mt'ay 3 ".".' . 79-" “3'" vw. “f7: z -,2 :.. v v - m ';. ‘4' ~ ““'-'.-| u w .J . . .4 .-A ~ ~- , ‘. . .. "u.“ .. ‘._ ._ , .... , . ‘ , ‘ PHYSICAL AND CATALYTIC PROPERTIES:OFsTHE'PURlN-E Z NUCLEOSIDE PHOSPHORYLASE-S. ISOLATED FROM; ,. VEGETATWE CELLS AND SPORES 0F BACILLUS CEREUS ' Thesis for the Degree’of Ph’. D. ‘ MiCH‘IGAN STATE UNWERSITY RICHARD WILLIAM GILPIN 1970 YHPQQ This is to certify that the thesis entitled Physical and Catalytic Properties of the Purine Nucleoside Phosphorylases Isolated from Vegetative Cells and Spores of Bacillus Cereus presented by Richard William Gilpin has been accepted towards fulfillment of the requirements for Ph.D. degree in Microbiology W JJJ JJJ Major professor[ Date May 14, 1970 0-169 LIBRARY Michigan State University ABSTRACT PHYSICAL AND CATALYTIC PROPERTIES OF THE PURINE NUCLEOSIDE PHOSPHORYLASES ISOLATED FROM VEGETATIVE CELLS AND SPORES OF BACILLUS CEREUS By Richard William Gilpin The purine nucleoside phosphorylase from vegetative cells and spores of Bacillus cereus T was purified to electrophoretic homogeneity by ion exchange chromato- graphy and polyacrylamide gel electrophoresis. The specific activity of these preparations was higher than those previously reported. Phosphate ion caused an increase in the thermal stability of both purified enzymes whereas inosine had no effect. The catalytic properties of the two enzymes were similar and indicated an ordered, sequential reaction mechanism where inosine was bound before phosphate. The Michaelis constants (inosine as substrate) for the vegetative cell and spore enzymes were A M and 7.0 x 10'“ M respectively. The Michaelis 4.6 x 10- constant for phosphate was 1.5 x 10'3 M for the vegetative cell enzyme and 1.3 x 10"3 M for the spore enzyme. Both enzymes had similar turnover numbers and the specific Richard William Gilpin activities of the enzymes appeared to be constant at varying protein concentrations. Negative homotropic effects were proposed since the Lineweaver-Burk plots with inosine as the changing fixed substrate and phos- phate as the variable substrate showed downward curvature at high phosphate concentrations. Sucrose density gradient studies indicated that spore purine nucleoside phosphorylase (PNPase) increased from a sedimentation velocity of 5.3 S in the absence of phosphate to a value of 5.7 S in the presence of 10 mM potassium phosphate. The increase in sedimenta- tion velocity may have been irreversible. The vegetative cell PNPase had a sedimentation velocity of 5.5 S in the presence or absence of phosphate ion. The calculated minimal molecular weight of spore PNPase was A6,300 in the absence of phosphate, and 92,600-in the presence of 10 mM phosphate. The vegetative cell enzyme had a molecular weight of 88,300 and 91,600 in the presence and absence of phosphate respectively. The subunit size for both enzymes was 2A,000‘:10%. The conformational structure of the spore enzyme was more compact in the absence of phosphate ion. Dimerization of the spore enzyme to a molecular weight approximating the size of the vegetative cell enzyme was accompanied by a change in conformation. Richard William Gilpin Although the synthesis of both the vegetative cell and spore enzymes was directed by the same genomic unit, the enzymes differed in their size and shape when iso- lated in the absence of phosphate ion. PHYSICAL AND CATALYTIC PROPERTIES OF THE PURINE NUCLEOSIDE PHOSPHORYLASES ISOLATED FROM VEGETATIVE CELLS AND SPORES OF BACILLUS CEREUS By Richard William Gilpin A THESIS Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Microbiology and Public Health 1970' ACKNOWLEDGMENTS I wish to thank my advisor, Dr. Harold L. Sadoff, for his excellent support and guidance during this investigation. I am grateful to George M. Stancel, Department of Biochemistry, for his assistance and counsel in the sedimentation equilibrium studies. I thank Andrew E. Seer, Jr., Department of Chemistry, for construction of the preparative disc-gel electrophoresis apparatus. Appreciation is also extended to James C. Johnson, Department of Biochemistry, for his informative discussions concerning subunit analysis. I also wish to thank Dr. Ralph N. Costilow, and other staff members of the Departments of Microbiology and Biochemistry for their guidance. 11 TABLE OF CONTENTS ACKNOWLEDGMENTS. . . . . . . . . . LIST OF TABLES . . . . . . . . . . . LIST OF FIGURES. . . . . . . . . . . INTRODUCTION. . . . . . . . . . . . LITERATURE REVIEW . . . . . . . . . . Metabolism and Sporulation . . . . Protein Turnover During Sporulation . Ordered Events During Sporulation. Heat Resistance. . . . . . . Purine Nucleoside Phosphorylase . . METHODS . . . . . . . . . . . . . Growth of Vegetative Cells and Spores of Bacillus cereus . . . . . . . Spectrophotometric Assay of Purine Nucleoside Phosphorylase . . . . . Measurement of Protein Concentration. . Protein Concentration by Ultrafiltration Preparation of DEAE-cellulose for Column Chromatography . . . Preparative Polyacrylamide Disc-gel Electrophoresis . . . . . . Analytical Polyacrylamide Disc-gel Electrophoresis . . Kinetic Studies of Vegetative Cell and Spore PNPase . . . . . . . Sucrose Density Gradient Centrifugation. Protein Absorption Spectrum of Spore PNPase . . Subunit Analysis by SDS- Polyacrylamide Gel Electrophoresis . . . High Speed Sedimentation Equilibrium Molecular Weight Study. . . . . . 111 Page ii vi \OO‘tU‘l-lrl': .C' |'-’ 13 l3 13 15 15 16 18 21 26 27 27 29 Page RESULTS . . . . . . . . . . . . . . 33 Purification of Purine Nucleoside Phosphorylase. . . . . . . . . . 33 Preparation of Crude Extract. . . . . 33 Streptomycin Sulfate Precipitation. . . 3A Ammonium Sulfate Fractionation of Crude Extract . . . . 35 DEAR-cellulose Column Chromatography . . 35 Preparative Disc-gel Electrophoresis .- . 37 Thermal Inactivation of Purine Nucleoside Phosphorylase. . A3 Analytical Polyacrylamide Disc-gel Electro- phoresis of PNPase . . . . A9 Ultraviolet Absorption Spectrum of Pure Spore PNPase . . . . 51 Effective Assay Range for PNPase Activity . 51 Subunit Activity of Spore PNPase . . . 5A Initial Velocity Analysis Vegetative Cell and Spore PNPase. . . . . 5A Subunit Size of Purine Nucleoside Phosphorylase. . . . . 66 Sucrose Density Gradient Sedimentation of PNPase . . . . . 68 High Speed Sedimentation Equilibrium Molecular Weights . . . . . . . . 74 DISCUSSION . . . . . . . . . . . . . 82 SUMMARY . . . . . . . . . . . . . . 99 LIST OF REFERENCES. . . . . . . . . . . 101 APPENDICES . . . . . . . . . . . . . 112 Appendix A--Materials Used in the Investigation . . 113 Appendix B--Derivation of the Fundamental Sedimentation Equation . . . 115 Appendix C--Ca1culation of Frictional Coefficients . . . . . . 120 iv LIST OF TABLES Table Page 1. Purification of vegetative cell purine nucleoside phosphorylase. . . . . . AA 2. Purification of spore purine nucleoside phosphorylase. . . . . . . . . . A5 3. Michaelis constants of the substrates for pure vegetative Cell and spore PNPases . 67 A. Sucrose density gradient centrifugation of pure vegetative cell and spore PNPase o o o o o o o o o o o o 73 5. Molecular weights of vegetative cell and spore PNPase by sedimentation equilibrium . . . a. . . . . . . 76 6. Calculated frictional ratios of veg. and spore PNPases. . . . . . . . . 81 Figure 10. ll. 12. 13. LIST OF FIGURES Elution profile of spore purine nucleoside phosphorylase from the preparative disc-gel electrophoresis column . . . Elution profile of vegetative cell purine nucleoside phosphorylase from the preparative disc-gel electrophoresis column. . . . . . . . . . . . Thermal inactivation of vegetative cell and spore PNPase at 60 C. . . . Analytical polyacrylamide disc-gel electrOphoresis of spore PNPase Ultraviolet absorption spectrum of purified spore PNPase. . . . . . Subunit activity of spore PNPase. Initial velocity of vegetative cell PNPase with inosine as the variable substrate Initial velocity of spore PNPase with inosine as the variable substrate. . . Initial velocity of vegetative cell PNPase with phosphate as the variable substrate Initial velocity of spore PNPase with phosphate as the variable substrate . . Determination of the molecular weight of the polypeptide chains of vegetative cell and Spore PNPases by SDS- polyacrylamide gel electrophoresis . Plots of In D vs. r2 for spore PNPase . . Plots of In D vs. r2 for vegetative cell PIJPaSeO O O O O O O O O I O 0 vi Page 39 Al A7 50 52 55 58 6O 62 6A 69 77 79 INTRODUCTION Members of the Bacillaceae are able to undergo a morphogenesis to form resting spores. Recently, this process has been compared to an abortive cell division (28). It is stringent with respect to both the metabolic state of the cells and their mineral nutrition. That is, certain molecular species of cations either enhance or inhibit sporulation (A0, 50, 55, 97). Spores are metab- olically dormant refractile bodies which are able to withstand extremes in environmental conditions much better than the vegetative cells from which they are derived. In suitable environments, spores germinate and resume vegetative growth. Thus, spore-forming organ- isms must be taken into consideration in processes which achieve sterilization by heating, drying, or chemical means. Scientific interest in spores was initially directed toward finding methods to decrease their survival during the processing of canned foods. More recently, because of the striking morphogenesis which occurs, sporulation has been studied as a model of cellular differentiation. Intact spores are heat resistant and thus it may be assumed that their most labile constituents, proteins, must also be heat stable in Xilgn Spore proteins would appear to be excellent models for study of the mechanism of heat resistance at the molecular level. In such studies, enzymes are usually the proteins of choice because they are easily detected by their catalytic properties and they can be purified to homogeneity. Comparisons between the physical and chemical properties of vegetative cell and spore enzymes have revealed dif- ferences which must be related to mechanisms by which spores achieve heat resistance. The significance of these differences in relation to spore properties is Just beginning to be understood. Gardner and Kornberg (20) found that the synthesis of the purine nucleoside phosphorylase (PNPase) enzymes from vegetative cells and spores of Bacillus cereus var. terminalis was directed by the same genomic unit. These authors concluded that the two enzymes had similar cata- lytic and physical properties. A careful examination of their data, however, showed that distinct differences existed between the two PNPases. Therefore, further studies were made in this laboratory. Engelbrecht and Sadoff (16) found that pure spore and vegetative cell PNPases differed in their physical and catalytic properties depending on the phosphate ion concentration of the surrounding environment. Further characterizations of the physio-chemical differences between the cell and spore enzymes are reported in this dissertation. LITERATURE REVIEW Metabolism and Sporulation In batch cultures, the sporulation of Bacillus species occurs at the end of exponential growth when r n cells are in a metabolic shift-down condition. Control of sporulation is not well understood, but the avail- ability of glucose and the buildup of metabolic products from the fermentation of glucose, such as acetate and poly-B-hydroxybutyrate, seem to be important. Glucose is a non-specific repressor of sporulation and may cause catabolite repression of the sporulation-specific enzymes (AA, 80). Acetate, derived from the metabolism of sub- strates or when added exogenously, induces the glyoxylate cycle in bacilli in the absence of glucose (25). The tricarboxylic acid (TCA) cycle enzymes have been found at higher levels in sporulating cells than in log phase cultures (6, 13, 89), which are usually devoid of aconitase activity (89). It would appear that these enzymes function in the sporangium during the course of sporulation and, when coupled to oxidative phosphorylation, supply energy for the sporulation pro- cess. Spore extracts have no detectable TCA cycle activity. Thus it would appear that this metabolic cycle is a well compartmentalized function during spor- ulation. Protein Turnover During Sporulation Protein turnover occurs during sporulation. In Bacillus subtilis sporulating in minimal medium (57) or in supplemented nutrient broth (8A), protein turnover during sporulation was 18% per hour. Proteins were con— served during exponential growth (8A) and little or no turnover occurred. Genetic (83) and kinetic data (2, 19, 6A, 93) have indicated that the protein turn- over occurs in the sporangium during sporulation. Ordered Events During_Sporulation Sporulation occurs in a temporal sequence of events after the derepression of the TCA cycle. The time course for both morphogenesis (66) and characteristic enzyme formation (13, 73) has been noted. Sporulation commences in bacilli at the end of exponential growth and is com- plete in 6 to 8 hours. Studies of particular interest have been made of the onset of protease activity (5), alkaline phosphatase activity (63), glucose dehydrogenase activity (3), refractility (72), synthesis