-gl uconate ——-- 2-keto-3-deoxy-6-phospho-D-gluconate-—-——-pyruvate + D-glyceraldehyde- 3-phosphate. This so-called Entner-Doudoroff pathway is known to be widely distributed amoung species of the Enterobacteriaceae and pseudo- monads (44); however, in members of the Enterobacteriaceae, the presence of an inducible kinase allows D-gluconate to be phosphorylated direct- ly to 6-phospho-D-gluconate. In §;.ggll (45) and Salmonella typhimur- 139 (46), D-gluconate is metabolized through the following pathway: D-gluconate---6-phospho-D-gluconate (6PG)---2-keto-3-deoxy-6-phospho- D-gluconate (KDPG)-—-¥-pyruvate + triose phosphate; an active hexose monophosphate shunt (HMP) pathway is also utilized by these bacteria to convert 6-phospho-D-gluconate to ribulose-S-phosphate and pentose intermediates. Pseudomonas fluorescens (pgtidg), has been found to possess both the Entner-Doudoroff (ED) pathway, HMP pathway, and an inducible D-gluconate kinase (converting D-gluconate to 6-PG) leading to the formation of pyruvate and triose phosphate (26): D-glucose -—-- D-gl uconate -—-- 6-phospho-D-gl ucona te —-- Z-keto- 3-deoxy-6- phospho-D-gluconate---pyruvate + D-glyceraldehyde-3-phosphate. The presence of 2-keto-D-gluconate and Z-keto-G-phospho-D-gluconate intermediates in Aerobacter cloaceae and A;.aerogenes (48,49), 51gb; sigllg_aerogenes (SO), and a variety of pseudomonads when grown on - either D-gluconate or 2-keto-D-gluconate has been well documented. In Pseudomonas fluorescens (47) the presence of a dehydrogenase, kinase, and reductase results in the following divergent pathway for D-gl ucose: D-gl ucose -——-D-gl uconate———-— 2-keto-D-gl uconate—— 2-keto-6-phospho-O-gl ucona te —- 6PG —-. KDPG —-- pyruva te + D-glyceraldehyde-B-phosphate. Formation of 2-keto-3-deoxy compounds through dehydration of the sugar acid before phosphorylation is quite common in the metabolism of other sugars besides D-glucose and D-gluconate. In the metabolism of the uronic acids D-glucuronic and D-galacturonic acids in g;_gglj_(43), D-mannonic and D-altronic acids are formed, respectively. Each are found to undergo dehydration by specific dehydratases to a common product, 2-keto-3-deoxy-D-gluconic acid (KDG), which is found to be further metabolized through a kinase and aldolase in the following manner: KDG ——-- 2-keto-3-deoxy-6-phospho-D-gluconic acid -—- pyruvate + triose phosphate. In Rhodopseudomonas spheroides (21) mutant cells that are able to grow on D-glucose are found to acquire D-gluconate dehydratase and enhanced levels of KDG kinase and 6-phospho- D-gluconate dehydrogenase activities. These activities allow D-glucose to be metabolized by way of the ED pathway. In addition, the presence of an "aldose dehydrogenase" system allows D-mannose and D-glucose to be degraded through the following pathways: D-mannose-——- D-manno-y-lactone —-mannoni c acid—KDG -—-- KDPG -—--pyruvate + triose phosphate; D-glucose-——-—-D-glucono-y-lactone --—-D-gluconic acid-—--KDG-—-—-KDPG-—-—-pyruvate + triose phosphate. Along these same lines, it has been found that growth of Clostridium aceticium (51, 25) and the Achromobacter-Alcaligenes 64-70% (G-C) subgroup (52) on D-gluconate induce the dehydratase responsible for the following series of reactions: D-gluconate —?-KDG ——--KDPG. The third route. dehydration of the sugar acid without phosphor- ylation, mainly concerns the metabolism of the sugars D-arabinose. D-galactose, D-galactonate, D-glucarate, D-fucose, L-arabinose, and L-fucose. In Pseudomonas saccharophila, the sugars D-arabinose and D-galactose are metabolized through two completely different routes than are generally found in the various bacterial classes. Mutant strains adapted to grow on D-arabinose (20) are found to possess the enzymes responsible for effecting the following reactions: D-arabino- furanose—D-arabino-v-lactone——-2-keto-3-deoxy-D-arabonic acid ——-—-pyruvate + glycolaldehyde. Galactose adapted strains, which cannot utilize either D-glucose or D-fructose are found to degrade D-galactose as follows: D-galactofuranose-—-—-D-galactono-y-lactone -———D-galactonic acid—~2-keto-3-deoxy-D-galactonic acid—— pyruvate + D-glyceraldehyde (19). This later scheme has also been found to function in the metabolism of D-galactose in Gluconobacter ligug; faciens (18) although cleavage is believed to take place through a phosphorylated intermediate, 2-keto-3-deoxy-6-phospho-D-galactonate, to yield the products pyruvate and triose phosphate. Recently in the K-12 strain of g; g911_(28,29) the direct catabolism of D-galactonic acid was found to precede as follows: D-galactonate-—-—-2-keto-3- deoxy-D-galactonate—-2-keto-3-deoxy-6-phospho-D-gal actonate -—- pyruvate + D-glyceraldehyde-B-phosphate. This strain was found to utilze D-galactonate as a sole carbon source for growth and to possess an inducible system for the catabolism of this aldonic acid, i.e., a D-galactonate dehydratase. a 2-keto-3-deoxy-D-galactonate kinase, and a 2-keto-3-deoxy-6-phospho-D-galactonate aldolase, when grown on the in- ducer D-galactonate. The manner by which D-gluconate is catabolized in Aspergillus giggg,(54) is markedly different than described for pseudomonad and 10 Escherichia species in that no phosphorylation occurs: D-gluconate —-—-KDG——-pyruvate + D-glyceraldehyde. Metabolism of the dicarbox- ylic sugar acid D-glucarate by §;_ggli_(40) and possibly other members of the family Enterobacteriaceae, is quite different than that which will be discussed for pseudomonads. D-Glucarate is initially dehydra- ted at each end of the compound to give a mixture of 2-keto-3-deoxy-D- glucaric acid and S-keto-4-deoxy-D-glucaric acid of which the latter predominates; both intermediates are degraded further by the presence of a 2-keto-3-deoxy-D-glucaric acid aldolase to pyruvate and tartronic semialdehyde. Tartronic semialdehyde can be further degraded to glyceric acid by a reductase. There is good evidence that D-glucarate is degraded in the same manner by the species Aerobacter aerogenes, Salmonella typhi- mggium, and Klebsiella pneumoniae (40). Concerning the metabolism of D-fucose in bacteria, a rare carbohy- drate, Dahms and Anderson (31) were able to isolate a pseudomonad which readily utilized the sugar as sole carbon source for growth. Elucidation of the pathway revealed that D-fucose is metabolized as follows: D-fucose---D-fucono-d-lactone and D-fucono-y-lactone-—-—-2-keto-3- deoxy-D-fuconate--—-pyruvate + D-lactaldehyde (31-34). The enzymes involved in the pathway were also found to be effective in the metabolism of the common sugar L-arabinose by a similar series of reactions: L-arabinose -—--L-arabono-y-lactone—L-arabonic acid—~2-keto- 3-deoxy-L-arabonic acid (KDA)-—-——-pyruvate + glycolaldehyde (23). Thus a new pathway for the metabolism of L-arabinose has been shown to occur in a pseudomonad involving an aldolase type cleavage at the level of KDA rather than dehydration followed by dehydrogenation. This latter 11 pathway has subsequently been found to be effective in the metabolism of L-arabinose in Rhizobium japonicium which use this sugar as a pre- fered carbon source (55). Along these same lines, the metabolism of L-fucose in pork liver (35-38) has been found to procede as follows: L-fucose——- L-fucono-d-l actone—— L-fuconate——- 2-keto-3-deoxy- L-fuconate———-2,4(or 5)-diketo-5(or 4)-monohydroxyhexonate -——-- two moles of lactate. Concerning the final reaction group, namely those pathways show- ing successive dehydration followed by either dehydrogenastion or de- carboxylation, the following sugars as found in pseudomonads are considered: D-glucarate, D-galactarate, and L-arabinose. Both D-gluc- arate and D-galactarate are found to be initially dehydrated to 4-deoxy- S-keto-D-glucarate which is further dehydrated and decarboxylated to 2-keto-glutaric acid semialdehyde followed by dehydrogenation to Z-keto-glutaric acid in most pseudomonads studied (39,56). LeArabin- ose, as exemplified by Pseudomonas saccharophila (57), is metabolized through the following pathway: L-arabinose-——-—-L-arabino-a-lactone 2-keto-3-deoxy-L-arabonic acid-—-—4-2-ketoglutaric acid semi- aldehyde-———-2-ketoglutaric acid. Strategies for Sugar Catabolism in the Enterobacteriaceae ' Klebsiella pneumoniae, being a member of the family Enterobacter- iaceae, should show characteristic pathways of the class for common sugars. The following carbohydrates and their pathways are considered for future reference when considering growth studies of this bacterium: D-arabinose, L-arabinose, L-fucose, D-galactose, L-lyxose, L-xylose, L-arabitol, 12 xylitol, glycerol, D-lyxose, D-allose, L-mannose, D-xylose, D-fructose, D-mannose, sorbitol, mannitol, galactitol, cellobiose, gentiobiose, sucrose, and L-sorbose. As described by Mortlock et al. (1), metabolism of various pen- titiols and pentoses in Aerobacter aerogenes (presently classified as Klebsiella pneumoniae) were found to procede through the HMP pathway intermediates D-ribose-S-phosphate, D-ribulose-S-phosphate, and D-xylu- lose-5-phosphate, and two rare intermediates L-xylulose-S-phosphate and L-ribulose-S-phosphate. Investigations showed that: (i) D-ribose can be converted to D-ribulose-S-phosphate; (ii) D-arabinose and ribitol to D-ribulose then to D-ribulose-S-phosphate; (iii) D-xylose, D-lyxose, D-arabitol, xylitol to D-xylulose then to D-xylulose-S-phosphate; (iv) L-arabinose to L-ribulose then to L-ribulose-S-phosphate; (v) and L-xylose, L-lyxose, and L-arabitol to L-xylulose then to L-xylulose- 5-phosphate. The metabolism of D-arabinose, as has been deduced from studies with §;_ggli_(59,60) and A; aerogenes (1), suggest two pathways. In A;_aerogenes and strain B/R of §;_ggli (61), which utilize D-arabinose but not L-fucose, the pathway is as follows: D-arabinose----D-ribulose ----D-ribulose-S-phosphate-—-—-D-xylulose—S-phosphate. In the K-12 strain of E;_ggli_(60), which does use L-fucose, the following reactions are found:' D-arabinose-—--—D-ribulose-—-——-D-ribulose-l-phosphate -—¥——-dihydroxyacetone phosphate (DHAP) + D-glyceraldehyde. It is be- lieved that the K-12 strain of E_._ c_ol_i makes use of the L-fucose enzymes in order to effect the isomerase, kinase, and aldolase cleavage reactions (1,60). 13 L-Arabinose metabolism in A;_aerogenes is as follows: L-arabinose --—-L-ribulose L-ribulose-S-phosphate-—-——-D-xylulose-S-phosphate (62). L-Fucose metabolism in g; £911 (63-65) has been studied and is believed to be similar in A;_aerogenes, Salmonella enterdidis, and several Shigella species: L-fucose-—-—-L-fuculose-—-——-L-fuculose- 1-phosphate-—-—-DHAP + L-lactaldehyde. D-Galactose metabolism in these bacteria is believed to take place strictly through the Le Loir pathway: D-gal actose——- D-galactose-I-phosphate—- UDP-gl ucose -—-——-D-glucose-I-phosphate (78). I In addition, the metabolism of L-mannose. D-fructose, sorbitol and mannitol, galactitol, and L-sorbose in A;_aerogenes are as follows: L-mannose-——- L-fructose —- L-fructose-I-phosphate —- DHAP + L-glyceraldehyde (11,12); D-fructose-—-—-D-fructose-I-phosphate .___—-D-fructose-I,6-diphosphate (66-68); mannitol and sorbitol--- —— mannitol-I-phosphate and sorbitiol-l-phosphate— D-fructose- 6-phosphate (69,70); galactitol-——--D-galactitol-6-phosphate-—-- D-tagatose-G-phosphate D-tagatose-I,6-diphosphate-——-DHAP + O-glyceraldehyde-3-phosphate (71); L-sorbose-—-—-L-sorbose-l-phosphate ---D-glucitol-6-phosphate-——-D-fructose-6-phosphate (72). The dissacharides, gentiobiose. cellobiose, and sucrose are all found to be cleaved to give characteristic glycolytic intermediates in §;_pneumoniae. Gentiobiose is metabolized through an ATP-dependent phosphorylation of the dissacharide to give gentiobiose monophosphate followed by hydrolysis to give D-glucose and D-glucose-G-phosphate (73). Cellobiose is similarly modified to give D-glucose and D-glucose-G- phosphate as hydrolysis products (74-76). Sucrose is not phosphorylated, 14 but hydrol ' yzed d1rectly to give D-glucose and D-fructose (77) SECTION I Elucidation of the Metabolic Fate of D-Fuconate 15 INTRODUCTION The pathway for the catabolism of D-fuconate in Klebsiella pneumoniae has not been previously determined. D-Fuconate is known to be an intermediate in the metabolism of D-fucose in a pseudomonad, but §;_ggeumoniae seems not able to be forced to use D-fucose as a sole carbon and energy source for growth. However, §;_pneumoniae will grow on D-fuconate after a lag of several days. The experiment- al results presented in this section will provide chemical, enzymatic, and genetic evidence for a pathway for D-fuconate in this bacterium as follows: D-fuconate-—-—-2-keto-3-deoxy-D-fuconate pyruvate + D-lactaldehyde. The gain mutation which facilitates growth of this bacterium on D-fuconate has been determined to be the constitutive production of a dehydratase that is normally induced by D-galactonate but not by D-fuconate. The aldolase which cleaves 2-keto-3-deoxy-D- fuconate is induced by this substrate, but not by D-galactonate or its metabolites. Thus, whereas D-fuconate is apparently a fortuitous _ substrate for D-galactonate dehydratase, the normal function for the aldolase is difficult to conceive. 15‘ MATERIALS AND METHODS Bacterial Strain, Cell Growth, and Pregaration of Cell-Free Extracts Bacterial Strain. Klebsiella pneumoniae PRL-R3,U', an auxotroph requiring uracil for growth, was used as the parental strain for the D-fuconate-positive mutants. Media, Broth cultures of cells were grown in a mineral medium containing 0.15% KH2P04. 0.71% MaZHP04, 0.3% (NH4)ZSO4, 0.01% M9502, and 0.0005% FeSO4-7H20, supplemented with 0.5% carbohydrate and 0.005% uracil, or in nutrient broth (Difco), prepared by mixing 8.0 g dehy- drated powder with 1.0 liter distilled water to which 0.005% uracil was added. Mineral-medium-agar plates were prepared by mixing sterile mineral salts and agar to give the same concentration of salts as in the broth, but with an agar content of 1.5%. All mediums were adjusted to pH 7.0 before used. . Cell Growth. All broth and agar plate cultures were grown in the dark at 30°C in a thermostatically controlled incubation room. Broth - cultures in 18 X 150 mm culture tubes were equipped with plastic clo- sure caps and contained 7.0 ml of media; cultures of loo-ml volume were grown in 250-ml erylenmeyer flasks equipped with cotton plugs. Aeration was accomplished by constant motion on either reciprocal or rotary shakers of the New Brunswick Scientific Company. Broth cul- tures were inoculated from fully grown (7.0 ml) cultures having an absorbance at 600 nm of 0.60 or better. 17 18 MonitoringgCell Growth. Cell growth was monitored with either a Coleman Junior Spectrophotometer, Model 6A, or a Gilford, Model 2400, Spectrophotometer at a wavelength of 600 nm. Cultures grown in 18 X 150 mm culture tubes could be directly measured in the Cole- amn Jr. using an uninoculated culture tube as a blank. A plot of corrected absorbance versus the percent of maximal growth was used when reporting turbidity measurements. This plot. prepared by correlating absorbance values from undiluted samples to absorbance values from diluted samples, permitted one to correct for deviations from Beers law at the higher cell concentrations. The Gilford spectrophotometer was used when sampling growing cultures directly with the aid of sterile pasteur pipets; measurements were made in microcuvettes (0.2-ml volume). Harvesting of Cells. Cell cultures were incubated at 30°C on appropriate shakers until an uncorrected optical density of 0.60 at 600 nm was reached on the Coleman Jr. or an uncorrected value of 2.0 at 600 nm with the Gilford spectrophotometer. The cells were suspended in 0.85% NaCl and centrifuged again at 12,000 X g_for 10 min. The - cell pellets could then be used within the hour or frozen at -20°C until needed. Centrifugation. All centrifugations were done in a Sorval refrig- erated centrifuge, model RC-ZB, at O-4°C. The rotor radius was either 4.34 or 5.75 inches. Preparation of Cell-Free Extracts. Cell extracts were prepared by suspending the harvested, washed cells in 0.05 M potassium phosphate 19 buffer (pH 7.0) and exposing the solution to sonic vibration (10,000 Hz) in a Raytheon sonic oscillator, model DF-101, for 30 min. Prior to sonication, cell pellets derived from 7.0-ml cultures were resus- pended in 0.5 ml of the sonication buffer and placed in 0.5 X 3.0 inch cellulose nitrate-centrifuge tubes equipped with rubber stoppers. These tubes were then placed in the chamber of the apparatus and cold water was added to the level of the solution inside of the tubes. Larger cell-suspensions were sonicated directly in the chamber. All sonications were done in the presence of glass beads and having the chamber temperature near 4°C with the aid of a circulating water-ice bath. Following sonic disruption of the cells, the suspension was centrifuged at 12,000 X g_for 10 min to remove whole cells and debris. The resultant supernatant was decanted as the crude extract. Protein Determination ~ Protein concentration was determined using the procedure of Lowry et al. (79). Bovine serum albumin was used as the protein stand- ard. §a§;Liguid Chromatography - Trimethylsilylated derivatives of aldoses and their respective aldonic acids and lactones were prepared by incubating 0.1 mg of the carbohydrate with 0.2 ml dry pyridine and 0.2 ml N,0-Bis-(Trimethyl- silyl) trifluoroacetamide (BSTFA) for 30 min at 50°C in a teflon-sealed 0.5-dram vial. Gas-liquid chromatography was performed on a Beckman Varian Aerograph, model 210 , employing a 3% 0V-17 column and temper- ature program from loo-230°C. Traces were recorded on unmarked chart 20 paper but calibrated in terms of temperature knowing the rate of temper- ature change during the trace. Retention times were normalized to the internal standard potassium D-gluconate; under the conditions employed, this standard had a retention time of 22.0 ininutes. Preparation of Ion Exchange Resins Dowex resins used as ion exchangers for purposes of desalting or column chromatography were prepared by preconditioning with either acid or base; the desired ion form was then prepared by treatment of the resin with the ion solution of choice through either a batch-wise or a column method. Cation exchange resins, Dowex-50, were treated first with 2.0 N NaDH then washed to neutrality with distilled water and then treated with 2.0 N HCl to give the acid form. Anion exchange resins, Dowex-1, were treated alternately with 2.0 N HCl, distilled water, 2.0 N NaOH, distilled water, and then either ammonium bicarb- onate in the preparation of Dowex-1 (bicarbonate) or 2.0 N formic acid, distilled water, sodium formate in the preparation of Dowex-1 (formate). All resins were washed with distilled water in the final form and air dried or used in packed form in a column of known dimen- sions. Colorimetric Assays, A Gilford spectrophotometer, model 2400, was used to measure absorbance units of sugar chromogens at their specific wavelengths. Cuvettes with a 1.0-cm path length were used in all cases. In cases where standard curves were developed, commercial-grade carbohydrates used were dried to constant weight over phosphorous pentoxide before 21 preparing the stock standard in solution. Blanks for all assays were made by substituting distilled water for the carbohydrate solution. Reducing Sugar Assay. The Phenol/H2504 assay of Dubois et al. (80) was used to quantitate reducing sugar solutions. Standard curves were prepared using between 10 to 70 ug of carbohydrate in the stand- ard assay. D-Glucose was used as standard for hexoses of which chromogens were read at 490 nm. Pentoses and uronic acids were quantitated using D-ribose and D-glucuronic acid, respectively, as standards which were read at 480 nm. Aldonic Acid Assay. Aldonic acids were quantitated by the alka- line hydroxylamine assay of Hestrin (81) after conversion to their cor- responding lactones. Lactonization was effected by heating an aqueous solution of the aldonic acid, 2 to 8 umol, in 2.0-ml volume, with 2.0. N HCl for 15 min at 90°C. Excess HCl, which interfers with the hydroxyl- amine formation, was completely removed from the samples by use of a Rotomix evaporator, temperature-regulated at 90°C. D-Galactono-y-lact- one and potassium D-galactonate gave identical standard curves in this procedure. The purple-brown chromogen produced was read at 540 nm. Aldehyde Colorimetric Assays. D-Lactaldehyde was determined as the tetraazopentamethine cyanine dye by the method of Paz et al. (82). Acetaldehyde, 0.05 to 0.10 umcl, was used to develop the standard curve. The blue chromogen which developed in the assay was read at 670 nm. The molar extinction coefficient for acetaldehyde of 60,000 (82) was used to quantitate the aldehyde solution. Alpha-Keto Acid Determination. All a-keto acids were determined as their semicarbazones by the procedure of MacGee and Doudoroff (15). 