/V/~3>3>3>3>3> Ul-P—‘WNH .6. Stopping Time Data Collection and Calibration Data Analysis Computer Graphics A. EXPERIMENTAL METHODS - AMP- AMINOHYDROLASE. Materials . . . . . . . Preparation of AMP—Aminohydrolase SDS Gel Electrophoresis Fluorescence Experiment Analytical Ultracentrifuge Experiments. Stopped-Flow Experiments. F.l. Scanning Experiments. F.2. Fixed Wavelength Experiments. F.2.l. Temperature Studies F.2.2. Preincubation Studies 5. RESULTS AND DISCUSSION AMP—AMINOHYDROLASE. Intermediates vi Page 38 38 A0 A0 A1 A3 A3 uu A6 A7 A7 A8 A8 A8 51 52 53 53 5A 5A 56 57 57 Chapter CHAPTER A. B. PART B. CHAPTER A. CHAPTER A.l. Fluorescence Study. A.2. UV-VIS Scanning Experiments Analytical Ultracentrifuge Studies. Preincubation Studies Temperature Studies D.l. Lag Times D.2. Shutoff Behavior. . . . . . . . D.3. Activation Energy 6. MODELS FOR AMP—AMINOHYDROLASE Model A The Monod, Wyman and Changeux Model (MWC) . . . . . . . . . . . . Phenomenological Model (Model B). STOPPED-FLOW STUDIES OF THE REACTION OF S-BENZYL-L-CYSTEINE WITH TRYPTOPHANASE 7. TRYPTOPHANASE - AN INTRODUCTION . . Historical. Enzyme Source Structure and Properties. Spectral Properties Mechanism of Tryptophanase Catalysis. 8. STOPPED-FLOW STUDIES OF THE REACTION OF S-BENZYL-L-CYSTEINE WITH TRYPTOPHANASE. Introduction. Materials and Methods vii Page 61 61 6A 69 73 75 88 91 93 9A 101 120 139 139 1A0 1A1 1A2 1A7 15A 15A 156 Chapter B.1 Reagents. B.2 Tryptophanase Preparation B.3 Scanning Stopped—Flow Experiments B.A Data Analysis . . . . . . . . C. Results and Discussion. 0.1. Absorbance Changes at 337 nm. C.2. Absorbance Changes at A20 nm. Absorbance Changes at A78 nm (508 nm). Absorbance Changes at 360 nm. 0.5. Absorbance Changes at 397 nm. 0.6. Correlations Between the Fast Phases at A20 and A78 nm (508 nm). . . . . . . . . . . C.7. Correlations Between the Slow Phases at 337 and A78 nm (508 nm). D. Preliminary Principal Component Analysis of the Reaction of S- -Benzyl— Cysteine with Tryptophanase . D.1. Introduction. D.2. Data Analysis . . . . . . . . . CHAPTER 9. SUGGESTIONS FOR FUTURE WORK A. AMP-Aminohydrolase. B. Tryptophanase APPENDICES APPENDIX A - Uncomplicated "Clean" Reaction and Its Independence of Wavelength APPENDIX B - Model A - Derivation APPENDIX C - MWC Model — Derivation REFERENCES. viii Page 156 157 157 158 159 159 159 16A 167 167 168 168 169 169 169 173 181 181 182 183 186 190 200 Table LIST OF TABLES Non-Michaelis—Menten Enzyme Models. Sedimentation Coefficients for AMP- Aminohydrolase at 20°C. Effect of Preincubating NHZ, IMP, 5'AMP and K+ on (dP/dt) Nine-Data Sets, Tested for Convergence by KINFITA. Phenomenological Model, Data-Fitting Results at 2°C. . . . . . . . . . . Phenomenological Model, Data-Fitting Results at 12°C Phenomenological Model, Data-Fitting Results at 22°C Rate Constants and Amplitudes for Ab- sorbance Changes at 337, A20 and A78 nm When 2.27 mg mL-1 Tryptophanase is Mixed with 1 mM S-Benzyl-L-Cysteine Rate Constants for 337, A20 and A78 nm As a Function of S-Benzyl—L-Cysteine Concentrations. ix Page 12 68 70 100 125 126 127 161 162 Table 8.2B Page Total Absorbance Changes at 337, A20 and A78 nm When 2.27 mg mL-1 Trypto- phanase is Mixed with S-Benzyl-L- Cysteine. . . . . . . . . . . . . . . . . . 163 Figure LIST OF FIGURES A General Allosteric Model. Flow Velocity Profile Mixing Efficiency Spectra Static Spectra of 5'-AMP, IMF and AMP— Aminohydrolase. Scanning Spectrum of 3.A uM AMP-Amino— hydrolase with 0.5 mM 5'-AMP. Fluorescence Spectra of AMP-Aminohydrolase in 0.5 M (CH3)uNC1 and 0.5 M KCl. Excitation and Emission Spectra of AMP-Aminohydrolase in 0.5 M (CH3)uNC1 Sedimentation Velocity Spectrum of 1.8 uM AMP—Aminohydrolase in 0.5 M KCl at 280 nm Sedimentation Velocity Spectrum of 1.8 pM AMP—Aminohydrolase in 0.5 M KCl at 260 nm Sedimentation Velocity Spectrum of 1.8 uM AMP-Aminohydrolase in 0.5 M (CH3)uNCI at 280 nm . . . . xi Page 10 A2 A5 59 6O 62 62 66 66 67 Figure Page 5.8 Sedimentation Velocity Spectrum of 1.8 uM AMP-Aminohydrolase in 0.5 M (CH3)HNC1 at 260 nm . . . . . . . . . . . . 67 5.9 Preincubation of NH: with AMP-Amino- hydrolase . . . . . . . . . . . . . . . . . 71 5.10 Preincubation of IMP, 5'-AMP and K with AMP-Aminohydrolase . . . . . . . . . . 72 5.11 Lag Time as a Function of 5'-AMP at 22°C. . . . . . . . . . . . . . . . . . . . 77 5.12 Lag Time as a Function of 5'-AMP at 12°C. . . . . . . . . . . . . . . . . . . . 78 5.13 Lag Time as a Function of 5'—AMP at 2°C . . . . . . . . . . . . . . . . . . . . 79 5.1A Lag Time as a Function of AMP—Amino- hydrolase at 22°C . . . . . . . . . . . . . 80 5.15 Lag Time as a Function of AMP-Amino- hydrolase at 12°C . . . . . . . . . . . . . 81 5.16 Lag Time as a Function of AMP-Amino— hydrolase at 2°C. . . . . . . . . . . . . . 82 5.17 HardCOpy Output for the Reaction of 3.6 uM AMP-Aminohydrolase with 0.625 mM 5'—AMP, up to 2A00 Milliseconds, at 22°C. . . . . . . . . . . . . . . . . . . . 83 5.18 Hardcopy Output for the Reaction of 3.6 uM AMP—Aminohydrolase with 0.625 mM xii Figure 5.19 5.20 5.21 5.22 5.23 5.2A 6.1 Page 5'—AMP, up to 500 milliseconds, at 22°C. . . . . . . . . . . . . . . . . . . . 8A Hardcopy Output for the Reaction of 0.9 uM AMP-Aminohydrolase with 0.625 mM 5'-AMP, up to 5000 milliseconds, at 2°C ... . . . . . . . . . . . . . . . . . . 85 Hardcopy Output for the Reactions of 2.5 mM 5'-AMP with 3.60, 1.80, 0.90, 0.A5 and 0.225 uM AMP-Aminohydrolase; the Curves Display (l/Eo)[(A’Ao)/(Aw’Ao)] Versus Time . . . . . . . . . . . . . . . . 86 Hardcopy Outputs for the Reactions of 2.5 mM 5'-AMP with 3.60, 1.80, 0.90, 0.A5 and 0.225 uM AMP-Aminohydrolase; the Curves Display (A-Am) Versus Time . . . 87 Lineweaver-Burke Plot for 0.9 uM AMP- Aminohydrolase Versus 5 mM 5'-AMP, in 0.5 M (CH3)uNC1, at 22°C and pH 6.5 . . . . 89 Lineweaver-Burke Plot for 0.9 uM AMP- Aminohydrolase versus 5 mM 5'-AMP, in 0.5 M KCl, at 22°C and pH 6.5. . . . . . 90 Arrhenius Plot for AMP-Aminohydrolase . . . 92 Model A, Two—Data Set Fitting; 0.225 uM AMP-Aminohydrolase with 0.3125 mM 5'—AMP, 22°C. . . . . . . . . . . . . . . . 99 xiii Figure Page 6.2 Model A, Two-Data Set Fitting; 0.225 uM AMP-Aminohydrolase with 5.0 mM 5'—AMP, 22°C. . . . . . . . . . . . . . . . 99 6.3 Model A, Nine-Data Set Fitting; 0.225 uM AMP-Aminohydrolase with 0.3125 mM 5'-AMP, 22°C. . . . . . . . . . . . . . . . 103 6.A Model A, Nine-Data Set Fitting; 0.225 uM AMP-Aminohydrolase with 1.25 mM 5'—AMP, 22°C. . . . . . . . . . . . . . . . 103 6.5 Model A, Nine—Data Set Fitting; 0.225 uM AMP-Aminohydrolase with 5.0 mM 5'-AMP, 22°C. . . . . . . . . . . . . . . . 103 6.6 Model A, Nine-Data Set Fitting; 0.90 uM AMP—Aminohydrolase with 0.3125 mM 5'-AMP, 22°C. . . . . . . . . . . . . . . . 105 6.7 Model A, Nine-Data Set Fitting; 0.9 uM AMP-Aminohydrolase with 1.25 mM 5'-AMP, 22°C. . . . . . . . . . . . . . . . 105 6.8 Model A, Nine-Data Set Fitting; 0.9 uM AMP-Aminohydrolase with 5.0 mM 5'-AMP, 22°C. . . . . . . . . . . . . . . . 105 6.9 Model A, Nine-Data Set Fitting; 3.60 uM AMP-Aminohydrolase with 0.3125 mM 5'-AMP, 22°C. . . . . . . . . . . . . . . . 107 xiv Figure Page 6.10 Model A, Nine-Data Set Fitting; 3.60 uM AMP-Aminohydrolase with 1.25 mM 5'-AMP, 22°C. . . . . . . . . . . . . . . . 107 6.11 Model A, Nine-Data Set Fitting; 3.60 uM AMP-Aminohydrolase with 5.0 mM 5'-AMP, 22°C. . . . . . . . . . . . . . . . 107 6.12 MWC Model for AMP-Aminohydrolase. . . . . . 109 6.13 MWC Model, Nine-Data Set Fitting; 0.225 uM AMP-Aminohydrolase with 0.3125 mM 5'-AMP, 22°C. . . . . . . . . . . 115 6.1A MWC Model, Nine-Data Set Fitting; 0.225 “M AMP—Aminohydrolase with 1.25 mM 5'-AMP, 22°C . . . . . . . . . . . . . . 115 6.15 MWC Model, Nine-Data Set Fitting; 0.225 uM AMP-Aminohydrolase with 5.0 mM 5'-AMP, 22°C . . . . . . . . . . . . 115 6.16 MWC Model, Nine-Data Set Fitting; 0.90 uM AMP-Aminohydrolase with 0.3125 mM 5'—AMP, 22°C. . . . . . . . . . . 117 6.17 MWC Model, Nine-Data Set Fitting; 0.90 uM AMP-Aminohydrolase with 1.25 mM 5'—AMP, 22°C . . . . . . . . . . . . . . 117 6.18 MWC Model, Nine—Data Set Fitting; 0.90 pM AMP-Aminohydrolase with 5.0 mM 5'-AMP, 22°C . . . . . . . . . . . . . . 117 XV Figure Page 6.19 MWC Model, Nine—Data Set Fitting; 3.60 pM AMP-Aminohydrolase with 0.3125 mM 5'-AMP, 22°C. . . . . . . . . . . 119 6.20 MWC Model, Nine-Data Set Fitting; 3.60 uM AMP-Aminohydrolase with 1.25 mM 5'-AMP, 22°C . ... . . . . . . . . . . . 119 6.21 MWC Model, Nine-Data Set Fitting; 3.60 pM AMP—Aminohydrolase with 5.0 mM 5'—AMP, 22"c . . . . . . . . . . . . . . 119 6.22 Phenomenological Model, Single Set Fitting; 1.8 uM AMP-Aminohydrolase with 2.5 mM 5'-AMP, 22°C . . . . . . . . . . . . 129 6.23 Phenomenological Model, Single—Set Fitting; 3.6 uM AMP-Aminohydrolase with 2.5 mM 5'-AMP, 12°C . . . . . . . . . . . . 129 6.2A Phenomenological Model, Single-Set Fitting; 0.9 uM AMP—Aminohydrolase with 1.25 mM 5'-AMP, 2°C . . . . . . . . . . . . 129 6.25 Phenomenological Model, [k2 + kl(S)] Versus (SO) at 22°C . . . . . . . . . . . . 132 6.26 Phenomenological Model, [k2 + kl(S)] Versus (SO) at 12°C . . . . . . . . . . . . 133 6.27 Phenomenological Model, [k2 + kl(S)] Versus (SO) at 2°C. . . . . . . . . . . . . 13A xvi Figure 6.28 6.29 6.30 8.1 Phenomenological Model, [k2 + kl(S)] Versus (E0) at 22°C Phenomenological Model, [k2 + kl(S)] Versus (E0) at 12°C Phenomenological Model, [k2 + kl(S)] Versus (E0) at 2°C. Interconversion of Tryptophanase Spec- tral Forms. A Mechanism for the Interaction of a Competitive Inhibitor, Ethionine to Form a Quinonoid. Changes in Tryptophanase Absorbance at 337 and A20 nm as a Function of Time After Mixing with S-Benzyl—L-Cysteine in 25 mM Kepps, 0.1 M KCl, pH 8.0, 1 mM KEDTA, 1 mM DTT and 5 uM PLP. After Mixing, Concentrations Were 1 mM S-Benzyl-L—Cysteine and 2.27 mg mL—1 Tryptophanase Biphasic Changes in Tryptophanase Ab- sorbance at A78 nm as a Function of Time After Mixing with S-Benzyl-L-Cysteine in 25 mM Kepps, 0.1 M KCl, pH 8.0, xvii Page 135 136 137 1A8 151 160 Figure Page 1 mM KEDTA, 1 mM DTT and 5 uM PLP. After Mixing Concentrations Were 1 mM S-Benzyl-L-Cysteine and 2.27 mg mL—1 Tryptophanase . . . . . . . . . . . . . . . 165 8.3 Overall Changes in Tryptophanase Ab- sorbance at A78 nm as a Function of Time After Mixing with S-Benzyl-L- Cysteine in 25 mM Kepps, 0.1 M KCl, pH 8.0, 1 mM KEDTA, 1 mM DTT and 5 uM PLP. After Mixing Concentrations Were 0.5 mM S-Benzyl-L-Cysteine and 2.27 mg mL-1 Tryptophanase . . . . . . . . . . . 166 8.A Experimental Surface, A, for the Re- action of 0.125 mM S-Benzyl-L-Cysteine with 2.27 mg mL-1 Tryptophanase . . . . . . 17A 8.5 Reconstructed Absolute Surface, A6, for the Reaction of 0.125 mM S-Benzyl-L- Cysteine with 2.27 mg mL-1 Tryptophanase. . 175 8.6 Reconstructed Absolute Surface, A5, for the Reaction of 0.125 mM S-Benzyl-L- Cysteine with 2.27 mg mL"1 Tryptophanase. . 176 8.7 Reconstructed Residual Surface, (A6-A), for the Reaction of 0.125 mM S-Benzyl-L- Cysteine with 2.27 mg mL-1 Trypto- phanase . . . . . . . . . . . . . . . . . . 178 xviii Figure Page 8.8 Reconstructed Residual Surface, (A5- A), for the Reaction of 0.125 mM S- Benzyl-L-Cysteine with 2.27 mg mL-1 Tryptophanase . . . . . . . . . . . . . . . 179 xix CHAPTER 1 INTRODUCTION An existing computer-interfaced rapid scanning stopped- flow system (9) was used to study the full time course kinetics of the catalytic reactions of two enzymes, AMP- aminohydrolase and tryptophanese. Results from these studies were analyzed in terms of new and existing models, consistent with the observed data, for AMP—aminohydrolase and tryptophanese respectively. Both enzymes can be activated by monovalent cations (50, 1AA,1A5,175). In the absence of monovalent cation acti- vators, AMP-aminohydrolase can also be activated by its substrate, 5'-AMP (50,1A6), while tryptophanase absolutely requires monovalent cations for activity (175). The first part of this work (Part A) describes stopped- flow studies of the activation of AMP—aminohydrolase by the substrate, 5'-AMP; while the second (Part B) describes such studies of the reaction of S—benzyl-L-cysteine with tryptophanase. PART A: AMP—AMINOHYDROLASE Non-linear steady state kinetics are often observed in enzyme systems (25,27-31,39,A0,A5,1Al). The Michaelis- Menten model alone is unable to account for the behavior of such kinetic patterns. Several more complex theoretical models have been proposed to explain such patterns (20-31, 3A,AA,A5,126), and most of those have been interpreted in terms of allosteric and/or hysteretic interactions (20—31, 39,A0,A5,126,127,1A1). In recent years, interest has been focused on the mode of action of regulatory enzymes (27-29,31,l27,1A1), the majority of which have been shown to exhibit both allosteric and hysteretic phenomena (31,127). Of interest among these metabolically relevant group of enzymes is AMP-aminohydro- lase (EC3.5.A.6), which catalyzes a simple reaction, namely, the irreversible deamination of 5'—AMP to IMP. The interest in AMP-aminohydrolase stems from the fact that in the presence of a monovalent cation activator, such as K+, the enzyme follows the Michaelis-Menten scheme; whereas, in the absence of any effectors except the sub- strate, 5'-AMP, this enzyme can display both allosteric (1A6) and hysteretic (27) phenomena in its deamination re- action, with the substrate doubling as a homotropic activa- tor. Such a simple system, with only enzyme and substrate, which can display allosteric and hysteretic behavior, both of which are removable in the presence of a monovalent cation activator, could be said to represent a most basic form of non-linear interactions in enzyme catalysis. Several mechanisms (27-29) were proposed to describe the homotropic activation of AMP—aminohydrolase by sub- strate, in the absence of monovalent cation activators, but none were reported to consistently fit the full time course data as a function of different initial substrate concentrations. The present study was undertaken in order to investigate for a mechanism(s) that would not only consistently fit the full time course data over a wide range of initial enzyme and substrate concentrations, at various temperatures, but also account for substrate depletion throughout the course used for measuring the kinetic behavior. The initial part of the study consisted of a search for Spectrophotometrically observable intermediates. The search, however, was unsuccessful. Later, sedimentation velocity experiments showed that the enzyme maintained its tetrameric configuration even in the absence of monovalent cation activators. This eliminated the possibility that polymerization or depolymerization effects were a cause for the observed allosteric and hysteretic phenomena. Preincubation studies of the product, IMP, with the enzyme indicated that while the product, at high concentra- tions, was able to partially remove the hysteretic lag, it was not substantially activating the enzyme as evidenced by the steady state regions of the full time course progress curves. Finally, a systematic stopped—flow study of the kinetics of deamination of 5'-AMP, in the absence of monovalent cation activators, was performed over a wide range of en- zyme and substrate concentrations, at 2°, 12° and 22°C, and at pH 6.5. As a precursory step to simulation and modelling, lag times in the pre-steady state regions, as well as the shut- off behavior of the progress curves were examined as func- tions of initial substrate and enzyme concentrations. The activation energy of the catalytic reaction was also de- termined, using data at the different temperatures used in the study. Three models were prOposed and fitted to the full time course data curves, at the three temperatures, using the generalized non-linear curvefitting program KINFITA (18). Substrate depletion over the time course used for measur— ing the kinetic behavior was incorporated in the three models tested. One common feature embodied in the mechanisms of these models involved an isomerization step(s). The first model which assumes the existence of an iso- merization step from an inactive enzyme-substrate complex, to an active enzyme-substrate complex, without invoking co- operative interaction phenomena in its scheme, is an ex- tension of the Suelter et a1. model (27). The second was the Monod et a1. model (20). While both models were able to describe the general behavior of the full time course data, they were unable to consistently fit all the progress curves as a function of initial enzyme and substrate con- centrations. The third model, which is a generalization of the mechanism proposed by Suelter 32 31. (27), is a phenomeno- logical model with apparent cooperativity effects in its scheme. This model was able to consistently fit the