of dipicolinic acid (69), calcium ion uptake (72), and heat resistance (9A). A metabolic or genetic block at any point in the sequence of sporulation blocks all subsequent sporulation events once the commitment to sporulation has been made. Heat Resistance Heat resistance is one property of bacterial spores and it is easily measured. Spore enzymes are usually heat stable in vivo but become labile when removed from the resting spore. A few exceptions have been found, however. The reduced nicotinamide-adenine-dinucleotide oxidase from spores of Clostridium botulinum (22) and the catalase from spores of B. cereus (75) appeared to be intrinsically stable. In general, the mechanism of heat resistance of spore proteins is unknown. Various authors, in attempting to explain heat resistance, have suggested that spore and vegetative cell enzymes of homologous activity are uniquely dif— ferent proteins (23, 2A). If this were the case, the cell genome would have to possess a large sporulation- specific complement. This "spore genome" has not been found. One specific cistron per enzyme directed the synthesis of both vegetative cell and spore PNPase (20 alanine dehydrogenase (56), and DNA polymerase (17). Presumably most spore enzymes are coded by the same segment of the chromosome which directs the synthesis of the corresponding vegetative cell protein and some mechanism exists to render them heat resistant (3, 5, Spore enzymes may be heat stable due to their attachment to particles in the intact spore. Alanine racemase of B. cereus was found to be stable when it ), A1). was attached to spore wall material (85, 86). Ribosidase was also thought to be stable for this reason (A, 71). Alanine racemase (85), catalase (58), and adenosine deaminase (71), which are active in the intact spore (A8), were found in either the spore coat or the exo- sporium. If this is the case, the aggregates must decom— pose on disruption or germination of the spores, rendering most of the proteins labile. The ionic environment of a protein must play a role in its heat stability. It is known that spores contain- ing high levels of divalent cations tend to be beat stable, whereas those grown in media deficient in these ions tend to be heat labile (9A). The effect of the ionic environment on heat resistance can also be shown with purified enzymes. Glucose dehydrogenase from spores of B. cereus was found to be heat stable due to a revers- ible dimer-monomer interconversion promoted by hydrogen ions, with the monomer showing more thermal stability (3, 76). This enzyme could be further stabilized by increasing concentrations of Group IA cations. The fructose 1,6-diphosphate aldolase from spores of the same organism was activated and stabilized specifically by increasing concentrations of calcium (77). A close relationship between protease activity and sporulation has been described by several authors (5, 51, 60, 61, 79, 82). The activity of protease may result in many enzymes being modified or degraded during sporulation. That is, proteolytic activity may not be restricted to protein turnover in the sporangium. The altered enzyme forms resulting from proteolysis of the "vegetative protein" may acquire the attribute of heat resistance in the environment of the spore. The fructose 1,6—diphosphate aldolase from vegetative cells of B. l“ cereus appeared to be cleaved by a limit—protease during sporulation to produce an enzyme which was smaller than its vegetative cell counterpart (77). The modified aldolase in the presence of calcium was heat resistant. Protease-negative mutants of B. Subtilis were either asporogenous or oligosporogeneous (57, 79, 82). No studies have been made of the heat resistance of enzymes from these types of mutants. When a protease inhibitor such as L-cysteine was added to the medium of a spor- ulating culture, the intracellular degradation of protein was delayed for many hours and the final yield of spores was much lower than normally found (57). When chloram- phenicol was added to a sporulating culture at levels that inhibited protein synthesis, protein degradation and sporulation were prevented (8A). The chloramphenicol may have prevented the synthesis of protease or prevented the synthesis of a protein activator for protease. Kornberg (A2) suggested that protein synthesis may be required during sporulation to depress the level of a protease inhibitor(s) such as L-cysteine. Once protein degradation started, the addition of chloramphenicol did not alter its rate although protein synthesis and sporulation were completely blocked (8A). Proteolysis might account for the size differences which have been noted between the vegetative cell and spore purine nucleoside phosphorylases found in B. cereus (l6). Purine Nucleoside Phosphorylase Purine nucleoside phosphorylase (PNPase, EC 2. A. 2. 1) occurs in vegetative cells and spores of B. cereus T, from which it can be readily isolated. This enzyme can also be induced to three-fold higher concentrations in vegetative cells by one of its substrates, inosine (20). PNPase catalyzes the reaction; purine nucleoside + orthophosphate ea-D-ribose-l-phosphate + purine. Interest in this enzyme was stimulated by its possible role in germination induced by inosine. B. cereus T spores can be germinated in sodium phos- phate buffer containing inosine or inosine plus L-alanine (7A). Krask and Fulk (A3) found a nucleoside phosphorylase in extracts of B. cereus that cleaved adenosine or inosine in the presence of inorganic phosphate to produce ribose- l-phosphate and purine base. A phosphoribomutase and an adenosine triphosphate ribokinase were also found. These authors concluded that these enzymes could function during germination by synthesizing phosphate esters which would 10 ultimately yield ATP. It has been shown that the viability of stored erythrocytes can be restored by the addition of nucleosides which leads to the resynthesis of phosphate esters and ATP (8). Spores of many bacilli can cleave adenosine to free base and ribose (A6, A7). Spores of B. cereus T were able to degrade adenosine, guanosine, inosine, xanthosine, adenylic acid, cytidine, and uridine (A6). Powell and Hunter (70, 71) found that other strains of B. cereus, which germinated more efficiently with inosine instead of adenosine, could hydrolyze ribosides at variable rates. These data suggested a role for nucleoside phosphorylase in spore germination. However, Gardner and Kornberg (20) using a mutant of B. cereus T containing low levels of PNPase activity found that germination was as rapid and complete in inosine as the wild type. This finding contradicts the hypothesis of Krask and Fulk (A3) and reopens the question of the role of inosine and PNPase in riboside-induced germination. The first nucleoside phosphorylase was discovered in rat lever (32, 33, 3A, 35). The enzyme catalyzed the reaction; hypoxanthine + ribose-l-phosphate #inosine + inorganic phosphate. The reaction equilibrium favored the production of inosine. PNPase which will degrade adenosine (1A, 39, A6, A7) or inosine and guanosine (27, 58) has also been isolated. This enzyme has been found in human erythrocytes (37, 92), chicken liver (65), ll yeast (97), Escherichia coli (58), and B. cereus (A6, A7, A8). The metabolic significance of this enzyme is unknown. The ease with which it can be purified makes it useful as a model enzyme in the study of protein differences between vegetative cells and spores of B. cereus. Gardner and Kornberg (20) compared the PNPases from vegetative cells and spores of B. cereus and con- cluded that the two enzymes were under control of the same genomic unit. Both enzymes appeared to have similar catalytic and physical properties, but there were subtle differences which prompted further study. Engelbrecht and Sadoff (16) found that pure spore and vegetative cell PNPases differed in their physical properties depending on the phosphate ion concentration of the surrounding environment. In the absence of phosphate ion spore PNPase had an estimated molecular weight of 87,000 which increased to 117,000 in the presence of 10 mM phosphate. Vegetative cell PNPase had a molecular weight of 110,000 regardless of the phosphate concentration. Spore PNPase also had a 7-fold higher thermal stability than the vegetative cell enzyme in the absence of phosphate. When the phosphate ion concentration was increased to 10 mM, the thermal stab- ility of the spore PNPase decreased to a value equivalent to that of the vegetative cell PNPase which did not 12 change. Engelbrecht and Sadoff (16) proposed that spore PNPase underwent aggregation in phosphate ion concen- trations greater than the Km for phosphate as substrate A (7.2 x 10- M). Vegetative cell PNPase was assumed to be unaffected by phosphate. METHODS Growth of Vegetative Cells and Spores of Bacillus cereus Bacillus cereus T (76) was grown in modified G- medium (26)., Three liters of exponentially growing cells were used to inoculate a 100 liter fermenter using the step—culture method of Sadoff, et a1. (76). Vege- tative cell cultures were grown at 30 C to mid- logarithmic phase (0.7 optical density at 620 nm on a Bausch and Lomb Spectronic 20) in A hr. Cultures grown for 21 hr yielded greater than 95% free spores. The cells and spores were harvested from the 100 liter fermenter by cooling the culture to 1A C and collecting the cells with a Sharples continuous flow centrifuge. The cell pastes were stored at -15 C. Spectrophotometric Assay of Purine Nucleoside Phosphorylase PNPase cleaves inosine in the presence of inorganic phosphate to yield hypoxanthine and ribose—l-phosphate., A coupled spectrophotometric assay of the enzyme was utilized (35) in which hypoxanthine was oxidized to uric acid by xanthine oxidase. The rate of increase in absorbance due to uric acid formation was followed at 13 1A 290 nm. The assays were performed at 37 C in a Perkin- Elmer Model 12A, double beam diffraction grating spec- trophotometer coupled to a Sargent Model SL recorder. Alternatively, a Beckman Model DU spectrophotometer coupled to a Sargent Model SL recorder was used. The standard assay system contained the following reactants: inosine, 2 mM; potassium phosphate buffer, pH 7.5, 30 mM; xanthine oxidase, 0.01 unit; and l to 10 units of PNPase. The final assay volume was 1 ml in 1 cm light path cuvettes. A11 essay components except PNPase were preincubated for 10 min at 37 C to remove trace amounts of hypoxanthine. For the kinetic studies all enzyme and substrate dilutions were made in 50 mM trishydroxymethylaminomethane (Tris)-HCl, 10 mM 2- mercaptoethanol buffer, pH 7.5. This buffer was used throughout the study and was designated the standard buffer. One unit of PNPase activity was that amount of enzyme needed to catalyze the hydrolysis of 1 umole of inosine per hr at 37 C. Specific activity was calculated as units of PNPase activity per mg protein. Xanthine oxidase was prepared from fresh un- pasteurized Jersey cow milk by the procedure of Gilbert and Bergel (21). The purified protein was kept at 2 C as a 65% ammonium sulfate precipitate in 200 mM potassium 15 phosphate buffer, pH 6.0, and only 35% loss of activity was found after ten month's storage. One unit of xanthine oxidase was that amount of enzyme needed to catalyze the oxidation of l umole of hypoxanthine per min at 25 C. Measurement of Protein Concentration, Protein concentrations were estimated routinely by the method of Warburg and Christian (95). The pro- tein contents of highly purified PNPase preparations were determined by the method of Lowry, et a1. (5A). Protein Concentration by Ultrafiltration Dilute solutions of PNPase were concentrated in a Diaflo ultrafiltration cell with type UM-lO ultrafiltra- tion membranes (Amicon Corporation, Lexington, Ma.). A positive pressure of 50 psi was maintained over the membrane with compressed nitrogen. Preparation of DEAR-cellulose for Column Chromatography One kilogram of Whatman DE-52 DEAE-cellulose was suspended in 5 liters of distilled water and the fine particles were removed by decantation. The exchanger was filtered over Whatman #1 filter paper in a Buchner funnel and suspended in A liters of 0.5 N HCl. The suspension was stirred for 30 min and then filtered l6 and washed with distilled water until the effluent pH was A.0. It was then stirred into A liters of 0.5 N NaOH for 30 min, filtered, and subjected once more to this treatment. The DEAE-cellulose was washed with distilled water until an effluent pH of 7.0 was obtained. It was then suspended in A liters of 2A0 mM NaCl and the pH of the mixture was slowly adjusted to 7.5 using 0.2A N HCl. The ion exchanger was then filtered and equi- librated in 2A0 mM Tris-H01, 10 mM 2-mercaptoethanol buffer, pH 7.5. Preparative Polyacrylamide Disc-gel Electrophoresis The apparatus used to purify the PNPase enzymes was constructed according to the design of Jovin, et a1. (31), by Andrew E. Seer Jr., Master Glassblower, Depart- ment of Chemistry, Michigan State University, East Lansing, Michigan. An important design modification was made consisting of a u—tube which was designed to hold the lower buffer and the anode. This simplified the operation of the device since it did not have to be partially immersed in