22 Semicarbazones of d-keto acids have Characteristic molar extinction coefficients of 10,200 at 250 nm (15). Standard curves were developed using sodium pyruvate as the standard. 2-Keto-3-0eoxy Sugar Acid Determination. The procedure of Weiss- bach and Hurwitz (83) was used to determine 2-keto-3-deoxy sugar acids. Carbohydrates were subjected to periodate oxidation in this assay which yields B-formyl pyruvate as reported (83), giving a characteristic chromogen with thiobarbituric acid absorbing maximally at 551 nm. Molar extinction coefficients for various sugar acids quantitated by this assay were determined using the semicarbazone assay to quantitate the a-keto acid content. As latter sections of this thesis will show, the following compounds have molar extinction coefficients at 551 nm in this assay of: 3,6-dideoxy-thrgg;D-hexulosonic acid (2-keto-3-deoxy- D-fuconate), 50,060; 3,6-dideoxy- rythro-D-hexulosonic acid (Z-keto- 3-deoxy-D-quinovonic acid), 10,930; 2-keto-3-deoxy-D-galactonate, 60,000. Preparation of Substrates Aldonic Acid Synthesis. Aldonic acids were prepared from respect- ive aldoses by the hypoiodate oxidation procedure of Moore and Link (84). The aldose (2.0 g) was initially dissolved in distilled water (4.0 ml) to which was added 25.0 ml absolute methyl alcohol. D-Galac- tose has to be heated into solution before the addition of the alcohol. The aldose solution was then added to a stirring solution of 5.7 9 re- sublimed iodine in 80.0-ml acetone-free methanol (absolute) temperature regulated at 40°C. A three-neck flask (500 ml) equipped with 23 ground-glass joints, a separatory funnel, CaSD4 drying tube, and thermo- meter heated in a water bath-magnetic stirrer/heater apparatus. Heat was removed once the aldose was added to the reaction mixture and a solution of 40% KOH in methanol (115 ml) was gradually added over a period of 30 minutes. The endpoint of the reaction was detected upon the development of a straw-yellow color, indicating that the iodine was consumed, and was stirred 20 additional minutes to assure complete salt formation of the aldonic acid. Recovery of the aldonic acids from the reaction mixture is depend- ent on their solubility in methanol/water medium. L-Arabonic, D-galact- onic, and D-gluconic acids precipitated out of the reaction mixture as their potassium salts. These salts could be collected by suction fil- tration, and washed with methanol and ether to give dry products easily weighed and handled. D-Fuconate was also obtained as a potassium salt, however the reaction mixture had to be kept at 4°C in order to initiate cyrstallization. After one day at this temperature, all of the salt could be collected by suction filtration and washed with methanol and ether. D-Lyxonic, D-mannonic, and D-xylonic acids were obtained as their barium salts by treating the reaction mixture with a suspension - of 20% barium iodide in methanol (20 ml). The salts were collected by centrifugation and washed several times with methanol and ether before drying. The potassium salts could be obtained by treatment of the barium salts first with Dowex-50H (H+) and then with Dowex-50w (K). In all cases, no less than an 80% molar yield was obtained, and all salt products were shown to be homogeneous by gas-liquid chromatography. Retention times for D-glucose, D-galactose, and D-fucose aldose, aldonic 24 acids, and acid-generated lactones as prepared by the this procedure are given (Table 1). In each case, only one peak is seen with the aldonic acid generated from the commercial aldose which form character- istically one lactone peak when prepared as in the hydroxylamine assay (see above). Preparation of Oxaloacetic Acid. Oxaloacetic acid used in the synthesis of 2-keto-3-deoxy sugar acids was prepared from commercial grade sodium diethyloxaloacetic acid (Sigma Chem. Co.) as described by Cornforth et al. (85). The solid oxaloacetic acid diethyl ester (200 g) was initially washed by resuspension in ether (400 ml) and filtered by suction filtration through fritted glass until a light yellow powder was obtained. The collected powder, 158.0 g, was then acidified by dissolving into a stirring solution of 2.0 N H2504 (2.0 liter) and ether (500 ml) at 4°C. A large erylenmeyer flask equipped with a rod stirrer was used to mix the solution. After the solid had completely dissolved, about 20 minutes, the ether and aqueous layers were separated and the aqueous layer was extracted with three (250 ml) portions of ice-cold sodium bicarbonate (5-101 w/v) until the washings ' no longer were colored. The ether extracts were then dried over anhydrous M9504 and dried to a syrup on a Spinco evaporator at 25°C under reduced pressure. The syrup was then distilled at 135-140°C at 15.microns Hg to a clear oil using Bantam-ware micro-distillation equiptment, a thermo- statically controled oil bath, and a HiVac diffusion pump. The re- sultant distillate, ethyl oxaloacetate, could be stored indefinitely 25 Table I. Retention times for carbohydrates determined by gas- liquid chromatography. The following aldoses, aldonic acids, and aldonic acid lactones were prepared as trimethylsilyl derivatives and subject- ed to gas-liquid chromatography as described in Materials and Methods. Reported aldoses were commercial grade; aldonic acids (K+ salts) were prepared as described in the text. Lactones of respective aldonic acids were prepared by heating in acid and evaporating to rid of excess HCl as described for the hydroxylamine assay. Retention times were all norm- alized to the retention time of D-gluconate, having a retention time of 22.0 minutes under the conditions as described in Materials and Methods. CARBOHYDRATE SUGAR FORM RELATIVE RETENTION TIME (min/22.0 min) D-FUCOSE ALDOSE 0.587, 0.618. 0.676 ALDONIC ACID 0.825 LACTONE 0.726 D-GALACTOSE ALDOSE 0.823, 0.858, 0.906 ALDONIC ACID , 1.00 LACTONE 0.965 D-GLUCOSE i ALDOSE 0.830, 0.850, 0.892 ALDONIC ACID 1.00 LACTONE 0.972 26 at -20°C, later to be hydrolyzed to the free acid. The yield of ethyl oxaloacetate from the diethyl ester (200 g) was 55.0 g; the oil den- sity was 1.108 g/ml. Free oxaloacetic acid was prepared by hydrolyz- ing aliquots of the ethyl oxaloacetate oil in concentrated HCl; 27.0 ml of the oil (30.0 g) was dissolved in 125.0 ml of HCl and left to stand for 2-3 days. The free acid that crystallized out upon stand- ing was collected by filtration through fritted glass giving a yield of 14.8 g of free oxaloacetic acid per 30 g of the oil or a 50% weight yield. The collected acid was then dried over KOH in a desiccator and stored at -20°C until needed. Quantitation of the synthetic product by either the semicarbazone assay (15) or with malate dehydrogenase gave identical results. Synthesis of D-Lactaldehyde. 'D-Lactaldehyde was prepared from L-threonine by the ninhydrin oxidative deamination procedure of Zagalak et al. (86). Commercial grade L-threonine (3.0 g) was rapidly added to a vigorously stirred solution of 9.1 g ninhydrin in 0.05 M sodium citrate buffer (pH 5.4) of 600-ml volume. Carbon dioxide was liberated in the course of the reaction so a large mixing vessel was used which permits maximal surface area. After 15 min stirring at 100°C, the resultant purple mixture was allowed to cool to room temperature. Once cooled, the solution was filtered through Vhatman #1 filter paper and collected by suction filtration. The filtrate was then treated alternately with Dowex-50H (H+) and Dowex-I-XB (bicarbonate) as describ- ed by Zagalak et al. (86). The final solution was shown to be clear at neutral pH and when quantitated for aldehyde by either the colori- metric assay (82) or with alcohol dehydrogenase a yield of 14.8 umol 27 product was found. Chemical Synthesis of 2-Keto-B-Deoxy-D-Fuconate. (i) Portsmouth Synthesis. 2-Keto-3-deoxy-D-fuconate was prepared by the alkaline aldol-condensation reaction of oxaloacetic acid and D-lactaldehyde as described by Portsmouth (87). This reac- tion is predicted to form two isomers, 3,6-dideoxy-thrgp-D-hexulosonic acid (2-keto-3-deoxy-D-fuconic acid; KDF) and 3,6-dideoxy-erythro-D- hexulosonic acid (2-keto-3-deoxy-D-quinovonic acid; K00), in a ratio of 1.0 to 2.0, respectively. In addition, the procedure is capable of completely resolving the two isomers by ion exchange chromatography. A solution of 8.0-ml volume containing D-lactaldehyde (2.8 mmol) was added to 20.0 ml of 0.05 M potassium phosphate buffer (pH 7.5) at 25°C and adjusted to the pH of the buffer with the aid of a Sargent. pH Stat with an 8.0 N KOH reservoir. Oxaloacetic acid (5.7 mmol) was then added gradually as the pH of the solution was maintained at 7.5. Once all of the oxaloacetic acid was dissolved, the reaction mixture was left to stand for a total of 16 hrs with continuous stirring. Assays of the reaction mixture at the end of this incubation by the procedure of Heissbach and Hurwitz (83) showed no further production of 2-keto-3-deoxy sugar acid compounds and thus the reaction was judged to be complete. To prepare the reaction mixture for ion exchange chromatography, the pH of the solution was adjusted to pH 4.0 with formic acid and de- gassed by gentle aspiration over steam. The sample in 70.0-ml volume was applied to a 3.8 X 50.0 cm column of Dowex-l-XB (formate), 200-400 mesh, at a rate of 140 ml per hour by gravity. Once the reaction 28 mixture was loaded, the column was washed in the following manner: (a) 2.0 liters of distilled water was passed through the column to wash products that did not stick to the resin from the ion exchanger, (b) 2.2 liters of a 0.23 M formic acid solution was passed over the resin to remove loosely bound products, and (c) 3.5 liters of 0.46 M formic acid solution was passed over the resin to elute the keto-deoxy sugar acids. Unreacted D-lactaldehyde was detected by the aldehyde assay of Paz et al. (82) in approximately 252 to 420 ml of distilled water wash; a recovery of 223 umol aldehyde was determined upon pooling and concentra- tion of the peak fractions. Assays of the eluate fractions from the 0.23 M formic acid washings failed to show any additional aldehyde or semicar- bazone-positive material. However, two well resolved peaks were found in eluate fractions subsequent to the 0.46 M formic acid wash; both peaks were found to give positive semicarbazone derivatives characteristic of arketo acids and reacted as keto-deoxy sugar acids in the thiobarbituric acid assay (83). The elution profile of the 2-keto-3-deoxy sugar acids (Fig. 1) following the 0.46 M formic acid washing was similar to that found by Portsmouth (87). The first peak, defined by Portsmouth as the erythro isomer (K00), and second peak, defined as the £01299. isomer (KDF), were- pooled separately and concentrated to syrups under reduced pressure at 40°C several times with distilled water to remove formic acid. These syrups were taken up in known volumes of distilled water, neutralized with KOH, and characterized by known colorimetric assays. Semicarbazone assays of each isomer gve 873 umol d-keto acid for K00 29 Figure 1. Elution profile of the 3,6-dideoxy-D-hexulosonic acids from the Portsmouth synthesis. The reaction mixture was prepared and treated as described in the text. Fractions of 21.0-ml volume were assayed for d-keto acid content by the semicarbazone assay (15) and for 2-keto-3-deoxy sugar acid content by the thiobarbituric acid assay (83) in eluate fractions of the 0.46 M fonmic acid wash. «(a é) (W 199) 3an9303911 O'O L O' Iva. gusts: zc_hu<¢m me. oo— mm— cm. me- o«— mn— amp P n n P pl - b b l @ adage O «Na b cw— m.. h s.— we- 7. r9 001 I DIDV 013N'VHdTV HTDNOUSIN 31 (fractions 110 to 130) and 425 umol d-keto acid for KDF (fractions 140 to 160), based on the molar extinction coefficient of 10,200 at 250 nm (15). Molar yields of these products were 67.2% and 32.8% for K00 and KDF, respectively. The ratio of KDQ to KDF, observed as 2.0 to 1.0, is comparable to that value reported by Portsmouth (87). In addition, quantitation of each product by the semicarbazone assay facilitated the assignment of molar extinction coefficients for the thiobarbituric acid assay (TBA) (83); values for KDF and KDQ, at 551 nm, were determined as 50,060 and 10,930, respectively. (ii) Dahms Synthesis. In contrast to the findings of Ports- mouth, Dahms (33) reported that one isomer, KDF, was produced by the aldol-condensation between oxaloacetic acid and D-lactaldehyde. This fact contradicts chemical theory, as the asymmetric carbonyl group of D-lactladehyde should permit both syp- and agti-facial attack by the oxaloacetic acid carbanion generated in the chemical reaction (87). The result of such an attack must be the formation of two isomers, unless steric hindrance prevented one from being formed; as the reac- tion does yield an isomeric mixture in the Portsmouth synthesis, steric hindrance is ruled out. After examining the protocol by which Dahms synthesized the com- pound KDF two possible postulates were drawn to explain why he failed to detect two isomers: (a) the alkaline conditions (pH 11) underwhich the synthesis was conducted resulted in the loss of K00, or (b) the method of chromatography failed to separate KDQ from KDF. when the Dahms' protocol was tried, a yield of thiobarbituric acid-positive material, comparable to that found by Portsmouth, was detected in the 32 reaction mixture and which did not diminish over time at pH 11 (see below). However, elution of the product with a gradient of 0.0-0.46 M formic acid from a Dowex-I-XB (formate) column (0.5 X 15.0 cm) did not result in two TBA-positive peaks. These results suggested that the column dimensions used in the Dahms procedure could not resolve the two isomers formed, rather than indicate that only one isomer was gen- erated. To prove this was the case, a reaction mixture was prepared according to the Dahms' protocol and was chromatographed by the pro- cedure of Portsmouth. The reaction mixture was prepared using a similar protocol as described for the Portsmouth synthesis; the main difference between the two procedures is the pH of the medium and the time of reaction. Oxaloacetic acid (3.4 mmol) was initially dissolved in 4.0 ml of 0.05 M carbonate-bicarbonate buffer (pH 10.7) to which was added 9.2 mmol of D-lactaldehyde (9.2 ml); the pH of the reaction mixture was adjusted to 11.0 with 8.0 N NaOH. The reaction was monitored for pro- ducts releasing B-formyl pyruvate by the thiobarbituric acid assay and was judged complete when further production was negligible (about _ 20-40 min at 25°C). The reaction mixture was treated with formic acid and degassed as done in the Portsmouth protocol and loaded onto another Dowex-I-XB (formate) column of the same dimensions as that used before. Approximately 1.16 mmol of unreacted D-lactaldehyde was recovered from the water wash. Elution of the column with 0.46 M formic acid revealed similar results as seen in the Portsmouth synthesis; two well resolved peaks were detected by both the semicarbazone and thiobarbi- turic acid assay (Fig. 2). The peak fractions were pooled separately 33 Figure 2. Elution profile of the 3,6-dideoxy-D-hexulosonic acids from the Dahms' synthesis. The reaction mixture was prepared and treat- ed as described in the text. Fractions of 21.0-ml volume were assayed for d-keto acid content by the semicarbazone assay (15) and 2-keto-3- deoxy acid content by the thiobarbituric acid assay (83) in eluate frac- tions of the 0.46 M formic acid wash. 5' 31.1 é) (um 199) 3311119210591: 1 0'1 5'0 0'0 .3 <3 <3 G i- Q Q Q Q I- G) <3 9 I- <3 3* <3 <3 G - <3 <3 <3 <3 <3 <3 G i- <3 A Q Q ' a Q - ‘9 I r T 7 fir . I 04 05 0? OS 02 01 O I 013V OlJX'VHdTV BTDNOUDIH 115 120 . 