data as a function of the different initial enzyme and substrate concentrations at the three temperatures. Further analysis of the results indicated that subunit dissociation was occurring in the enzyme at temperatures below 20°C. PART B: TRYPTOPHANESE Tryptophanase (ECA.1.99.1), a pyridoxal 5'-phosphate (PLP) dependent enzyme, catalyzes a,B-e1imination reactions of amino acid substrates, and requires monovalent cations for optimum catalytic activity (175). The binding of PLP to the enzyme results in a pH—dependent absorption maxima at 337 and A20 nm (18A). When tryptophanase interacts with competitive inhibitors, a stable, dead end quinonoid complex, characterized by an intense absorption band centered at 508 nm, is produced (18A,197,198). A mechanism of quinonoid formation was proposed by June 33 al. (197) based on their studies on the interaction of two competitive inhibitors, ethionine and alanine, with tryptophanase. Their stopped-flow studies (197) of the kinetics of formation of the quinonoid with the inhibitor ethionine showed that the A20 nm form of the enzyme was the active form. Their assignment was supported by the following observations: (1) the rate constant for disap- pearance of the A20 nm absorbance was the same as that of the fast phase of the biphasic growth of quinonoid at 508 nm; (2) the rate of disappearance of the 337 nm absorbance was the same as the rate of the slow phase of quinonoid forma- tion and was essentially unaffected by the inhibitor con- centration or the nature of the inhibitor, ethionine or alanine; (3) the average rate constant for the slow phase obtained with ethionine agreed closely with the rate con— stant for the conversion of the 337 nm form, conformation y, to the A20 nm form, conformation 8, following a rapid decrease in pH (195). Thus the authors (197) concluded that the 337 nm form did not form quinonoid directly, but was first converted to the A20 nm form before the a-proton of an amino acid inhibitor was removed to form quinonoid. It could be argued however that the interaction of tryptophanase with competitive inhibitors, which leads to a stable dead end quinonoid, may not represent the same interaction as with a substrate, where the quinonoid ap- pears only as a transient intermediate. The present work describes a stopped—flow study of the changes in absorbance at 337, A20 and 508 nm following the interaction of a substrate, S—benzyl-L-cysteine, with tryptophanase. The objective was that if the time progress at these wavelengths, for this substrate, could be inter- preted in the same fashion as was done with the inhibitors, ethionine and alanine (197), then this should confirm the scheme for the mechanism of quinonoid formation proposed by June et al. (197). Analysis of the data established the correspondence between the present results to those obtained by June et al. (197). Preliminary principal component analysis [M- analysis] (17,201-203) indicated that a minimum of six independent absorbers were necessary to reconstruct the experimental surface to within its random error. This was in agreement with the experimental data which showed the presence of six absorbers. Non-Michaelis-Menten Enzyme Models Plots of initial velocity versus substrate concentration are non-hyperbolic for a large number of enzymes, particu- larly those termed regulatory (31,A5,1Al). For such enzymes it is necessary to devise models more complex than the Michaelis—Menten scheme. These non-Michaelis-Menten models are frequently defined in terms of allosteric and hysteretic interactions. Recall that AMP-aminohydrolase is a regulatory enzyme which displays both allosteric and hysteretic be- havior. The following is a brief introduction to some proposed allosteric and hysteretic enzyme models: Allosteric Enzyme Models Allosterism has been defined in terms of deviations of the reaction velocity from the predictions of the Michaelis- Menten Scheme (128). Two "classical" limiting molecular models have been proposed to account for allosteric control mechanisms. One is due to Monod, Wyman and Changeux (20), the MWC model; the other is due to Adair, Koshland, Nemethy and Filmer (21—23), the AKNF model. Both models are based on the subunit structure of proteins and alterations in conformation coupled to ligand binding. Both are illustrated in Figure (1.1). #211 I --._ I l l is _% €53 88 $ i 88 E :23! 9E? £99 Figure 1.1. A General Allosteric Model From Hammes and Wu (25) and Eigen (2A). (Note: The figure illustrates a general allosteric model for the binding of substrate (S) to a four sub- unit enzyme. The squares and circles are different conformations of the subunits. The MWC model is shown by dashed vertical lines, and the AKNF model by dotted diagonal lines. The free substrate and arrows between the states are omitted for the sake of clarity. Some of the intermediates in which (S) can bind to either the square or the circle are pre- sented by one state.) 10 The MWC model predicts a concerted conformational change (all or nothing), while the AKNF model predicts a sequential conformational change for each subunit. The homotropic cooperative effects are always positive for the MWC model (i.e., the binding of the first molecule enhances the binding of the second, etc.), whereas they may be positive or negative in the AKNF model. The MWC model assumes the same conformational state is stabilized by a variety of ligands, whereas the AKNF model predicts that different ligands may induce different conformational changes. Both the MWC and AKNF models are limiting cases of a more general scheme, as shown in Figure (1.1), initially proposed by Eigen (2A). This is because states with more than a single ligand bound can take up different geometrical configurations; for example: @D/ O 11 Finally, a complete solution based on experimental evi- dence, to any of these models without any simplifying as- sumptions is quite difficult, as the number of unknown parameters becomes unrealistically large. The general model combining both the MWC and AKNF extremes, along with dissociation of the subunits, etc., has been analyzed (26), but the results are too complex for general use, and it is far more convenient to interpret experiments in terms of the MWC or AKNF limiting schemes. The MWC and AKNF models, which assume the existence of interactions among the enzyme subunits are not unique in describing allosteric behavior. Several other models have been prOposed (27,29-31,3A,AA,A5,126,127) and some of these are shown in Table (1.1). A common feature among these models is the existence of at least one slow step in their scheme Table (1.1), which appears to be sufficient in order for allosterism to be manifested even when cooperative interactions are not considered. The slow step(s) normally describes a confor- mational change, such as an isomerization of the enzyme and/or enzyme-substrate complex, from an inactive to an active form, with the active form having higher affinity for substrate. Allosterism then results from this inter- conversion of configurations, which are assumed to be slow relative to other catalytic steps, such that the different configurations are no longer in equilibrium with each other 12 "I II "IIOO'OIJ .... m in a .A m m 2: 552 a+.nm_x.||m.m TIIHW m.m¢||.|..vm+m r llllllll '8'---“'-"---0-|"-'L xaAJsfllxa 33v .882 _ a. .E’ .alflalqvn. Azmv Exam ...w flumwxflmmwfiw .... .wfiopo: meancm Gonzozlmflaowcoflzucoz .H.H ofiowe 13 Aomv .mm mm mnfimcfia Azav on can Ammo :mcofizz AANV .mm Mm tmsamsm xmx (m a...» _. =3... 3.4 Mm / \ am uflllllliv Ilv Illv a+x.m¢||xmm¢|l 3%ng rising: ......innss nnnnnnnn ..— .cmscflpcoo .H.._H mHQwB 1A during catalysis. The extensive reviews by Whitehead (A5) and Citric (1A1) describe a large variety of models which illustrate this point as well as showing that no one mechanism proposed is the right one, but as Whitehead (A5) pointed out, they are all true for some cases in their description of allosterism. Hysteretic Enzyme Models Frieden (29) defined hysteretic enzymes as those which respond slowly to rapid changes in ligand concentration. Hence, unlike those enzymes which obey the Michaelis-Menten scheme, where product formation is always linear with time, hysteretic enzymes exhibit an initial lag or burst in product formation as a function of time. Frieden (29) suggested that mechanisms which may be responsible for slow processes in hysteretic enzymes include: (1) isomerization processes; (2) displacement of a tightly bound ligand by another with a different effect on activity; and (3) polymerization or depolymerization processes. The majority of regulatory enzymes have been shown to be hysteretic (27-31,3A,AA,A5,126,l27,1Al). Blum (3A) was the first to describe such systems (Table 1.1). Later, Weber (126), Suelter et a1. (27), Rabin (AA), Ainslie e: 31. (30), and Ho (1A) also presented schemes describing hysteresis (Table 1.1). For extensive reviews of hysteretic models see Citri (1A1), Whitehead (A5), l5 Frieden (31) and Kurganov (127). It is interesting to note that models proposed for hysteretic enzymes (27-31,A5,127,1Al), including the Frieden model as analyzed by Pierce (32), not only include a slow step(s) in their scheme, but are also able to display (or were initially proposed to account for) allosterism. Only recently, with the availability of computers, which faci- litate numerical analysis of models by simulations (l8) and other techniques (32,33), has it been possible to show that with proper choice of rate constants, many schemes can be shown to display not only allosteric as well as hysteretic phenomena, but that it is also possible to reduce complex schemes to simpler ones which display similar behavior to the full model. For example, Pierce (32) using non-linear sensitivity analysis (33), analyzed the Frieden (29) and Ainslie gt gt. (30) models in terms of bursts, lags, and cooperativity. He found that with judicious choice of rate constant values, both models can display several different combinations of bursts, lags, positive and negative cooperativity. Furthermore, he was able to show that the Ainslie gt g1. model can be reduced to one which exhibits essentially the same behavior as their full model. While it will always be both interesting and necessary to examine theoretical models, the challenge still remains in finding a model(s) which can account for what is observed l6 experimentally. In his review article, Whitehead (A5) summed it up as follows: "In light of the above discussion, the various models on offer will perhaps not be seen as a set of rival and exclusive orthodoxies, but are all closely related. The simpler ones are special cases of more general ones. Choice between them amounts to asking which equilibria, which interactions, which rates can for practical purposes be forgotten as a determining factor in the overall effect? Thus not only may different models be applicable to different enzymes, but different models may be applicable to one enzyme." In conclusion, the ttgg mechanism for an enzyme may never be discerned, and one can only hope to provide plaus- ible models to help explain that which is observed. But this is not a strange phenomena to one who studies kinetics. The oft quoted phrase that "one can always disprove a mechanism", may in the end, prove to be the only true mechanism. PART A STOPPED-FLOW STUDIES OF THE ACTIVATION OF AMP-AMINOHYDROLASE BY 5'-AMP CHAPTER 2 AMP-AMINOHYDROLASE - AN INTRODUCTION This chapter introduces the background, physiological significance, chemical and physical properties of AMP— aminohydrolase; after which the reaction of the enzyme with 5'-AMP is explored in light of the work of previous authors. A. Background AMP-aminohydrolase (EC 3.5.A.6; 5'-adenylic acid amino- hydrolase; adenylate deaminase) catalyzes the irreversible deamination of 5‘-AMP (5'-adenosine monophosphate; adenylic acid) to form IMP (5'-inosine monOphosphate; inosinic acid) and ammonia: (page 18) It was first described by Schmidt (A6-A8), who separated it from adenosine deaminase from rabbit skeletal muscle, and demonstrated its specificity for 5'-AMP. The enzyme is widely distributed in animal tissues, and has been isolated from a variety of sources (A9-5A), including muscle (55-6A), calf brain (65), guinea pig and human epidermis (66), pig kidney (67), bakers yeast (68), marine bacteria (69), human erythrocyte (70), and rabbit cardiac l7 l8 NHZ OH k “ + +—9m+NHI | RIBOSE(P) RIBOSE(P) 5’-AMP IMP muscle (71). Lowenstein (72) reported that the specific activity of the skeletal muscle enzyme is more than ten times higher than that found in other tissues. The highest concentration of the enzyme is also found in skeletal mus- cle (A9,56,72), where it is reported to be closely as- sociated with myosin (A9,5l,57,58,73—77). Ashby and Frieden (76) reported that in rabbit skeletal muscle, binding of the enzyme to myosin in vitro appears to be specific, in the sense that there are approximately two moles of enzyme bound per mole of myosin, and that the interaction may be affected by the presence of cer- tain ligands. Shiraki gt g1 (7A,75) reported that in rat muscle, in vitro, AMP-aminohydrolase is bound to the myosin 19 molecule in a ratio of one mole of enzyme to three moles of myosin. Sections (B) through (F) present additional information on AMP-aminohydrolase, which in the author's Opinion is complimentary to the present study of the enzyme; however, a reader may gloss over these sections without loss in continuity. B. Physiological Role Although the physiological role of AMP-aminohydrolase remains obscure, it has long been recognized that the re- action catalyzed by this enzyme is the major source of ammonia in muscle (78,79), and that ammonia production is proportional to the work done by the muscle (80). The levels of AMP-aminohydrolase in skeletal muscle from humans suffering Duchenne muscular dystrophy have been shown (5A) to be lower than in normal patients. Pennington (81) earlier reported similar findings in mice. Recently, Suelter gt g1. (82) and others (82-85) have studied the activity of AMP-aminohydrolase in normal and dystrophic chicken. They reported that the activity of the enzyme was lower in dystrophic chicken muscle, as compared to normal chicken muscle. Raggi gt g1. (86,87) reported that in the rat, white muscles have a higher AMP-aminohydrolase content than red muscles; that different types of AMP- aminohydrolase exist in white and red muscles, and that 20 the higher enzyme activity in white muscles is accounted for by an enzyme form which is present in much lower amounts in red muscle. The activity of AMP-aminohydrolase in muscle contraction (88) and as related to muscle fatigue (89-91) has also been reported. Setlow gt gt. (61) proposed a scheme for the regulation by AMP-aminohydrolase, in the interconversion of purine nucleotides in mammalian organs, where adenosine triphos- phate (ATP) acts as an activator and quanosine triphos— phate (GTP) as an inhibitor. The levels of (ATP) and (GTP) necessary for activation and inhibition respectively, are in themselves complicated and further modulated by other nucleotides, as well as monovalent and divalent ions. Hence, a proposed control mechanism based upon observed changes in vitro, is subject to still undefined in vivo effects by other factors. In the cyclic scheme of purine nucleotide intercon- versions described by Lowenstein (61), the deamination of aspartate in muscle tissue also serves important regulatory functions where (5'—AMP) is generated as (IMP) is depleted. The net effect of the cycle is the deamination of aspartate to yield ammonia in skeletal muscle. aspartate + GTP + H2O + fumarate + NH3 + GDP + Pi Lowenstein (72) also proposed that the purine nucleotide 21 cycle consists of AMP-aminohydrolase, adenylosuccinate synthetase, and adenylosuccinatelyase, and that these en- zymes operate in the ammoniagenesis from amino acids in skeletal muscle. He also suggested that this cycle was closely linked to glycolytic oscillation (96). An interesting physiological role for AMP—aminohydro- lase is in the biochemical pathway of the purine catabolism cycle, whose major end product in humans and the higher apes is uric acid. The production of above normal levels of uric acid can lead to gout. Those afflicted with this disease have markedly elevated levels of uric acid in their blood and other body fluid systems. Hers gt gt. (103) have proposed that primary gout is caused by the presence of an abnormal AMP-aminohydrolase which is less sensitive to its physiological inhibitors, thus the degradation of adenine nucleotides in the liver is increased resulting in excess uric acid formation. C. On the Regulation of