a large Jar containing the lower buffer. The electrophoresis unit was operated in a 5 C cold room and was cooled by circulating ice water. At the current loads used for this study, the gel temper- ature was maintained below 5 C. 17 The gel system was a modification of that reported by Ornstein and Davis (68). The stock solutions used for preparation of the gels contained the following com— ponents taken to a final volume of 100 ml with distilled water: A B A.O ml conc. HCl A.0 ml conc. HCl 0.23 ml TEMED O.A6 m1 TEMED 36.3 g Tris 5.98 g Tris Final pH, 8.8 to 8.9 Final pH, 7.3 to 7.A CN DN 30.0 g acrylamide 1A.0 g acrylamide 0.8 g bisacrylamide 0.25 g bisacrylamide E Catalyst A.0 mg riboflavin 0.1A g ammonium persulfate The lower separating gel contained the following amounts of the stock solutions in a final volume of 100 m1; Parts Stock 1 A 2 ON A Catal. These were mixed, placed in the column, overlaid with distilled water, and allowed to polymerize at 5 C for 2 hr. The upper stacking gel contained the following DrOportions of stock solutions in a final volume of 50 ml; 18 Parts Stock 1 B 2 DN 1 E A H2O The water was removed from the top of the polymerized lower gel and the upper gel mixture was added. The stack- ing gel was photopolymerized for 2 hr at 5 C with four 15 watt fluorescent lights. The upper buffer consisted of 5.76 g glycine and 1.2 g Tris in a final volume of 2 liters, pH 8.3. The lower/elution buffer was made by adding solid Tris to 3 liters of distilled water containing 15 m1 of cone. H01, until a pH of 8.1 was obtained. A freshly prepared gel column was used for each electrophoresis run. Analytical Polyaorylamide Disc-gel Electrophoresis The analytical disc-gel electrophoresis technique of Ornstein and Davis (68) was used to determine the purity of a variety of PNPase preparations. The elec- trophoresis buffer cOntained 28.8 g glycine and 6.0 g Tris per liter with a final pH of 8.3. This buffer was diluted lO—fold with distilled water before use. The stock solutions used for preparation of the gels con- tained the following components taken to a final volume of 100 ml with distilled water: l9 A B A.0 m1 conc. HCl 2.8 ml 1 N HCl 0.23 ml TEMED O.A5 ml TEMED 36.6 g Tris 5.98 g Tris Final pH 8.9 Final pH 6.7 C D 20.0 g acrylamide 10.0 g acrylamide 80.735 g bisacrylamide 2.5 g bisacrylamide E Catalyst A.0 mg riboflavin 0.1A g ammonium persulfate F A0.0 g sucrose The lower separating gel contained the following proportions of the stock solutions; Parts Stock 1 A 2 C A Catal. 1 H2O These were mixed and 0.7 ml were placed in each 5 x 75 mm electrophoresis tube. The gels were overlaid with 0.1 ml of distilled water and allowed to polymerize for A0 min at 25 C. The upper spacer gel contained the following pro- portions of the stock solutions; 20 Parts Stock .ckamJH ’IJL'IIUCD These were combined and 0.15 ml of the mixture was placed over the polymerized lower gel after the water layer had been removed. The upper gel was photopolymerized under a 1 layer of distilled water with four 15 watt fluorescent lights for 10 min at 25 C. The gels were "pre- electrophoresed"aj 5 c:for 30 min at 2 ma per tube. (2 Protein (5 mg to 100 pg per tube) was introduced in 100 pl of an 8% sucrose solution. Approximately 0.2 ml of a 0.01% brom phenol blue solution in distilled water was mixed into the upper buffer reservoir to serve as an anionic marker. Electrophoresis was carried out at 5 C with a constant current of 1.5 ma/tube until the dye marker had migrated half-way through the upper gel. The current was then increased to 2 ma/tube and maintained at that level for the remainder of the run. The gels were; removed from their respective tubes and a 3 mm length of fine copper wire was inserted horizontally through each gel to mark the position of the dye marker. This was necessary because subsequent procedures led to loss of the marker. Duplicate gels were run for each protein sample. One gel was stained for protein and the other for PNPase activity. 21 The gels to be stained for protein were immersed for 10 min in 1% buffalo black dye in 7% acetic acid. The gels were destained by standing for A8 hr in 7% acetic acid. The gels to be stained for PNPase activity were washed with distilled water, placed in IA x 100 mm test tubes and covered with a solution consisting of 0.3 ml of 50 mM Tris-H01; 0.1 m1 of lOmM inosine; and 0.1 m1 of 50 mM sodium arsenate. This combination was incubated at 37 C for 15 min. The gels were then removed and washed well with distilled water. Each gel was covered with 5 ml of l N NaOH and 1 ml of 0.5% triphenyltetrazolium chloride solution and placed in a boiling water bath for a few minutes until a red precipitate started to form. Then they were washed with distilled water and stored in 7% acetic acid. Kinetic Studies of Vegetative Cell and Spore PNPase A standard Lineweaver-Burk treatment of the Michaelis-Menten equation was used in this study. The Michaelis-Menten equation for a single substrate is: Vmax (S) Km + (S) 22 where v, is the initial velocity; V is the velocity max’ obtained under saturating substrate (S) conditions; and Km, is the Michaelis constant (the concentration of substrate which produces half of the maximal velocity). This equation was rearranged by Lineweaver and Burk (53) to give: 1 K 1 l _ = __ + v V (S) V max max which was plotted as l/v vs. l/(S). The reaction mechanism of PNPase is bimolecular since two substrates were involved (inosine and phos- phate). Actual kinetic analysis indicated that a "ping-pong" or shuttle mechanism (10) was not involved. Cleland (11) has shown that the following rate equation will describe the initial velocity for a bi- reactant, sequential mechanism (where both substrates must be present on the enzyme surface before the reaction occurs) in the presence of all reactants: Vl A B K K + K 1a b b A + Ka B + AB where it is assumed that inosine (A) combines with the enzyme before the phosphate (B) is added. This assump- tion was based on the findings of Kim, et a1. (38) who 23 studied the kinetics of human erythrocyte PNPase and found that inosine did bind first. The terms are defined as follows: A = inosine concentration, Molar. B = phosphate concentration, Molar. Ka = Michaelis constant for A, B saturating. Kb = Michaelis constant for B, A saturating. K1a = dissociation constant for EA complex (E=enzyme). v = initial velocity, units per minute. Vl = maximal velocity, A and B saturating, in the forward direction. Vl/Et = turnover number, units per mg protein. Pure vegetative cell and spore enzymes were assayed at varying concentrations of both substrates. Standard Lineweaver-Burk plots were constructed for each enzyme with either phosphate as the changing fixed substrate and inosine as the variable substrate or with inosine as the changing fixed substrate and phosphate as the variable substrate.l lNotation used by Kim, et a1. (38) where the con- centration of the second substrate was held constant and the concentration of the first substrate was varied to produce one line on the Lineweaver-Burk plot. Another concentration of the second substrate was chosen and held constant while the concentration of the first substrate was varied to produce a different line on the Lineweaver- Burk plot and so on until a family of lines were produced. 2A The family of intercepts from each Lineweaver-Burk plot were replotted against the reciprocal of the chang- ing fixed substrate concentration. When the resulting line was extrapolated to the vertical axis, a value of V1 for that particular Lineweaver-Burk plot was obtained. The reasoning for this procedure was as follows. When kinetic data are plotted on a standard l/v vs. l/substrate plot, the intercept of the line at the vertical axis is the miximal velocity obtainable at saturation of the variable substrate in the presence of the fixed amount of the second substrate used. When the amount of the second substrate is changed, different maximal velocities (intercepts) are obtained. Different maximal velocities will be found until saturating amounts of the second substrate are used, after which the inter- cept will no longer change. The replot of the inter- cepts vs. the reciprocal of the changing fixed substrate when extrapolated to the vertical axis, gives the maximal velocity obtained at saturating concentrations of both inosine and phosphate. The family of slopes from each Lineweaver-Burk plot were replotted against the reciprocal of the chang- ing fixed substrate concentration. When the resulting line was extrapolated to the vertical axis, a value for the Michaelis constant divided by the maximal velocity was obtained. When this value was multiplied by the V1 25 value found from the previous replot, a Michaelis con- stant for the variable substrate being studied was found. The reasoning for this procedure was as follows. When kinetic data are plotted according to the Lineweaver—Burk relationship, the slope of the line is the Michaelis constant for the variable substrate divided by a particular maximal velocity. That maximal velocity represents the value obtained at saturation of the variable substrate in the presence of the fixed amount of the second substrate used. When the amount of the second substrate is changed, different Michaelis constants and maximal veloc— ities will be obtained. The slope will continue to change until saturating amounts of the second substrate are used. In this case, the slope will no longer change since the enzyme is at maximal velocity. The replot of the slopes vs. the reciprocal of the changing fixed sub- strate, when extrapolated to the vertical axis, gives the Michaelis constant divided by the maximal velocity under saturation conditions of both substrates. When this value is multiplied by the maximal velocity found in the presence of saturating substrates (V1) the true Michaelis constant for the variable substrate is found. This constant is independent of the changing fixed sub- strate concentration. An alternative method was also used for finding the Michaelis constant for the variable substrate. The 26 line obtained from assays near saturating levels of the changing fixed substrate was extrapolated to the hori- zontal axis on the Lineweaver—Burk plot. This intercept was equal to the reciprocal of the Michaelis constant. The procedure is less precise than the above method since extrapolation is less accurate at high concen- trations of the changing fixed substrate. Sucrose Density Gradient Centrifugation Analytical grade sucrose was used to make linear 5 to 20% gradients by the method of Martin and Ames (59). The “.55 ml gradients were made at 5 C in 1.27 x 5.08 cm cellulose nitrate tubes (Beckman-Spinco, Palo Alto, Calif.). The gradients were kept at this temper- ature for 3 hr before use. Each gradient was layered with equal parts of lactic acid dehydrogenase (LDH, 0.10 mg/ml) and PNPase (0.25 mg/ml) in a total volume of 100 ml. The gradients were subjected to centrifuga- tion in a Spinco Model L preparative ultracentrifuge (Beckman-Spinco, Palo Alto, Calif.) with a SW-39L rotor at 36,000 rpm for 20 hr at 5 C. Immediately after centrifugation, 37 to 38 3-drop fractions were col- lected from each gradient and assayed for LDH and PNPase activity. The LDH was assayed according to the procedure presented by Worthington Biochemical Corporation (98). 