125 I30 135 I40 I45 150 155 160 FRACTION NUMBER 110 35 and treated as before. Assays of the pooled neutralized fractions gave 621 umol d-keto acid for the first peak, and 420 umol o-keto acid for the second peak by the semicarbazone assay. As defined by the Portsmouth procedure, yields of 59.7% for K00 and 40.4% for KDF were observed. These results show that the mechanism of the reac- tion was the same in both procedures of synthesis as the products were produced in the same overall yield. As noted in the Portsmouth synthesis, molar extinction coef- ficients for K00 and KDF were determined to be approximately 11,000 and 50,000, respectively, at 551 nm. As these isomers were not separ- ated by the Dahms procedure of chromatography, the reported molar ex- tinction coefficient of 27,900 (33) should reflect that for the isomeric mixture. In fact, if the molar yields of each isomer (0.6 M for K00 and 0.4 M for KDF) are used to calculate the molar extinction coeffic- ient for the mixture, a value approximating that reported by Dahms is obtained: , 551nm 551nm MIXTURE EXT. COEF. 11.000KDQ (0.5 M) + 50,000KDF (0.4 M) 3 26.600 DAHMS' EXT. COEF. a 27.900 (iii) Summary of the Synthetic Methods. These results show that either synthesis (Portsmouth or Dahms) can be used to prepare iso- meric mixtures of 3,6-dideoxy-D-hexulosonic acids which can be separ- ated by methods of column chromatography. The procedure of Portsmouth should be used in purification of the isomers K00 and KDF as the Dahms procedure fails to effect separation. This latter point is critical 36 in the preparation of synthetic standards to be used as substrates in enzymatic reactions. With regard to the studies conducted by Dahms on the pseudomonad D-fuconate dehydratase and the KDF aldolase, misleading results may have been obtained. Although Dahms showed that his product was cleaved by periodate at a rate similar to that of 2-keto-3-deoxy-D- galactonate and slower than 2-keto-3-deoxy-D-gluconate (33) this does not prove the thggg configuration for this compound in light of the fact that his substrate contained an isomeric mixture. To determine the configura- tion of the chemically synthesized product Dahms conducted a periodate rate study using the thiobarbituric acid assay to measure the formation of a-formyl pyruvate (88). This assay has been found to be inhibited to a limited degree by acetaldehyde (89), a product of the periodate oxida- tion of 3,6-dideoxy-D-hexulosonic acids. It is likely that when rates were measured by Dahms using the isomeric mixture as reactant, the rate of color development fortuitously matched that of 2-keto-3-deoxy-D-galac- tonate and pure (enzymatic) 2-keto-3-deoxy-D-fuconate. In addition, substrate specificity studies on the pseudomonad KDF aldolase using the reaction mixture prepared as described by Dahms gave yields of one mole of D-lactaldehyde and pyruvate per mole of substrate (34); this indicates that each isomer is used by this enzyme. If this is the case, both reported substrate specificity and kinetic constants for the KDF aldolase are inaccurate. 37 (iv) Alkaline Lability of the Portsmouth Synthetic Pro- gggtg. Dahms (88) suggested that the failure to produce two isomers in his synthesis was a result of the pH used in the synthesis rather than the method of column chromatography. To dispute this premise, the isomers from the Portsmouth synthesis were subjected to similar conditions of pH as found in the Dahms procedure. Results (Fig. 3) show that within the restraints of the Dahms synthesis no loss of either isomer occurs over a 90 min period of incubation; therefore, losses of either isomer would not be expected to occur once they were synthesized as a result of the high pH used in the Dahms synthesis. Preparation'of other2:Keto-3-Deoxy Sugar Acids. 2-Keto-3-deoxy- D-galactonate (as well as 2-keto-3-deoxy-D-fuconate from D-fuconate) was prepared enzymatically from D-galactonate, using the purified D-galactonate (D-fuconate) dehydratase as described in Section II, Results. Purification of these products from the reaction mixture was accomplished using the chromatographic procedures as described in the Portsmouth synthesis (87). Quantitation of the o-keto acid content by the semicarbazone assay gave molar extinction coefficients of 50,060 for KDF and 60,000 for KDGal at 551 nm in the TBA assay. 2-Keto-3-deoxy-D,L-arabonate (D,L-KDA) and 4-hydroxy-2-keto- D,L-glutarate (KHG) were prepared by the aldol-condensation between oxaloacetic acid and glycolaldehyde and glyoxylate, respectively. The procedure of Stoolmiller and Abeles (57) was used in the synthesis and purification of KDA. The product of the reaction was detected using the TBA assay (83) both in the synthesis and subsequent 38 Figure 3. Alkaline lability of the chemically synthesized isom- ers of the Portsmouth synthesis. Stock isomers prepared and purified according to the Portsmouth procedure (87) were diluted in 0.05 M car- bonate-bicarbonate buffer (25 ml) at a pH of 10.7; 2-keto-3-deoxy-D- quinovonic acid (KDQ) was used at 5.2 mM concentration and 2-keto-3- deoxy-D-fuconic acid (KDF) was used at 1.9 mM concentration. The pH of the mixture was titrated to a pH of 11.0 with the aid of a Sargent pH stat and an 8.0 N KOH resevoir upon mixing of the components. Samples were taken every 5.0 min and assayed immediately by the thio- barbituric acid assay (83) to determine losses over time of the TBA- positive material. (A) Time of incubation study of 2-keto-3-deoxy-D- quinovonic acid at pH 11.0, and (B) time of incubation study of 2-keto- 3-deoxy-0-fuconic acid at pH 11.0. 39 KDQ A 9O 80 70 %:%8&%%Qe%%9o S<_eue_e_moa «up ez_z_<=u¢ ac ezuueua KDF _%8&Q:W%%&m; 20 3O 4O 50 6O 7O 80 90 TIME OF INCUBATIDN (minutes) 10 4O purification steps. Quantitation of the purified product by the semi- carbazone assay gave a molar extinction coefficient of 60,000 for the TBA assay at 551 nm. KHG was prepared by the procedure of Aronson et al. (90), using the colorimetric assay of Kramer et al. (91), to detect the disappear- ance of glyoxylic acid in the reaction mixture. In the column chromat- ography, eluate fractions collected were assayed for glyoxylate as mentioned, for pyruvate and oxaloacetic acid by lactate dehydrogenase and malic acid dehydrogenase, respectively, and the reaction product (KHG) by the semicarbazone assay. Quantitation of the purified product as the semicarbazone gave reported yields; this product cannot be cleaved by periodate to give B-formyl pyruvate in the TBA assay and therefore no extinction coefficient is reported for this product. 2-Keto-3-deoxy-6-phospho-D-galactonate (barium salt) was a gift from the laboratory of H.A. Hood. The barium salt was taken in the solid state and converted to the corresponding potassium salt by treat- ment with Dowex-50H (K+) in solution of known volume; quantitation of this phospho-ester was by the TBA assay and correlated to the weight amount for accurate concentration determination. ggzymatic Assays One unit of enzymatic activity is defined as the amount of enzyme that catalyzes the conversion of one micromole of substrate to product per minute at 30°C. Assays involving the oxidation or the reduction of pyridine nucleotides were monitored in microcuvettes with a 1.0-cm path length at 340 nm, using a thermostated Gilford spectrophotometer, 41 model 2400. Other assays that involved incubations were done in a temperature-controlled Precision Scientific Company water bath. The concentrations of the substrate added to an assay were determined on a weight basis of the added substrate, except in the case of carbohy- drate derivatives, which were assayed by specific colorimetric assays as described above. Dehydrogenase Assays (using D-Galactose, D-Galactonate, D-Fucose, or D-Fuconate as substrate). Assays of 0.2-ml volume consisted of 50 mM Hepes buffer (pH 7.0), 10.0 m 119012, 5.0 mN NAD+ or NADP+, 40.0 m D-galactose, D-galactonate, D-fucose, or D-fuconate, and rate-limiting amounts of cell extract. D-Galactonate (D-Fuconate),Dehydratase. Assays of 0.3-ml volume were composed of 50.0 nN Pipes buffer (pH 7.0), 10.0 mM MgCl2, 1.0 m EDTA (pH 7.0), 20.0 mM D-galactonate (K+) or 30.0 mM D-fuconate (K+), plus rate-limiting amounts of the dehydratase. The reaction mixture in 1.3 X 10.0 cm culture tubes were preincubated at 30°C before adding the enzyme to initiate the enzymatic assay. At appropriate time per- iods, the assay was stopped by the addition of 1.0 ml of semicarbazone reagent (1.0% semicarbazide-HCl plus 1.5% sodium acetate, in distilled- water) directly to the sample. Once added, the samples were incubated at 30°C for an additional 20 min to form semicarbazone products of the d-keto acids produced by the enzyme. Samples were then diluted to 5.0-ml volume with distilled water and centrifuged with the aid of an Inter- national clinical centrifuge to remove protein precipitates. Controls to correct for protein were minus substrate (D-fuconate or D-galactonate). 42 Semicarbazones for d-keto acids were quantitated using the molar ex- tinction coefficient of 10,200 at 250 nm (15). Under the conditions of the assay, an absorbance of 2.04 was equal to 1.0 nmol of 2-keto- 3-deoxy-aldonate. The reaction was linear in the range of 5.0 nmol to 5.0 nmol product. One unit of dehydratase was defined as the amount of enzyme that converted 1.0 nmol of aldonic acid to d-keto acid per minute. 2-Keto-3-Deoxy-D-Galactonate Kinasey(KDGal kinase). The reaction mixture of 0.2-ml volume consisted of 50.0 mM Hepes buffer (pH 7.0), 10.0 mM MgCl2, 0.25 mM NADH, 2.5 mM PEP, 2.5 mM ATP, 11.0 mM 2-keto- 3-deoxy-D-galactonate (enzymatic), excess pyruvate kinase and lactate dehydrogenase, and rate-limiting amounts of the enzyme. Controls to correct for ATPase and NADH oxidase were minus KDGal as substrate. 2-Keto-3-Deoxy-G-Phospho-D-Galactonate Aldolase (KDPGal aldolase). Assays of 0.2-ml volume consisted of 50.0 mM Hepes buffer (pH 8.0), 10.0 mM MgCl2, 0.25 mM NADH, 2.5 mM 2-keto-3-deoxy-6-phospho-D-galac- tonate (enzymatic), excess lactate dehydrogenase, and rate-limiting amounts of the enzyme. Controls to correct for NADH oxidase were minus KDPGal as substrate. 2-Keto-3-Deoxy-D-Fuconate Aldolase (KDF aldolase). The reaction assay of 0.2-ml volume consisted of 50.0 mM Hepes buffer (pH 8.0), 10.0 mM MgClz, 0.25 mM NADH, 1.0 mM EDTA (pH 7.0), 10.0 mM 2-keto-3- deoxy-D-fuconate (chemically synthesized or enzymatic), excess coupling enzyme and rate-limiting amounts of the aldolase. Lactate dehydrogenase was used as the coupling enzyme when determining pyruvate production 43 by the aldolase. Alcohol dehydrogenase (horse liver) was used as the coupling enzyme when determining aldehyde (D-lactaldehyde) production by the aldolase; this coupling enzyme shows a broad substrate specific- ity for aldehydes and has been shown to act very well on D-lactaldehyde (86). Controls were run without KDF to correct for NADH oxidase when crude extracts were assayed by either of these assays. End-Point Assays for the Determination of Pyruvate, D-Lactalde- hyde and Oxaloacetic Acid. The reaction mixture of 0.2-ml volume con- sisted of 50.0 nu Hepes buffer (pH 8.0), 10.0 «11119012, 0.25 mN NADH, 0.025 to 0.10 mM substrate, and excess coupling enzyme. A substrate blank was used to correct for dilution when the coupling enzyme was added to initiate the assay. The end-point of the assay was judged complete when no further NADH was oxidized, as measured at 340 nm. Pyruvate was assayed with beef liver lactate dehydrogenase (92), D-lactaldehyde with horse liver alcohol dehydrogenase (86), and oxalo- acetic acid with malic acid dehydrogenase from yeast (93). Quantita- tion of the oxidation of NADH in the assay was done assuming an extinc- tion coefficient of 62,200 at 340 nm. Selection of D-Fuconate- and D-Galactonate-Negative Mutants An overnight culture of §;_pneumoniae PRL-R3,U', strain CH-101 (see Results, this section), which is constitutive for growth on D-fuc- onate, was grown in nutrient broth and was harvested in a sterile centri- fuge tube. The cell pellet was washed once in mineral-salts medium and resuspended in 5.0 ml of mineral medium containing 0.2 M ethylmethane sulfonate (EMS) (94). After incubating the EMS-treated culture for 44 2.0 hrs at 30°C on a reciprocal shaker the cells were harvested and suspended in 100 ml of mineral-medium broth which contained 0.5% O-glu- case and 0.005% uracil. The culture was then incubated overnight, allowing for a ten-fold increase in cell number. Once grown, the culture was harvested and washed with mineral-medium. Hashed cells were then suspended in 0.5% D-fuconate (K+)-mineral-medium at a cell 8 cell/ml. Cultures were then incubated concentration of 2.0 X 10 until a doubling of-the cell number was observed, at which time the culture was made 10.0 mM in D-cycloserine, and penicillin-G was added at a concentration of 2000 units/ml (100). The cultures were then incubated for 6.0 hrs, allowing for 2-3 times the generation time on D-fuconate to occur. Cells which were unlysed by this treatment were enhanced in number by suspending them in 7.0 ml of mineral-medium plus 0.5% D-glucose and 0.005% uracil, after washing, and incubated for an overnight period of growth at 30°C. The D-glucose-grown cultures were then plated onto mineral-media agar containing 0.5% D-fuconate, 0.005% D-glucose, and 0.005% uracil. Plates were inoculated with about 200 viable cells per plate, incubated. at 30°C, and checked for the appearance of colonies after 24 and 48 hrs of incubation. Pin-point colonies were taken from the plates with a culture loop and transfered to D-glucose-mineral-medium (7.0 ml) where they were grown overnight. Colonies were screened for growth on D-fuc- onate and D-galactonate by streaking respective mineral-agar plates from these cultures. Growth was monitored visually upon incubation of the plates; positive growth showed a definite lawn of transluscent 45 cells, whereas negative or sparse growth showed no lawn or at best isolated colonies which most likely represented revertant colonies. Suspect mutants for either D-galactonate or D-fuconate negativity or both were then screened for normal growth on various carbohydrates by streaking onto appropriate plates. Enzymatic characterization of the mutants was done using stand- ard assay procedures for the dehydratase and aldolase of the D-fuconate pathway as described in this section. Nutrient broth cultures were used for crude extracts when testing for the dehydratase. Crude extracts for the KDF aldolase were prepared from nutrient broth cultures supple- mented with 0.5% D-fuconate; D-fuconate was added at half-maximal growth on nutrient broth and incubated for 3.0 hr in the presence of the aldonic acid. In all cases, a control for both the normal levels of the dehydra- tase and the aldolase were run using a culture of the strain CH-IOI grown under the same conditions. Bevertants for the Dehydratase-Negative Mutant Dehydratase-negative mutant (CH-103) was grown in nutrient broth overnight and plated onto 0.5% D-galactonate, 0.005% D-glucose, and 0.005% uracil mineral-medium-agar plates. Plates were then incubated . at 30°C and checked for growth at 24 and 48 hrs. Both pin-point and large colonies were picked and grown overnight in nutrient broth. Colon- ies were screened as described, for growth on D-galactonate and D-fuc- onate-mineral-medium plates. Crude extracts were prepared from nutri- ent broth cultures of colonies screened and assayed for the dehydratase by standard procedures. 46 IMViC Tests for the Bacteriological Classification of all Strains Standard procedures for IMViC tests used in this section were adapted from a published source (96). As controls, the parental strain, Klebsiella pneumoniae PRL-R3,U' (grown at 30°C) and Escherichia coli, strain B/r 353:2;(grown at 37°C) were used in all tests. Indole production was determined using the Ehrlich Rosendale reagent with bacterial strains having grown 2.0 days in peptone water. A positive test gave a rose pink color upon the addition of the reagent to the ether extracted culture. A negative test gave no color change upon the addition of the reagent. Indole, at a concentration of 0.05 M in peptone water, was used as the control. Quantitation of the color yield was not made. Acid production from 3-day-old glucose-phosphate-broth cultures of the strains was determined by the addition of a few drops of methyl red directly to the culture. Positive acid production in this test gave a definite red color. Negative test left a pale yellow color upon the addition of the dye. No quantitation of the color yield was made. A broth culture titrated to a pH of 3.0 was used as control. Voges-Proskauer tests, for the detection of acetoin or acetyl- methylcarbinol, were performed on 2-day-old glucose-phosphate-broth cultures of all strains tested. A positive test gave a pink color which developed upon standing after the addition of KOH directly to the culture. Acetoin at 0.05 M in the medium was used as the control. A negative test gave no color change. No attempt was made to quanti- tate the color yield. Growth on citrate was checked using Simon's citrate medium with 47 and without 0.005% uracil. Inoculations were made from peptone water cultures grown overnight. Cultures in the test, were incubated at 30°C or at 37°C on a reciprocal shaker and turbidity measurements were made at 24 hrs of growth. Positive growth gave an absorbance of 0.3 at 600 nm or better; a 0.5% D-glucose-mineral-medium-broth culture inocu- lated with the strain CH-101 was used as a positive control. Sources of Materials All reagents (including carbohydrates, enzymes, and assay co- factors) included in this section were obtained from the Sigma Chemical Company, unless noted otherwise. Special notice is given to the com- mercial sources of the following materials: Bacto-agar and nutrient broth, used for mineral-medium plates and for broth cultures were obtained in the dry state from Difco Laborator- ies. Glass beads used in the preparation of crude extracts were obtained as Microspheres (125-80 microns) from La Pine Scientific Company. Bovine serum albumin, used as the Lowry protein standard, was obtained from Sigma Chemical Company. N,0-Bis-(Trimethylsilyl)-trifluoroacetamide (BSTFA) was obtained from the Pierce Chemical Company. Packing material, 3% 0V-17 on Gas-Chrom 0, was obtained from Applied Science Laboratories Incorporated as a dry powder. Dowex resins were obtained from the Sigma Chemical Company as dry beads, which were washed with HCl, NaOH. and methanol to extract impurities of the commerical preparations before use. 3-Methyl-2-benzothiazolinone hydrazone hydrochloride monohydrate (MBTH) used in the colorimetric assay of Paz et al. (82) and acetalde- hyde were obtained from the Aldrich Chemical Company. Thiobarbituric 48 acid (TBA) used in colorimetric assays for the detection of 2-keto-3- deoxy sugar acids, and semicarbazide.HCl used in the determination of o-keto acids were obtained from Sigma Chemical Company; TBA was recryst- allized from hot ethanol after passing through aluminum oxide (BDH Lab- oratories) before use, as the commerical preparation has colored impur- ities which are exhibited in aqueous solution. Ninhydrin was obtained from the Pierce Chemical Company. Glycolaldehyde and glyoxylic acid were obtained from Sigma Chemical Company. Ethylmethane sulfonate was obtained from Eastman-Kodak. Indole and acetoin, used as standards in the IMViC tests were obtained from Sigma Chemical Company and Aldrich Chemical Company, respectively. Horse liver alcohol dehydrogenase was obtained from the Horthington Biochemical Corporation. RESULTS Selection of Mutant Strain CH-IOl (D-Fuconate-Positive) from Klebsiella Pneumoniae PRL-R3,U', and Gross Characterization Hhen cells of Klebsiella pneumoniae PRL-R3,U' from either nutrient broth or D-glucose-mineral-medium-broth cultures were inoculated into 0.5% D-fuconate-mineral medium plus uracil, initiation of growth took approximately 60 hrs at 30°C on a reciprocal shaker. If such cultures were then plated onto mineral-media-agar plates containing 0.5% D-fuconate plus 0.005% D-glucose and 0.005% uracil, and incubated at 30°C for 48 hrs, two main colony types developed: pin-point and large-size colonies. The large colonies could grow to an absorbance of 0.3 at 600 nm in approximately 8 to 10 hrs of incubation when transferred to mineral-medium broth supplemented with D-fuconate and Uracil, as above. However, when pin-point colonies were transferred to D-fuconate-mineral medium plus uracil growth took the normal 60-hr incubation as seen with_ the parental strain. Moreover, normal (i.e., parental strain) growth patterns were seen with both colony types on a variety of carbohydrates other than D-fUconate. These results indicated that a mutant strain with the augmented ability to use D-fuconate as the sole source of carbon for growth was selected from the parental strain. The large colonies on D-fuconate-mineral-agar plates were then designated D-fuconate-positive mutants. Selection of the strain CH-IOI was performed in a similar manner as follows. 