AMP-Aminohydrolase by ATP, ADP and GTP as a Function ofng Ranieri-Raggi gt gt. (10A), Ronca-Testoni gt gt. (105), and Raggi gt gt. (106) studied AMP-aminohydrolase (isolated from rat skeletal muscle) in order to clarify the effects of pH and salt on the activity of the enzyme and its sensitivity to modifiers. While they confirmed the general activatory and inhibitory effects of (ATP) and (GTP) 22 respectively, their results further indicated the modula— tion of the effects of these modifiers on the enzyme ac— tivity as a function of pH. They reported that whereas at acidic pH values, a hyperbolic substrate saturation curve is obtained at optimal KCl concentration, the curve is always sigmoidal at pH 7.1 regardless of the concentration of KCl employed. Sim- ilar observations have been reported by Wheeler and Lowen- stein (107,108). The former authors (10A-106) further pointed out that the study of the kinetic properties of the enzyme has mainly been restricted to the Optimal pH value of 6.5; a rather distant one from the physiological pH of 7.1. They interpreted their results by tentatively assuming the existence of two kinetically distinct states of AMP-aminohydrolase whose ratio is pH dependent. State (I), which corresponds to the active conformation of the enzyme and present at acidic pH, is characterized by the familiar hyperbolic substrate saturation curve at optimal KCl concentration. State (II), whose induction is trig- gered by an elevation of the pH above neutrality is a less active conformation of AMP-aminohydrolase showing sigmoidal kinetics even at optimal salt concentration. Relevant deductions for the in vivo activation pattern of the skeletal muscle enzyme can be inferred from the above for the enzyme regulation. In view of the alkaline pH in resting muscle, State (II) is likely to be the primary 23 conformation of the enzyme. Therefore, the activity of AMP-aminohydrolase should be very low under these condi- tions, and also because of the low 5'—AMP (0.1 mM) content of tissue (109), and the effective inhibition excreted by (ATP) at low concentrations (10 UM), whereas at concentra- tions higher than 50 uM (ATP) acts as an activator. This possibility is supported by the reported inability to detect ammonia formation in resting muscle (110). During sustained contractile activity the decrease of tissue pH may be the cause of activation of AMP-aminohydro- lase, causing a shift in the equilibrium towards State (I) which characteristically has a much higher affinity for 5'-AMP and a low sensitivity to inhibitors. The activation pattern of the enzyme is again confirmed by the circum- stances of ammonia production in muscle (105,110). Barasacchi, gt gt. (71) reported that the affinity of AMP-aminohydrolase for 5'-AMP, in rabbit cardiac muscle, was low in the absence of activators (Km = 3.1 mM), but in the presence of (ATP), it became as high as that of the skeletal muscle enzyme (KIn = 0.A mM). At pH 7.5-8.0, the cardiac muscle enzyme was maximally activated, whereas the skeletal muscle enzyme was maximally inhibited by (ATP). Thus the cardiac muscle enzyme may always be in the activated state (State 1), whereas the skeletal muscle enzyme was active only during exhaustive contractions. Coffee and Solano (111) also reported that in rat 2A skeletal muscle, the activity of AMP-aminohydrolase is very stringently controlled as evidenced by the fact that the enzyme is operating at only 10 to 15% of its capacity. The results of Coffee and Solano are in complete agreement with the suggestions of Ronca-Testoni gt gt. (105) who reported that the enzyme activity in resting muscle (or in muscle undergoing moderate contractions) may be in- hibited to a great extent. They suggested that maximal activity may be approached only under conditions of extreme stress. Chung and Bridger (112) present yet another compli- mentary view for the activation sensitivity of the rabbit cardiac muscle AMP-aminohydrolase towards (ADP) and (ATP). They reported that both nucleotides activate the cardiac enzyme, but much higher concentrations of ATP (10 mM) than ADP (1.3 mM), are required to produce a demonstrable effect. The authors estimated that at 0.1 mM AMP, Kés for ADP and ATP are approximately 0.65 mM and 0.9 mM respectively. Their results suggest that ADP may be an important physio- logical regulator for the activity of AMP-aminohydrolase in cardiac muscle. Tornheim and Lowenstein (113) have suggested that 5'- AMP deamination might protect indirectly against ATP de- pletion, by virtue of the fact that removal of 5'-AMP would cause net conversion of ADP to ATP via readjustment of the adenylate kinase system: 25 2 ADP 1 ATP + 5'-AMP Hence AMP-aminohydrolase may play the regulatory role in the adenylate kinase system, as a buffer against ADP accumulation in the contracting muscle. The allosteric properties of AMP-aminohydrolase thus provide for negligible activity at the physiological con- centrations of 5'-AMP until ADP has accumulated. High rates of conversion of ATP to ADP resulting from prolonged strong contraction of muscle will result in, activation of AMP-aminohydrolase, reduction of 5'-AMP, and readjustment of the adenylate kinase equilibrium with net conversion of ADP to ATP at the eXpense of the overall pool of adenine nucleotides. Such activation might be regarded as an emergency measure providing short term but immediate rescue from con— ditions favoring ADP accumulation. Uncontrolled enzyme activity would not be beneficial because of the eventual net consumption of energy required for the regeneration of 5'-AMP from IMP via the purine nucleotide cycle. Ranieri-Raggi gt gt. (10A) have concluded that the cor— respondence in the modulatory effects of both ATP and pH on muscle AMP-aminohydrolase can thus play an important role in protecting the muscle cell against the danger of unrestrained activity of the enzyme, which would ultimately lead to depletion of the adenine nucleotide stores. 26 It has also been suggested by Mockrin gt gt. (11A) that negative cooperativity Which ATP has with AMP-aminohydro- lase) with its inherent difficulty of saturating the active sites of the enzyme, can be a very advantageous solution for limiting the catalytic activity at high substrate levels. The connection between the mechanism of regulation sug- gested above for AMP-aminohydrolase from skeletal muscle, and the specialized metabolism of this tissue is under- lined by the remarkable differences in kinetic properties of the cardiac muscle enzyme (71). D. Structure and Composition Any realistic crystallographic structural determination for an enzyme as complex as AMP—aminohydrolase must begin with homogenous and highly active single crystals. In this regard Smiley gt gt. (115) reported on the prepara- tion of homogenous, highly active, hexagonal bipyramidal crystals of the enzyme. Earlier, Lee (55) was the first to report on the crystallization of AMP—aminohydrolase. Although Lee presented evidence that his preparation was homogenous, the ratio of the optical densities at 280 and 260 mu (nm) of about 1.2 indicated the presence of nucleic acid contaminants, whereas Smiley, gt gt. had a ratio of 1.8 or higher, indicating little or no contamina- tion by nucleic acids. However, no crystallographic 27 studies have been reported so far. Depending on the animal source and the tissue from which the enzyme was extracted, AMP—aminohydrolase has an average molecular weight close to 280,000 and is composed of four identical subunits (60,68,111,ll6-12A). The amino acid composition of the enzyme from a variety of sources has been reported (60,62,117,l20,125). The rabbit skeletal muscle enzyme has a normal amino acid content, i.e., no unusually small or large amounts of a particular amino acid was found. Zielke and Suelter (116) characterized AMP-aminohydro— lase, isolated from rabbit skeletal muscle, as a zinc .metalloenzyme. They reported that the apo-enzyme binds A gram atoms of zinc per mole of enzyme, while the native enzyme as isolated contains 2.6 to 3.2 gram atoms of zinc per molecule. Although the addition of the fourth zinc atom increased V this increase was only 28% of that max’ expected, assuming that a linear correspondence exists between Vm and gram atoms of zinc per mole of enzyme up ax to A gram atoms of zinc. They concluded that the fourth zinc atom does not appear to be directly associated with activity. They further suggested that the problem of assigning a precise integral stoichiometry for the zinc to protein ratio in vivo resides with the difficulty in defining the similarity of the isolated enzyme in vitro to that in vivo. Raggi gt gt. (129) reported that the 28 rat muscle enzyme contains 2 gram atoms of zinc per mole when isolated by the procedure of Smiley gt gt. (115). Isoenzymes having the same catalytic reaction as AMP- aminohydrolase, but differing in activity on the basis of differences in relative substrate specificities have been reported (7A,l22,130—132). E. Substrates AMP-aminohydrolase is a highly specific enzyme towards its native substrate 5'-AMP. Other naturally occurring nucleotides, and synthetic analogs either have very low Vmax or very high Km values when compared to 5'-AMP (133- 138). Zielke and Suelter (138) examined a total of 2A nucleotides as substrates for rabbit muscle AMP-aminohydro- lase. Their results showed that 5'-AMP is the preferred substrate, having the largest Vm and the lowest Km ax values. They also found that while adenosine and ADP were deaminated, albeit poorly, ATP with three phosphoryl groups was not. Substitution in the ribose moiety have marked effects on deamination. Indications that the phosphate group may be important for binding of substrate are obvious. 29 F. Activators and Inhibitors F.1. Inorganic The enzyme from many sources is activated by mono- valent cations such as K+, Na+, Li+, NHZ, Rb+, and Cs+(50, 105,139,1A0). Campbell and Suelter (1A7) reported that AMP-aminohydrolase from rabbit skeletal muscle is also ac- tivated by H+. They presented evidence which indicated that the binding of H+ and K+ was linked, in that the dis- sociation constant KA for K+ activation was reduced as the pH was lowered. Smiley gt gt. (115) reported that K+, Na+ and Li+ are the most effective activators, with K+ being the best for the rabbit muscle enzyme. However, the relative order of effectiveness is dependent upon the source of enzyme. For example Setlow and Lowenstein (65) reported that in the calf brain, Li+ is the most effective alkali metal ion. Smiley et al. (115) studied the stability of AMP-aminohydrolase in various concentrations of K+. The general role of monovalent cations in enzyme activation has been explored by Suelter (1AA,1A5). The same Vmax was ob- served at high concentrations of 5'—AMP in either the pres- ence or absence of activating cations (50). The Vm ax values in the presence of K+, Na+ or (CH3)uN+ are identical (1A6). The demonstration that the Vmax of the reaction is the same in the presence and absence of K+ eliminates the 30 possibility that the mechanism of activation involves a direct role of K+ in catalysis. Rather, the binding of K+ affects the active site indirectly (allosteric hetero- tropic activation), such that the apparent affinity for 5'-AMP is increased, presumably through a change in the protein conformation. Coffee and Solano (111) reported that for the rat muscle enzyme Km and Vm appear to be in- ax dependent of K+ concentration, in the absence of other effectors in the reaction. Many divalent cations are known to inhibit the enzyme. 2+ 2+ (65,1A8), Zn , Fe3+ and Ag+ (1A8), 2+ These include Cu Cd2+ and Ni2+ (65), and Hg (1A9). A variety of anions such as inorganic phosphate (lA8,l50-l53), sulfate and nitrate (15A), pyrophosphate (1A8,155), tripolyphosphate (155), and F' (1A8,152,153,156-l58) inhibit the enzyme to various degrees. F.2. Organic In the absence of monovalent cations, AMP-aminohydro- lase can be activated by its own native substrate 5'—AMP (27,28,65). Thus the substrate acts as an allosteric homo- tropic activator. Smiley and Suelter (1A6) reported that in the absence of K+ or Na+ and at pH 6.5, the enzyme from rabbit skeletal muscle can also be activated by ADP or ATP, and inhibited by GDP and GTP. Their data suggested that ADP was a better activator than ATP. Chung and Bridger 31 (112) reported the same finding for the rabbit cardiac AMP-aminohydrolase, except that much higher concentrations of ADP and ATP were required to demonstrate the activation effect in the cardiac enzyme. ADP activation is inhibited by GDP and GTP, while the ATP activation is inhibited only by GTP. Inorganic phosphates also strongly inhibit the activation by nucleotides Setlow gt gt. (61) reported that in the absence of monovalent cations and at a pH of 7.2, AMP-aminohydrolase isolated from a variety of rat tissues was activated by ATP and strongly inhibited by GTP, and that the GTP inhibition was relieved by the addition of ATP. Setlow and Lowenstein (65) found that the calf brain enzyme is also activated by ATP, and reported (155) the effects of several nucleo- tide triphosphates on the calf brain enzyme. They found (155) that ATP was the most effective activator, and GTP the most effective inhibitor. The inhibition by GTP was reversed by raising the concentration of ATP. The activa- tion of ATP was reversed by raising the concentration of GTP, and both ATP and GTP were very effective in protecting the enzyme against heat inactivation. Tomozowa and Wolfenden (159) reported that in the enzyme from rabbit skeletal muscle, binding of ATP and GTP are mutually inhibitory, and proposed two binding sites for GTP and a minimum of four binding sites for ATP, and that both act as allosteric effectors towards AMP-aminohydrolase. 32 Yoshino gt gt. (123) reported that the chicken eryth- rocyte enzyme was inhibited by concentrations of ATP and GTP below 100 uM and 250 uM respectively, but was activated at concentrations of ATP and GTP above A00 uM and 1500 pM respectively. They suggested that these unusual kinetics with respect to ATP and GTP could be explained by assuming two inhibitory and four activatory sites for each ligand. Ashby and Frieden (6A) reported that for the rabbit skeletal muscle enzyme, in 0.15 M K+ and pH 6.5, ATP was an inhibitor at low concentrations (10 uM) and an activator at high concentrations (200 pM). GTP also inhibits at levels below (20 pM) but the inhibition is decreased at its level is increased above (20 pM) and, at (100 uM), the initial rate is almost the same as that in the absence of GTP. Wheeler and Lowenstein (107) reported similar find- ings for the rat muscle enzyme. Barsacchi gt gt. (71) reported that AMP-aminohydrolase from rabbit heart was activated by ATP, and less efficiently by ADP. Ronca-Testoni gt gt. (105) studied the enzyme isolated from the skeletal muscle of frog, pigeon, guinea pig, rabbit and rat. ‘Coffee and Solano (lll) studied the rat muscle enzyme. Both groups attempted to focus more specifically on the role of K+ ions in the expression of the regulatory properties of the enzyme, and on the specific effects of ADP, ATP, GTP, Pi (inorganic phosphate) and creatine-P, on the kinetic parameters Km and Vm For instance, both ax' 33 teams suggested that at physiologically significant levels of these metabolites, and as pH 6.5, (25 uM) ATP and (10 mM) GTP inhibit the enzyme by reducing its affinity for 5'- AMP; i.e., they increase Km from 0.35 mM to about 0.6 mM, without significantly affecting the Vmax of the reaction. But at K+ concentrations of 5 mM (approximately twenty times less than the average in vivo levels in muscle, ADP and ATP both cause a dramatic increase in Km to about 1.0 mM; and a decrease of about A0% in the value of Vmax' Skladanowski gt gt. (160) reported a different view on the K+ dependent regulation by ATP and ADP of AMP-aminohydro- lase from beef heart. Their results showed that ATP was a much more efficient activator than ADP, both at low and high K+ concentration. Melander (161) reported that IMF and NH: both inhibit rabbit skeletal muscle AMP-aminohydrolase; however Ashby and Frieden (6A) reported that IMP substantially activates the enzyme, and found that when 0.5 mM IMP was preincubated with the enzyme at 20°C, pH 6.5, and 0.15 M KCl, the ac- tivity increased by about 8% over that with no IMP added. Suelter §£.