27 Protein Absorption Spectrum of Spore PNPase Protein spectra were measured in the standard buffer with a Shimadzu double beam diffraction grating spectrophotometer (Seisakusho Limited, Kyoto, Japan) incorporating an expanded absorbance scale meter. Absorbance readings weremade in 1 nm wavelength incre- ments from 290 to 3A0 nm with a 1 nm bandwidth. Expanded scale absorbance readings were made in 1 nm increments from 250 to 290 nm with a 0.5 nm bandwidth. Molar absorptivity was calculated using the standard Beer's law relationship (9): A E E = -log10 T = e l c where A, is the absorbance; E, is the extinction; T, is the transmittance; e, is the molar absorptivity; c, is the concentration in moles per liter; and l, is the length of the light path in cm. Subunit Analysis by SDS Polyacrylamide Gel Electrophoresis Subunit analysis of PNPase was performed by a modification of the method reported by Shapiro, et a1. (81). The subunit molecular weights of proteins which have been disaggregated in sodium lauryl sulfate (SDS) are proportional to their migration rates in acrylamide gels. The stock solutions used to make the 5.0% 28 acrylamide gels contained the following components taken to a final volume of 100 ml with distilled water: A 8.0 ml, 1 M sodium phosphate buffer, pH 7.1 0.23 ml TEMED C G 20.0 g acrylamide 0.1“ g ammonium persulfate 0.735 g bisacrylamide The stock solutions were combined in the following pro- portions; Parts Stock FMDRJH :EQOD To each of 12, 5 x 75 mm tubes was added 1.5 ml of gel mixture. The gel was overlaid with distilled water and polymerized at 25 C for 30 min. The electrophoresis buffer contained 0.1% SDS in 100 mM sodium phosphate buffer, pH 7.0. The gels were "pre-electrophoresed" at 7 ma per tube for 30 min at 25 C. The protein dissociation mixture contained equal proportions of the following components which were com- bined just before use: 10% SDS in distilled water; 29 10% (v/v) 2—mercaptoethanol in distilled water; and 100 mM sodium phosphate buffer, pH 6.8. The marker mixture contained 0.01% brom phenol blue, 10 mM dithiothreitol, and 50% (v/v) glycerol in 10 mM sodium phosphate buffer, pH 7.0. DNase I, RNase, and bovine serum albumin (1 mg/ml stock solutions) were used as subunit molecular weight markers. Their molecular weights after dissociation in SDS were 31,000 (52), 13,700 (96, 99), and 68,000 (87, 91) respectively. The protein stain was made by dissolving 250 mg of coomassie brilliant blue in 10 ml of distilled water. To this solution was added 5 ml of methanol and enough 10% trichloroacetic acid (TCA) in distilled water to bring the total volume to 100 ml. The solution was mixed well and kept overnight. Just before use, the stain was filtered through Whatman #1 filter paper. After the gels had been fixed in 10% TCA, they were placed in l“ x 100 mm tubes and covered with the stain solution. The staining procedure took 10 hr at 25 C, after which time the gels were destained by standing in 10% TCA for H8 hr. High Speed Sedimentation Equilibrium Molecular Weight Study A Spinco analytical Model E ultracentrifuge (Beckman— Spinco, Palo Alto, Calif.) equipped with phase-plate interference optics was used for this study. An AN—D rotor was used with standard double sector cells whose 30 solution column depths were 3 mm (110 pl of sample per cell column). Techniques were available to allow multicell operation with interference optics. The PNPase samples were subjected to centrifugation at 17,100 rpm for 2“ hr at 9.5 C. Simultaneous photographs of the interference patterns for each cell were taken. The fringe patterns, which are a measure of protein concentration differences within each cell, were measured with a Bausch and Lomb two-dimensional comparator. The photographic plate was aligned on the comparator X— coordinate and the displacement along the most defined fringe was measured along the Y—coordinate. Measurements were taken at intervals of 50 u on the X-coordinate. A partial specific volume of 0.717 ml/gm, obtained from an amino acid analysis of vegetative cell PNPase (16), was assigned for both the vegetative cell and spore PNPase enzymes. A pycnometer (29) was used to determine the density of the standard buffer (1.00022 g/cc) and that of the standard buffer plus 10 mM potassium phosphate (1.00163 g/cc) at 20 C. Complete statistical analysis of the data by the method of thantis (100) was performed using a computer program which was adapted to a Control Data Corporation 3600 digital computer by workers in the Department of Biochemistry, Michigan State University, East Lansing, Michigan. A discussion of this analysis is presented in 31 Appendix B. The data obtained from the computer printout are summarized below: Whole-cell average, weight-average molecular weight. Number-average molecular weight at zero concentra- tion (Mn), from the extrapolated slope of a molecular weight vs. protein concentration plot. Weight-average molecular weight at zero concentra- tion (Mw), from the extrapolated slope of a molecular weight vs. protein concentration plot. Plots of the weight-average molecular weight vs. protein concentration. Plots of the number-average molecular weight vs. protein concentration. Plots of the In of fringe displacement (protein concentration) vs. the squared distance from the center of rotation. All of these data were statistically analyzed and pre- sented with accompanying confidence intervals. The fundamental relationship between the molecular weight and sedimentation coefficient for a spherical protein under ideal conditions can be solved. By assign- ing the value 0.717 ml/gm for the partial specific volume of PNPase, the following equation yields the sedimentation coefficient for a hypothetical spherical PNPase molecule with the experimentally determined molecular weight: 2/3 3° = 3.79 x 10'3 M where 80, is the sedimentation coefficient in Svedberg units; and M, is the molecular weight (Appendix C). With this equation and the whole cell weight-average 32 molecular weight values obtained from the high speed sedimentation equilibrium study, sedimentation coef— ficients were calculated for both the vegetative cell and spore PNPase enzymes in the presence or absence of 10 mM potassium phosphate. Using equation I from Appendix C, minimal frictional coefficients (f0) were calculated from the molecular weight data and calculated S0 values. The actual fric- tional coefficients (f) were also calculated from the molecular weight data and the experimentally obtained sedimentation data. From the f/fO ratios an indication of the molecular asymmetry could be determined. A per- fectly spherical molecule would have a f/fO ratio of 1.0. Globular proteins have ratios in the range of 1.1 to 2.0. RESULTS Purification of Purine Nucleoside Phosphorylase The PNPase from vegetative cells and spores of B. cereus T was purified so that each enzyme could be studied in the absence of uncontrolled interactions between it and other cell components. The purification procedure was a modification of those used previously (16, 20). A11 purification steps were carried out at 5 C. Centrifugation was at 20,000.x g for 60 min. Both the vegetative cell and spore PNPase enzymes were purified by the same procedures except that the spores were heat treated before they were broken. A11 dialysis tubing was boiled 1 min in 10 uM ethylenediaminetetraacetate (EDTA) to remove trace metals before use. Preparation of Crude Extract Vegetative cells (500 g, wet weight) were suspended in 1.2 liters of the standard buffer and washed into an Eppenbach colloid mill. To the mill were added 1 kg of #110 glass beads and 3 m1 of antifoam B. The cells were ground at full speed for NO min at 10 C. This procedure resulted in over 80% cell breakage as judged by phase contrast microscopy. The mixture was removed from the 33 3“ mill and the extract was separated from the beads. The cell extract and 300 m1 of buffer used to wash the glass beads were then clarified by centrifugation. The super- nate was filtered through cheesecloth and assayed. This was designated fraction I. Washed spores (360 g, wet weight) were suspended in 800 m1 of the standard buffer and heated to 70 C for 30 min to inactivate any contaminating vegetative cell enzyme. The spores remained refractile under phase contrast microscopy after this heat treatment. The spores were quickly cooled in a -20 C alcohol bath and washed into the colloid mill along with an additional 400 ml of buffer. The spores were broken at full speed in the presence of 1 kg of #110 glass beads and 3 ml of antifoam B for 55 min at 10 C. Less than 15% of the cells remained intact after this procedure. The spore extract-was combined with 200 m1 of buffer used to wash the beads and clarified by centrifugation. The super- natant fluid was filtered through cheesecloth and assayed as spore PNPase, fraction I. Approximately U-times more enzyme was recovered from vegetative cells than from spores, based on equal starting wet weights of cells and spores. Streptomycin Sulfate Precipitation Nucleic acids were partially removed from the crude extracts by slowly adding 3.3 g of streptomycin sulfate 35 for each liter of extract. The suspensions were stirred for an additional 60 min and then allowed to settle at 5 C for 12 hr. The precipitate was removed by centri- fugation and the supernate containing the PNPase activity was filtered through cheesecloth to remove the lipid- 1ike material which did not sediment. Ammonium Sulfate Fractionation of Crude Extract The enzyme solution from the streptomycin step was taken to 50% saturation by slowly adding solid ammonium sulfate (288 g/liter) with stirring in an ice bath. The solution was stirred for an additional 60 min and then clarified by centrifugation.) The precipitated protein contained very little PNPase activity after dialysis and was discarded. The supernatant was taken to 80% satura- tion with an additional 197 g/liter of solid ammonium sulfate, stirred for 60 min, and'the protein precipitate was removed_by centrifugation. The pellet containing the PNPase activity was resuspended in the standard buffer and dialyzed against u liters of the same buffer for 16 hr. The enzyme was stored at -15 C with no appreciable loss of activity. This preparation was designated fraction II. DEAR-cellulose Column Chromatography After several unproductive purification attempts were made using gel filtration methods, it was discovered 36 that DEAE—cellulose chromatography was a more efficient process. Various combinations of pH and ionic strength were tested but the following procedure was the most effective. Spore PNPase fraction II was thawed and 67 ml (837.5 mg of protein) were placed on a DEAE-cellulose column (3.5 x 100 cm) which had been equilibrated with 2 liters of 2ND mM Tris-HCl, 10 mM 2—mercaptoethanol buffer, pH 7.5. The protein was eluted from the column with 2.5 liters of the above buffer and the column effluent was collected in 21 m1 fractions (25 min per fraction). This zero-gradient elution removed most of the protein from the column. A linear gradient was then started consisting of 1.5 liters of 290 mM Tris-HCl, 10 mM 2- mercaptoethanol buffer, pH 7.5, and 1.5 liters of 330 mM Tris-HCl, 10 mM 2-mercaptoethanol buffer, pH 7.5. The PNPase activity eluted after 1.6 liters of eluent had passed through the column. This corresponded to elution in 290 mM Tris. Ninety percent of the enzyme placed on the column was recovered with a 21—fold increase in specific activity for the peak fractions. This prep- aration was called spore fraction III. The vegetative cell PNPase fraction II was thawed and 110 m1 (1.375 mg of protein) were placed on a 3.5 x 100 cm DEAE-cellulose column and eluted with 2ND mM Tris-H01, 10 mM 2—mercaptoethanol buffer, pH 7.5. 37 Eighteen—milliliter fractions were collected (30 min per fraction). After 3.6 liters of buffer had passed through the column, a 3 liter linear gradient was started as before but no further enzyme activity was eluted off. The recovery was 87% of the enzyme activity. A 6-fold increase in specific activity was obtained for the peak fractions after they were pooled and concentrated. This preparation was designated fraction III. Vegetative cell PNPase differed from the spore enzyme by eluting at lower Tris concentrations. This may have been due to a difference in size and/or charge between the two enzymes. Preparative Disc-gel Electrophoresis The preparative disc-gel method had been previously used in this laboratory to purify PNPase (16). The total recovery of enzyme from this procedure was usually only 50% when a modification of the gel system reported by Jovin, et al. (31) was used. Further modifications of the separating gels during this investigation resulted in yields of up to 90%. Spore PNPase fraction III was made 8% (w/v) with respect to sucrose and 100 pl of a 0.01% solution of brom phenol blue in distilled water was added for anionic marker. The mixture was layered over the preparative polyacrylamide gel and a constant current of 15 ma was 38 applied to the column. The ammonium persulfate was eluted from the column with buffer at 0.25 ml/min. When the dye marker had migrated half-way through the upper stacking gel, the current was increased to 20 ma and kept at this level for the remainder of the run. As the marker entered the lower separating gel, the elution rate was increased to 0.5 ml/min. When the marker had moved to the lower extremity of the column, the elution rate was increased to 1.5 ml/min and fractions were collected every 10 min. The fraction containing the marker was designated tube 1. As shown in Figure 1, the PNPase activity came off the column with very little accompanying 280 nm absorbing material. Tubes 32 through 35 were pooled and concentrated. Eighty percent of the enzyme placed on the column was recovered with a 13-fold increase in specific activity for the peak fractions. Fraction III of the vegetative cell enzyme was placed on a similar electrophoresis column and run under identical conditions. The final elution rate was 1.2 ml/min. As shown in Figure 2, the activity was eluted from the column without a measurable 280 nm absorbing peak. Tubes 28 and 29 were combined and concentrated by ultrafiltration. Over a 90% recovery of enzyme activity was found with a 38-fold increase in specific activity for the peak fractions. a n .. 39 Figure 1. Elution profile of spore purine nucleo- side phosphorylase from the preparative disc-gel electro- phoresis column. Current was maintained at 20 ma. The elution rate was 1.5 ml/min and the fraction volumes were 15 ml. The ordinates are activity (u=units ojenyyme) and absorbance. See text for experimental details. 140 Dmmommbzom “N000 :3. 94‘\‘\.‘\6 I 2.5.2. 3 >t>io< .85.: 8 2 0 0 .l O 0. 0 0. l q d d d - d - I .l\\hv ‘\~\~\~\~\‘\‘\‘\~\‘\‘\ - [IR - b! - L b b n n n n b O O m 0 O O O c 3 2 I 30 20 IO TUBE NUMBER w—fiT U1 Figure 2. Elution profile of vegetative cell purine nucleoside phosphorylase from the preparative disc-gel electrophoresis column. Current was main- tained at 20 ma. The elution rate was 1.2 ml/min and the fraction volumes were 12 ml. The ordinates are activity (u=units of enzyme) and absorbance. Experi- mental details are given in the text. 42 bmmommbznm ANQO 33v Q\~\‘\a 8 6 O «4‘ 4 2.0 ~ 0.0 «0.0 40.04 “ 0.02 .0 1“ d d 141 50 I 2.5:... 