49 50 The parental strain, §;_pneumoniae PRL-R3,U-, was inoculated into fresh 0.5% D-fuconate-mineral broth, plus 0.005% uracil, and incubated at 30°C on a reciprocal shaker for approximately three days. Upon devel- opment of a turbid culture, aliquots of the culture were diluted into mineral-salts medium and used to inoculate several 0.5% D-fuconate- mineral-medium-agar plates, plus 0.005% uracil, at about 50 cells per plate. These plates were then incubated at 30°C and one colony which developed after about 48 hrs was then picked and grown overnight in nutrient broth. The nutrient-broth culture was then preserved by trans- ferring cells to nutrient-agar slants which could be stored at 4°C after the normal overnight incubation. These slant cultures were then used for inoculants of the strain CH-101 in all subsequent studies noted in this thesis. To show'gross growth characteristics of both the parental strain and the D-fuconate-positive mutant strain, CH-101, these strains were grown on the aldonic acid D-fuconate and its analogue D-galactonate using standard mineral-salts medium. D-Glucose was used as control for positive growth with both bacterial strains. As the growth curve for the parental strain shows (Fig. 4), both D-glucose and O-galactonate were used readily as carbon sources, with a short lag period, whereas growth on D-fuconate began long after the other two carbohydrates were exhausted. Growth of the mutant strain, CH-101, on either D-glucose- or D-galactonate-mineral medium showed similar patterns as seen with the parental strain (Fig. 5), but growth on D-fuconate occured much sooner; approximately 3.5 hrs were required to induce this strain to use D-fuconate. From the growth curves, generation times for the 51 Figure 4. Growth of the Klebsiella ppeumoniae PRL-R3,U' (paren- tal) strain on D-glucose, D-galactonate, and D-fuconate. An overnight culture of the parental strain in nutrient broth was used to inoculate three 7.0-ml cultures containing 0.5% 0-glucose-, D-galactonate-, and D-fuconate-mineral broth (plus 0.005% uracil). Cultures were incubated at 30°C on a reciprocal shaker and turbidity measurements were taken at various times by sampling with sterile pasteur pipetes and reading the resultant absorbance at 600 nm. Plots were made of corrected absorb- anCe versus the incubation time. Generation times were estimated as the time required for the culture to double from an absorbance of 0.15 to 0.30 at 600 nm. ABSORBANCE (600 nm) 52 O O 0000-4 N l O u l 0.03-1 0.0 D-GLUCOSE A o D-GALACTONATE [X D-FUCONATE '__Jv glr I f 6 62 64 TIME (hours) 66 68 53 Figure 5. Growth of the D-fuconate-positive mutant strain, CH-IOI. on D-glucose, D-galactonate, and D-fuconate. Conditions are described in figure 4. ABSORBANCE (600 nm) 54 D-GLUCOSE A ° O-GALACTONATE U‘IO‘NWWO 14111 O C) 00000-4 0'2 ‘ a D-FUCONATE LjeL 19> a 0‘ O O m a 0.00 TIME (hours) 55 parental strain on D-glucose (0.72 hr), D-galactonate (0.89 hr), and D-fuconate (2.20 hr) and the strain CH-IOI on D-glucose (0.78 hr), 0-galactonate (1.06 hr), and D-fuconate (1.22 hr) were obtained. In addition, when strain CH-101 was grown on equimolar concentrations of D-glucose, D-galactonate, or D-fuconate in mineral broth, equal total growth was observed (Table 2). These results show that the O-fuconate-positive strain can readily assimilate the aldonic acid D-fuconate as the sole carbon and energy source for growth. . In summary, the process of obtaining a strain capable of us- ing D-fuconate as a growth substrate apparently involved the selec- tion of a D-fuconate-positive mutant in the normal bacterial popula- tion. As seen from the growth curves, the parental strain required a longer pre-incubation before using D-fuconate, but once growth had begun, the generation time was similar to (about one-half) that of ‘ the D-fuconate-positive strain. In this regard, serial transfer of the mutant strain on D-fuconate-mineral broth, as done by Mayo and Anderson (12) with Aerobacter aerogenes on L-mannose, failed to pro- duce subsequent strains with shorter generation times on D-fuconate. If strains with shorter generation times were found in the case where §;_pneumoniae acquires the ability to use D-fuconate for growth, than factors other than the single-step selection of a D-fuconate-positive mutant would have been considered. 56 Table 2. Growth yield of strain CH-101 on D-glucose, D-galact- onate, and D-fuconate. Broth cultures of the mutant strain CH-IOI were grown at 30°C for 48 hrs (see Materials and Methods for details) to exhaust the carbon sources. Corrected optical density readings (absorbance) were taken for each carbohydrate at the indicated con- centrations at the end of this incubation. CARBOHYDRATE CARBOHYDRATE CONCENTRATION CORRECTED ABSORBANCE pg, 600 nm D-GLUCOSE 2.8 0.37 5.5 ‘ 0.73 D-GALACTONATE 2.8 0.35 5.6 0.72 D-FUCONATE 2.8 0.35 5.6 0.69 57 Elucidation of the Enzymatic Reactions Involved in the Catabolism of D-Fuconate Enzymatic Reactivity of D-Fuconate. Several possible avenues were considered as candidates for the catabolic pathway of D-fuconate. Those involving initial dehydration or dehydrogenation of D-fuconate were considered the most plausible and were examined first. In add- ition, dehydrogenases for D-galactose and D-fucose were looked for with the idea that their absence would indicate that the D-fuconate pathway was unique rather than part of the pathway for the metabolism of an aldose. Other possible reactions, such as D-fuconate epimeriza- tion, would be examined only if D-fuconate dehydration or dehydrogena- tion could not be detected. For the following studies, extracts were prepared from freshly harvested cells. The postulated dehydrogenase reactions were tested using either NAD+ or NADP+ as cofactor at both neutral and basic pH ranges. I was not able to detect (less than 0.001 nmol product X min'lx mg protein'l) such activities on the substrates D-fucose, D-galactose, D-fuconate, or D-galactonate (see Materials and Methods for conditions) using cell extracts from D-galactose-, D-galactonate-, or D-fuconate-grown §;_ppgg;, mggigg, strain CH-IOI. However, D-fuconate dehydratase activity (0.07 1x mg protein'l) was detected in all of the to 0.22 umol product X min' above extracts. This activity was dependent on the carbon source used for growth as determined by the semicarbazone assay for a-keto acid production (15). The formation of d-keto acid was found to be linear with time (Fig. 6) and proportional to protein concentration (Fig. 7) 58 a l m l E 2 " l j; ‘ 120.0 mg ' a 60.0 L19 ‘6 = 8 r: 3 “.- g 3 _ 30.0 119 - a. V F! ”i O 8 o 61’ '4- __ 15.0 119 .. .9 52 A. O I I l I O 10 20 30 40 50 Time (minutes) Figure 6: Linearity of the D-fuconate dehydratase assay with respect to time. The ug amounts for each curve indicate the amount of protein (crude extract) used in the assav. LO 1 1 put A LI- 9 8 v 0 l “A c-I ‘- 00 F0 CH O: U: 3'!- LBS I\ O'H- >~§ x L §v m... I "PE 00 H‘- U ‘f N Q 1 I O 50 100 150 Protein (pg) Figure 7. Proportionality of the D-fuconate dehydratase activ- ity with protein concentration. The data are taken from Fig. 6. 59 when crude extracts of the strain CH-IOI, grown on D-fuconate, were assayed using 30.0 mM D-fuconate and 1.0 mM EDTA in the reaction mix- ture. Dehydratase activity was also detected with D-galactonate as 1 1); this substrate (0.19 to 0.64 umol product X min' X mg protein- activity was also dependent on the carbon source used for growth and was linear with time and proportional to protein concentration (data not shown). The inital reaction involving D-fuconate was therefore a dehydra- tion leading presumably to 2-keto-3-deoxy-D-fuconate (KDF). A detailed study of this dehydratase is presented in Section II of this thesis. gpzymatic Reactivity of Z-Keto-3-Deoxy-D-Fuconate. The enzymatic breakdown of 2-keto-3-deoxy-D-fuconate in crude extracts by either an aldol-cleavage or an oxido-reductase reaction was next examined. 2-Keto- 3-deoxy-D-fuconate aldolase activity was detected in extracts of strain 1 CH-101 grown on D-fuconate (0.11 nmol product X min' X mg protein-1) but not on D-glucose or D-galactonate (less than 0.001 umol product 1 ‘1). X min‘ X mg protein Assays of crude extracts prepared from cells grown on any of these three substrates with either NAD+ or NADP+ showed no oxido-reductase activity on KDF (less than 0.001 umol product X min"1 X mg protein'l). To clearly demonstrate the existence of a 2-keto-3- deoxy-D-fuconate aldolase in extracts of the D-fuconate-positive strain, the products of the aldolase-catalyzed cleavage were quantitated enzy- matically. The production of pyruvate was measured using the standard reac- tion mixture and beef heart lactate dehydrogenase (LDH) as the coupling 60 enzyme; LDH prefers pyruvate as substrate having a rate of reduction over ten-fold higher than other d-keto and a,Y-di8tO acids (92). D-Lactaldehyde production was measured using horse liver alcohol dehy- drogenase (HLADH) as the coupling enzyme in the reaction mixture; HLADH possesses a broad substrate specificity for aldehydes, among which are acetaldehyde, glycolaldehyde, formaldehyde, propionaldehyde, butyraldehyde, valeraldehyde, isobutyraldehyde, glyceraldehyde, and lactaldehyde (86). Results of a study using either chemically synthe- sized KDF or enzymatically prepared KDF (see Section II, Results) with these coupling enzymes (Table 3) show that not only are both pyruvate and an aldehyde (presumably D-lactaldehyde from the chemical structure of KDF) produced from the substrate Z-keto-3-deoxy-D-fuconate, but that both products are produced at the same rate. These data confirm that an aldolytic activity for KDF is present in crude extracts of the strain CH-101 when grown on D-fuconate. Further investigation of the KDF aldolase from extracts of the strain CH-IOI grown on D-fuconate revealed a pH optimum of 8.0 (Fig. 8). In addition, a divalent metal was absolutely required for activity as demonstrated by complete inactivation of the enzyme with 1.0 mM EDTA and restoration of maximal activity with 10.0 mM MgCl2 (Fig. 9). Exam- ination of the effect of substrate concentration on the velocity of the reaction revealed that the Km of the enzyme for 2-keto-3-deoxy-D-fucon- ate was approximately 0.25 mM. Using 2.5 mM substrate, the assay for the aldolase was found to be constant with time (Fig. 10) and propor- tional to protein concentration (Fig. 11). Crude extracts from D-fuc- onate grown cells of this strain were found to possess additional 61 Table 3. KDF aldolase reaction products: measurement of the reaction products with appropriate coupling enzymes. The aldolase was assayed as described in the text with lactate dehydrogenase and alcohol dehydrogenase (HLADH) as coupling enzymes to measure the production of pyruvate and D-lactaldehyde, respectively. A crude extract of the strain CH-101 grown on D-fuconate was used. Standard assays for the KDF aldolase were followed according to established protocols using either chemically or enzymatically prepared 2-keto-3-deoxy-D-fuconate (KDF) as substrate. Assays of the chemically synthesized KDF contained 1.59 munits of activity and assays of enzymatic KDF contained 1.12 munits of activity. COUPLING ASSAY USED SPECIFIC ACTIVITY OF THE ASSAY ON SYNTHETIC KDF ENZYMATIC KDF nmol NAD+/mina nmol NAD+jmin° LACTATE DEHYDROGENASE ONLY 1.64 2‘. 0.08 . 1.10 1 0.08 ALCOHOL DEHYDROGENASE ONLY 1.53 _+_ 0.11 1.13 i 0.03 LACTATE DEHYDROGENASE a 3.42 1 0.06 2.37 i 0.08 ALCOHOL DEHYDROGENASE a these values are averages from three determinations : 5.0. 62 Figure 8. pH optimum of the KDF aldolase. Crude extracts of the strain CH-IOI were grown on 0.5% D-fuconate-mineral-medium broth and pre- pared as described in Materials and Methods. Maleic aCid was used below pH 7.0; Tris-maleate buffer between pH 7.0 and 8.0;,and glycine buffer above pH 8.0 at concentrations of 50.0 mM in the assay. Maximal activity was 0.40 umol product X min.1 in the assay at pH 8.0. Hepes buffer at pH 8.0 gave the same velocity as Tris-maleate (not plotted). 63 a 8 q u a 1 as 8 cm 3 >:>:u< 35x5. ”3 h5g5: °d 64 Figure 9. Metal ion requirement of the KDF aldolase. Crude ex- tracts of the strain CH-IOI grown on 0.5% D-fuconate-mineral broth were prepared as described in Materials and Methods. Crude extracts were then treated with 1.0 mM EDTA and chromatographed on a column of Sephadex G-25 (0.5 X 10.0 cm) equilibrated with 0.05 M potassium phosphate buffer (pH 7.0). KDF aldolase activity was assayed using 2-ketO-3-deoxy-D-fuconate as substrate in the lactate dehydrogenase coupled assay with the MgClz.concentration varied. In all trials, reaction mixtures were made 10.0 mM in KCl to assure activity of the coupling enzyme. Maximal activity was 0.40 umol product X minute‘l. 65 a cop cm on as oo cm cc. on cu o— o >p_>_hu< a5 v m 0 15 30 Protein (pg) Figure 11. Proportionality of the 2-ket0-3-deoxy-D-fuconate aldolase activity with protein concentration. These data are taken from Fig. 10. 67 activities on the substrates 2-ketO-3-deoxy-D-quinovonic acid, 2-keto- 3-deoxy-D-galactonate, and 2-keto-3-deoxy-D,L-arabonate when tested in the aldolase assay. These activities showed a similar inactivation by EDTA and divalent metal requirement (Table 4) as that found with the substrate 2-keto-3-deoxy-D-fuconate suggesting that the aldolase of the D-fuconate pathway had a varied substrate specificity. Hhen mixtures of these substrates were assayed, rates were found to be competitive rather than additive (Table 5) suggesting that one enzyme acted on all four substrates. Inducibility of the D-Fuconate and D-Galactonate Pathway Enzymes on Various Carbohydrates Subsequent to the initiation of studies on the catabolic path- way of D-fuconate, a pathway for the catabOlism of D-galactonate was discovered in both the parental strain and the D-fuconate-positive strain, CH-101, of §;_pneumoniae. In much the same manner as described for the D-fuconate pathway, activities for a D-galactonate dehydratase, a 2-keto-3-deoxy-D-galactonate (KDGal) kinase, and a 2-ketO-3-deoxy- 6-phospho-D-galactonate (KDPGal) aldolase (but not a 2-keto-3-deoxy- D-galactonate aldolase) were found in crude extracts of the strain CH-101 grown on D-galactonate. Due to the analogous nature of these two aldonic acids, it was of interest to determine how closely related their pathways were. Investigations of the inducibility of the path- way enzymes proved to be useful in this regard. Hhen crude extracts prepared from both the parental strain and strain CH-101 grown on various carbohydrates were examined for pathway 68 Table 4. Effect of MgCl2 on aldolase activities in EDTA- treated extracts. The following substrates were assayed using standard protocol for the KDF aldolase assay and EDTA-treated crude extracts from D-fucon- ate-grown cells (see figure 9, for conditions). The reaction mixture consisted of 0.2-ml volume and the following components: Hepes buffer, 50.0 mM. pH 8.0; MgClz. 10.0 mM; NADH, 0.25 mM; excess lactate dehydro- genase; EDTA, 1.0 mM; and the following keto-deoxy sugar acids: 2-ket0- 3-deoxy-D-fuconate (KDF), 10.0 mM; 2-keto-3-deoxy-D-galactonate (KDGal), 5.0 mM; 2-keto-3-deoxy-D,L-arabonate (KDA), 5.0 mM; 2-keto-3-deoxy-D- quinovonic acid (K00), 10.0 mM; and 4-hydroxy-2-keto-D,L-glutarate (HKG), 10.0 mM. SUBSTRATE ACTIVITY MINUS MgCT2 PLUS Mgc12 mm mun. KDF < 0.02 0.378 KDGal < 0.02 0.113 KDA < 0.02 0.251 KDQ < 0.02 0.175 HKG < 0.02 < 0.020 69 Table 5. Additive enzymatic activity studies on keto-deoxy sugar mixtures with KDF aldolase. Limiting amounts of D-fuconate-grown strain CH-IOI crude extract were assayed for aldolase activity using the follow- ing substrates and established assays. The components of the assay were: Hepes buffer, 50.0 mM, pH 8.0; MgClz, 10.0 mM; EDTA, 1.0 mM; NADH, 0.25 mM; excess lactate dehydrogenase; and KDF (10.0 mM), KDGal (5.0 mM), KDA (5.0 mM), and K00 (10.0 mM). MIXTURE TRIAL SUBSTRATE RESULTANT ACTIVITY unitsimla _KDF & KDA KDF 0.378 1 0.004 KDA 0.251 g 0.004 KDF G KDA 0.346 :_0.050 KDF a K00 KDF 0.378 :_0.004 KDQ 0.175 :_0.005 KDF 3 K00 0.183 1 0.016 KDF & KDGal KDF 0.378 1 0.004 KDGal 0.113 :_0.011 KDF & KDGal 0.298 1 0.005 a these values are averages from three determinations 1.5.0. 