§l° (27) reported that products of the reaction have no activating or inhibiting effect. Zielke and Suelter (116) reported that the rabbit skeletal muscle enzyme was sensitive to inhibition by di- and tricarboxylic acids such as succinate, maleate, fumarate and citrate. The ADP activated enzyme was more sensitive 3A to these inhibitors than enzyme assayed in the presence of activator K+. A more complete list of activators and inhibitors for AMP—aminohydrolase is found in Zielke's dissertation (162) and reference (50). G. Reactions with 5'-AMP In the presence of monovalent cation activators, such as K+, and at pH 6.5, AMP-aminohydrolase displays normal Michaelis-Menten kinetics (27,28,50,6A,1A6). At 30°C, cal- culated values for Km range from 0.16 mM to 0.7 mM; Vmax values from 1000 to 1500 umoles 5'-AMP deaminated per minute per milligram enzyme (28,6A,10A,lll,ll5,lA0,lA6,l63). The Hill slope (19,16A) is equal to 1.1, as reported by Smiley and Suelter (1A6). Smiley and Suelter (1A6) investigated the allosteric prOperties of AMP-aminohydrolase prepared from rabbit skeletal muscle. Their results showed that a univalent cation can act as an allosteric activator for the enzyme, but that K+ is not absolutely required for the catalysis. The same authors (1A6) also demonstrated that the enzyme had homotropic effects with its substrate 5'-AMP. In the absence of monovalent cation activators, a lag phase in the deamination reaction, which occurs on the stopped—flow time scale (27,28) clearly indicates activation by the substrate, 5'-AMP. Suelter gt gt. (27), using 35 stopped-flow techniques, studied the time dependence of activation of muscle AMP-aminohydrolase by substrate and K+. They observed (27) that when substrate or substrate plus activator K+ were mixed with the enzyme, there was an initial increase in the rate of deamination of the sub- strate, 5'—AMP. They interpreted the time course of this change in rate to reflect the progress of activation by substrate or K+, in that the enzyme is initially inactive, and that the rate—limiting step in the reaction was the ac- tivation of the enzyme by substrate or K+, i.e., the activa- tion process involves a slow conformational transition from an inactive to an active form. The same authors (27) pro— posed a scheme consistent with their data and capable of explaining the sigmoid dependence of initial velocities versus substrate concentrations observed in steady state kinetics, without involving the concept of cooperative interaction between catalytic sites. Their scheme is represented as follows: : ““““ ‘ """""" “1 v 1 ka é ks kp 3 E + szx + szssx —-—>EX + P k k -a —s + where X is 5'—AMP, H+, K or other activators; k k and s’ -s kp are assumed to be fast compared to ka and k-a' They further proposed that their scheme, like the schemes pro- posed by Rabin (AA) and Weber (126) [see Table 1.1], might 36 explain sigmoid velocity-substrate curves, even with only one catalytic site per enzyme molecule, and that the scheme is able to account for the time course of product formation even under conditions far from steady—state situations, as observed with AMP-aminohydrolase at high concentrations. The authors (27) analyzed data via their proposed scheme when K+-free enzyme was mixed with substrate con- taining saturating levels of K+. Hence both substrate and K+ were simultaneously activating the enzyme, and activa- tion effects due to substrate alone were not analyzed via the scheme. Later, Ellis and Sturtevant (28), also using stopped- flow methods, studied the activation of AMP-aminohydrolase by 5'-AMP in the absence of monovalent cation activators. They reported that it was possible to interpret their data in terms of the Monod gt gt. model (20). Although they employed a wide range of initial substrate concentrations, including levels well above Km, their conclusions were probably incorrect, since they did not take into account substrate depletion in their analysis. Furthermore, they fitted their data to the Monod gt gt. model with the assump- tion that the enzyme was composed of six identical sub- units; whereas in fact the enzyme has been shown to be tetrameric (60,116). Ashby and Frieden (6A) attempted to analyze the full time course of the catalytic reaction, in the absence of 37 monovalent cation activators, by computer simulation, using mechanisms which assumed substrate depletion. They reported (6A) that none of the mechanisms attempted would consistently fit the data as a function of different initial substrate concentrations. The reason for this, they explained, was that in the absence of K+, their data indicated that one of the products of the reaction, IMP, was substantially activating the enzyme. No further studies on the mechanism of homotropic ac- tivation of AMP-aminohydrolase, in the absence of mono- valent cation activators, have been reported so far in the literature. CHAPTER 3 INSTRUMENTATION AND DATA ANALYSIS A. Stopped—Flow Studies The stopped-flow method was introduced by Roughton (l) in 193A, and greatly improved by Chance (2) some six years later. Further modifications were made by Gibson (3), Sturtevant (A), Dye (5), Dye and Feldman (6), Dewald gt gt. (7), and others (8—13). The technique is one of several fast kinetic methods available to investigators involved in the study of enzyme mechanisms. Essentially the method involves the rapid (a few milliseconds) and efficient mix— ing of two reactants, stopping the flow, and at a convenient point after mixing, the progress of the reaction is moni— tored spectrophotometrically. Modern stOpped-flow systems are generally computer, interfaced for data acquisition. Whether data are col- lected in a scanning or fixed wavelength mode, instrumenta- tion can be complex. In order to extract useful as well as accurate information from such a system, several charac- teristics have to be either known, or calibrated prior to each experiment performed. A thermostated, computer-interfaced, double beam 38 39 scanning stopped-flow system built by Guan—Huei Ho (1A) was used in this research. The entire flow system is made up of inert material such that reagents contact only Kel- F, quartz (mixing and observation cell), polypropylene and Teflon tubing. The entire flow system, including reagent reservoirs and pushing syringes, is housed inside a water-tight five gallon bath which permits circulating water to be pumped from the bottom to the top of the bath, and allowing for tempera- ture controlled experiments. In absorbance measurements, flexible quartz fiber optics transmit light dispersed from the scanning monochromator through the water bath, to the observation cell window, and out to the photomultiplier tubes. The use of heavy- walled precision bore pushing and stopping syringes, with a small cross section (Trubore, I.D. = 0.396 inch, ACE Glass Inc., Vineland, NJ) helps reduce the volume of solutions re— quired per push to about 0.25 ml for each of the two re- actants. To insure complete mixing of reagents prior to observation, a quartz double four-jet mixing chamber is used. The sample and reference cells are constructed from precision bore quartz capillary tubing, and each provided with two path length options (0.2 cm and 1.86 cm). The mixing chamber is fused directly to the entrance of the sample cell. Details on the construction of the ob- servation cells and the mixing system are found in Guan- Huei Ho‘s dissertation (1A). A0 Some of the essential characteristics of this stopped- flow system are as follows: A.l. Scanning Mode A Perkin-Elmer Model 108 Scanning Monochromator is able to repeatedly scan from 3 to 150 complete spectra per sec- ond, in the 220 to 1200 nanometer (nm) range, using the ap- propriate photomultiplier tubes (PMT). The scanning mode advantages are three-fold. First, precious enzyme is con— served, since a complete spectrum is scanned repeatedly in one push. The need for several fixed-wavelength mode pushes at different wavelengths is thus eliminated. Second, it minimizes reproducibility problems associated with per- formance of the same experiment at different wavelengths. Third, the problems of long-time baseline drift are mini- mized. The disadvantages in the scanning mode include noise introduced by the rotating (nutating) mirror of the monochromator, and the inability to follow reactions whose half-lives are less than about 15 milliseconds. A.2. Fixed Wavelength Mode Both the disadvantages associated with the scanning mode are eliminated in the fixed wavelength mode. The study of reactions whose half—lives are in the order of 5 milliseconds is possible. In this mode, the progress A1 of a reaction can be followed at any desired wavelength simply by setting the scanning monochromator to that par- ticular wavelength. The absence of noise introduced by the rotating mirror results in the enhancement of the signal- to-noise ratio (S/N). A.3. Flow Velocity The flow of solutions should be fast enough to insure turbulence which is necessary for efficient mixing. The flow velocity can be calculated by measuring the flow time, which is the time required for the stopping syringe to travel the distance between the start flag and the stop flag. A measure of turbulent flow is the Reynold's number (R): = B R (d)(v)(n) where (d) is the diameter of the tube in (cm); (v), the flow velocity in (cm sec—l); (p), the density of the medium, and (n), the viscosity of the medium, for water (n) is 1.002 1 sec-l). poise (gm cm- Figure (3.1) shows the velocity profile of the aqueous solution as a function of time, at 35 pounds per square inch (psi). For the 2 mm I.D. tubes used, the calculated value of 556.A cm/sec (corresponding to 17.A8 ml/sec) is A2 609‘ 500) M van!“ ‘WM- Gmukag 3901 deal ””1 V 1 40 milliseconds Figure 3.1. Flow Velocity Profile. A3 well above the velocity of 200 cm/sec required for tur- bulent flow. A.A. Dead Time (tdt The dead time of a stopped-flow system is defined as the time required for a solution to be transferred from the mixing point to the observation point. It can be estimated by the equation, where (V), the dead volume in (ml), is the volume from the mixing point to the end of the observation cell window; and (v), is the average flow velocity in ml/sec. The dead volume was computed by Ho (1A) to be 0.059 ml for the short path length, and 0.1A6 ml for the long path length. This resulted in a dead time of 3.38 msec and 8.35 msec for the short and long path lengths respectively. A.5. Mixing Efficiency The mixing efficiency of the double four—jet mixer was tested by using stoichiometric amounts of NaOH and HCl. 0.01 M NaOH with phenolphthalein, and an absorbance of 0.83 at 536 nm, was pushed against 0.01 M HCl (colorless). Data collection was triggered with the start flag, which initiates AA data collection before the flow stops. The resultant solu- tion had a constant absorbance of 0.25 at 536 nm, during and after the flow had stopped. Since protonation reac- tions of acid-base indicators are diffusion-controlled, the color change of the phenolphthalein should be complete within the 8.35 msec dead time of the system. Figure (3.2a) shows the spectral changes which occurred upon mixing the acid and base solutions with phenolphthalein as the in- dicator. Figure (3.2.b) shows the time progress of the absorbance at 536 nm collected in the fixed wavelength mode. Figure (3.2c) is a lOO-fold expansion of Figure (3.2b). The constancy of the absorbance as shown in the last two figures clearly indicates complete mixing was achieved before the arrival of the solutions at the obser- vation point. A.6. Stqpping Time Stgpping time is defined as the time required for the mixed solutions to come to complete rest, after the flow has been stopped. It was obtained by Ho (1A) who de- termined its value by studying the fast reaction, + 2+ (0.02M)Fe3 + (0.02M)CNS- + FeNCS in 0.2M H010“, and monitoring the absorbance changes at A56 nm. He found the stopping time to be reproducible A5 . (loo/I) (C) Jeamds (III) (5)__ 4m 435 536 (N M) (a) Spectral changes upon mixing 0.01 M HCl and 0.01 M NaOH with phenolphthalein as indicator. (b) Time progress of absorbance at 536 nm col— lected in the fixed wavelength mode. (c) 100—Fold expansion of (3.2b). Figure 3.2. Mixing Efficiency Spectra. A6 and less than 0.5 msec under a pushing pressure of 50 psi. More complete mechanical, electronic, and computer- interfacing details are found in the doctoral disserta- tions of Guan-Huei Ho (1A), Nicholas Papadakis (15), and Richard B. Coolen (16). B. Data Collection and Calibration A PDP8/I computer interfaced to the stopped-flow system was used for data acquisition. The raw data collected were subsequently stored on floppy disks for later calibration and analysis. Data stored on floppy disks can also be retrieved and displayed on a Tektronix Model 610 storage display scope, connected to a Tektronix Model A601 hard- copy unit. Figures (3.2a, b and c) are photocopies of out- put from the hard copy unit. This capability in the system allows for great flexibility during the performance of experiments, in that it gives the advantage for one to examine results from an experiment immediately after data collection is ended, or anytime thereafter. Data stored on floppy disks were then transferred to the Michigan State University CYBER7000 computer. Cali- bration of raw data was performed as described in the Ph.D. dissertation of Robert N. Cochran (l7). A7 C. Data Anatygis An existing general non-linear curvefitting computer program KINFITA was used for all data calculations. KIN- FITA is a modified version of program KINFIT originally developed by Dye and Nicely (18). Individual or multiple progress curves were fitted to derived equations in order to extract rate and equilibrium constants. Models proposed for a reaction were first tested by simulation with KINFITA. When the simulated progress curves followed the progress of the experimental data, the program was then used to fit the actual data to the derived rate equations, in order to find the "best" parameters to fit the experimental results. D. Computer Graphics Two dimensional computer plots were obtained from two sources; the plotting subroutine of KINFITA on the CYBER 7000, and the Program MULPLT on a PDP-ll computer in the Department of Chemistry. The latter program was developed by Dr. Tom Atkinson at Michigan State University. Three dimensional plots were obtained using Program GEOSYS. It is available on the Michigan State University CYBER 7000. CHAPTER A EXPERIMENTAL METHODS — AMP-AMINOHYDROLASE A. Materials Frozen rabbit skeletal muscle, Type 1, deboned, was purchased from Pel-Freez Biological, Inc., Rogers, Arkansas 72756. 5'-AMP, IMP, MES (anhydrous, 2[N-morpholino]ethane sulfonic acid), TRIS (TRIZMA base, TRIS (Hydroxymethyl) aminoethane), 2-mercaptoethanol(2-hydroxyethyl-mercaptan; -mercaptoethanol), SDS (sodium dodecylsulfate), acryl— amide, and ammonium persulfate were all purchased from the Sigma Chemical Company, St. Louis, Missouri. Bisacryl- amide (N,N'-methylenebisacrylamide) was purchased from Canalco. Cellulose phosphate was purchased from Whatman, and (CH3)uNCl (Tetramethylammonium chloride) from Aldrich Chemicals. Distilled water was passed through a mixed ion exchange resin bed prior to use. B. Preparation of AMP-aminohydrolase The enzyme was prepared according to the method of Smiley et al. (115). Five hundred grams of partially thawed rabbit skeletal muscle were cut into small pieces A8 A9 and placed, along with 2 liters of phosphate buffer at pH 6.5, in a large Waring Blendor. The mixture was homog- enized for 15 seconds at high speeds, and the process repeated every 15 minutes for a total of one hour. The resulting slurry was transferred to polyethylene bottles and centrifuged at high speeds (about 7000 x g) for 15 minutes, in a refrigerated (A°C) centrifuge. The super— natant fraction from each bottle was poured through two layers of cheese cloth to remove lipid particles. All succeeding steps were performed at room temperature. About 5 grams (dry weight) of treated cellulose phos- phate was added to the pooled extract. The suspension was stirred for 30 minutes after the cellulose phosphate was allowed to settle. At least 90% of the enzyme was bound to the cellulose phosphate at this stage. The reddish supernatant solution was siphoned off and discarded. The remaining cellulose phosphate slurry was then transferred to a sintered glass suction filter where it was washed repeatedly with approximately two liters of extraction buffer in 200-300 ml portions. During this process, the cellulose phosphate pad was not allowed to become dry. It was then washed with 1 liter of 0.A5 M KCl adjusted to pH 6.5 with 50 mM MES-TRIS buffer, again in 200-300 ml portions. This salt solution and all following solutions contained 1 mM 2-mercaptoethanol. The cellulose phosphate was transferred to a (2 x 10 cm) polyethylene column, and 50 the enzyme eluted with l M KCl adjusted to pH 6.5 with 50 mM MES-TRIS buffer. Each fraction was assayed for protein at 280 nm; e (280 nm) = 0.913 ml/cm-mg,[from Zielke and Suelter (165D. Fractions with readings below one absorbance unit were dis— carded. Those with readings higher than one absorbance unit were pooled into a screw-capped polyethylene tube, and purged with nitrogen gas prior to closure. About 50 milligrams (mg) of pure enzyme were obtained per 500 grams of muscle, with an average enzyme concentra- tion of between 2.5 and 3.5 mg/ml. An average value of 1350 micromoles 5'-AMP deaminated per minute per milligram of enzyme was obtained for the specific activity. The difference in the extinction coefficient of 5'-AMP and IMP, As, is equal to 0.12 x 103M'1cm'l at 290 nm, 30°C and pH 6.5 (27)- Related to the specific activity, S.A., the turnover number, T.N., is defined here as the number of 5'—AMP molecules deaminated per active site of enzyme per second. Hence, at saturation conditions (S >> Km) for AMP-amino- hydrolase, with a molecular weight of 278,000 (116) and four subunits, one S.A. unit is equivalent to 1.16 T.N. units. To remove K+ ions, the enzyme solution was routinely dialyzed against 0.5 M (CH3)uNCl, which contained 50 mM MES-TRIS buffer at pH 6.5 and 1 mM 2-mercaptoethanol. The 51 final solution, designated as K+—free enzyme throughout this present work, contained 10 uM or less K+. C. SDS Gel Electrgphoresis In order to investigate the purity of the enzyme pre- pared according to the method described above, SDS gel electrophoresis experiments were conducted on several en- zyme preparations. Enzyme isolated from newly purchased (one month or less) frozen rabbit muscle gave only one band when analyzed in 7% SDS gel electrophoresis. This confirms the findings of Smiley gt gt. (115). However, enzyme isolated from frozen rabbit muscle which was about six months old, yielded two additional minor bands. Both minor bands constituted about 10% of the total absorbance. However, the 8.A. of the enzyme remained high, ranging from l250-lA00. Several enzyme preparations from this same muscle were treated via a modified process (as recommended by Professor C. H. Suelter, Department of Biochemistry, Michigan State University, East Lansing, Michigan) in order to remove these minor impurities. Instead of eluting the enzyme from cellulose phosphate with 1 M KCl, it was eluted with a linear gradient between 0.A5 M and 1 M KCl. However, the contaminants were not removed. 