3 C53: 352.. TUBE NUMBER “3 The protein concentrations for all of the pure PNPase preparations were below the measurable range of assay by the Lowry method. The enzymes were concentrated by putting them into small dialysis bags, covering them with dry Sephadex G-25 and dehydrating for 12 hr at 5 C. The dialysis bags containing the concentrated PNPases were then dialyzed against the standard buffer and stored at -15 C. These preparations were called fractions IV. The rapid migration of this enzyme under the con- ditions used in this procedure made the purification of relatively large quantities of enzyme possible. A sum- mary of the overall purification procedures for the vegetative cell and spore PNPases are presented in Tables 1 and 2 respectively. Thermal Inactivation of Purine Nucleoside Phosphorylase A previous study indicated that the spore PNPase was stable but became thermal labile in the presence of phosphate ion (16). An effort was made to determine the basis for the effect of phosphate on heat resistance. Two—milliliter samples of spore and vegetative cell PNPase from the DEAE-cellulose column (fractions III) were each dialyzed for 24 hr against 200 m1 of the standard buffer at 5 C. An enzyme solution (200 pl) was diluted with 200 pl of the standard buffer and placed in a 14 x 100 mm test tube. After a zero time assay sample UM .uxmp map 2H cmppoamp one mafimpme Hmpcmsammmxm .HH coapomnm mo puma mace pom: mca>m£ mom empompnoo .ompsmmmpoea mam macapomam xmma cmaooa one masom mm oma.fi o.oom.m ooa.mm mmfimmnonaonuomam Hewlomfio .thm .>H m: on o.mmm oo:.ms messaoo mmoasaamoumEouQmppm .HH ooa . H :.e ooo.HsH pomspxm mespo .H m Ufiom mE\mpHc5 mufics spa>fipo< xsm>oomm soapmoHMHhsm muH>Huw< mEmNcm coapompm oamaomam Hence .mwmamponomogu eeflmomflozc mgapsa flame m>fiumummm> mo coapmofihansmuu.a mamm£ pom wouomppoo .Umpcmmmndmn mam mCOfipompm xmma cmfioon on» macom em osm.© o.ose.e oo:.HH «mammnonaoppomam Howaomfic .Qmpm .>H mm mfim o.osm ooa.na maesHoo mmoasaflmoum¢mm .HHH am am m.©m oom.em emusamae .Aaownomv mumuHSm Seacoasm + mummasm. CHo>EOpamppm .HH OOH H H.H oo=.om pomppxm mouse .H R each mE\mpHss means. spa>fipo< msm>oomm coaumoHMHssm zpfi>fipo< mezncm coapompm oauaomam Halos .mmmapo£QmonQ mofimomaosc mcHMSQ macaw mo coaumofihapsmll.m mqm._..>:.o< PZMUIMQ 20‘- “9 Samples of vegetative cell and spore PNPase (frac- tions I) were then dialyzed in the standard buffer con- taining 10 mM CaCl The half—lives of the vegetative 2. cell and spore enzymes were 10 min and 3.0 min respec- tively. The calcium ion labilized the vegetative cell PNPase and appeared to stabilize the spore enzyme. When 20 mM potassium phosphate was added to these preparations, the vegetative cell PNPase increased to a half-life of 30 min and the spore PNPase increased to a half-life of 10 min. Therefore, the addition of phosphate produced an increase in heat stability for both enzyme prepara— tions in either the presence or absence of calcium ion. Analytical Polyacrylamide Discegel Electrophoresis of PNPase Samples of purified vegetative cell and spore PNPase (fractions IV) were checked for electrophoretic homogeneity using the analytical disc—gel electro- phoresis technique. Both the vegetative cell and spore PNPase enzymes from the preparative polyacrylamide electrophoresis columns behaved in an identical manner. A single protein-staining band which migrated with the PNPase activity was found for either enzyme indicating that each preparation was electrophoretically homo- geneous. The mobilities of the vegetative cell and spore enzymes were similar. The results of a typical run are shown in Figure u. 50 , E E i {f I V K: r , 5? 'Wi Figure 4.--Analytica1 polyacrylamide disc—gel electrophoresis of spore PNPase. Tube #1, was stained for PNPase activity. Tube #2, was stained for protein. The control tube (C), contained no protein and was a control on marker migration. A control gel without protein (not shown) did not produce a band when stained for PNPase activity. See text for the experimental procedures. 51 Ultraviolet Absorption Spectrum of Pure Spore PNPase An absorption spectrum of pure spore PNPase was measured to determine whether the protein contained bound cofactors, unique amino acid structures (viz. thiazoline rings), or bound inosine which might be detected by this procedure. Two concentrations of pure spore PNPase (1&2 and 700 ug/ml) in the standard buffer were measured for absorption in the ultraviolet region of the spectrum. A molar absorptivity of 4.7 x 10“ liter M.1 cm-1 was calculated from the maximum absorption peak at 279 pm. No unusual absorption peaks were found (Figure 5). Effective Assay Range for PNPase Activity The effective assay range was determined so that subsequent initial velocity studies would be a true reflection of the kinetics involved in the enzyme reaction mechanism. A series of ten—fold dilutions of pure spore and vegetative cell PNPase were made in the standard buffer. Aliquots (100 ul) were assayed in the standard assay system at 37 C. The optimum enzyme con- tent per assay for both enzymes ranged from 0.2 to 2.0 U8 0f protein and no lag in initial velocity was noted. 52 Figure 5. Ultraviolet absorption spectrum of pure spore PNPase in the standard buffer where the absorbance is plotted vs. wave length. Also included is an expanded scale plot over the absorbance range of 2H — 38 x 10 -3, See text for the experimental details. 53 EXP. SCALE ABS. (xno3) .--.. QCDQ'NOCD‘D'Q' ”MMMMNNN ' ' rIIfi FTII 09" 0.8»— 0.!— 0.2 ~— 1 J l l N (D ID d; «3 o’ 0' o' o o’ --- BONVBHOSSV 280 300 320 340 WAVE LENGTH (nm) 260 240 Subunit Activity of Spore PNPase A previous study indicated that in the absence of phosphate ion (16) the spore PNPase was smaller than the vegetative cell enzyme. An effort was made to deter— mine whether a concentration—dependent aggregation of protein was a necessary prerequisite for spore PNPase catalytic activity. Pure spore PNPase (fraction IV) was assayed at various protein concentrations in the standard assay system. The calculated specific activity was then plot- ted against the protein concentration used for each assay. As shown by Figure 6, the slope was essentially zero over a broad concentration range. Initial Velocity Analysis of Vegetative Cell and Spore PNPase Kim, et a1. (37, 38) studied initial velocities and reported unusual kinetics for the PNPase from human erythrocytes. A previous study of vegetative cell and spore PNPase in this laboratory also indicated non- linear Lineweaver—Burk plots (16). The kinetics of both enzymes were therefore studied at various concen- trations of each substrate in order to ascertain the extent of the catalytic differences between the two' proteins. The pure vegetative cell and spore enzymes (frac- tions IV) were diluted in the standard buffer to 0.95 F). 5, .mafimpmn Log uxmp mom .mmmmm some pom pom: coapmppcmocoo samuopd mcp pmcfimwm Umuuoaa was Ampficsusv zufi>fipom camaomdm UmumHSOHmo on» new Emummm memmm osmocmpm one CH mCOHmep ucmocoo Camposd msofipm> um pmmmmmm was A>H :ofipomsmv mEmNcm Umfimfissm .ommmZm whoam mo mufi>apom pacsosm .m mpswfim 56 as nose Emma mum .ozoo zmfioma mv N¢ mm on ¢N m. N. m 0 _ |4 q i A 4 1— _ O. (Ow/n €01 X) '10V 'leBdS 57 and 1.15 ug/ml respectively. All dilutions of the phos- phate buffer and inosine stock solutions were made in the standard buffer. All kinetic data were collected using the same enzyme dilutions and reagents on the same day in order to minimize variations in the results. All assays were performed at 37 C after the assay mixtures had been equilibrated for 10 min in the cuvette changer. The reactions were started by adding the enzyme. With phosphate as the changing fixed substrate and inosine as the variable substrate, the Lineweaver- Burk plots (Figures 7 and 8) were linear for both the vegetative cell and spore enzymes. Since the plots intersected to the left of the vertical axis, a sequential reaction mechanism was indicated (11). Using the termi- nology of Cleland (10) for a bisubstrate reaction, the average Michaelis constants for inosine (Ka) were “.6 x -N M and 7.0 x 10"Ll M for the vegetative cell and spore 10 PNPase enzymes respectively. There was a 1.5-fold dif- ference in the Ka values for the two enzymes. With inosine as the changing fixed substrate and phosphate as the variable substrate, the Lineweaver- Burk plots (Figures 9 and 10) showed a downward curvature at low inosine concentrations. Kim, et a1. (37, 38) described a similar downward curvature for the PNPase from human erythrocytes. When the intercepts were replot- ted vs. the reciprocal of the inosine concentration, the .pxmu one Ca emppoams mam mHHmpoU Hmpcmefismdxm .2 tea x m.: .mm "ma Apmncfiv mpoadms cam poaa man» Eopm Umpmgfipwe mmpmsmnmq oameHx one .mCOprchmosoo mumnamoca mo mwcmu m nm>o mCOHpmnucmocoo mnemocfi no Hmoomofiomp on» .m> mafiooam> mo Hmoohaaomn mo poam .mpmnpmnsm magmapm> on» we enamosfi nuaz mmmmzm flame e>fipmpmwm> mo mufiooam> HmeHCH .n madman 58 llufl‘lvlllll.ll )illall. A29; wz.moz.\_ 59 o. m o c u o ~n . . . . , V. . t8 ..\\\\ \\ on .2... upsamozaz. 3. 3 o.~ 0.. 0.. so to too .IA a . q o 18 m.— i Lou. 18. on. Lee 1 >2 12~ Loom o I N 12h 0...... a noon 8% d to :03 to»... :2 mpo mcowpmhpcmocoo enamocfi mo Hmoosdaomp on» .m> zpfiooam> mo Hmoopofioep mo poam .mumpmeSm efinmfism> one we mnemocfi cues mwmmzm macaw mo mpaooam> HmeHCH .m esswwm b O 61 .25. wz.moz_: o. o m c u o N- _ . H 1d i - .1 a \|\ V) LON. 100. low. >2 JO.N L OVN 1 CNN ‘0 L Don L Onn . SE wh<3mmozm b p P P - 2E whdimmoxax. n.~ o.N 0.. 0.. n6 I o 1 l o o N - 1d30831Nl L O '0 .mossveoopd Hmpcmefisedxm pom uxmp exp mom .2 mIoH x m.H .nx ”ma Apmmcfiv wuofiamm ecm head wasp Eopm ompmsfipme pmpmemsma ,h ofipmcfix one .mcoapmpuceocoo mcfimosa mo mwcmp m pm>o mCoau Imspcmocoo mpm£QmonQ mo Hmoopafioma ecu .m> mpfiooam> mo Hmoos Iofioms mo pon .mpmppmndm memHsm> one mm mumndmond aha: mmmmzm flame m>Hpmpmwe> mo mpfiooam> HmHuHcH .m mpzwfim 63 O 4113.; O ..o :5 .uzaoz. _ . N .25. mhdxaworat 0.. N.. _ . 1.1.11! 0.0 J ¢.o q \‘Ilu O ‘1lll .‘w o m cm Jom ION. 100. 1.00. >2 IO_N IO¢N LOhN J Con i onn ¢.OI 0.00 «.7. q a SE wz.moz.2 0.0wVN { — q d 1 d OO O 0 IO N 1.630831!!! 0 V 6U .fll‘l lilqlllll ‘ In‘I . .pxmp we» Ca omppoaop mam mHHmumU Hmpceefipmdxm .2 mloa x m.H «ox umH Apmmcfiv muoaqmp new poaa was» Eopu nonmefiumm smpmewsmm ofiumcfix one .mCOHumsucmocoo mcfimoca mo mwcms m pe>o macapmnpcmocoo mumsamona mo HmOOAQHomp one .m> muaooam> no Hmoosafiomp mo uoam .mpmhmeSm manmfipm> on» we mpmcdmonq Spas mwmmzm macaw mo zpfiooam> HmeHsH .oa mhsmfim 65 SE. mh\. IO_N IO¢N lohN 1 con 1 on» v.0..- O. ’ SE mz.moz.\. t N # 66 results were not linear. This may have been due to either subunit interaction (30) or to negative cooperative effects between the two substrates (15). The Michaelis constants for phosphate (Kb) were 1.5 x 10-3 M and 1.3 x 10_3 M for the vegetative cell and spore PNPases respec- tively. The turnover numbers (VI/Et) for the vegetative cell and spore enzymes at saturating levels of both substrates were 8,320 and 8,690 units/mg protein respec— tively. A summary of the kinetic data is presented in Table 3. Subunit Size of Purine Nucleoside Phosphorylase Although cell and spore PNPases appear to be synthesized under the direction of the same genomic unit (20), a previous study indicated that the ratio between their molecular weights was unusual (16). This phenomenon could have been the result of a pro- teolytic conversion of vegetative cell to spore PNPase where a portion of the vegetative enzyme was lost. A precedent for this mechanism existed in the conversion of vegetative to spore aldolase in B. cereus (77). This kind of proteolysis might lead to an unequal modification of subunits. Therefore, an investigation was made of the vegetative cell and spore enzyme sub- unit sizes. 67 TABLE 3.--Michaelis constants of the substrates for pure vegetative cell and spore PNPases. a Parameter Spore PNPase Veg. PNPase -u -A Inosine, Ka 7.0 x 10 M 9.6 x 10 M Phosphate, Kb 1.3 x 10’3M 1.5 x 10‘3M Turnover numberb 8,690 units/mg 8,320 units/mg aAverage Michaelis constant values are given from the replots of the slopes and intercepts of the Lineweaver-Burk plots (Figures 7-10) and from the direct plots. bThe turnover numbers were calculated from (Vl/Et); where Et, was the concentration of enzyme (mg) in the assay and V1, was the units of enzyme activity/min in the assay. Experimental details are given in the text. 68 Duplicate samples of pure vegetative cell PNPase (100 pl, 0.38 mg/ml), spore PNPase (100 pl, 0.U6 mg/ml), and three marker proteins (100 pl, 1.0 mg/ml) were each added to separate 6 x 50 mm tubes. To each tube was added 50 ul of the dissociation mixture and the samples were incubated for 3 hr at 37 C. A marker-glycerol mixture (20 ul) was added to each tube and 10 to 30 ug of protein was introduced onto the top of polyacrylamide gels. The proteins were "electrophoresed" at 7 ma/tube for 30 min at 25 C. The gels were fixed in 10% TCA, stained and decolorized. The protein band migration distance was measured from the top of each gel. A plot of the log of molecular weight vs. migration distance was linear (Figure 11) and the subunit size for both the vegetative cell and spore PNPases was 2u,ooo i10%. This experiment was repeated with a current of u ma/ tube and the same results were obtained. Sucrose Density Gradient Sedimentation of PNPase The sedimentation properties of spore PNPase are affected by phosphate ion (16). This behavior was investigated at varying phosphate concentrations to determine whether subunit association could be detected at high phosphate concentrations. It was also of interest to determine whether the phosphate effect was, reversible. .uxmp one CH emucmmmsa one messomoopm HmucmEHpmdxm .mmmmaosconfih use .H mmeefloscopfismxoee .cfissnfim Essen mcH>0b who: poms mcfimPOLQ sesame smash mze .mflmmsosoospooae How mefismfizpomzaoalmmm an mmmmmzm whoam use flame m>Humpwmm> mo mcamco mUHmeQ>H0Q one no pcwflmz smflsomaos exp mo coapmcfiesmpmo .HH mssmfim 6 9 ‘4 )1 4!. an 70 EB. >._._.:mo.2 m m e. m . o 3 _ q a O. W O _| 3 3 n w Lemma M B -Omm II.