7D enzymes involved in the catabolism of D-fuconate and D-galactonate the following trends were observed (Table 6): (i) Both the parental strain and strain CH-IOI possessed D-galactonate dehydratase, 2-keto-3-deoxy— D-galactonate kinase, 2-keto-3-deoxy-6-phospho-D-galactonate aldolase, and D-fuconate dehydratase activities on D-galactonate-mineral broth. However, these activities were elevated in the strain CH-IOI over the parental strain when grown on this substrate. (ii) The parental strain showed absolute inducibility of all the D-galactonate pathway enzymes and the D-fuconate dehydratase by D-galactonate and the KDF aldolase by a D-galactonate/D-fuconate mixture. Hhereas, the strain CH-101 ex- hibited constitutive production of all enzymatic activities noted above except for the KDF aldolase; this latter enzyme apparently reqUired D-fuconate as an inducer. (iii)The D-fuconate dehydratase appeared in both Klebsiella strains at about one-third the activity found for the D-galactonate dehydratase regardless of the carbon source used for growth. Both the D-fuconate and D-galactonate dehydratase activities showed similar patterns of induction in both strains. (iv) The KDF aldolase inducibility suggested that the dehydratase product from D-fuconate is the actual inducer of this enzyme as the parental strain ' required a mixture of both D-fuconate and D-galactonate to exhibit such activity. (The KDF aldolase activity was not detected on any other carbon source but D-fuconate; testing for this activity on the sugar acids D-glucarate, D-galactarate, D-glucuronic acid, and D-galacturonic acid did not reveal Specific activities higher than that found for either strain on D-galactonate.) 71 Table 6. Inducibilities for enzymes of the D-fuconate and D-gal- actonate pathways. Crude extracts were prepared from cultures grown overnight in 0.5% carbohydrate-mineral-medium broth (plus 0.005% uracil) at 30°C and 7.0-ml volume, as described in Materials and Methods. . 18888888 05 x 1888885 x 8888888 883: 8oc.o 88:8 8888 88828888 88888888 88 o:_8> 8 n 882 H g 72 . . 8888 888.8 888888-8 8888 888 8 2282228 2282288 888 8 88 88.8 28 888288222-8 88882888882 2282228 888.8 888.8 2288228 888 8 88228882828-8 8888 88mmim8mm88 8888 .8 888.8 888.8 888.8 888.8 888.8 2.8 88828888-8 a 00 MCOHUQ P81: 882 888.8 888.8 888.8 888.8 28828 82882882 888-28 882 888.8 888.8 888.8 888.8 888882828-8 282288 882 888.8 888 8 888.8 888.8 8888888-8 882 888.8 888.8 888.8 888.8 882288288-8 -8.82-8mm . 888.8 888.8 888.8 888.8 888.8 88228888-8 8888828888 .2 882 888.8 888.8 888.8 888.8 88228882828-8 . . . . . - 282288 888 8 888 8 888 8 888 8 888 8 8xwu8nwmuuwum “ 82828228 882 882 882 882 882 28828 82882882 -2.82-8mm 882 888.8 888.8 888.8 888.8 88228882828-8 8882888888 .2 iimmmmmmammm.rmmm:ouocpccmox ouceouucpcwiaiax 88828888-: 88888888888-a_ z8<¢8m 88288882 882282 88282288288 118 2888828 828288828 88828882 88888828 . _ 73 These data are consistent with the following suppositions: Firstly, that the acquired ability of 5;.ppeumoniae to utilize D-fuc- onate as a sole carbon source for growth is involved with the constit- utive production of the D-fuconate dehydratase. Secondly, that the dehydratase activities on D-fuconate and D-galactonate, in both the parental strain and the strain CH-101, were the result of one enzyme. Lastly, that the D-fuconate pathway is independent of the D-galactonate pathway beyond the initial dehydration reaction. Support for these suppositions was provided by an analysis of D-fuconate- and D-galacton- ate-negative mutants derived from the strain CH-IOI. Mutant Analysis and Verification of Enzyme Deficiency 5; pneumoniae, strain CH-IOI, was subjected to mutagenesis with ethylmethane sulfonate, screened for cells impaired for growth on D-fuconate by treating with a mixture of penicillin-G/D-cycloserine, enhanced by growing in D-glucose-mineral broth, and then plated onto mineral-medium supplemented with 0.5% D-fuconate, 0.005% D-glucose, and 0.005% uracil (see Materials and Methods, this section). 0f five plates inoculated (about 200 colonies per plate), 11 pin-point colonies were found. Screening of cultures from these pin-point colonies on D-fuconate- and D-galactonate-mineral-agar plates revealed that one exhibited impaired growth on both D-fuconate and D-galactonate. 5 Showed impaired growth on D-fuconate, and 10 showed normal growth on D-galactonate (Table 7). All strains showing impaired growth on D-fuconate and/or D-galactonate were screened for growth on a variety of carbohydrates to determine if other pathways were affected by the mutagenesis. Strain CH-102, exemplifying mutant strains normal on 74 Table 7. Screening of mutagenized cultures for growth on D-fucon- ate and D-galactonate. The strain CH-IOI was treated as in the text. Pin-point colonies, numbered arbitrarily, which developed from D-fucon- ate plates supplemented with growth-limiting amounts of D-glucose were grown overnight in 0.5% D-glucose-mineral-medium broth. Plates were checked for positive growth using an untreated culture of the strain CH-101 on D-glucose as control after 48 hrs of incubation at 30°C. BACTERIAL STRAIN GROWTH* ~ D-GLUCOSE D-GALACTONATE D-FUCONATE CH-101 + + + CH-IOZ + + - CH-103 + - - CH-104 + + + CH-105 + + - CH-106 + + + CH-107 + + - CH-108 + + + CH-109 + + - CH-110 + + + CH-111 + + - CH-112 + + + * (+) is positive growth; noticeable bacterial lawn. (-) is negative growth; no bacterial lawn, at best isolated colonies ' (revertants). 75 D-galactonate but impaired for growth on D-fuconate, and strain CH-103, impaired for growth on both aldonic acids, showed normal growth patterns on all carbohydrates but D-fuconate and/0r D-galactonate when compared with the parental strain and strain CH-IOI (Table 8). Strains CH-104, CH-106, CH-108, CH-IIO, and CH-112 were no different than the parental strain CH-101 when checked for growth on D-fuconate or D-galactonate and therefore were not studied further. To check mutant strains for deficiences in enzymes of the D-fuc- onate pathway, crude extracts were prepared form cultures grown under conditions which normally exhibited these activities in the strain CH-IOI (Table 9). Enzymatic assays confirmed that the strain (CH-103) which did not exhibit growth on either D-fuconate or D-galactonate was in fact deficient in the dehydratase for either of these substrates. This strain was classified dehydratase-negative. All other strains having normal growth on D-galactonate (CH-102, CH-105. CH-107, CH-109, and CH-Ill) were found to have normal levels of both the D-fuconate and D-galactonate dehydratase (specific activities of 0.40 and 0.13 1 1, respectively). umol product x min' X mg protein' 0f the strains showing no growth on D-fuconate only one strain (CH-102) was shown to be deficient in the KDF aldolase (specific act- 1X mg protein'l). This strain ivity less than 0.001 nmol product X min' was classified as KDF aldolase-negative. Stains CH-IOS, CH-107, CH-109, and CH-111 were found to possess normal levels of the aldolase despite their negative growth on D-fuconate; these strains must have reverted to being D-fuconate-positive. A summary Of the geneology and pheno- type of these bacterial strains is given (Table 10). 76 m>888mom m>888moa oo.8 c~.c 08.8 o~.8 08.8 No.8 om.H no.9 m881:u z~888mcm m>~h~h~h<¢mz «c.c mc.c mm.c mm.o m°.~ mm._ cm.~ cc.“ moaizu z__h_mC¢ u>~h~mcm cc.— em.c om.o mm.c cm.o no.6 om._ mm.o “omizu z~<¢hm ¢zom¢8uuoa8oc .8888835 88888858 283888 ouccooaeiaxouccouuepcmio 828 88888888-: 888 mcmizu can Ncuizu c—asum 5: sea 88 882858888883 888888888 .88: m8 888 882828 8888888088 8 :8 coon 88 28888 538883188888»: 188888 8>\:V nm.c :8 czocm ago: macaw—nu .mouocuxzoaccu 888888> :8 888288888 8:882: .8888888828 8.38 no 828 8:8 88 8883 one: Acouoeouosaocuoonm .88 cesspouv .m 0—888 77 Table 9. Enzyme activities in mutant and revertant strains. Crude extracts were prepared from Strains grown overnight in broth when determining D-fuconate dehydratase, D-galactonate dehydratase, KDGal kinase, and KDPGal aldolase activity deficiencies. Crude extracts were prepared from cultures grown to half-maximal absorbances (0.3 0.0.,600nm) on nutrient broth and then made 0.5% in D-fuconate, allowing 3.0 hr to induce, when determining KDF aldolase activity deficiencies. Strains CH-101, CH-IOZ, CH-103, and CH-113 are defined in Table 10. 78 888mm“ 88.882888”. “mum. 888$“ 88%.. mm<484< mhgouzuia 888.8 888.8 888.8 888.8 882282 88882 8 28828 82882822 888.8 888.8 888.80 888.8 88228828-8 888.8 888.8 888.8v 888.8 88228882828-8 88282288288 888.8v 888.8v 888.8v 888.8v 882 888.8 888.8 888.8 888.8 .888282 88288882 888.8 888.8 888.8 888.8 882282 88882 28828 82882822 888.8 888.8 888.8v 888.8 88228828-8 888.8 888.8 888.80 888.8 88228882828-8 88282288288 mdgizu z_<¢hm Nemizu z~<¢hm mcuizu z~<¢hm "681:9 zu<¢hm 8882888228 888288882 88888828 mz>sz >88 8888 +888 .+88¢ -mgvzgmc 8888:8888-av maacouuapumuo mc8-=u .828885 «>8uommcuwmuu -888 .uuaa unguxgmc .88888u88-ov 88828888888-o 888-:u 888 .+88@ .888825 «>888mmcummupcv88 max 888-29 .mmsz~=m 8832888 «88:88 .uupum-c 88:88 888 omaangu>zmu 888828 888 .+88@ -028-av massages—88-8 888 8088828088: -=.m2-822 .288888 -888 .+888 8228-888: 828 88 228288288 888828 82-822 + m¢>hozmzm um>hOZmu hzu¢m¢ mmzuav “2282 $58.. “3:8“. “3:8; “:28: 275 58.58% .m Az_<¢pm hz_p_p_h_moa m>_p_moa m>_p_mca u>_h_»=uov u>_p_h_moa u>_p_mca u>_h~moa u>_h_h_p_moa m>_p_moa u>_p_moa u>_p_p<¢uz m>_p_moa u>_p_moa m>_»_mca u>_h_h_h_h_mca m>_p_h_moa N-~E~ E\m __ou.4m - u_8<¢= + 4.8<¢= zo_h<~_u_p= mp<¢h_u mucoz_ ¢u= owe 4>=puz z_<¢pm 4<_¢mpuwgmc ecu =.aeum -=.m¢-4¢a ou.=os=o=m a__m.mga_x as“ to» momma u.>:_ usaueaum .~_ «peep 85 strains only when the medium was supplemented with 0.005% uracil; §;_ggli did not use citrate in the presence or absence of uracil. Kauffman (99) reports Klebsiella species as not forming indole, give positive Voges-Proskauer tests, give a negative methyl red test, and usually grow on ammonium citrate; Escherichia species usually form indole, give a negative Voges-Proskauer test, give a positive methyl red test, and do not use ammonium citrate. Variations from the class norm, in the IMViC tests for the Klebsiella strains derived from 5; pneumoniae PRL-R3,U’, must be indigenous to the strain. Thus, the only significant differences in the strains was in their ability to grow on D-fuconate or D-galactonate. DISCUSSION The results of this section showed the presence of a pathway involving the dehydration of D-fuconate to 2-keto-3-deoxy-D-fuconate, which is then cleaved to pyruvate and D-lactaldehyde (Fig. 12) in the D-fuconate-positive mutant strain (CH-101). (The fate of D-lactaldehyde was not pursued, but is presumed to be oxidized thru lactate to pyru- vate.) D-Fuconate and D-galactonate dehydratase activities as well as a 2-keto-3-deoxy-D-fuconate aldolase activity were detectable in crude extracts prepared from this strain grown on D-fuconate. The absence of aldose dehydrogenase activities leading to D-fuconate or 0-galactonate suggested that these aldonic acids were only used direct- ly as carbon sources rather than as intermediates of aldose pathways. In addition, the gain mutation which facilitates growth of 5; pneumoniae on D-fuconate has been determined, by enzyme induction and genetic studies, to be the constitutive production of a dehydratase that acts on both D-fuconate and D-galactonate. Comparision of the dehydratase levels in the parental strain and the D-fuconate-positive strain grown on D-galactonate, the inducer of the D-galactonate pathway (found to be present in both strains), revealed a D-galactonate to D-fuconate activity ratio of 3:1. Other enzymes of the D-galactonate pathway were found in these strains at about the same level as the dehydratase except for the KDF aldolase, which is lacking in both strains grown on this substrate. Growth of these strains on non-inducers of the D-galactonate pathway (i.e., nutrient broth, 86 87 .zusum m_;u :_ umumcpuapo mm znguma muucousmio on» .- wg:m_u mo>zmo4xoma-m-omv_-~ 52 8282-: 0:0 a_u< u_>=a>a 88 D-glucose, D-gluconate, etc.) did not result in the production of the D-galactonate pathway enzymes in the parental strain; however, such activities were found in the D-fuconate-positive mutant, when grown on these same carbon sources, in approximately the same level as seen for the parental strain grown on D-galactonate. As shown by comparative growth studies, the parental strain would not use D-fuconate as a growth substrate immediately, whereas the D-fuconate- positive strain readily uses D-fuconate for growth. Levels of the D-galactonate pathway enzymes, the D-fuconate dehydratase, and KDF aldolase in the D-fuconate-positive strain grown on D-fuconate were found to be comparable to those found in cells grown on the various non-inducers of the D-galactonate pathway. The parental strain, however, exhibited the KDF aldolase activity only when grown on a mixture of D-galactonate and D-fuconate. Analyses of mutants negative for the D-fuconate pathway, i.e., D-fuconate dehydratase-negative and KDF aldolase-negative mutants, revealed that both lost the ability to grow on D-fuconate with an accompanying loss of the respective enzymatic activities. KDF aldolase- negative mutants retained the ability to use D-galactonate whereas the D-fuconate dehydratase-negative mutants did not. In addition, normaT levels (strain CH-101) of both the D-galactonate and D-fuconate dehydratase activities were found in the KDF aldolase-negative mutants whereas both activities were lost in the dehydratase-negative mutants. Examination of a revertant of the dehydratase-negative mutant on D-galactonate revealed that both the D-fuconate and D-galactonate dehydratase activities were restored. 89 These results are consistent with the following conclusion: D-fuconate serves as a fortuitous substrate for the dehydratase of the D-galactonate pathway which is constitutively produced in the D-fuconate- positive strain (constitutivity results in the production of D-galacton- ate dehydratase. D-fuconate dehydratase, KDGal kinase, and KDPGal aldolase activities in the strain CH-101 without the aid of D-galacton- ate as inducer; in the parental strain these enzymes are apparently induced coordinately (100) ). This constitutive production of the dehydratase allows D-fuconate to be assimilated for growth, as the aldonic acid does not induce the D-galactonate pathway. The further catabolism of the dehydratase product from D-fuconate employs an aldolytic activity independent of the D-galactonate pathway. The origin of this enzyme is unresolved, but some postulates have been considered, 'the testing of which was outside the scope of this thesis, and may be expounded. Many bacterial aldolases have vary limited substrate specificities (101). The KDF aldolase, however, seems to possess a broad substrate specificity for keto-deoxy sugar acids; of five keto-deoxy sugar acids tested, 2-keto-3-deoxy-D-fuconate, 2-keto-3-deoxy-D-quinovonate, Z-keto- 3-deoxy-0—galactonate, and 2-keto-3-deoxy-D,L-arabonate were found to serve as suitable substrates. Such varied substrate specificities have been shown with aldolases of mgtgfcleavage pathways in catechol-metabo- lizing pseudomonads (102). Pseudomonads are known to possess catabolic pathways for the breakdown of phenol and cresols to hydroxy-phenolic compounds which can then serve as substrates for subsequent reactions of a meta-cleavage pathway. One such compound, para-hydroxy phenol 90 (hydroquinone), if formed, could give rise to the product 2-keto-3- deoxy-D,L-arabonate if acted on by a hydrolyase. As noted, this com- pound seems to serve as a good substrate for the KDF aldolase. It is possible that the aldolytic activity found in 5; pneumoniae on KDF is one in unision with a meta7cleavage pathway for catechol as seen in pseudomonads. Although Klebsiella species are not normally associated with the ability to metabolize catechol, the fact that pseudomonads possess the enzymatic mgtgycleavage pathway enzymes on transmissible plasmids (103 a,b,c) suggests the possibility that by transduction 5; pneumoniae could acquire this trait. Testing for a catechol pathway, using induc- tion of the KDF aldolase as a gauge when growing §;_pneumoniae on sub- stances (phenol, 2.5 mM. and sodium benzoate, 5.0 mM. in 0.5% sodium citrate) known to induce mgtgycleavage pathway enzymes in pseudomonads (102), failed to show.such activity. However, when procedures of plasmid isolation (104) were followed, the presence of high molecular weight DNA in crude extracts of both the parental strain and the D-fuconate-positive strain grown on nutrient broth was observed (pre- _ liminary data not included in this thesis). If not found as artifact- ual, this nucleic acid material may represent a plasmid (105 a,b) pos- sessing the genetic information responsible for the noted KDF aldolase activity. Such a finding would be one of the few known cases where a catabolic pathway for a carbohydrate is located on a plasmid. SECTION II Purification and Characterization of D-Galactonate (D-Fuconate) Dehydratase 91 INTRODUCTION In section I of this thesis, the constitutive production of D-fuconate dehydratase was shown to be the key event in the mutational acquisition of the ability of §;_pneumoniae to use D-fuconate as a carbon source. This enzyme initiates D-fuconate catabolism, and the product of the reaction induces the next enzyme in the pathway, KDF aldolase. Mutant analysis and other studies suggested that the dehydra- tase was also instrumental in initiating the catabolism of D-galactonate. Such a dual specificity for a dehydratase has not been previously demon- strated. Literature surveys of aldonic acid dehydrases as compliled by Dahms (106) and Hood (107), reveal that (i) dehydration results in the formation of an a-keto acid, (ii) the configuration of the aldonic acid substrate at the dehydration site is trans, (iii) a divalent metal ion is commonly required for best activity, (iv) thiols are required if a metal ion is not, (v) the optimal pH range is from 7.0 to 8.0, (vi) Km values range