52 Another batch was processed through two cellulose phos- phate columns. 1 M KCl was used to elute the enzyme from the first column, and then a linear gradient between 0.A5 M to 1.0 M KCl was used to elute it from the second column. The contaminating impurities were still evident, but to a lesser degree. Still another batch was processed through three columns of cellulose phosphate. The enzyme solutions from each column were eluted successively with l M KCl; a gradient between 0.A5 M and 1.0 M KCl; and finally, a gradient be- tween o.As M KCl and 0.u5 M KCl with 50 mM K+ pyrophos- phate. This process yielded one band in SDS gel electro- phoresis. D. Fluorescence Experiment Two AMP—aminohydrolase solutions were prepared. One contained 0.97 mg/ml enzyme in 0.5 M KCl, the other 1.2 mg/ml enzyme in 0.5 M (CH3)uNCl. Both solutions also con- tained 50 mM MES-TRIS buffer at pH 6.5. After adjusting the spectrofluorometer for maximum emission, the spectrum of 1 ml of each sample was recorded. Both samples were maximally excited at 305:2 nm, and emitted at 36Ai2 nm. 0.5 ml of 5 mM 5'-AMP was added to each of the above enzyme samples, and their spectra recorded. ‘v EfiY-u -"_E. r“-' 'n'.’ 53 E. Analytical Ultracentrifuge Experiments Sedimentation velocity experiments were performed on samples of enzyme which contained either KCl or (CH3)uNCl. Prior to these experiments, SDS gel electrophoresis performed on the same enzyme batch indicated the presence of one band. The specific activity of the enzyme in KCl was 150A, and that in (CH3)uNCl was 1A52. Three different enzyme concentrations (1.8, 0.9 and 0.7 uM) were examined. At each concentration, two samples were run in parallel, one contained 0.5 M KCl, the other 0.5 M (CH3)uNCl. Both contained 50 mM MES—TRIS buffer at pH 6.5. The ultracentrifuge was run at 32,000 rpm and 20°C. Two wavelengths, 260 and 280 nm were used to monitor the progress of the sedimentation velocity profile. F. Stppped-Flow Experiments Unless otherwise indicated, all stopped-flow experi- ments on AMP—aminohydrolase were conducted in the presence of 0.5 M (CH3)uNCl, 50mM MES-TRIS at pH 6.5, and 22°C. It is also important to note here that in all the stopped-flow experiments, concentrations of enzyme and substrate re- ported are the final values after mixing. This is because upon pushing enzyme against substrate, the final volume of the solution exactly doubles. 5A F.l. Scanning Experiments Scanning experiments were performed to determine if intermediates could be detected during the reaction of K+- free enzyme with 5'-AMP. The wavelength region scanned was from about 230 to 330 nm. Several combinations of enzyme and substrate concentra— tions were used. For the long path cell (1.86 cm), concen- trations of enzyme ranged from 0.36 to A.3 uM, while sub- strate values ranged from 0.05 to 5 mM. At 0.5 to 5 mM levels of 5'-AMP, only the region from about 285 to 330 nm was scanned, due to the high absorbances of 5'-AMP and IMP at the lower wavelengths. For the short path cell (0.2 cm), concentrations of enzyme ranged from 1.8 to 5 micromolar, and substrate levels from 0.05 to 0.5 mM. At these levels of 5'-AMP it was possible to record the complete spectrum from about 230 to 330 nm. Scanning pushes were performed at 2°, 12° and 22°C. F.2. Fixed Wavelength Experiments F.2.l. Temperature Studies Experiments were performed at 2°, 12° and 22°C. All initial enzyme solutions contained 1 M (CH3)uNCl, and 50 mM MES-TRIS buffer at pH 6.5. All initial substrate solutions contained 50 mM MES-TRIS buffer at pH 6.5. 55 Upon mixing the enzyme and substrate solutions in the stop- ped-flow system, the final (CH3)uNCl concentration was 0.5 M, whereas the MES—TRIS concentration remained unchanged. The resultant ttpgt enzyme and substrate concentrations were hence half the initial. In each of the reactions performed at the three tempera— tures, five ttpgt enzyme concentrations were employed: 3.6, 1.8, 0.9, 0.A5, and 0.225 “M; while the ttpgt substrate concentrations used were: 5.0, 2.5, 1.25, 0.63 and 0.31 mM. Each of the enzyme concentrations was pushed against all the five substrate concentrations. The highest and lowest concentrations of enzyme and sub- strate were chosen such that the preinduction time lags were completely covered, with minimal loss in signal to noise ratio (S/N). In other words, the maximum enzyme concentration (3.6 uM) when pushed against 5 mM substrate, displayed no detectable lag; whereas the minimum enzyme concentration (0.225 pM) versus the minimum substrate concentration of 0.3125 mM, produced a very large lag, but also resulted in a total absorbance change (substrate con- verted to product), that still had a high signal to noise ratio of more than 30. Obviously, the combination of lowest enzyme and substrate concentrations should present the lowest S/N ratios. This is because the rate of product formation, and the total absorbance change would be at their lowest values. 56 F.2.2. Preincubation Studies Unless otherwise indicated, all experiments were conducted in 0.5 M (CH3)uNCl, 50 mM MES-TRIS (pH 6.5), and at 22°C. Enzyme at a concentration of 0.A5 uM was preincubated with either 5'-AMP, IMP or NH; (each at either 2.5 mM or 0.625 mM), prior to reaction with 1.25 mM 5'-AMP. For com- parison, two additional enzyme samples were pushed against 1.25 mM 5'-AMP; one sample had no preincubated material, while the other had 0.5 M KCl instead of 0.5 M (CH3)uNC1. CHAPTER 5 RESULTS AND DISCUSSION - AMP-AMINOHYDROLASE A. Intermediates The Michaelis-Menten (M-M) rate law is often used to describe the overall rate of an enzyme catalyzed reaction. However, the detailed mechanism of an enzymatic reaction is generally quite complex. Steady state methods are fre- quently employed, and are often helpful in predicting the general pathway of the reaction. But as the number of in- dependent rate constants increases in a proposed mechanism, it becomes increasingly difficult to verify reliably the magnitudes of these constants without additional experimental evidence. A good "handle" often used to verify a mechanism, or part of a mechanism, is the study of transient inter- mediates detected during the course of a catalytic re- action. Detecting intermediates in enzyme catalyzed reactions involves the premise that during the course of a reaction, some groups on the enzyme, enzyme-substrate complex, or substrate, undergo changes in their environment in going from one form to the next. Techniques which can be used to monitor such changes include stopped-flow, UV-VIS 57 58 difference spectra, luminescence, ORD and CD. Transient intermediates fall into one of two categories, on or off enzyme. Off-enzyme intermediates have kinetics that are independent of enzyme concentration, and while they may yield some information about the reaction, they generally do not provide useful data in predicting a mechanism for the formation and interaction of the enzyme- substrate complex. On the other hand, on-enzyme inter- mediates are dependent on enzyme concentration, and are thus useful in mapping a more detailed and verifiable mechanism for the catalytic action of an enzyme. In order to detect such on-enzyme intermediates for enzymatic reactions which involve only enzyme and sub- strate, with no added cofactors, as with AMP-aminohydrolase, several factors should generally be satisfied. Among them are: (1) high concentrations of enzyme should be used, since the enzyme spectrum must be recorded, (2) the enzyme- substrate complex should have a low dissociation constant, generally in the order of a fraction of millimolar or less, (3) the substrate concentration should be high enough to insure as complete a saturation of the enzymatic sites as possible, and (A) the spectrum of the enzyme, or the enzyme-substrate complex must be at wavelengths other than those of either substrate or product. Figure (5.1) shows the spectra of AMP-aminohydrolase, 5'-AMP and IMP, and Figure (5.2) shows a scanning spectrum of 3.A uM AMP- aminohydrolase versus 0.5 mM 5'-AMP. 59 awn a: -Erafiill- - . {can - 1LT: till I o u C - o I . ~ A I . . . . K ‘ , . 9 , . O a . l- ;‘-i , 354nm: "$415”; ; "i‘i‘i—T‘““ ; g i .. i: ' s g i . M '53+5.K ‘ékaxh i i l ;l _ _ _ 1 I _ . .. .----_11_1- I I ....L... .. nocrnnomn< . _ . . . '0.0 Wavelength (nm) 260 270 i 250 240 Absorption spectra of 0.053 an AM? and 9054 m“! IMP in 0.05 H Tris-cacodylatc pH 6.5. IMP and AMP-Amino— Static Spectra of 5'-AMP, hydrolase. Figure 5.1. 60 [.50- I‘l l I ll"I WA aw. LOO- ‘ '1': g : I 3 In" 2 I" ‘ I O C Smw- I O < I I ' I '00! I I I .152 1‘0 281 (NM) (Reaction medium: 0.5 M (CH3)uNCl; 50 mM MES-TRIS, pH 6.5; 22°C.) Figure 5.2. Scanning Spectrum of 3.A nM AMP-Aminohydro- lase with 0.5 mM 5'—AMP. 61 A.l. Fluorescence Study Since neither 5'-AMP nor IMP, possesses a fluorescence spectrum whereas AMP-aminohydrolase does, an experiment was performed to investigate the fluorescence properties of the enzyme in the presence and absence of K+ and 5'- AMP, in order to assess the feasibility of conducting stopped-flow studies in the fluorescence mode. The two enzyme samples both contained 50 mM MES-TRIS buffer at pH 6.5, except that one sample was in 0.5 M KCl and the other in 0.5 M (CH3)uNCl. As can be seen in Figure 5.3, the overall shapes of both emission spectra are the same. Figure 5.A shows the excitation and emission spectra of the enzyme in 0.5 M (CH3)uNCl. The difference in the absolute intensity is due to the difference in the enzyme concentration of the samples. The results clearly indi- cate that the fluorescence characteristics of active and inactive AMP-aminohydrolase are the same; hence, it ap— peared unlikely that any fluorescent intermediates would be detected during the course of the reaction. A.2. UV-VIS Scanning Experiments Scanning stopped—flow experiments for the reaction of K+-free AMP-aminohydrolase with 5'—AMP were performed in order to determine if any transient intermediates could be detected during the course of enzyme activation 62 - in) Figure 5.3. Fluorescence Spectra of AMP-Aminohydrolase in 0.5 M (CH3)uNCl and 0.5 M KCl. «19! ’ i :00 1.. mung-n (an) q.- Figure 5.A. Excitation and Emission Spectra of AMP-Amino- hydrolase in 0.5 M (CH3)uNCl. 63 by substrate. Each subunit of the enzyme contains 33 residues of phenylalanine, 5 residues of tryptophan and 30 residues of tyrosine. All three amino acids absorb in the UV, with phenylalanine having a maximum around 260 nm; tryptophan and tyrosine around 280 nm. Hence these amino acids are probably the major constituents responsible for the enzyme spectrum at these wavelengths. The assumption was that if any spectral changes occur around these wavelengths, during the activation of the enzyme, they should be due to changes in the environment around these three amino acid residues. This would then indicate changes in the enzyme conformation from the in- active to the active form, during the course of activation. More specifically, one or more of these amino acids would then have to be present in the vicinity of the active and/or activator sites of the enzyme, since these sites should undergo the most changes when the enzyme undergoes a conformational change. In all cases examined, as described in Chapter A, Section (F.l), the only spectral changes observed were those due to substrate depletion with concurrent product growth (Figure 5.2). No further attempts were made to search for intermediates. 6A B. Analytical Ultracentrifuge Studies Sedimentation velocity studies were conducted in order to investigate three points. The first was to ascertain the purity of the K+ activated enzyme, the results of which would complement those obtained from gel electro- phoresis measurements conducted earlier on the same enzyme preparation which showed only one band. The second was to determine if the K+-free enzyme, present in (CH3)uNCl, undergoes any subunit dissociation, and if so, to measure the extent of this dissociation, and thus answer the ques- tion as to whether activation of the enzyme by monovalent cations might be associated with a change in the aggrega- tion state of the enzyme. The third was to examine whether the constitution of the enzyme is concentration dependent. Results from the last two points would then be used in the formulation of a full—time course model, which would en- compass activation of the enzyme by substrate, and include or preclude subunit association-dissociation phenomena. Three different enzyme concentrations (1.8, 0.9 and 0.7 uM) were used to determine if subunit dissociation was concentration dependent. Examination of the spectra revealed that the sedimen- tation velocity profiles were similar in all cases, and indicate no major variations from one sample to the next. The spectra for the K+-activated enzyme showed gpg moving boundary, which indicated the absence of contaminants, 65 and also that the active enzyme existed in the tetrameric configuration, as expected, in the presence of monovalent cation activators (115). Figures 5.5 and 5.6 show the sedimentation velocity spectra of 1.8 uM AMP-aminohydrolase in 0.5 M KCl, at 280 and 260 nm, respectively. When the K+-free enzyme results were compared to those of enzyme with K+, it was found that all the spectra were quite similar in their profiles for all the three concen- trations, except that the spectra for the K+-free samples had a slightly broader boundary than those for the enzyme samples with K+. For example, compare Figure 5.5 to 5.7, and Figure 5.6 to 5.8. This slightly broader boundary may be the result of one or two factors. The first factor could be a viscosity effect on the sedimentation, since the vis- cosity of 0.5 M KCl at 20°C is about 0.99 poise (relative to water at 1.00), while the viscosity of 0.5 M (CH3)uN01 at 20°C is about 1.15 poise (gm cm’lsec_l). Thus, the protein molecules in the 0.5 M (CH3)uNCl solution would experience slightly greater drag in their migration, which would then result in a slightly broader boundary than that manifested by those in the 0.5 M KCl solution. The second factor for the broadening in the 0.5 M (CH3)uNCl solution could be the result of some minor subunit dissociation of the en- zyme, but not to the extent that one could readily measure for these conditions at 20°C. Sedimentation coefficients at 20°C in aqueous solution 66 I- ...