— .on 0. look». .00 10h . F . _ 71 Pure spore and vegetative enzymes (fractions IV) wmecflhmed in standard buffer to a concentration of 0.25 mg/ml and dialyzed for 214 hr at 5 C. Each sucrose gmflhmtwas layered with 100 pl of enzyme sclution containing 125 ug of either vegetative cell or spore The gradients PNPase combined with 50 pg of LDH marker. In the stand- vmrespmiat 36,000 rpm for 20 hr at 5 C. ard buffer system the sedimentation velocities for the \mgeuudve cell and spore enzymes were 5.5 S and 5.3 S _+_ 0.1 8 respectively. The sedimentation coefficients of UnePNPase preparations were calculated by assigning a sedimentation value of 7.6 S to the LDH marker (36, 62). Studies using LDH in sucrose density gradients employing buffer systems similar to those used in this investigation indicated that its sedimentation coeffi- cient did not change in the presence of phosphate ion (George M. Stancel, Department of Biochemistry, Michigan State University, East Lansing, Michigan; personal com- munication). The peak PNPase activity fractions were pooled, concentrated and dialyzed for 18 hr at 5 C against the standard buffer containing 1 mM KH2POA’ at pH 7.5. The enzyme solutions were mixed with the marker enzyme, layered on sucrose gradients prepared in the same buffer as before. The vegetative cell system and sedimented enzyme had a sedimentation coefficient of 5.5 S and the 72 spore PNPase had a value of 5.“ S i 0.1 S. The vegetative cell enzyme sedimentation velocity did not change but the value for the spore enzyme increased (Table A). In another study, pure vegetative cell and spore PNPase solutions were diluted to give a protein concen— tration of 0.25 mg/ml. Three aliquots of each enzyme were then dialyzed against the standard buffer for 24 hr at 5 C. Two aliquots of each enzyme were then dialyzed against the above buffer containing 10 mM KH2POu, pH 7.5 for 2“ hr. At this time, one aliquot of each enzyme was returned to the standard Tris buffer, while the other aliquot remained in the Tris + phosphate buffer. Dial- ysis was continued for an additional ua hr with several changes of buffer. The enzymes were then sedimented through sucrose gradients containing the same buffer in which they were finally dialyzed. The results are tab— ulated in Table 9. A study was made to determine whether there was a protein concentration-dependent change in sedimentation velocity for the spore PNPase in the standard buffer. A pure spore PNPase preparation (fraction IV) was dialyzed against the standard buffer and three different concen— trations (“0 pg, 150 ug, and 230 ug) were each mixed with the LDH marker and layered on separate sucrose gradients. The enzymes were sedimented through the gradients at 36,000 rpm for 2H hr at 5 C. The PNPase activities 73 TABLE “.—-Sucrose density gradient centrifugation of pure vegetative cell and spore PNPases. Sedimentation coefficients Buffera Spore PNPase Veg. PNPase S20,w 10.1 s $20,WIO.1 5 Standard buffer 5.3 5.5 Standard buffer + 1 mM Phos. 5.“ 5.5 Standard buffer + 10 mM phos. then standard buffer 5.6 5.5 Standard buffer + 10 mM Phos. 5.7 5.5 3The standard buffer contained 50 mM Tris-HCl, 10 mM 2—mercaptoethanol, pH 7.5. The other buffers had a pH of 7.5. See text for the experimental procedures. 7“ sedhmnmed at the same rate relative to the LDH marker regamfless of the PNPase protein concentration. The calcuhned sedimentation velocities ranged between 5.3 S and 5JJS i 0.1 S. Therefore, no protein aggregation ommuwed within the range of protein concentrations ltested. High Speed Sedimentation_Equilibrium Molecular Weights Accurate molecular weight determinations for both the vegetative cell and spore enzymes were needed to exploit the findings of previous investigators (16, 20). These data could then be used to calculate frictional coefficients, substantiate the homogeneity of the enzyme preparations, and to demonstrate subunit association in the presence of phosphate. Both the vegetative cell and spore enzymes (frac- tions IV) were dialyzed and subjected to centrifugation in the standard Tris buffer or in that buffer containing 10 mM potassium phosphate. A previous sedimentation veltuxity study in the Model E ultracentrifuge indicated that tflue spore enzyme did not undergo aggregation in tflue'rris buffer system. The enzymes were placed in separate double sector cells and spun at 17,100 rpm for Simultaneous photographs of the inter- 2“ hr at “.5 C. All of ference patterns for each cell were then taken. tflqe nmflhecular weight data were collected from the same 75 experiment so that external variables were more con- trolhaL The results given in Table 5 include a complete sfififlstical analysis at the 99% confidence level. The plots of the natural logarithm of the fringe chsplacement (In D) vs. the square of the distance from the center of rotation (r2) were linear except near the top of the sector cells containing the spore enzyme (Figures 12 and 13). The enzyme preparations appeared to be homogeneous. The limiting slopes near the top and bottom of the sector cells as well as the overall slopes are tabulated in Table 5. The size of the vegetative cell PNPase did not change in the presence or absence of 10 mM phosphate. The spore enzyme was smaller in the Tris buffer and appeared to increase to a size similar to that of the vegetative cell enzyme in the presence of phosphate. The f/fO ratios (for calculations see Appendix C) indicated that the spore enzyme was quite spherical in Tris buffer and then assumed the shape of a more globular protein in the presence of phosphate (Table 6). This indicated that the molecular weight increase of the spore PNPase in the presence of phosphate ion probably prmxhiced a.change in shape as well as a change in size. 76 .m.s mo .opmzomoso ESHmmwp0d SE 0H .Homlmfise SE om oochucoo pseudo A.mosm + mHHBV mg» one mm.> mo .Hocmnpmoudmopmfilm .pxou one CH so>Hw mCOHpHosoo one soon: omHUSum one omswompo who: mmszuco ommmzm macaw one flame o>Hpmpowm> ousmm :5 OH .Homumase 2s om eoeamscoo seamen Amuse. was .Hocmnpmoudmoposlm :8 OH ems.H Hme.fi ems.a com H ooe.mm ooo.m H oom.mm 00: H oom.mm .monm + muse :mm.H smm.H :mm.H oom H oom..m ooo.m H ooa.:a so: H oom.aa mane ommmzm .mo> osm.fi mme.fi swo.m coo H oom.sm oom.m H ooo.moa cow H oom.sm .moem + muse oem.H aes.H mmm.H oom H oom.mm oom.m H oom..e com H 000.00 mate ommmzm maoom Hammo>o soupom QOB o u o .pszms o u o .uszez Hmmmwwmwe amazomaoe amazomaoe mwmho>m mommsolmezmsm . . m I m I ma m> Q Ca mo mooam o mhm>m hmnssz m mhm>m panmS Hamelwaon3 .EsHmoHHHsom am macaw ecm HHoo o>vapomo> Ho mpanmz amazomaos II.m mqm a na no mpon .mH onsmHm 78 N2... 8 ~ L 0.00 0.00 - W ' o\ 0.? 0.? (ma 0| ¢.N 0.N 79 .pxep en» nH oepnmmeno ene mHHepeo HepneEHneoxm 4m xHonoQa¢ mom .. OIIIIIIIIIOV epenowono EsHmmepoo SE 0H + nommsn oneonepm nH eEmNnm .A0 0. nemmsn oneonepm nH mazenm .ememzm flame e>Hpepewe> pom mm .m> O CH mo mpOHm .mH endem In 15 H. .E .20. «n 80 4 5 _ d a v.2. «do ado 0.3. o... 1 1 4 L 0.0 0.0 0.0 O. n 0.0 0.0 N.» 0.» Ci 0.“ 0.0 NJ IIIGU ( 81 HepneEHpmdxe pom axon mom .mHHepmo .Am xHoneQQen oasoz uanes man mo nHepond e noan uneHoHomeoo nOHuepnmEHoew enp no oomen .panoz seasoefioe ewenm>e Haeoleaonz one Eonm oopeHSOHeo mes om Lou esae> one o .m.s mm amuendmond EsHmmeQOQ SE 0H mead mpnenOQEoo o>one enp oenHepnoo nemmsn m as OH + mHne .m.e mo .HocenooOpaeonmsum as OH .Homumane 2s om emeaeocoo panama wanes :.H m.m oom.me e 2e OH + wane :.H m.m ooe.Ha mane . mmemzm e>Hpepewe> :.H e.m oom.sm a as OH + mane m.H m.m oom.me mane ememzm en>qm onmamz o . H.0H3.o neHSOmHoz - . . m SJI. n\o HHmoanonz a; on I essaem .mem mHOMm wHHmo Hoeoome> no mOHpen Henofipoan -l sewaHaOHae--.m mq_ae (I. DISCUSSION The cell and spore PNPases had been purified by slightly different procedures in previous studies (16, 20). In this investigation both enzymes were purified by the same procedure in order to minimize any effects which might be due to methods of enzyme preparation. The only significant difference in the purification of the two enzymes was the heat treatment of the spores before they were broken. It was assumed that the PNPase protein within the Spores would be protected from thermal denaturation whereas the vegetative cell PNPase would be inactivated. The heat treatment did not seem to initiate germination since no loss in refractility was detected when the spores were examined by phase microscopy. The vegetative cell-derived crude extract had approximately “-times more PNPase activity than the spore extract, based on equal starting weights. 'A preliminary study indicated that the level of PNPase activity increased in vegetative cells toward the end of exponential growth and remained at this level in the spores. The lower yield of spore PNPase activity found in this investigation may be the result of protein denaturation during the mechanical breaking of the 82 83 spores. Spores are particularly resistant to rupture and require a more prolonged grinding than do vegetative cells. Alternatively, the heating may have denatured some of the spore enzyme. Engelbrecht and Sadoff (16) reported that the spore enzyme was more stable than the vegetative enzyme n in the absence of phosphate and possessed the same low thermal stability as the vegetative enzyme in the pres- ence of phosphate. This was not found with the enzyme 1 preparations used in this study. B Both of the partially purified vegetative cell and spore PNPase proteins were labile in the absence of phosphate at 60 C. The enzymes used in the previous investigation were not electrophoretically homogeneous preparations and may have contained some unknown sta- bilizer which was stripped from the enzymes used in this work during the DEAE-cellulose purification step. This possibility was supported by the thermal inactiva- tion studies employing crude extracts of vegetative cells and spores in which the vegetative cell crude extract contained PNPase which was more stable than the extract containing the spore enzyme. After both preparations were purified by DEAE—cellulose chroma- tography their thermal stabilities were similar. The differential effect of phosphate ion on the thermal stability of cell and spore PNPase was also found 8“ to be different from that reported previously (16). The presence of phosphate ion stabilized both the vege- tative cell and spore PNPase proteins from either crude extracts or from the electrophoretically homogeneous preparations. The reason for this difference was not immediately apparent. The spore enzyme previously studied may have possessed some heat—stabilizing ion which was subsequently removed in the presence of phos— phate. The enzymes used in this investigation were routinely kept in a sulfhydral-protective environment (10 mM 2—mercaptoethanol) whereas the preparations used by Engelbrecht and Sadoff (16) were not. The thermal stability of spore PNPase from crude extract was not affected by the presence or absence of 2-mercaptoethanol since similar half-lives were obtained at 60 C when the enzyme was tested in either 50 mM imidiazole buffer, pH 7.5 or in the standard buffer. The maintenance of sulfhydral bonds during the purification of these enzymes could have affected their thermal stabilities. The absence of a sulfhydral protective reagent could have resulted in oxidation and the formation of some disulfide bridges which would permit a conformational change within the spore enzyme so that it could become more heat stable (7)- The spore PNPase preparation which was used in the previous thermal inactivation study (16) was eluted from 85 a calcium phosphate gel column before it was dialyzed and tested for thermal stability. It is possible that this enzyme preparation contained residual amounts of calcium ion which could have affected its stability. Vegetative cell and spore PNPases from crude extracts were tested in the presence of 10 mM CaCl The spore 2° . enzyme was slightly stabilized and the vegetative cell enzyme became labile. Heating the spores to 70 C for 30 min could have resulted in their activation and subsequent germination y had the proper agents been present in the buffer. Germination materials could have been derived