from 0.1 to 8.0 mM. and (vii) the dehydration reaction is essentially irreversible. Further characterization, as far as physical studies are concerned, has been lacking in the literature. Recently, however, a specific D-galactonate dehydratase from a pseudomonad was purified and character- ized more fully (97). Although most dehydratases exhibit mono-substrate specificities, exceptions are the D-fuconate dehydratase in a pseudo- monad which acts both on L-arabonate and D-fuconate (33), and the 92 93 D-glucarate dehydratase of §;_ggli_which acts on both D-glucarate and D-idarate (108); neither of these enzymes have been character- ized by physical means. This section describes the isolation and characterization of the D-galactonate (D-fuconate) dehydratase. MATERIALS AND METHODS All materials and methods not described in this section have been described in Section 1. Bacterial Strain and Culture Conditions Bacterial Strain. A nutrient broth agar slant of Klebsiella pneumoniae PRL-R3,U’, strain CH-101, a mutant which is derepressed for the D-galactonate (D-fuconate) dehydratase (see Section I, Results), was used for the preparation of the dehydratase. Medium, The bacteria were grown in 0.65 z K+ D-galactonate- mineral medium plus 0.005% uracil; this carbon source was found to produce the highest specific activity of the enzyme (Section I, Results). Mineral medium was prepared directly in the fermenter chamber (10.0- liter capacity) and sealed before autoclaving at 20 psi and 121°C for 75 min; measures were taken to prevent back pressure in the chamber due to autoclaving and possible contamination of the fermenter contents. D-Galactonate and uracil were autoclaved separately and added aseptic-- ally to the sterile contents of the fermenter. Starter Culture Preparation. A 1.0-liter exponentially growing culture of the strain CH-101 in D-galactonate-mineral medium was pre- pared for inoculation of the 10-liter culture. To prepare this inoc- ulum, a 7.0-ml culture, grown overnight in nutrient broth, was trans- ferred to a flask containing 250 ml of fresh 0.65% D-galactonate-mineral medium plus uracil. The resultant culture was then incubated on a 94 95 rotary shaker until a turbidity of 0.60 0.0. at 600 nm was reached. Serial transfer of this culture (250 ml to 500 ml, and 500 ml to 1.0 liter) in fresh D-galactonate-mineral medium was performed to pre- pare the final 1.0-liter starter culture. Cell Growth. D-Galactonate-mineral medium in a New Brunswick Scientific Co., Inc., Microferm fermenter was temperature-equilibrated at 30°C. Starter culture was then added, aseptically, with aeration and stirring turned off, to the sterile fermenter contents. Vigor- ous aeration (10.0 lb/inz) and mixing (200 rpm) were resumed to pro- 1 mote optimal growth. Growth was monitored by withdrawing 2.0-ml samples with sterile pasteur pipetes and making absorbance readings on a Gilford, model 2400, spectrophotometer at 600 nm. Samples were diluted to obtain corrected absorbance values and plotted versus the time of growth. Cells were harvested when the corrected absorbance value no longer changed with time; usually, a final absorbance of 2.0 at 600 nm was reached after 5.0 hrs of growth. Harvesting Cell Cultures. Bacterial cultures grown in the micro- fermenter were harvested with the aid of a Sharples air-driven centri- fuge cooled to 4°C with Freon and run at a speed of 40,000 rpm. The cells were weighed in tared plastic beakers and stored at -20°C until needed. Protein Determination. Protein concentration was determined either by the procedure of Lowry et al. (79) using bovine serum albumin as the standard or by the aid of a nomograph using absorbance values of the protein solution at 280 nm and 260 nm above the buffer absorbance. The Lowry chromogen 96 was measured on a Gilford 300 N colorimeter equipped with a red filter at 600 nm. A Gilford spectrophotometer, model 2400, was used to meas- ure in the ultraviolet range. D-Galactonate (D-FuconateliDehydratase Assay_ The assay for the D-galactonate (D-fuconate) dehydratase was per- formed as described in Section I, Materials and Methods, unless other- wise indicated. D-Galactonate was used as the substrate at a concentra- tion of 20.0 mM in the assay when assaying column fractions during a given purification step and when reporting activities of this enzyme under various assay conditions, unless noted differently. Eggyme Purification Procedures DEAE-Cellulose Chromatography. DEAE-cellulose was prepared by washing it sequentially with 1.0 N NaOH. 1.0 N HCL, and 1.0 N NaOH as described by Peterson and Sober (109). washed cellulose ion exchanger was de-fined by decantation. When not in use, the exchanger was stored at 4°C in 0.02: sodiun azide and 1.0 nfl EDTA to prevent bacterial growth. Packed columns were Judged equilibrated when both the pH and conductance of the eluate matched that of the buffer. Protein loading was done by‘ passing the protein sample through the column and washing with the appropriate buffer until the protein absorbance was no longer observed in the eluate. Adsorbed protein was eluted with a linear gradient of KCl. Once the resin was used in an enzyme purification step, the cellu- lose column was washed with 1.0 M KCl to remove all protein and was unpacked to store as above. 97 Sepharose A-SM Chromatography. Commercial Bio-Gel Sepharose A-SM (Bio-Rad Laboratories), which has an exclusion limit of 5.0-million- molecular weight, was swollen in 0.05 M potassium phosphate buffer (pH 7.0) at 4°C for 24 hours. Columns were packed under a 16-cm pres- sure head to establish linear flow rates of 5.0 ml per hr with the aid of a Gilson Multiplus-Z peristatic pump. Packed columns were equilibrated by washing with at least four bed-volumes of the same buffer. After use, the columns were washed with 0.02% sodium azide . to prevent bacterial growth. The volume of the protein sample applied to the columns were kept to 2.0% of the column bed-volume. Calibra- tion of the column for void volume (V0) and inclusion volume (Vt) was done using Blue Dextran (Sigma Chem. Co.) and K3Fe(CN)6, respectively (110). Hydroxyapatite Chromatography. Bio-Gel HT hydroxyapatite (Bio- Rad Laboratories), pre-swollen in 1.0 mM sodium phosphate buffer, pH 6.8, was used to prepare chromatographic columns. Pre-swollen hydroxyapatite was specified to contain 0.175 mg hydroxyapatite mater- ial per ml of well mixed slurry. To determine the capacity of the material for the dehydratase, at given purification steps, several mini-columns were used. Mini-columns of hydroxyapatite were prepared using standard 17.8 cm pasteur pipetes. They were packed with 2.01nn of glass wool, upon which a 2.0-mm bed of celite and various amounts of the hydroxyapatite material were layered. Columns ranging from 44 to 175 mg of hydroxyapatite were packed under gravity and equili- brated with 1.0 mM potassium phosphate buffer (pH 6.8). Protein was applied to each column, washed with the equilibration buffer, and eluate fractions were assayed for dehydratase activity. Adsorbed 98 dehydratase was eluted with 0.4 M potassium phosphate buffer (pH 6.8). The amount of dehydratase bound to a given amount of hydroxyapatite was determined by plotting the percent of bound dehydratase versus mg protein per mg hydroxyapatite. This graph (not shown) was used to calculate the optimal amount of protein to add to the column. (Similar preliminary studies were conducted in determining the capac- ity of the DEAE-cellulose ion exchanger resin noted previously.) The hydroxyapatite slurry was first diluted five-fold with 1.0 mM potassium phosphate buffer (pH 6.8) and loaded onto a 0.5-cm bed of celite by gravity flow. Before application of the sample, protein was diluted to give a potassium phosphate concentration approx- imating 1.0 mM and a pH of 6.8; these conditions were found to be opti- mal for the binding of the dehydratase protein. The column was washed with the equilibration buffer until no protein could be detected in eluate fractions. The column was then eluted with a linear gradient of potassium phosphate buffer (pH 6.8). Sequential washes of the column with 1.0 M KCl, 1.0 M potassium phosphate, and 0.02% sodium azide were done to condition the column for storage. Sephadex G-200 and Bio-Gel P-300 Chromatography. Materials obtained in the dry state were swollen in the appropriate buffer for 2.0 days at 4°C. Conditions of packing, equilibration, sample loading, and calibration were followed as described for the Sepharose A-SM gel fil- tration material. Pressure Dialysis Concentration of Protein Pressure dialysis was performed using an Amicon Diaflo apparatus, 50-ml capacity, equipped with a 1-liter pressure bell jar and a PM-30 99 membrane filter, having a 30,000-molecular-weight range cut-off. Con- centration of the protein was performed under nitrogen pressure (50 psi) as to establish flow rates no less than 20.0 ml per hour. When not in use, the filter was stored at 4°C in 95% ethyl alcohol after first washing it with distilled water. Conductivity Measurements Column fractions from purification steps were assayed for salt concentration using a standard conductivity meter with variable conduct- ance control. Samples of 0.05-ml volume were diluted 100-fold with de-ionized water and read against a buffer blank. The instrument was calibrated using the highest salt concentration of the gradient in the appropriate buffer. Polyacrylamide Gel Electrophoresis General Procedures. Slab gels were prepared using a Bio-Rad vertical slab gel apparatus, model 220, equipped with rectangular glass plates and appropriate spacers and combs. All slab gels prepared had dimensions of 100 X 140 mm; thickness, as determined by the spacer used, was either 1.5 or 3.0 mm. Plates were prepared by cleaning with chromic acid solution, neutralizing with sodium thiosulfate, and then rinsing with distilled water before air-drying before use. Components of the gel mixture were degassed over ice prior to mixing. Plates were sealed at the sides using appropriate spacers or with 2.0% Bacto- agar. Electrophoresis was performed using an ISCO, model 492, Electro- phoresis Power Supply (150 mamp, 1000 volt range) at constant current. After electrophoresis, slab gels were removed by first removing the spacers, followed by lubricating the gel-glass space with a stream of 100 water, and then by prying the plates apart with the aid of a spatula. All slab gels prepared consisted of a 9.0-cm high running gel and a 2.0-cm high stacking gel. The stacking gel was used to make 1.0-cm deep wells for convenient application of the protein sample. Preparation of Native Gels. Slab gels of 10.0 cm X 15.0 cm X 3.0 mm were poured. Running gels were filled to a height of 9.0 cm with a solution containing 5.0% acrylamide, 0.13% bisacrylamide (N,N'- methylenebisacrylamide), 0.025% TEMED (N,N,N',N'-tetramethylethylene- diamine), 0.045% ammonium persulfate (prepared fresh each time and added just prior to pouring the gel), and 0.375 M Tris-HCl buffer (pH 8.8). Once hardened, a stacking gel containing 5.0% acrylamide. 0.13% bisacrylamide, 0.025% TEMED, 0.045% ammonium persulfate, and 0.063 M Tris-HCl buffer at a pH of 6.8 was poured over the running gel, with a three-channel comb in place. The poured stacking gel was overlaid with a thin layer of 1-butanol to generate a level surface, and allowed to harden over a period of 20 min (as was done with the running gel). The final slab gel was attached to the electrophoresis apparatus by the use of clamps and was then pre-electrophoresed at a constant current of 20 milliamps at 4°C for 8.0 hrs to remove unreac- - ted amoniun persulfate and to condition the gel for use at this temper- ature. The electrophoresis buffer contained 0.2 M glycine, 0.05 M Tris- (hydroxymethyl)aminomethane (Tris), 0.06% thioglycolic acid with a final pH of 8.8. Prior to the application of the protein sample, 0.2 ml of 0.05% bromophenol blue dye in 10% glycerol was applied to each channel and run into the stacking gel at 5.0 milliamps current. The current 101 was stopped before application of the protein samples at 0.2-ml volume to each channel. Once protein samples were applied, the current was restored to 5.0 mamps until the dye marker entered the running gel, then increased to 10.0 mamps for an interval of 20ininutes. At the end of this time, the current was increased to 20.0 mamps, and allowed to run at this current for a total of 4-6 hrs at 4°C; the dye marker at this time was within 0.5 cm of the bottom of the slab gel. Gels were stained for protein by placing into a solution of 0.1% Coomassie Brilliant Blue R, 10% trichloroacetic acid (TCA), and 27% (v/v) isopropanol and allowed to stain overnight at room temperature. Dye bands were marked by the insertion of a small piece of wire before destaining, as the latter process removes all traces of the dye band. Destaining was achieved by inmersion in 10% acetic acid with three changes of destaining solution or until a faint blue background was obtained. Slab gels, once destained, could be dried for photographic purposes by transferring the gel to a sheet of filter paper (treated initially by soaking in 1% glycerol and 10% acetic acid for 30-45 min) and drying with the aid of a Model 224, Gel Slab Drier (Bio-Rad Labora- tories) under 733 m Hg vaccum for 45 minutes. The dried gel shows no - shrinkage once treated as described and is easily preserved in a note- book. Protein profiles of the stained channels were made by scanning the gel with a Gilford Gel Scanner at 600 nm and plotted on graph paper with a Sargent recorder. Slab gels were laid length-wise on their edges in quartz boat cuvettes such to allow measurements to be made at the center of the gel; distilled water was used to cover the gels while 102 scanning as to prevent dehydration. Traces prepared in this manner could be calibrated in terms of true gel length by taking into con- sideration the speed of the gel scanner and the recorder. Preparation of SDS Gels. Slab gels of 10.0 cm X 15.0 cm X 1.5 mm were prepared as described above. Running gels of 9.0 cm heigth were comprised of 10.0% acrylamide, 0.26% bisacrylamide, 0.025% TEMED, 0.045% amonium persulfate, 0.1% SDS, and 0.375 M Tris-HCl buffer at a pH of 8.8. The stacking gel consisted of 5.0% acrylamide, 0.13% bisacrylamide, 0.025% TEMED, 0.045% ammonium persulfate, 0.1% SDS, and 0.0625 M Tris-HCl buffer (pH 6.8). Both gels were allowed 20 min for polymerization to occur at room temperature. Appropriate combs were used to give the desired number of channels in the stacking gel. A layer of 1% SDS was used to generate a flat surface for both the running gel and the stacking gel. The electrophoresis buffer was com- prised of 0.2 M glycine, 0.05 M Tris(hydroxymethyl)aminomethane (Tris), and 0.1% SDS at a final pH of 8.8. Pre-electrophoresis at 5.0 mamps preceded sample application at room temperature to rid the gels of excess ammonium persulfate. Samples were prepared by diluting 0.05 mg of standard protein or - dehydratase preparation in 0.2 ml of 1.2% 505, 0.29 M Tris-HCl buffer (pH 6.9), and 10% glycerol. Immediately prior to electrophoresis, the sample solutions were made 2.0% in 2-mercaptoethanol and then heated at 100°C for 5.0 minutes. After heating, the samples were made 0.05% in bromophenol blue dye, and 0.04 ml sample was added to each channel. Slab gels were subjected to electrophoresis at 5.0 mamps through the stacking gel and 10 to 20 mamps through the running gel. Electrophoresis 103 under these conditions required about 4.0 hrs at room temperature. Gels were stained and destained, scanned for protein stained areas, and dried as described for native gels. Isoelectric Focusing Isoelectric focusing of the D-galactonate (D-fuconate) dehydra- tase was performed in a 110-ml capacity LKB-Produckter (Stockholm, Sweden) vertical column equipped with a Lauda-water bath cooled to 4°C with a 10% ethylene glycol solution. The procedure of Vesterberg (111) was used as rationale for the preparation of the electrophoretic medium. Solutions were loaded and eluted by the aid of a Gilson Multiplus-Z peristaltic pump and gradients prepared using a 110-ml- capacity conical gradient maker equipped with a tapered rod stirrer for proper mixing. The electrophoresis medium, comprise mainly of sucrose (high concentrations at the anode end, bottom; and low concen- trations at the cathode end, top), Pharmolyte ampholytes (Pharmacia, pH 4.0 to 6.5), and protein sample was generated by the fbllowing solutions, loaded in this order: (i) dense electrolyte solution, 23.54 ml, comprised of 58.94% sucrose and 0.043 M sulfuric acid; (ii) dense electrolyte junction buffer, 5.0 ml, comprised of 55.24% sucrose and 1.07% Pharmolyte ampholytes (pH 4.0 to 6.5); (iii) an 80.0-ml linear gradient of 51.53 to 4.99% sucrdse (negative gradient) in approximately 1.0% Pharmolyte ampholytes (pH 4.0 to 6.5), plus about 2.0 mg of protein sample; (iv) a light electrolyte junction buffer, 5.0 ml, of 2.5% sucrose and 1.0% Pharmolyte ampholytes (pH 4.0 to 6.5); and (v) a light electrolyte solution of 0.18 M NaOH. 104 Protein was subjected to isoelectric focusing at 4°C for a period of several days at constant voltage using an ISCD Electrophoretic Power Supply, model 492, which has an upper limit of 1000 volts capac- ity. Eluate fractions were collected from the base of the column and assayed for both enzyme activity by the standard dehydratase assay (above) and pH with a Beckman pH meter equipped with microelectrodes. Enzymatic Assays for Molecular Weight Standards Calibration Standards. Ferritin, blue dextran, and K3Fe(CN)6, used to calibrate gel filtration columns, were determined by their characteristic absorbances at 230 or 280 nm, 630 nm, and 420 nm, res- pectively. Lactate Dehydrogenase. Beef heart lactate dehydrogenase was assayed using the coupled assay procedure described in Section I fol- lowing the oxidation of NADH with time in the presence of sodium pyru- vate at 340 nm. Pyruvate Kinase. Rabbit muscle pyruvate kinase was assayed for the production of pyruvate from phosphoenol pyruvate (PEP). The reac- tion mixture (0.2 ml) contained 7.5 mM MgCl2, 0.3 mM ADP, 0.25 mM PEP, _ 50.0 mM potassium phosphate buffer (pH 7.5) and non-rate-limiting amounts of lactate dehydrogenase; the temperature was 30°C and the rate of aborb- ance change was measured at 340 nm. Alkaline Phosphatase. §;_ggli_alkaline phdsphatase was assayed in accordance with standard procedures following the production of pynitrophenol from the substrate prnitrophenyl phosphate (112). Ali- quots of the enzyme sample (0.4 ml) were inoculated in a 1.0-ml solu- tion consisting of 50.0 mM glycine buffer (pH 10.5) and 5.5 mM 105 penitrophenyl phosphate, for 30 min at 37°C. At the end of this time, 10.0 ml of 0.02 N NaOH was added to the samples, the solution was mixed, and absorbance readings were made at 405 nm against a reagent blank. Catalase. Bovine liver catalase was assayed using a 1.5-ml solu- tion of freshly prepared 0.5% hydrogen peroxide in 0.05 M Tris-HCl buffer (pH 7.5). Initial velocities were determined after blanking the instrument against the buffer alone at 240 nm. Rates were measured as the absorbance drop from 0.45 to 0.40 0.0. at this wavelength (113). Quantitation of the amount of hydrogen peroxide lost was made using the absorbance values reflected by various stock solutions of the substrate (114). Fumarase. Fumarase was assayed according to the procedure of Racker (115) following the conversion of L-malate to fumarate at 240 nm and 25°C. A reaction mixture of 1.0 ml was used with various aliquots of the enzyme sample. Methods for the Determination of Molecular Weight Sucrose Density Gradient Centrifugation. Sucrose density gradient centrifugation was run in 1.3 X 5.0 cm cellulose nitrate centrifuge tubes containing 4.6-ml volume of 5.0 to 20.0% sucrose in 0.05 M Tris- HCl buffer (pH 7.5). Gradients were poured at room temperature and cooled to 4°C over a period of several hours before use. Centrifugation was performed in a model L-2 preparative Beckman centrifuge with an sw-39 rotor for 15.0 hrs at 35,000 rpm at 4°C. Following the centri- fugation, tubes were punctured and fractions were collected from the bottom. 