-n ,- - : --I- I iii» - H II I- LI 2:... I I II ...... b 0.011.. I. . II .133: 331.231.“... / I - I III .I-I, H- .. H I... Hm...- -I.II-H--I 1 -.III -_.I III _ a _ H. m l M - I . II I- OI- . H In wh-H L u H . w W IITI I II...III.I I I II III mI. I I II II n , I- I.. | I- I I III III -I I .I-IFT “.111- ,I- III II III I -.---I|II II mIWII. I WIHIf-I II-mI I; I a I II. -I :I II I u,I.4III.III.I ,. 5IT+IJI I II.- I I . | I, II J. I I Via-I u ..L. ..., W - - - p_ I. I- ..I...I- - I» .1: 1 - - I- I I I I 'Néwmrs INF- . I l I I "‘.j§.‘““““““"“"""-_°‘_”‘°-‘"-““"‘ Sedimentation Velocity Spectrum of 1.8 uM P-Aminohydrolase in 0.5 M KCl at 280 nm. Figure 5.5. Sedimentation Velocity Spectrum of 1.8 uM AMP—Aminohydrolase in 0.5 M KCl at 260 nm. Figure 5.6. 67 (omuSA ‘ SCHILLER PARK, ILL I \n I I I I I I I I I I I rIINom-II-III:E ........ I. II .- Tweak-3313.. Vitae.- .. I - I I I ..l...- ...III- III- I. I-.- h. -..II : IL. I L I I I . _ . . . _ . , . III . IlldpllliI III I III... IIIVIIIIIIIWIII . IL I. .aIIWII .0! J IIIIH'IIL I A It. 1 , . II.I.I.I-I .I. II- . . .I I.... ....-.II:I I II I. .I ..III..... iq ,IIIII . . -.fi- II, I I- I I+ .-- uInI I--.I-I I I I II I I IIII I II. : IT-.... . — I 7+- I I I I I I Y ‘ T I I I I I I I ECKMAN INSTRUMENTS INC. I 0 O o I I . ~+—4 .... . ... . ‘ I I I I I .....W... I 4. ...» I I I . I l I ...;fi~.,.-.o.... ,' -9 __.g - I " I Sedimentation Velocity Spectrum of 1.8 uM AMP- Aminohydrolase in 0.5 M (CH3)uNCl at 280 nm. . I I I I I I . II Figure 5.7. -..._-.._.-.-...—...--... . . .- Sedimentation Velocity Spectrum of 1.8 uM AMP- Aminohydrolase in 0.5 M (CH3)MNCl at 260 nm. Figure 5.8. 68 (S ) were calculated for all the samples, and the re- 20,w sults are shown in Table 5.1. The 820 w values shown are 3 in Svedberg units. Again the values of S20 w in the table 3 clearly indicate that all samples were sedimenting in an equivalent fashion. Table 5.1. Sedimentation Coefficients for AMP—Aminohydro- lase at 20°C. 820 w values Calculated from Sedimentation Velocity Profiles at 280 nm. The Ultracentrifuge Velocity was 32,000 rpm. S20,w (Svedbergs) Enzyme Conc. (uM) In 0.5M (CH3)uNCl in 0.5M KCl 0.7 10.0 10.“ 0.9 10.0 10.7 1.8 9.2 10.U The S2O w values shown in Table 5.1 compare favorably with those obtained by Smiley et al. (115), who reported a value of 11.5 S for their enzyme preparation. In summary, the sedimentation velocity results indicate that the composition of AMP-aminohydrolase is that of the tetramer, at 20°C, whether the enzyme is in 0.5 M KCl or 0.5 M (CH3)UN01, and that this composition is not concen- tration dependent down to 0.7 pM. 69 C. Preincubation Studies Stopped flow experiments were conducted to explore the effect of preincubating the components of the catalytic reaction with enzyme prior to reaction with substrate. 5'-AMP, IMP and NH1, were each premixed with AMP-amino- hydrolase, after which each mixture was reacted with 5'- AMP. The reactions were performed in 0.5 M (CH3)uNCl, 50 mM MES-TRIS buffer at pH 6.5, and at a temperature of 22°C. In addition, one enzyme sample contained no pre- incubating agent, while another contained 0.5 M KCl instead of 0.5 M (CH3)QNC1. The effect of preincubating NH; with enzyme is shown in Figure 5.9. As can be seen from the figure, NHZ at concentrations up to 2.5 mM has little or no inhibitory effect on the reaction progress. Calculated values of (dP/dt) for the steady-state region, corresponding to a maximum rate in product formation, are shown in Table 5.2. The effects of IMF, 5'—AMP, and K+, are shown in Figure 5.10. Table 5.2 shows the calculated values of (dP/dt) for the steady-state region. IMP with concentrations up to 2.5 mM has essentially no effect on the steady-state region. However, the lag is slightly removed by 0.625 mM IMF, and more so by 2.5 mM IMP. 0n the other hand, 0.625 mM 5'—AMP increases the rate of product release at the steady-state region by approximately 17%, and 2.5 mM 5'-AMP increases it by about 52%. Furthermore, the lag 70 Table 5.2. Preincubation Studies.l 0.U5IJM (dP/dt), Steady-State Region Enzyme Preincubated With ' (uM IMP/Second) nothing 845 0.625 mM NH: 777 2.5 mM NH: 778 0.625 mM IMP 837 2.5 mM IMP 836 0.625 mM 5'—AMP 990 2.5 mM 5'—AMP 1283 0.5 M KCl (NO (CH3)uNC1) . 2121 1All samples contained 0.5 M (CH3)uNCl (unless otherwise indicated), and 50 mM MES—TRIS buffer (pH 6.5). The reac- tions were conducted at 22°C. The AMP-aminohydrolase con— centration is 0.45 uM; 5'—AMP concentration is 1.25 mM. 71 .mmmfiopesnocfieQOCHE1 1 d 1 a J. L ... .0 co. m .OV .. m .3 2.. . as: -3: a... on ... o a: inqu :36qu ... fl ... v .... W o. 3 co. n “6 0. %:\$ 0.. o o o o o o O O 0 can“ 4 n6 814 .oomm pm .mucoommfiaafie com ow a: amz onpso map mmwmaopuzsosfis A8pso mpg mmmmflogchzocfis ommHOLUzcocHEmo3mCfiA .mm.m opsmfim 2 M-Oct‘d AMWV M ? 9O 16. I31 )x n” (4’ m an: 8. V 5&5(pg) (infirm) “0 Figure 5.23. Lineweaver-Burke Plot for 0.9 uM AMP-Amino- hydrolase versus 5 mM 5'-AMP, in 0.5 M KCl, at 22°C and pH 6.5. 91 shows the plot for a K+ activated AMP-aminohydrolase. Km was calculated from Figure 5.22 to be about 0.36 mM, while 1 the turnover number was about 7A3 sec- , while from Figure 5.23, Km was about 0.“ mM, and the turn-over number about 796 sec-l. D.3. Activation Energy The Arrhenius relationship was used to calculate the activation energy (Ea) for the reaction of AMP—aminohydro- lase with 5'-AMP. Turnover numbers calculated from four different temperatures (2°, 12°, 22° and 30°C) were used. Figure 5.2A, shows the Arrhenius plot of lnk versus l/T. The calculated value of Ba was 16,500 calories per mole. The turnover number at 30°C was calculated from an enzymatic reaction which contained 0.15 M KCl and 5 mM 5'-AMP, and assayed on a Beckman DU Spectrophotometer. The turnover numbers at the other three temperatures were cal- culated from the full time progress curves, obtained from the stopped-flow reactions of AMP—aminohydrolase pushed against 5 mM 5'-AMP, in 0.5 M (CH3)uNCl. 92 .ommaoppzcocfieSOCHEmImE< HQHQfiCH pom wumo "madmfim CH czocm mpazmom :BHmZHx mo pofim .zeHmsz ho mocomso>coo Lou Umpmoe muom mumalmcaz .H.o magma 101 for R2; Figures 6.3 through 6.11 show the KINFITA plots of the l 0.9 r 0.19 mM for Km; and 36u9 : 171 sec"1 for kp final results. Values of 0.9 mM for Km and 36U9 sec- for kp, are both higher than those calculated from experi- ment. With these nine-data sets, the "overall" fit is no longer as good as it was for the two data set. Compare Figure 6.1 with 6.3, and Figure 6.2 with 6.5. Such re- sults are not unexpected if the model does not adequately describe the experimental behavior of the system. Further refinements of Model A were attempted. The primary one involved using the product as an additional activator for the enzyme; however, these attempts did not improve upon the original model, and were in fact less systematic than Model A. B. The Monod, Wyman and Changeux Model (MWC) The MWC model is an attractive "classical" model which has been shown to describe the action of many biological systems, of which the most notable is the allosteric bind- ing of oxygen to the hemoglobin molecule (20). This concerted (all or none) model accounts for positive cooperativity in a very simple and elegant manner. One of the assumptions made by the MWC model is that the same conformational state is stabilized by a variety of ligands. This point can further add to what was presented earlier in this chapter, where it was argued that whether activated 102 Figure 6.3. Model A, Nine—Data Set Fitting; 0.225 uM AMP- Aminohydrolase with 0.3125 mM 5'-AMP, 22°C. Figure 6.A. Model A, Nine—Data Set Fitting; 0.225 uM AMP: Aminohydrolase with 1.25 mM 5'-AMP, 22°C. Figure 6.5. Model A, Nine-Data Set Fitting; 0.225 0M AMP- Aminohydrolase with 5.0 mM 5'-AMP, 22°C. Figure 6.A 103 JI‘I SI I! tv'FII I It- *5 I IIlolI (0 III'I'.II.VI[U( II In! LIII: .o‘~I-3: IIIUI II tut nlnul : .'-:(or: .|\(I[ar\l x .9sgtoaa .;- MI I' II:.II OI IIIID. IIL;I II Int I)? t .ICUl-IO uILuI II In! B .5! I-zc .I.CIL' (\I : . (-)2 I -¢uus la (IUlII'I~IIL IC|nl. 5 ’[Ok‘ I (IL(L[IIID rolul Is uSIn qus '(SIO Is PLOIIZOI x -(.us a» In!(A|‘IhlI\ Azo (AL(UIIl[o Iona: lit I5 ["1 5'1 3(III I I! LIIII v i....§-.-.5....5.-..‘....5..-.5....\-...-,....5-.--§...-',....5...-‘l....\....5..--\.-..5.n-.5..-.5-....‘i I I I ‘ I X 0 I '. ° " I I I 1 . ' . . l 3 'I I I I I 0 I I I I I I 5 3 I ‘I I I I I I I I I I l a I f. I I I I I I I I I 5 I .‘ I I I I I 0 I I I 5 5 I l I I I I I I o I . u I I l I 0 l : l I I I 3 " l l ‘ : . . I I 5 l .2 I I I I I I 0 I | :I I $=I 3 I....5-..-5..--,.-.-3....5....3....g....5.-..5..-.3.-.-5...-5.-..5....5...-5-...5-.-.5-...,....g-...-,| SIII It! HU‘IEO 7 I2153S Is III.|I ol IIIVII.|I u at :I tut It!!! . 3!(- -o:. volut II Int III"! I .25 IZ-I7.l~clln$ul : .zssf; II ~EI=I I I O I ) I 3 I I I I I D O I I r O 1 I l I 5 CI rl . 'p .-..§.-.-5..-.5. 5 g g g 5 .-g..-.5.-..5-..-5--.-5.-..5-...5....5....5..-.3.-..g....5I )IIB St! nu at! 9 I:ILSI I\ III.II OI III II “In ”I :If II Int LIIII I~I-°I.IIIu( II Int IIcuI I . H0 ( u:.Iu(It-I9I I .II‘I-OO .I-IIcag I' II). II o- ”l u. ALu II Iu( In» I JII IooI.vIIUI II In! .-4IIu .I%.I In .I'Aclllth .I -II | 'l‘k2 ‘2 (I "(II‘INIIL '3 lul. O “(I N! (ILLUIIIIO Io HI (out! a I! USID hnIR' Itslu IS 'LOIItDI I Ilus n (I '(III-ulu no (“01qu :0!!! II! I‘ III “II. )ILII I it OILII .00.,-...,-n.0,....,...O‘OOO0,00”,COO0,00IC’...a,...as....,....’oooos....‘oono’cooo’00-.‘0000’00003 0 J I I I I I I l l 0 I O I o 0 I I O I II I I 0 I I I I o l I N i 0 I I I_. I I l 0 I I 4 9 I I I I I .' ' : O I I IO ' = I J I I I0 I :8 '1 . 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 5| ;III st! nu 95 " .' ' C ..5 ' gt . ”(a 9 at l 9(000 I l‘s I. III I. “I II In I VIIU n I I o o - oE-II in: I”: I: 2.? on I “5:: Au 2‘" Inl‘Ib‘ I .iIaIt 3.” “In I In! N! Iol I JIu-OMI-‘IIII' ”I I oily"! I III as :1 [I OIIIIIIIOL Oilvl. I Izoqs I (I Cut-Ito oDII-I IonII I IS ustn wtn ltSIO I8 rIOII(n| I III hi In IDIIIIIDIIL IMO (ILCUIIIIu voIn III In Int .InI JILII I "I PILI I l ...-3 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 ‘ 2 ‘ ‘2--'°2---'3 0 O O I l I I I I I I I I . o I I O I O ’ I I I I O l . I | .. l I O I I '0 I I ' I I 0 I I I 5 'I I O I o I I I) I) 0 :9 I Ito : = = 2 g = 3 3 g g = g g 5 g 9....9....5 lO 5 106 Figure 6.9. Model A, Nine—Data Set Fitting; 3.60 uM AMP- Aminohydrolase with 0.3125 mM 5'-AMP, 22°C. Figure 6.10. Model A, Nine-Data Set Fitting; 3.60 uM AMP- Aminohydrolase with 1.25 mM 5'-AMP, 22°C. Figure 6.11. Model A, Nine-Data Set Fitting; 3.60 uM AMP- Aminohydrolase with 5.0 mM S'-AMP, 22°C. lO7 jIIII {II nur'Il I [P IIIII OI II:IIO.|I III”! II In. ‘llv- .l‘1I-Q.‘.'lIUIAIIIII .I :u' -. '. ' . . II:II'I l’ IIJ-II I" ‘II . :Ilul II In-I I'n' .Io.‘I.eI.vnIu a: In! "I.” , ’::,~.,‘?;-h':::::',‘: ’ ..;IH.:")'.°. ' 'V‘VI I" II'IIIIIIIII PSIII I -' ~I I 'IItuIIIru IoIlI Io'.[v : I. , a NO I - p ,. I Il|i$ Is (IDIII~I~III I~D (IllUIIIlD vol"! III I! lnl :III 3! III I or :I:II"I "ID l' l” ' 0' .-..1.-..5....\-...5.-..5..--s..-.s--.-s..--5.-..s.-.-5....5....5..--5-...5.-.-5...-5.-.-,...-3-...-I II I I I I I g I I.) o 0 N O O B I ° 5 3 I ' . . I I I . I u' Fi i gure . . - , I 6 o O ' I o ‘ l I o I ' s I O I . " I, o I 8 . l I CI 0; 'p'l 'l y 2 2 2 2 2 2----2-- -2 2 2 2 2 2 3 5 1 5....9....5. :III S(' “u III I I.I:;I II II |III CI II:IIIIII.III0( II' Inf I" 'II 83‘(- 0?. I510 ur II In( one"! I .I: 00: .luctt-III I .IO‘I'O. -£*'11IL IS II;- II 3‘ ilsl a. IILu' II . 3v 2 .ISOIIJIoIIIuI II Int no on I .lIII'II-Iuc-I-III I .IIOI-Jl I DIIas Ah (I'll HI-III I01~I 0 PI: WC III(0 vol-I Ionlv : I! usIo qu~ .(Slo I) Igolllol - ‘II\S I'. ("1' “I II“ “-0 (IICUIIIIO 'Ollul I( I. “II III-I .‘I III I 6' (III l....5.-..‘....5....5...-5....5....3..:.5..:.5-...5..--5....5...-I.....g.o-.g..-.5....g....g....g.-..I: I I I o O I I l a] I I I o I I I O I I o s I o I I 3 I I o. l 5 a I I l 1 I l I 2 5 I ° . Fi ure . I I I 5 H i I I I 6 10 I : O ‘I 0 u l I I l I I I 5 I I ° I I 3 D I I I I o I l 2 I OI I I .' I ' 1 l o- I I I o. I . 'I ...-.5...-5..-.,....5. - I, -, I, I, g -, 5 5 g 3 7, g.-..3..-.§....5..-.5; :III '.'I ')-"I 2 I.‘::' I‘ III.II or IIIvII.II: :IIyt II I-( gift! . II-o).vIIu II IvI IIcnI 2 II 7'-II.I~cII-(~I : .IIII-II OIII|(II II .II cl Ilslc. IIU. II In! '3' 2 .Ilzl- :II.IIIUI I Int IuIIo- : .IIII00IIIIIII'IQI I .IIII-II l rtnms In [III-IIIIIII OOIII. I III": I (0; “II oln II IflhII 2 IS usIo autn IIIIo l5 'IOIIIOI x ntnns In (I'(II~IIIII two (I ICUIIIIO Io It I! la «I '.III OIIII I IV OIIII v ----2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2~- 2 2 '1 I I l I I l l I ' I O I . J I O I I o I Figure ' ' ' 6 ll - . l O I I I I ' . 0 I . 0 I | l 0 I . I i IO I. I I I '0 1:4. ..I-.--g - - . . l....5....9....3‘ 1.... ....1. ...5...-g....5- - .. -...3....5. --. .- , , , , , QCOOIODOOOIOIIOIOOOOII. lull':( OIII III'II. 'I~(" ILI$I I" I1 0' I‘I‘UI 108 by K+ or 5'- MP, the final catalytically active conforma- tion of AMP-aminohydrolase is the same. Ellis and Sturtevant (28) reported that they were able to fit their experimental data, for activation of AMP- aminohydrolase by substrate, to the MWC model. However, as mentioned earlier, they assumed the enzyme to be composed of six subunits, and also neglected to account for sub- strate depletion in their analysis. In this present work, it was felt that the MWC model merited further investiga- tion, but with the "correct" assumption that AMP-amino- hydrolase is tetrameric, and that substrate depletion be accounted for during the entire course of the reaction. The simplest MWC scheme with positive cooperativity involves one active, and one activator site per subunit. This was adopted here as the working model for AMP-amino- hydrolase. Therefore, a tetrameric enzyme such as AMP—aminohydro- lase would have the MWC scheme shown in Figure 6.12 (page 109): In Figure 6.12, k's represent rate constants; K's, dis- sociation equilibrium constants; E, inactive enzyme; E', active enzyme; 8, substrate; ESn, in active enzyme-substrate complex; E'Sn, active enzyme substrate complex, with n = 0,4. It is assumed that only the active enzyme manifold can process substrate to product. The total "active" enzyme uS+E MK ES“ Figure 6.12. 109 k0 \ ‘7i E' + us k-o A iE-K' W kl \ ‘— E'S + 38 k—o 4h 2 . —3—K v k2 \ E'S + 23 ‘ 2 k-o Ah 3 . —2—K k w 3 \ E'S +s ‘ 3 k-o Ah 1W W kn \ E'S \ u k-o MWC Model for AMP-aminohydrolase. llO concentration, E6, is: E6 = E! + E's + E'S2 + E'S3 + E'Su (l) The total "inactive" enzyme concentration, E0, is: E0 = E + ES + E82 + E83 + ES“ (2) Then, the initial enzyme concentration, (E0)t’ is: = v (E0)t 50 + E0 (3) Following enzyme activation by substrate is the Michaelis-Menten scheme: K m kp E6 + S c——— E68 ———9 E6 + P (u) where (Km) is the Michaelis-Menten constant; (kp), the turnover number; (S), the substrate and (P), the pro- duct. The rate of product formation is then (E'SiHS) (5) (LID. ff *0 II ”a? II M r: O The working equation for KINFITH (see derivation in 111 Appendix C) is: , _ -k't 21—? _ kp(EO)w(l e ) (6) dt (1+K /S) m where, k' = B + k—O (7) and (B)(E) , _ O t (E0)w ’ B+k_ (8) with B - k —————l+S/K' u (9) ‘ O 1+S/K and, s = 80 — p (10) Thus the familiar Michaelis-Menten equation for (dP/dt) is now modulated by (E6)m and an exponential. This model has six unknowns. To determine this many parameters, it was necessary to use the multiple data set 112 version of KINFITU from the start. Nine-data sets were thus simultaneously fitted to Equation (6), after the simulations, and the fittings of simulated data tests were carried out. The enzyme and substrate concentrations for these nine-data sets were the same ones used for Model A, and are listed in Table 6.1. One set of initial estimates of the unknowns, for which convergence occurred in KINFITN were: 1 x 10‘“ sec"1 1 K II k_O= 0.1 sec K' = 0.1 mM K = 1.0 mM K = 0.1 mM k = 1000 sec-1 Thus initially at equilibrium, when no substrate is pres- ent, the conformational change of the enzyme from inactive to active, was assumed to proceed at a very slow rate (k0); whereas the reverse rate (k_0) was assumed to be fast. The dissociation equilibrium constant for the active manifold (K') was calculated from the half-life of the activation, and was found to be about the same order of magnitude as Km. The dissociation equilibrium constant for the inactive manifold (K) was initially set to be lO-fold greater than (K'). (K) must necessarily be greater than (K') in order 113 for activation to occur. Cooperativity occurs as a result of the statistics imposed by the model. Results for the best convergence obtained were as follows: k0 = 0.18 x 10‘3 i 0.26 x 10'3 (sec-l) k_0= 0.95 x 10‘1 1 0.20 x 10‘1 (sec-l) K' = 0.077 i 0.039 (mM) K = 1.29 i 0.3M (mM) Km = 0.83 1 0.19 (mM) kp = 3066 i 162 (sec-l) Figures 6.13 through 6.21 show the KINFITA plots of the final results. As can be seen from the plots, the fits are not good for all the concentrations employed, indicating that while the (MWC) scheme used here is able to display the general trend of the experimental data, it is not ac- ceptable overall. Still, the (Km) value predicted by this model was in the same order of magnitude as the experi- mental (Km), while the turnover number (kp) predicted was the same as that obtained from experiment for the tetra- mer. Some refinements for this MWC scheme were attempted. The primary ones involved using the product as an addi- tional activator for the enzyme. However, the attempts proved inconclusive, and did not improve upon the original model. Figure 6.13. Figure 6.14. Figure 6.15. 