from residual material remaining after the spores were washed. If some germination of the spore population did occur, the extracted PNPase could have possessed a lowered heat stability. However, the spores were all refractile before they were placed in the colloid mill as were the remaining spores after the cell grind- ing procedure. The denaturation of any vegetative cell PNPase present in the spore crop was more important to the goals of this study (separation of cell and spore PNPases) than the possibility of activation effects. No unusual components were detected in the ultra- violet absorbance spectrum of spore PNPase. Therefore, the differences in molecular weight between the spore and vegetative cell PNPase enzymes previously reported 86 U6)wmm not due to the binding or release of a cofactor orouwreasily characterized ultraviolet-absorbing com- ponent. Hm spore enzyme appeared to be catalytically adfivein its lowest molecular weight configuration. Thepflotof specific activity vs. protein concentration vms Lhmar and essentially had a zero slope. Had a [notehiconcentration-dependent aggregation been nec- essamyibr activity, the plot would have shown a drop in specific activity at the low protein concentrations. It is possible that the presence of phosphate in the assay mixtures could have promoted aggregation of PNPase even at low protein concentrations. The kinetic analysis indicated a complicated reac- tion mechanism. The data suggest that an ordered, sequential.reaction occurs since the Lineweaver-Burk plcnxs.for phosphate as the changing fixed substrate and :LnosiJue as the variable substrate (or reverse) inter- In this sented to the left of the vertical axis (11). the first substrate, inosine, binds Peac tion mechanism , CC) true enazyme and is followed by the second substrate, This reaction mechanism was proposed by Kim, IDbKDSIDkuatea. (:38) from inhibition studies with the PNPase €3t aLl. The reaction mechanism derived from human erythrocytes. is graphically represented by; Hx-R Pi R-l—P Hx I + + f 'Hx-R E E'Hx-R E E'Hx E 'R-l-P where E, is the enzyme; Hx-R, is inosine; Pi, is inor- ganic phosphate; R-l-P, is ribose—l-phosphates and Hx, is hypoxanthine. The Michaelis constants for inosine were different for the vegetative cell and spore enzymes. The value for the spore enzyme was 1.5—fold greater than that found for the vegetative cell enzyme and both enzymes had greater Michaelis constants than that found for the erythrocyte enzyme (38). These differences may reflect subtle structural differences between the cell and spore proteins. The concave downward curvature of the Lineweaver- Burk plots for both enzymes with inosine as the changing fixed substrate and phosphate as the variable substrate was unusual. This indicated an increase in maximal velocity and/or an increase in the Michaelis constant for phosphate at low levels of inosine. Plots of initial velocity vs. phosphate concentration were not sigmoidal but were rectangular hyperbolas. Therefore, the pos- sibility of a classical allosteric effect was not sup- ported. An allosteric enzyme has an effector binding 88 site, different from the active site, which non- competitively binds molecules other than substrate. This binding may decrease or increase enzyme activity. If the assumption is made that the maximal velocity for the vegetative cell and spore PNPases does not change (ie., the line on the replot of the intercepts vs. the reciprocal of the inosine concentration is linear) then the downward curvature would be the result of an increase in the Michaelis constant for phosphate. This would seem to support the theory that inosine must bind to the enzyme before phosphate. In this case, the binding of inosine may cause a conformational change on the PNPase enzyme which allows more efficient binding of the phosphate. At low inosine concentrations this conformational change may not occur on every PNPase molecule in the solution. This would result in a greater average Michaelis constant for phosphate over the entire enzyme population. High phosphate concentrations could also sterically hinder the binding of limiting amounts of inosine if the binding sites for the two substrates were proximally located. This hinderance would cause a decrease in the maximal velocity. The reverse effect was obtained, however. Kim, et a1. (38) attributed the non-linearity of the Lineweaver-Burk plot to a possible cooperative interaction between subunits. Two types of subunit 89 interaction have been reported which could explain these results. In the first type, an enzyme's catalytic activity varies with the number of subunits per active molecule due to changes in subunit association- dissociation equilibria (30). The activity of the individual subunits is assumed to be different from that found when the subunits are combined. If this were the case for the vegetative cell and spore PNPase enzymes, the plot of specific activity vs. protein concentration would have shown curvature and the initial velocity of the reaction would not have been proportional over a wide range of enzyme concentrations. In the sec- ond type of interaction, the binding of one substrate at one site would decrease the binding affinity of the remaining sites due to negative homotropic interactions between active centers (15). Therefore, each enzyme molecule would have more than one active center per subunit; or more subunits, each having one active center. The resulting rate of the catalytic reaction would be maintained at a moderate level in the presence of low phosphate concentrations because of the rapid combina- tion of the first phosphate molecule. As the phosphate concentration was increased the rate of the reaction would increase rapidly. Engel and Dalziel (15) stated that, "the effect is as if the number of binding sites increases with the (phosphate) concentration, the 90 successive sites having larger Michaelis constants for the (phosphate) and perhaps greater maximum turnover rates.". The results obtained during this study are consistent with this mechanism. Both the spore and vegetative cell PNPase enzymes had similar Michaelis constants for phosphate. These values were 5—fold greater than that obtained for human erythrocyte PNPase (3.2 x lO-u M) (38). It would be anticipated on the basis of the Michaelis constants that phosphate binding to the cell and spore PNPase proteins would be very weak. Yet, a profound effect of phosphate ion was noted in the heat resistance and molecular configuration of the enzymes. It would appear that phosphate ion binds at sites on the PNPase other than the catalytic site. Preliminary sucrose density gradient studies indicated that the crystalline human hemoglobin used as the sedimentation marker in the previous study (16) apparently underwent a decrease in sedimentation velocity with increasing concentrations of phosphate ion. Field and O'Brien (18) reported that the sedimentation coef- ficient of hemoglobin decreased with a decrease in the hydrogen ion concentration. This drop in sedimentation velocity was due to a decrease in average molecular weight without accompanying configurational changes (90). Since there was some question of the usefullness of 91 hemoglobin as a marker in sucrose density gradient sedi- mentation studies, rabbit muscle lactic acid dehydrogenase was chosen. This enzyme could be easily assayed and had a reported sedimentation coefficient of 7.6 S (62). Data obtained from the sucrose density gradient sedimentation studies of vegetative cell and spore PNPase enzymes, in the presence or absence of phosphate, did not coorelate with those reported previously (16). This dis- crepancy may be explained by changes in the sedimentation coefficient of the hemoglobin marker previously used. However, the general trend toward higher sedimentation velocities for the spore PNPase in phosphate was con- firmed. The sedimentation velocity of the vegetative cell enzyme did not appear to change regardless of the phosphate ion concentration. It was found that the increase in sedimentation velocity for the spore enzyme in the presence of phos— phate ion may be irreversible. Spore PNPase which was dialyzed against buffer containing 10 mM potassium phosphate and then dialyzed against the standard buffer did not have a sedimentation coefficient corresponding to the value obtained in the standard buffer. The unique binding of phosphate which produced an enzyme with a greater sedimentation coefficient may be quite strong. This was in contrast to the phosphate binding which occurred in the catalytic reaction, with a Michaelis constant of about 1.3 x 10'3 M. .mihumagnd‘xm -. '- x I 92 The striking result of the sucrose gradient studies was the comparatively small difference in sedimentation coefficients between the vegetative cell and spore enzymes in the standard buffer. Since it was known that the spore enzyme was significantly smaller than the vegetative cell enzyme, the similar sedimentation coef- ficients could only be explained by large conformational differences between the two enzymes. This conclusion was borne out by noting the constancy of the shape factor (f/fo) for the vegetative cell-derived PNPase, in contrast to the increased value of the same parameter for the spore PNPase in 10 mM phosphate ion. The high speed sedimentation equilibrium molecular weight study was made to obtain accurate molecular weight data. The plots of the natural logarithm of the fringe displacement vs. the squared distance from the center of rotation were nearly linear. Thus, the data were con- sistent with expected results and the molecular weight values can be accepted with certainity. If dissociation had occurred at the lowest concentrations, the plots would have shown significant curvature. The preparations also appeared to be nearly homogeneous since the exper- imental points along each plot did not show large devia- tions from the slope. In the standard buffer (50 mM Tris-HCl, 10 mM 2-mercaptoethanol, pH 7.5), the spore PNPase had a lower . .L-R'S-W tr» . 93 whole—cell average molecular weight than the vegetative cell enzyme. After the spore enzyme was dialyzed against buffer containing 10 mM potassium phosphate it had an average molecular weight which approximated that found for the vegetative cell PNPase. The nature of the phosphate—dependent molecular weight change for the spore enzyme can be found by examination of the weight- average (Mw) and number—average (Mn) molecular weight data. The spore PNPase preparation, in either the presence or absence of phosphate, contained more than one molecular weight component since the Mw values were greater than those for Mn. Only one molecular weight component was present in the vegetative cell preparation since the Mw and Mn values were nearly equal. A previous sedimentation equilibrium study per- formed under similar conditions with a greater protein concentration of spore enzyme indicated that the prep- aration was not purely disperse. In a disperse system, the enzyme solution would have contained polymers that were not in chemical equilibrium with each other. In that case, the resulting plot of the 1n of the fringe displacement vs. (radius)2 would have shown a curvature that differed from that found at the lower protein con- centration (100). The slope, however, was similar and was shifted toward the top of the sector cell. This result would be expected for an associating system. 9“ Since the spore PNPase preparations were not dis- perse and the protein concentrations at the meniscus were negligible (no changes in refractive index near the tops of the sector cells), an estimation of the molecular weight of the smallest species present in the solutions could be made. If sharp differences had been found between the slopes of 1n D vs. (r2) near the 3 top and bottom of the sector cells the size of the 1 smallest species could have been obtained by direct observation. Direct estimation is useful only if the % solutes contain the smaller molecular weight component contaminated with polymers greater than trimer (100). Therefore, the plots obtained in this investigation suggested the presence of a polymer consisting of less than four of the smaller molecular weight components. The size of the spore PNPase enzyme in the absence of phosphate ion could be estimated from the relationship derived by thantis (100); j (Mn) - Mw Molecular weight = j - 1 where j, is the number of the smaller molecular weight components in the polymer; Mw, is the extrapolated weight-average molecular weight at zero protein concen- tration; and Mn, is the extrapolated number—average molecular weight at zero protein concentration. The 95 weight fraction of the polymer was assumed to be less than 25% near the top of the sector cell. By assigning various values for j ranging from 2 to “, different minimal molecular weights were calculated: At j = 2, the molecular weight was “6,300; at j = 3, the minimal weight was 50,000; and at j = “, the weight was 50,300. Since the whole—cell average molecular weight for spore r1 PNPase in the absence of phosphate ion was 69,900 it. t was apparent that small amounts of polymer were present in the preparation. A measure of the spread of polymer B distribution was obtained from the following relation- ship (90); Mw/Mn = l + p where p, is the extent of polymerization in a two com- ponent system. The value for p was l“%. Since the whole—cell average molecular weight for spore PNPase in the presence of 10 mM potassium phosphate was 9“,800 it appeared that dimerization had occurred. When j was assigned a value of 2, the resulting calculated minimal molecular weight corresponded to a dimer having a molecular weight of 92,600. When higher values were assigned to j, the resulting molecular weights were greater than 100,000. The whole-cell molecular weight for the vegetative cell PNPase, in the absence and presence of phosphate, ranged between 88,300 and 91,600. 