106 Sedimentation Velocity and Sedimentation Equlibrium Analysis. Sedimentation velocity and equilibrium studies were performed using a Beckman Model E Ultracentrifuge equipped with ultraviolet absorption optics as described by Lamers et al. (116). Protein concentration was observed as a moving boundary of absorbance above the buffer and was quantitated by Beers' law, as this absorbance is linear within the confines of the experiment. (i) Sedimentation Velocity. For sedimentation velocity analysis, centrifugation was performed in an aluminum An-G six-cell- capacity rotor at 20°C and at 32,000 rpm. Sedimentation of each cell was determined by making scans of the moving boundary at regular time intervals, determining the half height of the boundary (therefore, the distance from the center of rotation), and using these values in appro-. priate equations (see Results, this section) to calculate the sedimen- tation constant. (ii) Sedimentation Equilibrium. Sedimentation equilibrium was performed using the meniscus depletion method of thantis (117) using a four-sector AnF titanium rotor with standard double-sector cells. Protein sample (0.12 ml) was layered onto 0.01 ml of FC-43 ° silicon oil to prevent convection disturbances; the total sample volume was restricted to one-fourth the cell capacity to facilitate ease of equilibrium measurements. The centrifugation was done at 20°C and 11,000 rpm allowing 24 hrs for equilibration between trials. Measure- ments of absorbance and migration distance at the given absorbance, above background, were made at equilibrium for various distances from the center of rotation to determine the concentration/radius factor 107 needed for the molecular weight calculation at equilibrium (118) (see Results, this section, for calculations). Amino Acid Composition Determination All protein samples prepared as described in the Results, this section, were analyzed by Doris Bauer on an automated Beckman, model 121-8, Amino Acid Analyzer. Known standard amino acids were used to calibrate the column systems used to make the analyses. Quantitation of the amino acids was done by determination of the peak area as plot- ted on a multipoint recorder and by dividing by a predetermined color constant for that amino acid. All amino acids were converted automatic- ally to their ninhydrin derivatives in the course of the analysis. Norleucine was used as the internal standard for each amino acid deter- mination, unless noted otherwise. Sources of Materials D-Galactonate was prepared from D-galactono-y-lactone (Sigma Chemical Company) by neutralization with stock 1.0 N NaOH. Membrane filters (PM-10 and PM-30) used in pressure dialysis concentration of protein samples, were obtained from the Amicon Corporation. Phenol reagent (Folin and Ciocalteu) used in Lowry protein determinations was obtained from the Harleco Corporation. DEAE-cellulose as Cellex-D (anion exchange cellulose) in the hydroxide form, and Bio-Gel P-300 (100-200 mesh) were obtained from Bio-Rad Laboratories. Sephadex G-200 (median grade) and Blue Dextran were obtained from Pharmacia Fine Chem- icals“ Dowex-1-X8 (200-400 mesh) was obtained from the Sigma Chemical Company. "Enzyme-grade" ammonium sulfate was obtained from the Mallinc- krodt Company. Acrylamide was used without recrystallization from the 108 Ames Company, bisacrylamide from Miles Laboratories, and sodium dodecyl sulfate (SDS) from the Pierce Chemical Company. Norleucine and other amino acid standards were obtained from Calbiochem and the Pierce Chem- ical Companies. Specially prepared hydrolysis tubes were obtained from the MSU Glass-blowing Laboratory for acid hydrolysis treatment of protein samples for amino acid composition determinations. Cellulose nitrate centrifuge tubes for sucrose density gradient sedimentation studies were obtained from the Beckman Company. All chemicals and enzymes not mentioned were obtained from the Sigma Chemical Company or of better comercial grade. RESULTS Purification of D-Galactonate (D-Fuconate) Dehydratase During the course of the purification of the D-galactonate (D-fuconate) dehydratase, some techniques or conditions were found to be ineffective or counterproductive. Substantial losses of enzy- matic activity resulted from dialysis against 0.05 M potassium phos- phate buffer (pH 7.0), despite the fact that the enzyme was stable in this buffer upon storage at 4°C or by freezing. Losses of the enzyme activity occurred at pH ranges below 5.0 and above 9.0; therefore, pH precipitation was eliminated as a viable method. Bentonite treat- ment at 60 mg dry material per ml crude extract (15.0 mg/ml protein concentration), as used in other enzyme Purification procedures, did not adsorb the dehydratase. Carboxymethyl-cellulose did not bind the enzyme at pH ranges required for enzyme stability (pH 5.0 to 9.0). The following scheme was reproducible and yielded an electrophoretic- ally homogeneous enzyme preparation. Cell Extract. Frozen packed cell, 111.5 g wet weight, from two consecutive 10.0-liter D-galactonate-mineral~medium cultures of the strain CH-101 were thawed in cold 0.05 M potassium phosphate buffer (pH 7.0) to give a final cell slurry of 400 ml. The cell extract was prepared by sonic treatment in the presence of glass beads (125 to 88 microns). All purification steps were carried out at 00 to 4°C. A summary of the purification is given (Table 13). 109 110 uoom an :_E eon uoauoca cu ouncouoe—oaun mo pee: o.“ muco>cou «egg oexuco we «enase one m_o:ao u.:: 956 AN we.“ Ame em mu co~-o xoeegeom mm no.5 Nae mm as __ omo_=p_ou-m¢m>oum¢ >p_>_pu< u_e_uuom >p_>_pu< S zc_~u<¢e .omouaevagou Amueccoawrav mueeouuepamia ea :opuau.m_g=¢ .m~ open» 111 Protamine Sulfate Fractionation. The crude extract was diluted to 780 ml (to 10.0 mg/ml protein concentration) with the sonication buffer, to which was added 20.3 g of well ground ammonium sulfate to make a 2.6% solution. To the solution was added 156 ml of 2.0% prota- mine sulfate (pH 5.0) slowly while stirring. After 30 min stirring, the solution was centrifuged at 12,000 X g_to sediment the precipitate. The supernatant was collected by decantation. Ammonium Sulfate Precipitation. To 908 ml of supernatant from the protamine sulfate step was added slowly 182.7 g of ammonium sul- fate to bring the concentration to 40% of saturation. The solution was stirred for 30 min, and then centrifuged at 12,000 X g_for 10 min- utes. The 955 ml of supernatant was decanted, and 180.1 9 more ammonium sulfate was added to bring the concentration to 70% of sat- uration. After 30 min, the solution was centrifuged once again. The supernatant was discarded, and the pelleted material was suspended in 200 ml of 0.05 M potassium phosphate buffer (pH 7.0) for further purification. DEAE-Cellulose Chromatography I. The pooled 40 to 70% ammonium sulfate fraction was diluted to 800 ml with 0.05 M potassium phosphate- buffer (pH 7.0) and loaded onto a 4.0 X 18.5 cm DEAE-cellulose column, pre-equilibrated with the same buffer at pH 7.0. A 3.6-liter linear gradient of 0.0 to 0.35 M KCl in the same buffer was used to elute the protein from the column (Fig. 13). Fractions 74 to 115, containing- the peak of activity, were pooled for further purification. Pooled fractions showed no activity (less than 0.001 unit/mg) for the KDGal kinase or the KDPGal aldolase. 112 Figure 13. DEAE-Cellulose chromatography 1. Fractions of 20.0-ml volune were collected at a flow rate of 2.0 ml per minute. All other details are given in the text. Symbols: 0 D-galactonate (D-fuconate) dehydratase, Aprotein absorbance (280 nm), +salt gradient concentra- tion. 113 Axe. aux cow mNN cow mN— an. m- cop mm an .ee ocNV uuzeaeomae z_upc¢e =._ o.o e.o ~.c e.c m.c ..c n.c ~.c P p b P 80 95 110 125 140 155 FRACTION NUMBER 65 SO q G GS cm oe om c~ zc__8<¢a\m__z= 35 114 Sepharose A-SM Chromatography. The pooled fractions from the DEAE-cellulose column were concentrated to 34.0 ml by pressure dialy- sis as described in Materials and Methods, and loaded onto a 2.6 X 80.0 cm column of Sepharose A-5M equilibrated with 0.05 M potassium phosphate buffer (pH 7.0); the sample was divided into two portions, 17.0 ml each, and loaded sequentially with buffer washes in between runs. Protein was eluted from the column using the same buffer. Elu- tion profiles of each run, as exemplified by one such run (Fig. 14), showed peak dehydratase activity in fractions 52 to 76; these fract- ions were pooled from each run and combined for further purification. Hydroxyapatite Chromatography. The combined sample from the Sepharose A-SM chromatography step, 224 ml, was diluted to 0.5 unit per ml with 1.0 mM potassium phosphate buffer (pH 6.8) and loaded onto a 7.9 X 2.0 cm hydroxyapatite column equilibrated with the same buffer. (Capacity studies with the enzyme at this stage of purifica- tion gave a loading capacity ratio of 2.4 mg protein per ml packed hydroxyapatite, resulting in better than 95% of the dehydratase bound.) Dehydratase activity was eluted with a 2.4-liter linear gradient of 4.0 to 400 mM potassium phosphate buffer (pH 6.8). Fractions 21 to 42, containing the peak activity, were pooled for further purification (Fig. 15). DEAE-Cellulose Chromatography II. The pooled hydroxyapatite frac- tions were loaded directly onto a 1.5 X 15.5 cm DEAE-cellulose column equilibrated with 0.05 M potassium phosphate buffer (pH 6.0). Elution of the enzyme was done employing an 400-ml linear gradient of 0.0 to 0.35 M KCl in the above buffer. Fractions 52 to 72, containing the 115 Figure 14. Sepharose A-5M chromatography of the pooled DEAE-cell- ulose I fractions. Fractions of 4.0-ml volume were collected. All other details are given in the text. Symbols: .0-galactonate (D-fuconate) de- hydratase, Aprotein absorbance (280 nm). ‘ no al.. 1.». nee cam. uozemeomae z_u_o¢a em o.~ .3 o; are c... b 1+ 1 cc Om ON @— z¢_~u<¢u\mh.za 9... Si 35 45 55 75 85 95 105 FRACTION NUMBER 25 117 Figure 15. Hydroxyapatite chromatography of the combined Sephar- ose A-SM pooled fractions. Fractions of 22.0-ml volume were collected at a flow rate of 200 ml per hour. All other details are given in the text. Symbols: 0 D-galactonate (D-fuconate) dehydratase, A protein ab- sorbance, + salt gradient concentration. 118 :3 «as. 8N SN 8. 8. 8. 8. 8. 8 8 8 8 a - pi (- p (p b [- p . L i o I h L . c2 83 3.2282 238: as 3. .3 e... m... N... I. .5 — n n L p - p q 1 fi 1 1 7 1 q q i 8. 8 8 2 8 S 3 S 8 2 zc_hu<¢u\mh_z: SO 20 3O 35 40 45 FRACTION NUMBER 15 10 119 peak activity, were pooled for further purification (Fig. 16). Sephadex G-200 Chromatography. Fractions pooled from the second DEAE-cellulose column were concentrated to 26.0 ml by pressure dialysis and loaded onto a 4.0 X 95.0 cm column of Sephadex G-200 equilibrated with 0.05 M potassium phosphate buffer (pH 7.0). Protein was eluted from the column using the same buffer. Fractions 52 to 70 (Fig. 17), containing the peak activity, were pooled and concentrated by pressure dialysis to a final volume of 23.0 ml to constitute the purified dehydra- tase protein used for all kinetic and physical studies in this section. Determination of Purity of the Dehydratase Preparation As criteria for homogeneity both native and denaturing polyacryl- amide gels were used to judge the purity of the final purification step used in the dehydratase preparation. SDS Polyacrylamide Geliglectrophoresis. Slab gels comprised of 10% acrylamide were prepared as described in Materials and Methods. Two such gels were run. 0ne gel shows the dehydratase electrophoresed in the presence of known molecular weight standards (Fig. 18). From the molecular weights of catalase (60,000 MN), malate dehydrogenase (35,000 MN), tryptophanase (55,000 MN), aldolase (40,000 MW), and ovalbumin (45,000 MN), a subunit molecular weight for the dehydratase of approximately 45,000 was determined. The second gel (Fig. 19) shows that only one protein band can be observed visually or with the aid of a gel scanner. Hhen crude extracts of this strain were run on the same SDS gels one observed a protein band corresponding to the band observed in the purified dehydratase preparation; this band of protein was observed throughout the purification. 120 Figure 16. DEAE-Cellulose II chromatography of the hydroxyapatite pooled fractions. Fractions of 4.0-ml volume were collected at a flow rate of 90.0 ml per hour. All other details are given in the text. Symbols: 0 D-galactonate (D-fuconate) dehydratase,Aprotein absorbance (280 nm), +salt gradient concentration. 121 new mNP om— mmp :0. mm on mm o P D n E p A25 .9. . n E n If Ase cm~v uuzh_>_ho< OJ 1 1. 136 Figure 23. Lineweaver -Burk plot using D-fuconate as sub- strate in the dehydratase reaction mixture. Purified dehydratase (Sepha- dex G-200) was assayed in 0.3-ml volume reaction mixture comprised of 10.0 mM MgCl2, 1.0 mM EDTA (pH 7.0), 50.0 mM Pipes buffer (pH 7.0), and varying amounts of K D-fuconate. Standard procedures for the formation of semicarbazone derivatives of the dehydratase product was used in mea- suring enzymatic activity. Initial velocity (V) is in micromole product formed per minute, and substrate concentration (S) is in millimolar D-fuconate concentration in the reaction mixture. cm. mm.— co.— P m~.c ’ cm.c b m\— m~.o ’ cc.c m~.c- P om.c- P m~.c- ac..- AU A/L 138 Figure 24. Lineweaver-Burk plot using D-galactonate as substrate in the dehydratase reaction mixture. Purified dehydratase (Sephadex G-200) was assayed in 0.3-ml volume reaction mixture comprised of 10.0 mM MgCl2, 1.0 mM+EDTA (pH 7.0), 50.0 mM Pipes buffer (pH 7.0), and vary- ing amounts of K D-galactonate. Standard procedures for the formation of semicarbazone derivatives of the dehydratase product were used in measuring enzymatic activity. Initial velocity (V) is in units of micro- mole product formed per minute, and substrate concentration (S) is in millimolar D-galactonate concentration in the reaction mixture. 139 cm. ma.— om.c mm.c . P on.o r m2 mo.o c~.c- L m¢.c- o~.c- ma.c- D D b cm..- AU :9... 140 respectively. The relative Vmax value for D-galactonate was 3.33-fold higher than for D-fuconate. Divalent Cation Activation in the Presence of EDTA. Preliminary studies of the effect of divalent metals on the dehydratase-catalyzed reaction showed that a rate existed in the absence of metal added to a reaction mixture comprised of 20.0 mM D-galactonate and 170 mM Tris- HCl buffer (pH 8.0). This rate increased with time and was completely inhibited by 1.0 mM EDTA (Fig. 25). Addition of MgCl2 (10.0 mM) to the EDTA-treated mixture not only restored activity but resulted in a rate that was constant with time (data not shown) and proportional to enzyme concentration (Fig. 26). These data suggested that the presence of EDTA in the assay function to remove deleterious metal(s) that inhibit the enzyme at lower protein concentrations. Examination of the trials without EDTA (Fig. 26) show that proportionality of activity with pro- tein concentration is found only at the higher protein concentrations, suggesting that the unknown metal(s) are depleted from the reaction mixture if enough protein is added. Support for the presence of such deleterious metal(s) in the enzyme preparation is found when the dehydra- tase was dialyzed against 1.0 mM EDTA (Table 14); although considerable enzyme activity was irreversibly lost by this dialysis, complete removal of the activity that can be at least partially restored by MgCl2 was the result of this treatment. To test the effectiveness of various divalent metals to reverse the effect of EDTA, the dehydratase was pre-incubated in a reaction mixture comprised of D-galactonate, EDTA, and Tris-HCl buffer. Metals were then added to the reaction mixture and the course of the reaction was 141 Figure 25. Effect of EDTA 0n the dehydratase in the absence of metal in the enzymatic reaction mixture. The enzymatic assay was scaled up to 1.0-ml volume consisting of 20.0 mM D-galactonate, 167 mM Tris-HCl buffer (pH 8.0), and dehydratase. In one trail, EDTA was added to the above reaction mixture to make a 1.0 mM solution; in the other trial, components_were left as above. Reaction mixtures were incubated at 30°C for 10 minutes to temperature equilibrate. Protein was added to initiate the enzymatic reaction in both trials; 51.0 09 protein was added to the EDTA-minus trial, and 510 ug of protein was added to the EDTA-plus trial. Samples of 0.05-ml volume were taken from each reaction mixture at reg- ular time intervals and added to 0.2 ml of semicarbazone reagent (1.0% semicarbazide-HCl and 1.5% sodium acetate) and incubated at 30°C for 15 min as to terminate the reaction and to form semicarbazone derivatives of the dehydratase product. Absorbance readings were taken at 250 nm and converted to micromole o-keto acid as to plot the micromole amount of product formed versus the time of incubation. 