114 MWC Model, Nine-Data Set Fitting; 0.225 uM AMP-Aminohydrolase with 0.3125 mM 5'-AMP, 22°C. mwc Model, Nine-Data Set Fitting; 0.225 UM AMP-Aminohydrolase with 1.25 mM 5'-AMP, 22°C. MWC Model, Nine-Data Set Fitting; 0.225 uM AMP-Aminohydrolase with 5.0 mM 5'-AMP, 22°C. 115 ‘3'] 'Il ~‘.- I ' x I . n I. |I co lII'l II. II..IIuI II In! LI'I’ 1!- -o; 'IIUI II In' 'Idul = .‘>-'~aJ.|t(II-I~I I .I‘IIIII I"|I'II I1 II ‘III a! 'I’ III. III.' ll In! tor I .‘l\: H}, II UI II IuI ufiIIII - .leI-;-.In'uq-(ul 1 .0’9'-:) 0 'Vl‘.‘, I'I IIDIII'I'IIIL P'II'II. 9 I In'. (IL’UIII ID '”I\ I‘D L' II LUI‘ IIII‘A '(IIB I! PLPIIZDI v 'IIH: :v II’I'I'IHIOL I‘D :aItuIIIIc :olnv III IV III 'I-I )IIII I ‘I 'IIII v i.no.5....g....q....g....g....g..--5-.--fi....1.-.-5-.--5...-5....g.o..5.-..§-...5..-.9.-.-5..-.s....I| I I I I I O I I o I . s I I I I I I I I I I 5 8 l 0 I I I I I I I I s I ‘ I o ' I l I Figure . I I O I 13 ‘ ' ‘ I I O I I I I I I . l I l I I I 0 I I \ V 1' ' ' . ! I 0 I ‘ C l ' ' I I I I I I s I o ' I I l I I I ' I 0 I I ' o ‘ g l I I o I I o I I IIO I \--r ' I.L.-5....5.-..g..-.5....9....g....§-...5....9....5.-..5..-.5.---5---.3.-.-5....5....g-...9...-\-...5| :III va 'J. vI .rkl-s I' IIIoII (I IIIIII.II.IILuI II IuI LIIII ¢)¢I Ill 1 II I» I . z * - -. ' . .t"IrII I' -I:.II 9- ”I 0. VII” InI I1» - .‘I?'-IO.:::VI :1 v I 0:! It:.! ,|;I:£gf?l.::::::I' ' ‘ o.;;,:g’°. ' 'I‘-‘ IV (""I'(N'll ‘JIQI 0 .IH$ 0 fl Id 0' 2 ’0 5' ”O I x ~(st an 'l’I'l‘ (hIIL awn (AL(ULIIID ralnI .5'1'": tIr l SILIA'E ‘V‘EI rut”! IIIJ " 'IflIlItl !....s....5.--.5..--‘-.-.5--..9-...5....:....§--.-9..-.s...-\....5..--5....‘....5.-..q..-.q....s..-u I I I I 2 I I I . , J l n 1 I I l ' 0 I I I I R I I ' I F' 1 ° igure . . 'I I 0 . 6.14 . I . '. ° ' I I I I I l I s o ‘ ' I I I I I Q I " I 0 I . I I I I I s I ‘ | n I I I I 1 n I I I I a o ” ' . g I | :I I fix: ‘ I‘ .' a. g, A. = z 3 3 I. g g I, g g -.3-...juooos....5..oos.c..QI Cal: 3'! uu-F'I I .c ... ,« ...... c- .1.va..|'.'l[ul II Iur III. .I. I:.Iouur II IuI II‘NI I .III Io :.IALII-¢~I I -I(’(°Il l"'I(l| " ,..~.I. (. .m u. “U! .l I"! I v ; .|3| ILI:|::ILUI II III( PO'IOI '.‘ .0: I.(' 0. II‘C.(‘(‘I I 0“: ”' ‘ -|“s ‘ ('3'. I“~'.L '3 ~' . utlns ‘ f‘ (Ut‘Ilo 'fllH' IO’IL' : I‘ U‘I" out” .($|° " "0"(°. I 'Inus I (I’I' I-(IIIL II CIItUIIIIO Ooln II II In( SIP I 0(lIl I II OIIII I .‘. l----I--~-s---~1----s--- 1 a 1 s 2 = , . -s----a----I~---s----I----s-o--s---~9' I l 9 3 o I l I I I I l l I C I I I z I Figure , l I 6.15 I I I I I I I I I I . l I I : I ' I i i ' ' I . g s - 5 I . I I . ‘ I ' I I I o I I I I I l I I II I I I: ' ?:O:Os..-O,OOOI,D.....I..,....‘OO.C‘O...,I...‘....,O.0.5....‘COOO,QODO’COIO,CI..,..O.,IO..’O.ODSOOOO,l Figure 6.16. Figure 6.17. Figure 6.18. 116 MWC Model, Nine—Data Set Fitting; 0.90 uM AMP-Aminohydrolase with 0.3125 mM 5'-AMP, 22°C. MWC Model, Nine—Data Set Fitting; 0.90 uM AMP—Aminohydrolase with 1.25 mM 5'-AMP, 22°C. MWC Model, Nine-Data Set Fitting; 0.90 uM AMP-Aminohydrolase with 5.0 mM 5'-AMP, 22°C. ll7 IIII {II ,--I I I5“ ' I‘ I«!.II @- II-II I.II. II I II Iur {(711 .3'l( cI ugt . I ’ I I ' VI 5' 'Il . I I; 'I ‘I I 'I- II F' 'I‘ I‘- U' I I~I II' - .IIrv-rl.vifuI II In I ;O,,$§", , .9;;I:3F-I:}:E:I:I~: ' ;;3?3§$" 3 '-“ ‘1 'I 'II I'III vats -I A : PI I r ' "-“I I" II‘I‘I’I‘UI m cucuuuo run ”I I.“ IEPIJINHIH': 3‘33“” "no I'. unvum I.-..q.-..I....-..-. --.. .-.. .... -..- ..-. .-.. --. - I . 5 3 S 5 5 5 S UJ.-..‘..-.,--.-‘.O..:QOI.s-OCO’OOOO,I.QO‘OOOQ‘| I 0 o I I ° ; O I ° 2 : 0 II 3 I I ' ‘ i 0 I I I I I ' I I . g o I Figure - ' . 2 \ 6 16 I ' I o I 3 I I I I L 1 I l I I I ' C I ‘ I a I o ' ' ‘ I . , I I . I I ' I I I ‘ I I I I ' ‘ o I I I I 3 ' I l c I I I I I 2 I I." 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I Is U.In qu' IIIIO II PIOIIIOI v IIIVS I! IIIIIl-I all L IV. {ILCUIIIID fflln I. INK 'III OII II I Q! oII I..--O.-..o‘c-oo‘.o..1..o~\...o\.---\--.os.-.-.Soooo‘ooou‘n-u-‘conosooo:CU0.-\-c-cscoo.\-...\oo-0‘--Oo:I l o I o o a I I I ' 'I l 0 I I I I l l I ‘ I I I l O l i ' I 5 I I I I N 5 o | l I . l I I I ' ? l I I I I II 1 I - I I ° ' - I I I 0 I I . I R Q o I I I I I - I ° I an I " I 7’ m I 1.. S I--..3....5....,....g...-5... : g 5 9 9....5....,...-q..-.,....g....g| 'l .0. (I? or I'PUV Outilfltt 0".|= 120 The difficulty one encounters in attempting to pro- pose detailed models for enzymatic mechanisms is that the number of parameters (unknowns) to be determined becomes too large, even for relatively simple models. In addition, the assignment of proper initial values for these parameters is often difficult, due to the lack of experimental evidence to support such assignments. Enzyme systems such as AMP-aminohydrolase present yet additional difficulties, since no detectable intermediates are available, from which additional information could then add some degree of certainty to a proposed mechanism. C. Phenomenological Model (Model B) Modelling thus far has included one model (Model A) whose scheme did not address cooperativity effects, and one which did (MWC Model). While both models were suc- cessful in describing the overall experimental behavior for the catalytic reaction of K+-free AMP-aminohydrolase with 5'-AMP, they were not able to exactly fit all the individual reactions for the various enzyme and substrate concentra- tions employed. The phenomenological model (Model B) which includes apparent cooperativity effects in its scheme, is a gen- eralization of the Suelter et al. (27) model, with an ex- ponential turn on. In this model, the enzyme is assumed to exist in two manifolds, inactive Ei’ and active Ea' 121 Each manifold may exist in a number of enzyme forms, which may or may not have substrate bound to it. The key to the first step (activation) is that sub- strate binds to inactive enzyme to activate it in a pseudo first order process, which may have a complex dependence on [S] and/or [E]. Likewise, the reverse process (de- activation) is assumed to occur in a similar fashion by a pseudo first order process. Model (B) is represented schematically by: 1 Step 1 E31 + S ——45 Ea Km kp Step 2 Ba + SéeEaSQ Ea + P in which R1 and R2 are phenomenological rate constants which may depend on [S] and [E], and where Ei is the in- active enzyme; 8, the substrate; Ea the active enzyme; EaS, the active enzyme—substrate complex; P, the product; Km, the Michaelis-Menten constant; and kp, the turnover number. Hence, Step 1 involves the apparent cooperative activa- tion of the enzyme, while Step 2 is the Michaelis-Menten scheme. The initial enzyme concentration (E0) is E = E. + Ea + EaS (l) 122 Let the total active enzyme concentration (E2) be: E: = Ea + EaS (2) then so = Ei + E: (3) and d = k (E )(S) - k (Et) (u) dt 1 i 2 a Substituting the value of (E1) from (3) into (u) yields: d(E:) t "EE“ = kl(EO)(S) - [k2 + kl(S)] (5) Define, k! k2 + kl(S) (6) Then an; , . dt = kl(EO)(S) - k (Ba) (7) The rate of product formation is: 123 t d? _ kp(Ea) *3 - (8) <1 + Km/s> and S = SO - P (9) where (7), (8) and (9) are the working KINFITu equa- tions. Apparent cooperativity effects are thus embodied in the first order rate constant (k'). The behavior of Model B is described by the four param- l’ 2’ m when employed in Equations (7), (8) and (9) should then eters k k K and kp. Values for these four parameters map out the progress of product as a function of time, if the scheme is to successfully interpret the experimental data. Experimental evidence presented earlier in this work has indicated that at 22°C, the Michaelis-Menten constant (Km) is in the order of 0.35 mM, while that of the turnover number (kp) is about 3000 sec-l. The values of (kp) at 12°C and 2°C were thus calculated from the Arrhenius rela- tionship, by using the activation energy value of 16,500 calories per mole determined earlier. Thus the value of (kp) at 12°C was calculated to be 1158 sec-l, while at 2°C 1 it was 379 sec- These values were subsequently used in 12H fitting the experimental data. The value of the Michaelis- Menten constant (Km) was not assumed to vary appreciably over the small range in temperature (2°C-22°C) at which the experiments were performed. This is because (Km) is made up of a ratio of rate constants, and thus small changes in temperature should leave its overall value unchanged for the three temperatures. Using the values of (Km) and (kp) shown above, it was then possible to reduce the number of unknowns in Model B to only two, namely (kl) and (k2). Individual files for all the pushes were fitted with KINFITU. In all, twenty- five different AMP-aminohydrolase and 5'-AMP concentra- tions at each of the three temperatures (hence, a total of 75 combinations) were fitted to (kl) and (k2), while hold— ing (Km) and (kp) at the levels indicated in the preceding paragraph. Initial estimates were as follows: 1 -1 1000 sec- M 1 k 1 R2 2 sec- All the data fittings were successful. Tables 6.2, 6.3 and 6.“ show the results obtained. Figures 6.22, 6.23 and 6.24 are typical representations of KINFITU plots, for the fits obtained using Model B. Results from Tables 6.2, 6.3 and 6.4 were used to 125 Table 6.2. Phenomenological Model, Data-Fitting Results at 2°C. E0 sO kl k2 k' 5.00 u2.5(1.5)a 0.232(0.011) 0.uu5 2.50 57.6(2.1) 0.226(0.013) 0.370 3.60 1.25 73.0(u.1) 0.309(0.022) 0.u00 0.625 56.3(2.8) 0.307(0.017) 0.392 0.313 35.2(1.7) 0.37u(0.019) 0.385 5.00 M9.6(2.0) 0.158(0.008) 0.u07 2.50 63.u(1.5) 0.151(0.006) 0.310 1.80 1.25 71.3(1.7) 0.118(0.006) 0.207 0.625 78.3(3.2) 0.191(0.010) 0.2u0 0.313 u3.8(1.1) 0.216(0.006) 0.230 5.00 18.2(0.7) 0.061(0.003) 0.151 2.50 36.7(0.8) 0.099(0.003) 0.191 0.90 1.25 53.0(1.2) 0.114(0.003) 0.180 0.625 72.9(2.7) 0.166(0.007) 0.211 0.313 39.6(1.2) 0.257(0.008) 0.269 5.00 16.8(0.9) 0.0uu(0.003) 0.128 2.50 18.6(0.7) 0.035(0.002) 0.081 0.u5 1.25 ul.2(0.6) 0.065(0.001) 0.116 0.625 62.1(1.0) 0.102(0.002) 0.1u1 0.313 36.0(0.8) 0.161(0.00u) 0.172 5.00 5.6(0.3) 0.025(0.002) 0.053 2.50 9.2(0.5) 0.025(0.002) 0.0u7 0.225 1.25 11.3(0.u) 0.021(0.001) 0.035 0.625 39.6(0.u) 0.067(0.001) 0.092 0.313 32.6(0.9) 0.095(0.003) 0.105 2°C; kp = 379 sec-l. Km = 0.35 mM; E0 (uM); SO (mM); 1 kl (sec- M-l); k2 (sec-l); k' (sec-l). 8Numbers in parantheses are marginal standard deviation estimates. 126 Table 6.3. Phenomenological Model, Data-Fitting Results at 12°C. so so kl k2 k' 5.00 L117(16)a 1.061(0.051) 3.15 2.50 619(12) 0.637(0.035) 2.17 3.60 1.25 711(19) 0.88u(0.050) 1.77 0.625 63u(17) 1.u39(0.0u7) 1.84 0.313 062(27) 1.235(0.080) 1.38 5.00 239(8) 0.602(0.026) 1.80 2.50 338(8) 0.u61(0.020) 1.31 1.80 1.25 512(9) 0.638(0.018) 1.28 0.625 558(12) 0.796(0.022) 1.10 0.313 u35(20) 0.909(0.0u8) 1.0a 5.00 92(u) 0.300(0.016) 0.76 2.50 165(u) 0.327(0.013) 0.74 0.90 1.25 298(5) 0.437(0.010) 0.81 0.625 37u(16) 0.697(0.033) 0.93 0.313 151(2u) 0.593(0.098) 0.6a 5.00 517(28) 0.2u7(0.016) 2.83 2.50 112(2) 0.256(0.001) 0.5a 0.A5 1.25 161(3) 0 270(0 007) 0.u7 0.625 287(12) 0.035(0.020) 0.61 5.00 542(26) 0.257(0.01u) 2.97 5.00 15(12) 0.100(0.07u) 0.17 2.50 26(2) 0.065(0.007) 0.13 0.225 1.25 69(2) 0 126(0.005) 0.21 0.625 113(8) 0.191(0.01u) 0.26 0.313 62(5) 0.133(0.012) 0.15 12°C; k = 1158 sec-l. Km = 0.35 mM; Eo (0M); so (mM); - -1 -1 . -1 kl (sec M ); k2 (sec ), k' (sec ). aNumbers in parantheses are marginal standard deviation estimates. 127 Table 6.4. Phenomenological Model, Data-Fitting Results at 22°C. Eo so kl k2 k' 5.00 941(50)a 1.93(0.17) 6.63 2.50 906(86) 3.18(0.37) 5.45 3.60 1.25 513(26) 1.96(0.14) 2.60 0.625 340(15) 1.93(0.10) 2.15 0.313 179(5) 1.76(0.05) 1.82 5.00 1298(44) O.60(0.05) 7.09 2.50 962(9) 0.80(0.02) 3.20 1.80 1.25 622(14) 1.07(0.05) 1.84 0.625 375(10) 1.14(0 04) 1.37 0.313 162(19) 1.40(0.17) 1.45 5.00 2.50 1173(28) 0.55(0.03) 3.48 0.90 1.25 797(12) 0.68(0.02) 1.67 0.625 526(10) 1.06(0.02) 1.39 0.313 205(6) 1.58(0.05) 1.64 5.00 - 2.50 1220(24) 0.47(0.02) 3.52 0.45 1.25 956(16) 0.58(0.02) 1.78 0.625 745(14) 0.76(0.02) 1.22 0.313 398(23) 1.08(0.06) 1.21 5.00 1059(29) 0.47(0.01) 5.77 2.50 748(19) 0.35(0.01) 2.22 0.225 1.25 725(14) 0.39(0.01) 1.30 0.625 697(17) 0.65(0.02) 1.08 0.313 458(158) 0.90(0.32) 1.04 o. - ‘l - . . . 22 C, kp - 3000 sec . Km — 0.35 mM, E0 (uM), SO (mM), kl (sec—lM-l); k2 (sec-l); k' (sec-l). aNumbers in parantheses are marginal standard deviation estimates. l28 Figure 6.22. Phenomenological Model, Single-Set Fitting; l.8 uM AMP-Aminohydrolase with 2.5 mM 5'- AMP, 22°C. Figure 6.23. Phenomenological Model, Single-Set Fitting; 3.6 uM AMP-Aminohydrolase with 2.5 mM 5'— AMP, 1200. Figure 6.24. Phenomenological Model, Single-Set Fitting; 0.9 uM AMP-Aminohydrolase with 1.25 mM 5'— AMP, 2°C. Figure 6.22 Figure 6.23 129 II 1|. vi; 1 “' " " “'lul' \' "lVl‘oIIIIllUI II In. ((v1~ I)": a! II - . , '- ' LU! l' I"! 'ICHI 7 ' III . - I III II I o...lu ha suzI). l.\W' II Iu' [qr . ..I.¢.a.,.pfu. .. Inf "q,,". , .o;,'oo;°::g"""'“' ":?}1§| II 0 I < u .A -o l'. [lP‘I'Il'I‘ an I , . . “no. 0‘. ' ”'I'I‘hlt ."l’l (In. I run” [IUD lul'l' Iumv H u In ...-.q I] '0 H “ovum I (IltuIIIIfl VOINI II I I" In! 3"! Hill. 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I5- - {EL 0' IW’J' 130 produce Figures 6.25, 6.26 and 6.27, which show plots of the initial substrate concentrations (80) versus [k2 + kl(S)] at 22°, 12° and 2°C, respectively. Figures 6.28, 6.29 and 6.30 show plots of the initial enzyme concentra- tions (E0) versus [k2 + kl(S)] at 22°, 12° and 2°C, respec- tively. It is clearly evident from Figure 6.25 that the pseudo first order rate constant (k') is dependent on (SO) at 22°C, whereas it is independent of (SO) at 12°C and 2°C, as shown in Figures 6.26 and 6.27 respectively. The reverse is true however, for the enzyme dependence. Figure 6.28 shows that (k') is independent of (E0) at 22°C, but dependent on (E0) at 12°C and 2°C, as shown in Figures 6.29 and 6.30 respectively. Hence the apparent cooperativity effects embodied in (k') are manifested by their dependence on (SO) but not (E0) at the higher temperature (22°C); whereas these ef- fects are completely reversed at the lower temperatures (12° and 2°C), where they are dependent on (E0) but not (so). The mechanistic interpretation of these phenomena can be ascribed to the constitution of the tetrameric enzyme - as related to the association-dissociation equilibria of its four subunits - as a function of temperature, where the independence of the rate on (E0), at 22°C (Figure 6.28), indicates that the enzyme maintains its tetrameric 131 Symbols for Figures 6.25 - 6.27 Initial AMP-Aminohydrolase concentration (UM): Symbols for Figures 6.28 - 6.30 Initial 5'-AMP concentration (mM): . 5.00 I A‘l V 0.625 O 0.313 132 .OONN pm Aomv wsmpm> mAmvHx + mag .Hmeoz HQOHwOHocmEOQmsm .mm.m magmas e: noiomv n V n N p o .Ep-Pbp—ph-b—rnub-Ln-p—pp-n-b-np-h-pn—Lhnp-hh-n—pbhp-thL’ 11[IllllLllllllJllLlllLlllllll111111llllllllllljlllllllllllllllllllllllll {$143203 WW'Ql 0 fl 0495 I" Z >H-(s) l>1 (Pass) 0 n. TTUIIIIIIIII1U'IIII'UIIUIIIIIIUVTT“IIT'IIII'I'II'IIIIIITIIIIIII'IIIU'TI1I ' r . . . d‘qj‘Wd‘qd-ded-ddq4fld114-did+dqddddd¢ddudqud‘A-ddliqddddfi-Idl n t n N p O Summons; T8. Sonfloneac .985 coma 3821...... inmvi .m> om 133 .Ooma pm Aomv mumpm> mAmVHx + mxu aHoooz HmofimoaocmEocmcm .mm.o mpzwfim 3 nciomv 0 o v n N — DPF-rbLbn—npub—hhbn-nthP-h-n-nphn—..LL-Lbbb-nth—bh-n-pb-h lllkLlLlllLlLlllellLLILIAIllLLLlL k? > <| < [Ii 89:87:“ N'M'Ol O O I I“ r x. I l ) r S t ( - w r Z f ) IN a 1 3 x . i It ... ( r t In 111 om o 13“ q a com um Aomv mzmpo> mAmvHx + mxg .Hmooz HmofionocoEocmzm 3v noiomo —.0 q 0 '1 o '2 o 3 Inlumnflfix 7.00: annuunngc 5255.; UON tunings“. 22$ I .m> om 0 n v n N — o D rr’p D h n h - b h p b - b n n p — n b b n — n b n b - I DP h - p b b n b n n h b - D P b b - b b P D- D I xP 0.0 I I I I > I > II J <1 < . u u I I I I.o I I I I I I l I I I "I I I II I I ) 1 .- NO 5 I I . I\ . . + I I n I I z I I l I I I I T I l ) 1 I 8 .I ll ".0 a 1 u . I I I. I I ( I l L l L .I I I I I I I. 0.0 L I I I I I I T l l I I J I L V l n . I . . 