96 Thus, the spore PNPase appeared to undergo dimerization in the presence of phosphate ion to a size which approx— imated that of the vegetative cell enzyme. The subunit analysis of the vegetative cell and spore enzymes indicated that both preparations had a subunit size of 2“,000 i 10%. Therefore, the process which converts vegetative cell PNPase to spore enzyme does not appear to be the result of gross proteolysis, but rather the rupture of a relatively few bonds in the vegetative cell protein. The subunit size agreed well with the calculated molecular weights of each enzyme. The subunit data indicated that spore PNPase in the absence of phosphate (had two SDS-dissociable polypeptide chains whereas the vegetative cell PNPase had four. The calculated f/fO ratios indicated that spore PNPase in the standard buffer was more spherical (f/fO = 1.2) than it was in the presence of phosphate ion. In 10 mM phosphate the ratio was the same as that found for the vegetative cell PNPase in the presence or absence of phosphate (f/fO = 1.“). The axial ratios (78) of the spore monomer and dimer were about “ and 8 respectively. This was consistent with the subunits joining end-to-end, if large conformational changes did not occur during the dimerization process. Laskowski (“5) has stated that, ". . . a protein with about 500 97 amino acid residues could form one single glop of roughly spherical shape and still have enough surface to accom- modate its charged residues; beyond this size spherical proteins appear impossible." The increase in the f/fO ratio associated with dimerization of the spore PNPase is consistent with this View. Proteins may be more heat stable when they are in r a compact conformation Since interactions between the I protein and the surrounding environment would be reduced (90). Hydrophobic bonding may be the mechanism for t this increased heat resistance. Hydrophobic bonds result from the negative affinity of non-polar protein side chains for water. This results in a tight packing of these non-polar groups into the interior of the protein molecule where they are stabilized by hydrophobic bonds. There is some question whether these bonds are a separate entity in themselves or actually the result of Van der Waals forces between closely associated side chains. Other bonds, such as hydrogen and ionic bonds, may contribute to protein stability but they do not seem to be a major factor in protein denaturation by agents such as urea. The ip yiyp heat resistance of spore PNPase may be related to the phosphate effects which have been observed ip yippg. Nelson, et a1. (67) have shown that very little free phosphate is present in resting spores. 98 Therefore, the PNPase in spores may exist as nearly spherical monomers. SUMMARY The purine nucleoside phosphorylase from vegetative cells and spores of Bacillus cereus T was purified to electrophoretic homogeneity by ion exchange chromatog- 7 raphy and polyacrylamide gel electrophoresis. The specific activity of these preparations was higher than 5 those previously reported. Phosphate ion caused an g increase in the thermal stability of both purified enzymes whereas inosine had no effect. The catalytic properties of the two enzymes were similar and indicated an ordered, sequential reaction mechanism where inosine was bound before phosphate. The Michaelis constants (inosine as substrate) for the vegetative cell and spore enzymes were “.6 x 10_u M and 7.0 x 10‘” M respectively. The Michaelis constant for phosphate was 1.5 x 10_3 M for the vegetative cell enzyme and 1.3 x 10-3 M for the spore enzyme. Both enzymes had similar turnover numbers and the specific activities of the enzymes appeared to be constant at varying protein concentrations. Negative homotropic effects were proposed since the Lineweaver- Burk plots with inosine as the changing fixed substrate and phosphate as the variable substrate showed downward curvature at high phosphate concentrations. 99 100 Sucrose density gradient studies indicated that spore purine nucleoside phosphorylase (PNPase) increased from a sedimentation velocity of 5.3 S in the absence of phosphate to a value of 5.7 S in the presence of 10 mM potassium phosphate. The increase in sedimentation velocity may have been irreversible. The vegetative cell PNPase had a sedimentation velocity of 5.5 S in the presence or absence of phosphate ion. The calculated minimal molecular weight of spore PNPase was “6,300 in the absence of phosphate, and 92,600 in the presence of 10 mM phosphate. The vegetative cell enzyme had a molecular weight of 88,300 and 91,600 in the presence and absence of phosphate respectively. The subunit size for both enzymes was 2“,000 : 10%. 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Microbiol. 12:“36-““l. ‘llr”“ Worthington Enzymes for Research, Worthington Biochemical Corporation, Freehold, New Jersey, p, 1.1.1.27. VanHolde, K. E. and R. L. Baldwin. 1967. Rapid attainment of sedimentation equilibrium. J. Phys. Chem. 62:73“-7“3. thantis, D. A. 196“. Equilibrium ultracentrifuga- tion of dilute solutions. Biochemistry 32297-317. APPENDICES 112 APPENDIX A MATERIALS USED IN THE INVESTIGATION Acrylamide; N, N'-methy1enebisacrylamide 5 (bisacrylamide); glycine; 2—mercaptoethanol; and N, N, N'-tetramethy1ethy1ene diamine (TEMED) were purchased from Eastman Organic Chemicals. Inosine, hypoxanthine, E xanthine, and dithiothreitol were obtained from Calbiochem. Rabbit muscle lactate dehydrogenase (LDH, specific activity of 150 units per mg), streptomycin sulfate, riboflavin, and trishydroxymethylaminomethane (Tris) were products of the Nutritional Biochemical Company. RNase and DNase I were purchased from Worthington Biochemical Corporation. Analytical reagent grade sucrose was prepared by Mallenckrodt, and Antifoam B emulsion by Dow Corning. Sodium lauryl sulfate (SDS) was a product of Sigma Chemicals. The #110 glass pavement marker beads were purchased from the Minnesota Mining and Manufacturing Company. Sephadex G-25 was a product of Pharmacia Fine Chemicals. The 2, 3, 5-tripheny1tetrazolium chloride was from General Biochemicals, and Reeve Angel supplied the Whatman DE-52 microgranular DEAE-cellulose. The coomassie 113 11“ brilliant blue was from Mann; the brom phenol blue was a product of Difco and the buffalo black was from Allied Chemical. All other materials used were of the highest grade obtainable. APPENDIX B DERIVATION OF THE FUNDAMENTAL SEDIMENTATION EQUATION The sedimentation equilibrium method for molecular weight determination is based on the theory that at equilibrium the flow of solute due to sedimentation (Jsed) at every point in the cell is balanced by the counterflow due to diffusion (Jdiff). Jsed = velocity + solute concentration and, Centrifugal force = me2r' Taking into consideration the boyant force exerted by the solvent (1 - V p); Centrifugal force = m (l - Vo) e2r' where m, is the effective mass of the solute particle; e, is the angular velocity; r', is the distance from the center of rotation; V, is the partial specific volume displaced by the solute; and p, is the density of the solvent. Since effective mass (m) is equal to the molecular weight of a single solute molecule (M) divided by Avogadro's number (N), the centrifugal force equation can be rewritten: 115 116 M (l - V0) (w2r') Centrifugal force = ( I-) N and, Flow velocity = dr/dt = centrifugal force/f (11.) where f, is the frictional coefficient of the solute particle. The frictional coefficient is defined as; f = 6flnr where n, is the viscosity of the solvent; and r, is the effective radius of the protein. Combining equations (1.) and (11.) gives the flow velocity: _ 2 ' dr M (1 - v p) (e r ) (it N f Therefore; Jsed = flow velocity x concentration of solute (c), - 2 , M (l — V o) (w r ) (111.) or, Jsed = ~ ° c N f From Fick's first law of diffusion; Jdiff = - D dc/dr (IV.) 117 where D, is the diffusion coefficient of the solute; and dc/dr, is the solute concentration gradient; and from the Einstein diffusion equation; D ll '21 *3 (V.) 2 H: where R, is the universal gas constant; and T, is the absolute temperature, the force due to diffusion is derived. Combining equations (IV.) and (V.) gives: Jdiff = — —"— '-—— (VI.) At sedimentation equilibrium, Jsed = Jdiff. There- fore, combining equations (111.) and (VI.) gives; . 2 I do M (1 - V p) w r c -—-= (V11.) dr R T the fundamental sedimentation equilibrium equation for a single solute in an ideal solution. Determination of Molecular Weights From Actual Data From the integration of the fundamental equation (V11.), it follows that: M (1 — v p) .321“? 1n 0 = + c (VIII.) 2 R T 118 Given that: n = n k c (1X.) where n, is the refractive index of the solute; no, is the refractive index of the solvent; k, is a constant; and c, is the solute concentration, the interference fringes from each sector cell give an indication of the protein concentration at each point in the cell. By combining equations (VIII.) and (1X.) the relationship between protein concentration and molecular weight at equilibrium is established: M (1 - V p) w2r'2 1n (n - no) = + c 2 R T If a plot is made of 1n (n - n ) vs. r'2, the slope is 0 equal to the following; M (1 -Vp) 002 2 R T which allows a solution for average molecular weight. The computer program was designed to take five equally spaced adjacent data points along the X-coordinate and determine the least square straight line through these points. The slope of this line was equal to the slope at the central point given by fitting a least square quadratic through the same five points. The unweighted least squares 119 procedure, using five points at a time, produced an average least square straight line through the average slopes obtained for each set of five points over the entire cell. The molecular weights obtained by this procedure were very accurate. The computer was also programmed to analyze the data and compute weight-average (Mw) and number-average (Mn) F molecular weights at every data point in the cell. From these calculated values plots were constructed of either Mw or Mn vs. protein concentration. When the points on each plot were connected by a least square line and V extrapolated to zero protein concentration, the values for Mw and Mn of the smallest protein moiety were obtained. The mathematical operations used in these calculations are reviewed by thantis (100). APPENDIX C CALCULATION OF FRICTIONAL COEFFICIENTS The f/fO ratios derived from the molecular weight and sedimentation coefficient data were calculated in the following manner using the fundamental relationship between molecular weight and sedimentation velocity; o M (l - V p) S = (1.) N f‘ From Stoke's law; f = 6 flnr which is defined in Appendix B. From the volume of a sphere (V); o V = “/3 flrJ /3 V‘ 1/3 r = ———- (11.) “ fl and from the volume of a spherical protein (V1); vHierwy, 120 121 M V V' = -——- (III.) N the radius of a spherical protein (r), can be derived by combining equations (11.) and (111.): 1/3 3 M V r = (1V.) U N fl Combining equations (1.) and 1V.), gives the fundamental relationship between molecular weight and sedimentation coefficient for a spherical protein under ideal conditions: M (l - V p) S = (V.) 1/3 3 MV 6 N fl n U Nfl This equation was solved, assigning a value of 0.717 ml/gm for the partial specific volume (V) of PNPase. The values for the solvent density (0) and for the viscosity of the solvent (n) were known. The solution was; 0 = 3.79 x 10‘3 M2/3 122 Using this equation and the whole—cell average molecular weight values obtained from the sedimentation equilibrium study, sedimentation coefficients were calculated for both the vegetatiVe cell and spore PNPase enzymes in the presence or absence of 10 mM potassium phosphate. These calculated sedimentation coefficients were the values which would be expected if the PNPase enzymes were spherical molecules. Using a rearranged form of equation (1.); M F(l-Vo)] r: . i S L. N the minimal frictional coefficients (f0) were calculated from the whole—cell average molecular weight data and the calculated S0 values for each enzyme. Frictional coef- ficients were also calculated from the molecular weight data and the experimentally determined sedimentation coefficients to give the actual frictional coefficients (f) for each enzyme preparation. From the f/fO ratios, an indication of the molecular asymmetry could be found. A spherical molecule would have a f/fO ratio of one. Most proteins have ratios in the range from 1.0 to 2.0. llilllllltlllli 4444 u “ ll" ”II n “ u II" III HI 293 03061 31