142 Amwuacwsv mz_p cop om om ON on on cc cm as c~ .fllllllldi .IJV. . I44 ,1 J 1 A, e i x. x e a e, emu \ a x I ~ ’ “23:5 _ Tm mmzuc mbzua a.» .7 i.nv (sniun 639411949) AIIAIIDV 157 seconds. A plot of the average sedimentation constant determined for each protein concentration used (Fig. 31) show an extrapolated zero protein 13 seconds for 5. Little dev- concentration value of 13.22 :_0.09 X 10' iation from this value was found for any protein concentration used. Using both the derived sedimentation constant and Stokes radius from the gel filtration experiment (see Table 16, for calculations) a molec- ular weight of 317,300 was obtained. (iv) Sedimentation Eguilibrium. The same protein sample of the D-galactonate (D-fuconate) dehydratase used in the sedimenation velocity experiment was used for sedimentation equilibrium. Dehydra- tase was diluted with 0.05 M potassium phosphate buffer (pH 7.0) to a protein concentration giving a low absorbance (less than 0.1) at 280 nm and was subjected to ultracentrifugation as described in Materials and Methods. Calculation of the molecular weight was performed using the following equation: 2 RT I d (Log C) MN = _x 2.303 x (1.5.) .2 a ( r L J L J where: MN is the molecular weight, R is the gas constant 7 1 (8.314 X 10 ergs,°C' , mole’l), T is the absolute temperature of the centrifuge run (293.2°K), p is the density of the solvent at the given temperature (0.999 g/ml), u is the angular velocity in radians per second ( m = 2RPM/60), 5 is the partial specific volume of the protein, assumed to be 0.733 g/ml3 (that for an "average" protein), r is the 158 Figure 31. Sedimentation velocity determination of the sedimenta- tion constant for the D-galactonate (D-fuconate) dehydratase. The Sephadex G-200 purified D-galactonate (D-fuconate) dehydratase, equili- brated against 0.05 M potassium phosphate buffer (pH 7.0), was subjected to ultracentrifugation as described in Materials and Methods. Five concentrations of the protein, 0.03 mg/ml, 0.06 mg/ml, 0.20 mg/ml, 0.60 mg/ml, and 1.50 mg/ml, were run synchronously and absorbance traces were made for each at regular time intervals using a heat-sensitive plotter. Values for the half height of the absorbance boundary and corresponding migration distances from the meniscus were measured for each trace. An average of seven such measurements were made for each protein concentra- tion. Calculations of the sedimentation constant were as described in the text. For each protein concentration, calculations were made varying the r1 and t1 values to give a good statistical average for the sedimentation constant (5). Measurements of r are made knowing that the first reference hole of the double sector cell marks both the beginning of the absorbance trace and a distance of 5.7 cm from the center of rotation; while the second reference hole marks the end of the absorbance trace, a distance of 1.6 cm from the first reference hole. 159 c.~ m.— P a; r A.e\as. zc_e<¢_:mo=cu z_uec¢a 4.. . P N.— a; P m5 P f8 O 21 1' (“vas€l_01) fl r 9 lNVlSNOO NOI 1V1N3NI OBS 160 distance from the center of rotation in centimeters, and C is the pro- tein concentration at the given r value. Determination of the [d (Log C) / d (r2)] factor (the slope) from a plot of the Log C versus r2 (Fig. 32) from three separate centri- fuge runs, gave an average molecular weight of 293,200 :_3900 for the dehydratase. Subunit Molecular Neight. SDS polyacrylamide gels, 10%, were pre- pared as described in Materials and Methods. D-Galactonate (D-fuconate) dehydratase, Sephadex G-200 purified protein, was electrophoresed with known molecular weight standards catalase (60,000 MN), trypsin (23,000 MN), and lysozyme (14,300 MN). From a plot of the Log (MN) versus mo- bility (Fig. 33) a subunit molecular weight of approximately 46,000 was interpolated for the dehydratase. Summary of BothpNative and Subunit Molecular Neight Determinations. Through either single-step experiments or by combination of molecular weight parameters, an average molecular weight of 289,100 was determined for the dehydratase native enzyme (Table 16). The dehydratase subunit molecular weight, of about 46,000, suggested that the native enzyme was comprised of six identical subunits. Amino Acid Composition of the Dehydratase Sephadex G-200 purified protein was used for all studies involving the amino acid composition and dry weight analysis of the D-galactonate (D-fuconate) dehydratase. For amino acid composition, the protein was equilibrated against 0.05 M potassium phosphate buffer (pH 7.0) by pas- sing over a Sepharose A-5M gel filtration column of appropriate 161 Figure 32. Sedimentation equilibrium Log C versus r2 plot. The Sephadex G-200 purified D-galactonate (D-fuconate) dehydratase, equili- brated against 0.05 M potassium phosphate buffer (pH 7.0), was subjected to ultracentrifugation to establish equilibrium as described in Materials and Methods. A protein concentration of 0.09 mg/ml in the above buffer was run against a buffer blank and absorbance traces were made using a heat sensitive plotter. Values for both the absorbance at 280 nm and corresponding distances from the center of rotation were made as described for the sedimentation velocity run. Baseline absorbances for these distances were subtracted from absorbance measurements as they originated from buffer absorbances only, not protein concentration. .Baseline absorbances were established by pelleting the sample to the bottom of the cell, running at 24,000 rpm for 24 hrs, and recording absorbance values at regular distances from the center of rotation. Additional equilibrium measurements, not shown, were obtained by setting the speed back to 11,000 rpm after pelleting and allowing for a 24 hr re-equilibration. (The process of pelleting and restoration of equili- brium was repeated several times accounting for the three separate trials mentioned in the text). Simplification of the molecular weight calcula- tion was made by substituting corrected absorbance (280 nm) values for the concentration factor (C), as the absorption optical system used‘ followed Beers' law at protein concentrations used in these ultracentri- fuge runs. 162 ‘ oo.c m—.o- om.oi m¢.ou ca.o- m~.o- oa.o- mo..- Asa om~ .uo=< mpom 38.2. .. 5:29.. mace, 35 a: 8.52538 2m 3..-: com.~_m m.e.=.=.m.z n 3: za_oo.o> =o_uou:ms_umm nee =o_aoeu_.» .oa Romu.~v copueucme_umm . m.e.=.w.c.z n 3: ago—coca xu_m:ou omoeozm com mam tee copuocu_pw _ou Eta—3...... a- oo~.mm~ we msmem> Auv mod copuaucospuom me x «Anzv n ~Azzv =o_uua:oe_uom gem—uoga .. oac.om~ «\m Admv xupmcou omocusm .. coo.mm~ oe=_o> mama—o mamco> Azzv mod eo.umcu—_e poo u_=:a=m osx~=u o>—umz p:a_m3 ¢<4=uu4ox .omoumcuxgmu Amuecouzmiov oueeouuo_emio one we agape: co—aumpcs ecu cow zmhuz<¢ Fe2+ > Mn2+) are an absolute requirement for activity as demonstrated by complete inacti-- vation of the dehydratase with EDTA and restoration of the activity by the addition of the above metals. Such specificities for divalent metals have been shown with other aldonate dehydratases (25,33,56,130,131). D-Galactonate (D-fuconate) dehydratase does not process detectable KDGal kinase, KDF aldolase, or KDPGal aldolase activities in the purified enzyme preparation. The further metabolism of KDF is dependent on aldolytic cleavage, whereas that of KDGal is dependent on phosphorylation and cleavage, as described in the preceding section. REFERENCES 197 10. 11. 12. 13. 14. 15. 16. 17. 18. REFERENCES Mortlock, R.P., Fossitt, P.P., and Hood, N.A., Proc. Nat. Acad. Sci. (U.S.A.) s1, 572 (1965). Camyre, K.P., and Mortlock, R.P., J. Bacteriol. 29, 1157 (1965). Tanaka, S., Lerner, S.A., and Lin, E.C.C., J. Bacteriol..g§, 642 (1967). Torriani, A., and Rothman, P., J. Bacteriol. 8;, 835 (1961). Lerner, S.A., Nu, T.T., and Lin, E.C.C., Science 136, 1313 (1964). NU, T.T., Lin, E.C.C., and Tanaka, S., J. Bacteriol. 26. 447 (1964). Rigby, P.N.J., Burleigh, Jr., 8.0., and Hartley, B.S., Nature 251, 200 (1974). Rigby, P.N.J., Gething, M.J., and Hartley, 8.5., J. Bacteriol. 125, 728 (1976). Clarke, P.H. (1974) in Evolution in the Microbial Norld (Carlile, M.J. and Skekel, J.J., editors), pp 183-217, Cambridge Univeristy Press, London. aackiag, A.J., Aquilar, J., and Lin, E.C.C., J. Bacteriol. $36, 522 1978 . Anderson, R.L., and Allison, 0.P., J. Biol. Chem. 240, 2367 (1965).- Mayo, J.N., and Anderson, R.L., J. Biol. Chem. 243. 6330 (1968). Mayo, J.N., and Anderson, R.L., J. Bacteriol. $99. 948 (1969). Hegeman, G.V., and Rosenberg, S.L., Ann. Review of Microbiol. g5, 429 (1970). MacGee, J., and Doudoroff, M., J. Biol. Chem. 210, 617 (1954). Entner, N. and Doudoroff, M., J. Biol. Chem. 196, 852 (1952). Ashwell, G., Methods. Enzymol. 3, 73 (1957). Stouthamer, A.H., Biochem. BiOphys. Acta 48, 484 (1961). 198 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40. 41. 199 DeLey, J., and Doudoroff, M., J. Biol. Chem. 227, 745 (1957). Pallerone, M.J. and Doudoroff, M., J. Biol. Chem. 223, 499 (1956). Szymona, M., and Doudoroff, M., J. Gen. Micro. 22, 167 (1960). Neimberg, R., J. Biol. Chem. 234, 727 (1959). Dahms, A.S., and Anderson, R.L., Biochem. BiOphys. Res. Commun. 36, 809 (1969). Elzaiag, T.A., Hassan, M.M., and Allan, A.M., J. Bacteriol. 114, 457 1973 . andreasen, J.R., and Gottschalk, G., Arch. Mikrobiol. 62. 160 1969 . Narrod, S.A. and Hood, N.A., J. Biol. Chem. 229, 45 (1966). Gunsalus, T.C., Horecker, B.C., and Hood, N.A., Bact. Rev. I2. 79 (1955). Deacon, J., and Cooper, R.A., EEBS Letters 11, 201 (1977). Cooper, R.A., Arch. Microbiol. 118, 199 (1978). Cooper, R.A., FEBS Letters 193, 216 (1979). Dahms, A.S., and Anderson, R.L., J. Biol. Chem. 247, 2222 (1972). Dahms, A.S., and Anderson, R.L., J. Biol. Chem. 251, 2228 (1972). Dahms, A.S., and Anderson, R.L., J. Biol. Chem. 247, 2233 (1972). Dahms, A.S., and Anderson, R.L., J. Biol. Chem. 247, 2238 (1972). Yuen, R., and Schachter, H., Can. J. Biochem. 59. 798 (1972). Nwokoro, N.A., and Schachter, H., J. Biol. Chem. £59, 6185 (1975). NwokOro, N.A., and Schachter, H., J. Biol. Chem. 259. 6191 (1975). Chan, J.Y., Nwokoro, N.A.,_and Schachter, H., J. Biol. Chem. 254, 7060 (1979). Dagley, S., and Tridgell, P.N., Biochem. J. 95, 48 (1965). Blumenthal, J., and Fish, O.C., Biochem. BiOphys. Res. Commun. 11, 239 (1963). Blumenthal, J., and Jepson, T., Biochem. BiOphys. Res. Commun. Ll, 282 (1964). 42. 43. 44. 45. 46. 47. 48. 49. 50. 51. 52. 53. 54.. 55. 56. 57. 58. 59. 60. 61. 62. 63. 200 Merrick, J.N., and Roseman, S., J. Biol. Chem. 235, 1274 (1960). Ashwell, G., Nabba, A.J., and Hickman, J., J. Biol. Chem. 235, 1559 (1960). Kerster, K., and DeLey, J., Antonie von Leeweenhoek 34, 293 (1968). Eisenberg, R.C. and Dobrogosz, N.J., J. Bacteriol. 23, 941 (1967). Fraenkel, D.G. and Horeaker, B.L., J. Biol. Chem. 239, 2765 (1964). Frampton, E.N. and Hood, N.A., J. Biol. Chem. 236, 2571 (1961). DeLey, J., Enzymglogia lg, 99 (1957). Frampton, E.N. and Hood, N.A., J. Biol. Chem. 236, 2578 (1961). Neijssel, O.M. and Tempest, D.N., Arch. Microbiol. 105, 183 (1975). Andreesen, J.R., and Gottscalk, G., J. Bacteriol. 107, 570 (1971). Kersters, K., Khan-Matsubara, J., Nelen, L., and DeLey, J., Antonie von Leewenhoek 31, 233 (1971). Palleroni, M.J., and Doudoroff, M., J. Bacteriol. 14, 180 (1957). Allam, A. M., Hassan, M. M., and Elzaing, T. A., J. Bacteriol. 124, 1128 (1975). Pedrosa, F.0., and Zancan, G.T., J. Bacteriol. 119, 336 (I974). aeffcaat, R., Hassel, H., and Dagley, S., Biochem. J. 115, 969 1969 . Stoolmiller, A.C., and Abeles, R.H., J. Biol. Chem. 24;, 5764 (I966). Touster,0 .(1969) in Com rehensive Biochemistr (Florkin, M., and' Stotz, E.H ., editors) Vol. I7, pp 219- 240, Elsevier Publishing Company, New York. LeBlanc, D.J., and Mortlock, R.P., J. Bacteriol. 196, 82 (1971). LeBlanc, D.J., and Mortlock, R.P., J. Bacteriol. 106, 90 (1971). Boulter, J., Gielow, 8., McFarland, M., and Lee, N., J. Bacteriol. 1_Il, 920 (1974). Simpson, F.J., Nolin, M.J., and Hood, N.A., J. Biol. Chem. 230, 457 (1958). Heath, E.C., and Ghalamor, M.A., J. Biol. Chem. 237, 2423 (1962). 64. 65. 66. 67. 68. 69. 70. 71. 72. 73. 74. 75. 76. 77. 78. 79. 80. 81. 82. 83. 84. 201 Ghalamor, M.A., and Heath, E.C., J. Biol. Chem. 237, 2427 (1962). Eagon, R.C., J. Bacteriol. 82, 548 (1961). Hanson, T.E., and Anderson, R.L., Proc. Nat. Acad. Sci. (USA) 6;, 269 (1968). Sapico, V., Hanson, T.E., Nalter, R.H., and Anderson, R.L., J; Bacteriol. 26. 51 (1968). Hanson, T.E., and Anderson, R.L., J. Biol. Chem. 241, 1644 (1966). Egan, J.B., and Morse, M.C., Biochem. Biophys. Acta 21, 310 (1965). aiss,)M., Horwitz, 5.8., and Kaplan, N.0., J. Biol. Chem. 231, 1342 1962 . Markwell, J., Shimamoto, G.T., Bissett, 0.L., and Anderson, R.L., Biochem. Biophys. Res. Commun. 1;, 221 (1976). Kelker, N.E., Simkins, R.A., and Anderson, R.L., J. Biol. Chem. 247, 1479 (1972). Palmer, R.E., and Anderson, R.L., J. Bacteriol. 112, 1316 (1972). Palmer, R.E., and Anderson, R.L., Biochem. BiOphys. Res. Commun. 15, 125 (1971). Palmer, R.E., and Anderson, R.L., J. Biol. Chem. 247, 3415 (1972). Palmer, R.E., and Anderson, R.L., J. Biol. Chem. 247, 3420 (1972). Kelker, N.E., Hanson, T.E., and Anderson, R.L., J. Biol. Chem. 245, 2060 (1970). LeLoir, L.F., Biochem. J. 9;, 1 (1964). Lowry, 0.H., Rosenbrough, N.J., Fair, A.L., and Randall, R.J., gt; Biol. Chem. 12.3.. 265 (1951). Dubois, M., Gilles, K.A., Hamilton, J.K., Rebers, P.A., and Smith, F., Anal. Chem. 28. 350 (1956). Hestrin, S., J. Biol. Chem. ;§Q, 249 (1949). Paz, M.A., Blumenfeld, 0.0., Rojkind, M., Henson, E., Furfine, C., and Gallop, P.M., Arch. Biochem. Biophys. $92, 548 (1965). Neissbach, A. and Hurwitz, J., J. Biol. Chem. 234, 705 (1959). Moore, 5. and Link, K.P., J. Biol. Chem. 133, 293 (1940). 85. 86. 87. 88. 89. 90. 91. 92. 93. 94. 95. 96. 97. 98. 99. 100. 101. 102. 202 Cornforth, J.N., Firth, M.E. and Gottschalk, G., Biochem. J. 68. 57 (1958). Zagalak, 8., Frey, P.A., Karabatsos, G.L., and Abeles, R.H., J; Biol. Chem. 241, 3028 (I966). Portsmouth, P., Carbohydrate Research 8, 193 (1968). Dahms, A.S., Ph.D. Thesis (1969) Metabolism of D-Fucose and L-Arabinose in a Pseudomonad, pp 185-186. Narren, L., J. Biol. Chem. 234, 1971 (1959). Aronson, L.0., Rosso, R.C., and Adams, E., Biochim. Biophys. Acta. l3_2. 200 (1967). Kramer, D.N., Klein, N., and Baselia, R.A., Anal. Chem. 3_1_, 250 (1959). Pese, A., McKay, R.H., Stolzenbach, F., Cohn, R.D., and Kaplan, N.0., J. Biol. Chem; 239, 1753 (1964). Hohorst, H.S., and Reim, M. (1963) in Methods of Enz atic Anal sis (Bergmeyer, H.U., editor), pp 335-339, Academic Press, New York and London. . Loveless, A., and Howarth, S., Nature 185, 1780 (1959). Bissett, 0.L., Ph.D. Thesis (1975), Lactose and D-Galactose Metabolism in Staphylococcus aureus. . ., p 179. . Baker, F.J. (1967) in Handbook of Bacteriolo ical Techni ue (Baker, F.J., and Breach, M.R., editors) pp -2 2, App eton-Century-Crofts Publishing Company, London. Donald, A., Sibley, 0., Lyons, D.E., and Dahms, A.S., J. Biol. Chem. 2.5.4.. 2132 (1979). . Simkins, R.A., Ph.D. Thesis (1975), L-Sorbose-I-Phosphate Reductase ‘tggm_5grgpacter Aero enes: Its Purificatio , Characterization and Role in thgvMetabolism of L-Sorbose and D-Fructose, pp 103-105} Kauffman, M.D. (1966) in The Bacteriolo of Enterobacteriaceae, pp 19-41 (Escherichia) and pp - 3 K ebsie a), Nilliams and Nilkins Company, Baltimore. Hayes, N. (1968) in The Genetics of Bacteria and Their Viruses, pp 711-718, John Niley and Sons, Inc., New York. Rutter, N.J., Fed. Proced. 23, 1248 (1964). Feist, C.F. and Hegemon, 6.0., J. Bacteriol. 100, 869 (1969). 103. 104. 105. 106. 107. 108. 109. 110. 111. 112. 113. 114. 115. 116. 117. 1180 119. 120. 121. 122. 203 a.) Norsey, M.J., and Nilliams, P.A., J. Bacteriol. 123, 7 (1975). b.) Nilliams, P.A., and Murray, K., J. Bacteriol. yep, 416 (1974). c.) Nong, C.L., and Dunn, N.N., Genet. Res. 2;. 227 (1974). Revzin, A.R., Private Communication. a.) fialacaaudhun, S., and Chakrabarty, A., J. Bacteriol. 126, 410 1976 . b.) Duggleby, C.J., Bayley, S.A., Norsey, M.J., Nilliams, P.A., and Broda, P., J. Bacteriol. 130, 1274 (1977). Dahms, A.S., Ph.D. Thesis (1969) Metabolism of D-Fucose and L-Arabinose in a Pseudomonad, pp 258-262. Hood, N.A. (1972) in The Enzymes (Boyer, P.D., editor), Vol VII, 3rd Edition, pp 281-301, Academic Press, New York and London. Blumenthal, N.J., Methods Enzymel. g, 660 (1966). Peterson, E.A., and Sober, H.A., Methods Enzymol. §, 3 (1962). McGuinness, E.T., J. Chemical Educ. eg, 826 (1973). Vesterberg, 0., Methods Enzymol. ge, 389 (1971). Linhardt, K., and Nalter, K. (1963) in Methods of Enzymatic . Analysis, (Bergmeyer, H.U., editor), pp - , Academic Press, New York and London. Luck, H. (1962) in Methods of Enzymetic Analysis, (Bergmeyer, H.U., editor), pp 885-894, Academic Press, New York and London. Crouthomil, C.E., Meek, H.V., Martin, 0.5., and Banks, c.v.,,1e_ Amer. Chem. Soc. 1;, 3031 (1949). Racker, E., Biochim. Biophys. Acta.fi. 211 (1950). Lamers, K., Putney, F., Steinberg, J.Z., and Schochman, H.K., Arch} Biochem. BiOphys. egg, 379 (1964). thantis, 0.A., Biochemistry g, 297 (1964). VanHolde, Fractions 1, (1967). ((Iaren3 A., and Levinthal, 0., Biochim. BiOphys. Acta §§, 470 1960 . Kanarek, L., Marler, E., Bradshaw, R.A., Fellows, R.E., and Hill, R.L., J. Biol. Chem. 239, 4207 (1964). Morawiecki, A., Biochim. Biophys. Acta 35. 604 (1960). Steinmetz,M, and Deal, Jr., N.C., Biochemistry §, 1399 (1966). 123. 124. 125. 126. 127. 128. 129. 130. 131. 204 Martin, R.G., and Ames, 8.N., J. Biol. Chem. 236, 1372 (1961). Brewer, J.N., Pesce, A.J., and Ashworth, R.B. (1974) in Experimental Techniques in Biochemistry, pp 195-197, Prentice-Hall, Englewood, Cliffs, New Jersey. Penke, 8., Ferenczi, R. and Kovacs, K., Anal. Biochem. e9, 45 (1974). Erestgield, A.M., Moore, S. and Stein, N.H., J. Biol. Chem. 238, 622 1963 . Malamud, 0., and Drysdale, J.N., Anal. Biochem. pg, 620 (1978). Reeck, G., Handbook of Biochemistry and Molecular Biology (3rd edt. , (G.D. Fasman, ed.), "Proteins“, Vol. III, pp 504-519 (1976), Press). Dahms, A.S., Fed. Proc. pg, 1900 (1974). Smilely, 0.0., and Ashwell, G., J. Biol. Chem. 235, 1571 (1960). Bender, R., and Gottschalk, G., Eur. J. Biochem. 59, 309 (1973).