414‘4 - d d d d d 11 ‘ 1- < d I 4 —‘ A q 1 «l. d 1 dl d d d 1 1 d — d 4J1 d - d d d d d d d 1 I‘Id 4 d d 1 d4 4 no 0 n ¢ n N p a .Nm.m madman WtO‘fl WWt VS. 22°C Turnover numbar=3000 soc“ Km=.35E—3M Full-Model 135 h on n c- n N .- o . * IVIIIIIIILILIIIIIIIIlIlIIllIIIlLlIIIIIIIIIIIIIILIJIIIIIIIIIIlIIIIlIIIllILLl ' q . - P < P . ' ' o . - r- 4 L i , 4 . n - — n 4 . . .- . . .. _ A . " 2 d - N .1 \ — N % d .. .- ‘ , ., _ 2‘?) d l - e . .- -I — . . . .. d . . . C- d I- P a i c h d b .4 .- 4 .. 1 . a . d . P 4 n- O -O IIIWIIIIIU'lI—IIIIIIII'IIIIIII'TIIIIWIIII'IIII'IIII'IIIUIIIIIITIIIIIIII vs a on v n N - o (L_338) Zn+(s) 1x ”$23“ W-Ot Phenomenological Model, [k2 + kl(S)] versus (E0) at 22°C. Figure 6.28. 136 .ooma um nomv mampm> mfimvax + mxg .Hmooz HmOfiwOHocoEococm .mm.m ogzmfim A3 moiouv lMWLI WWL v n N p O o BED-b-pn—h-pp-Phhh—nunP—yh-be—ythb—P-hb o L I I I .1 1| I I I I —l [p L I n 1 T It ) L r S 1 o 1 l\ I I z L \ I 1 - o 1 I I I ) NI rlN s 1 I an“ .. n I . I Y I I l ( IL I L r I I I I L I n1 In I I o ' L I . T Jddqfiqqu< «14‘4J4‘fi—dduu-4Jqa—uq-Jdfiudq V n—i N p O znlunn.nEx «Icon amp—"505:5: .3053 UON— .ovOZIaau 2+3va .m> om 137 .oom pm Aomv msmpm> mAmvHx + mxg aHmooz HMOHwoaocmEocmnm 3 moiouv V n N 0.0 PPDD-erLFbpup-l-rbhr-phhb_brbbh b? h p I. I— - b p p b —.0 N6 nd Q6 IIIIlLIILlIILLIILLIIIIIIlILIIlIIIIIJIILlIlLIiILLL TII'TIfilIIII’lIIIIIITIIIIrIIIIIIIIIIIIIIIII'IIII no Nildfi‘ld‘ddd‘fiqq-qdd1d1dIJ‘dJJlT—‘4ddd‘I4JWJ1 v n N p o anunH.HEx pluoo mhnnconEac ~30:ng UON 3321.3... 22$; .m> om Od pd N6 n6 '6 0.0 .om.m opzmfim Z >1+(s) l>1 ( l__:>os) 60W“ WWI 138 configuration at this temperature even in the absence of monovalent cation activators. This is in agreement with the sedimentation velocity results presented earlier in this work. Dissociation to subunits occurs at the lower temperatures, as indicated by the dependence of the rate on the initial enzyme concentration, (E0), at these tem— peratures (Figures 6.29 and 6.30). PART B STOPPED—FLOW STUDIES OF THE REACTION OF S-BENZYL-L-CYSTEINE WITH TRYPTOPHANASE CHAPTER 7 TRYPTOPHANASE — AN INTRODUCTION A. Historical In 1875, Kuhne (166) and Nencki (167) reported on the production of indole during putrefaction of protein. In 1903, Hopkins and Cole (168) showed that indole was formed by bacterial decomposition of the newly isolated (168,169) amino acid, tryptophan. Happold and Hoyle (170) in 1935 identified tryptophanase (ECA.1.99.1) as the enzyme respon— sible for the production of indole in bacterial cultures. Happold and Struyvenberg (171) demonstrated that trypto- phanase action required the presence of NH1, K+ or Rb+, and was inhibited by Na+ or Li+. Wood 33 31- (172) dis- covered that pyridoxal-P (PLP; pyridoxal 5'—phosphate), a coenzyme, was required for the activity of tryptophanase, and that indole, pyruvate and ammonia were formed in stoi- chiometric amounts from L-tryptophan. Thus by 195A, the re- action catalyzed by crude, cell-free preparations of trypto- phanase could be summarized as follows: Tryptophanase L-Tryptophan + H O 4%blhdole + Pyruvate + NH 2 + 3 PLP, NH“ 139 1A0 Reviews by Happold (173), Wada (17A) and Snell (175) effectively cover what was known about tryptophanase up to 1975. B. Enzyme Source Snell (175) had suggested that the enzyme plays a catabolic role in animal metabolism, by regulating the intracellular concentrations of the essential amino acid tryptophan. Tryptophan is known to induce the formation of tryptophanase in a variety of bacterial cultures (175). However, the amount, physical and catalytic properties of the induced enzyme are dependent on the bacterial source (175). In order to produce a consistently pure form of trypto- phanase, free of a contaminating tryptophan synthetase, Newton and Snell (176) turned to Escherichia coli B/lt7 as a source of tryptophanase. This organism is auxotrophic* for tryptophanase, because it completely lacks the genes which code for tryptophan synthetase (177). It retains an inducible tryptophanase (175), and was made constitutive for this enzyme by selecting spontaneous mutants that grew on minimal medium lacking tryptophan but supplemented with indole (176,177). * AuxotrOphic mutants are defective in a biosynthetic path- way, but revert to normal growth when provided with the normal product of the pathway. 141 The resulting strain, designated E. coli B/lt7-A, pro- duces large amounts of tryptOphanase [up to 10% of the solu— ble intracellular protein (179)] under proper cultural con- ditions (179,180), and it was from this strain that crystal- line tryptophanase, homogeneous by electrophoretic and ultracentrifugal criteria, was first obtained by Newton and Snell (179). C. Structure and Properties Isolated from Escherichi coli B/lt7-A (179), trypto- phanase has a molecular weight of 220,000, and is composed of four subunits (179,181,182). Reversible association- dissociation processes are highly dependent on concentra- tion, temperature, ionic environment, pH, and the presence or absence of the coenzyme, pyridoxal-P (169). In the absence of pyridoxal-P (PLP), the apo-enzyme is catalyt- ically inactive, and has the spectrum of a simple protein with an absorption maximum at 278 nm. Equilibrium dialysis studies showed that the apo-enzyme binds four molecules of PLP per molecule of the native tetrameric enzyme (181,183). The resulting holo-enzyme shows characteristic pH—dependent absorption maxima at 337 nm and “20 nm (184) in the presence of K+ or NH: ions, which are essential for catalytic activity. At equivalent concentrations of Na+ or imidazole (0.05-0.1 M), only the A20 nm band is formed, and the enzyme has little 142 or no catalytic activity. Suelter and Snell (188) however presented evidence which indicated that Na+ was able to appreciably activate tryptophanase if employed in sufficient amounts. Their data showed ahat 0.5 M Na+ elicited nearly 30% of the activity observed with 0.05 M NHZ. Snell (175) suggests that PLP in holotryptophanase is apparently present in azomethine linkage with the e-amino group of a particular lysine residue at the active site, as found for all other PLP dependent enzymes investigated (185), and that the four binding sites are equivalent (175, 186). Snell further proposes (175) that this equivalence is most simply explained by assuming the presence of one binding site on each of the four identical subunits of holotryptophanase. A large conformational change that results in a substantially more compact structure, ac- companies the binding of PLP by apotryptophanase (182,187). Snell (175) points out that as a result of its more compact structure, holotryptophanase is more resistant than apo- tryptophanase to denaturation by sodium dodecyl sulfate (182), by heat (187), and by changes in pH. D. Spectral Properties Morinoarfl.Snell (184) reported that in the presence of K+, holotryptophanase showed two pH dependent absorption bands with maxima at 337 nm and 420 nm, which they attributed to the active and inactive forms of the 1A3 enzyme respectively. Their conclusions were based on the following experimental evidence: (a) At acidic pH, the A20 nm band predominated, whereas the 337 nm band was more prominent at alkaline pH, where holotryptophanase displayed maximal catalytic activity (above pH 8.0). Similar spectral changes with pH were observed when K+ was replaced by NHZ; (b) when 0.1 M K+ was replaced by 0.1 M Na+ or 0.1 M imi- dazole (neither of which replaces K+ as an activator), there was a heightened absorption at “20 nm and little or no absorption at 337 nm, even at pH 8.0;and (c) the spectra of these enzymatically inactive forms of holotryptophanase resembled that obtained in the presence of K+ at pH 6.1 where the inactive species predominated. Later, and in contrast to the above conclusions of Morino and Snell (18”), June 32 al. (197) presented experi- mental kinetic evidence from which they concluded that the A20 nm band corresponded to the active form of the enzyme, while the 337 nm band was attributed to the inactive form. Their analysis suggested that this kinetic interplay was manifested by complex pH and structural effects. Further discussion on their findings will be presented in the fol- lowing section of this chapter. It has been suggested from model studies (189) on pyridoxal-P aldimines that the band at A20 nm is due to the following structures: 1AA Lfime LYSINE .. H /N\H /N{ 5 5‘ 0’ <——-> 9" + H* P P Johnson and Metzler (190) and Metzler et al. (192) have shown that the state of protonation at the pyridinium nitrogen does not greatly alter the position of the absorp- tion maxima of pyridoxal-P Schiff bases although the extinc- tion coefficients differ. Davis and Metzler (191) and Snell (175) suggested the following as possible structures for the 337 nm band (page 1A5): June et al. (193) reported that the interconversion of the 337 and A20 nm bands following a change in pH or K+ concentration, occurs on the stOpped-flow time scale. The authors analyzed results from three experiments performed LYSINE LYSINE r' /N~..H /N- If P .JJHP bl H+ N at 2A°Cz (a) a pH drop from 8.53 to 6.72; (b) a pH jump from 7.38 to 9.30 and (c) a sudden change (K+-jump) from 0.1 M Na+ to 0.05 M Na+ and 0.05 M K+ at pH 8.0. They presented the following scheme for the process: EH+ E'H+ k1 (abrupt) fl 1H+ ‘I A iH+ H (abrupt) k 2 E E' A20 nm 337 nm (low pH—form) (high pH-form) where k1 = 0.56 i 0.03 sec-l, and k2 = 0.A3 i 0.03 sec-l. 1&6 The scheme assumes rapid protonation and deprotonation (involving one or more protons) of the two slowly inter- convertible enzyme forms. The authors attributed the abrupt (<6.5 msec) spectral changes observed in their ex- periments, to reflect the rapid protonation and deprotona- tion of the E and E' forms, while the slow spectral changes (which required several seconds) reflected the intercon- version of these forms. Their K+-jump experiment also showed slow conversion of the “20 nm peak to the 337 nm absorption, but the kinetics were not clearly first order. An approximate first-order rate constant of 0.3-0.5 sec-l could be obtained from their data at short times, but the analysis was complicated by continuing changes in absorbance. The authors concluded that their results indicated the oc— currence of either enzyme conformational changes in the interconversion process or slow protonation-deprotonation steps. Followup experiments on the pH-dependent interconversion of holotryptophanase spectral forms, studied by scanning stopped-flow spectrophotometry, were analyzed by June et al (194). The experiments which involved rapid incremental increase (jump) or decrease (drop) in pH over the range of enzyme stability in 0.2 M KCl at 2N:0.3°C, showed three distinct time-dependent phases. They were: (1) an abrupt phase which is complete in less than 6.5 msec; (2) a fast first order interconversion of the 337 and U20 nm absorbances, 147 and (3) a slow first order process involving growth at 355 nm, coupled to two decays centered at 325 and 430 nm, in the incremental pH jumps and decay at 355 nm with concom- mitant growth at H30 and 290 nm in the incremental pH drop experiments. The authors (19“) analyzed the results of these experi- ments in terms of Scheme I (Figure 7.1), involving enzyme forms Ea’ EB’ EBH+’ By, EyH+ and E5. They proposed that the Ed form predominated in the absence of activating mono- valent cations and absorbs at 420 nm. Those in the B— manifold, EB and EBH+, also absorb at M20 hm, while those in the y—manifold, EY and EYH+’ absorb at 337 nm. They assigned the changes at 355 nm to the E6 form. EBH+ and EyH+ represent the protonated form of the enzyme in each manifold. The points of entry for the a- and y-forms in their proposed Scheme I are still unclear, and presently under study. Further details on the above work appear in the doctoral dissertation of David S. June (195). E. Mechanism of Tryptophanase Catalysis Morino and Snell (184) reported that on addition of substrates or of competitive inhibitors such as L-alanine, tryptophanase exhibited an absorption band with a maximum near 500 nm, which disappeared as the substrates decomposed, but remained unchanged in the presence of inhibitors. 1M8 Scheme I Interconversion of Tryptophanase Spectral Forms (19a) Conformation Low UK High pH USWE H i H / 5' x . \H‘ 8 'f 1d 6. 6 8d a 5.4m” (Egglrfi) P O lst phaseP H* B‘ 8 2nd phase (fast) kH ----—--—----- 1.. k —--------- BY 3 ‘Y8 BY H “Y H y-manifold w? L KKH \ ‘3 +H+ ’3 V I (3J7 nm) P O lst phasep O (Abrupt) N H“ (E. H+) (E ) Y Y “_| 3rd phase _ ______ __ i (slow) koy ikYé 6—manifold (355 nm) j 0’ P 0 N4. 7 H (L16) Figure 7.1. Interconversion of Tryptophanase Spectral Forms. 1&9 Their exchange experiments in the presence of D20 and T20 showed that the a-hydrogen of L-alanine was labilized under these conditions. On these and other grounds, they as- cribed the 500 nm band to a quinonoid intermediate that lacks the a-proton of the bound amino acid. Results of their experiments, and from previous kinetic data, prompted them to conclude that the B-elimination re- action pgt gg was the rate limiting step. Later, Suelter and Snell (188) found that labilization of the a-proton was the rate limiting step in the reaction, when S-orthoni- trophenyl-L-cysteine (SOPC) was employed as the substrate. Furthermore, Hillebrand gt gt. (196), also using SOPC as a substrate, reported that the enzyme-aminoacrylate complex decayed to yield either a-iminoprOpionate or the carbinol— amine pyruvate as a transient intermediate, along with the regenerated active enzyme. The transient intermediate then decayed in a non-enzymatic reaction to yield pyruvate and ammonia. June gt gt. (197) investigated the interaction of the amino acid inhibitors, ethionine and L-alanine, with trypto- phanase. Their stopped-flow studies on the kinetics of quinonoid formation with ethionine led them to conclude that the “20 nm band of the enzyme was the active form. They supported this assignment by the following observations: (1) the rate constant for disappearance of the M20-nm ab- sorbance, l8.0:2.2 sec-l, was the same as that of the fast 150 phase of the observed biphasic growth of quinonoid at 508 nm, l5.0:l.2 sec-l; (2) the rate of disappearance of the 337-nm absorbance, 0.56:0.08 sec-l, was the same as the rate of the slow phase of quinonoid formation, 0.63:0.06 sec-l, and was essentially unaffected by inhibitor concen- tration or the nature of the inhibitor, ethionine or L- alanine, and (3) the average rate constant for the slow phase obtained with ethionine, 0.51:0.14 sec-l, and L- alanine, 0.H7:0.l5 sec-l, agrees closely with the rate constant, 0.66:0.4 sec-l, for the conversion of the 337- nm form, conformation y, to the U20—nm form, conformation 8, following a rapid decrease in pH (19“). These and other results (193,19u,197) prompted them to infer that the 337- nm form apparently does not form quinonoid directly, but is first converted to the U20-nm form before the d-proton of an amino acid inhibitor is removed to form quinonoid. Furthermore, deuterium isotope effects on the kinetics of formation of quinonoid with L-[a-lH] alanine and L-[a-2H] alanine led them to argue that the rate-limiting step of quinonoid formation is the abstraction of the a-proton. Based on their results, they proposed Scheme II (Figure 7.2) as a partial mechanism for quinonoid formation. The authors (197) presented the following additional factors upon which they based their proposed scheme, Scheme II (Figure 7.2), for the reaction of ethionine with tryptophanase: 151 Scheme II H+ + k0 KQH SSH: ESQ + H+\——-‘ EQH+ k -Q I xSB V SH + + KBH + BH \ " B + H H Kay Kev K + YH + YH ‘ ‘ Y + H KY6 a Figure 7.2. A Mechanism for the Interaction of a Competitive Inhibitor, Ethionine (SH), to Form a Quinonoid (EQ and EQH) (197). 152 (a) the 337-nm form of tryptophanase (y-conformation) does not form quinonoid directly but is first converted to the U20-nm form (B—conformation), the active form of the enzyme, which forms a Schiff base from which the a- proton of the amino acid inhibitor is removed. Thus the B-conformation is the one poised for reaction. Further- more, according to the scheme, the protonated form of the B-conformation is the one which proceeds to form the quinonoid. This is attractive because formation of the quinonoid requires a protonated pyridinium nitrogen, (b) Values of the first order rate constant, kl, for disappearance of the U20-nm absorbance (which were the same as those for the fast phase of the biphasic growth of the quinonoid at 508 nm) exhibited a hyperbolic de- pendence on ethionine concentration. This hyperbolic de- pendence is in accord with the results of previous inves- tigators (175,182,184,l88) who reported that the mechanism of interaction between holotryptophanase and a suitable amino acid substrate or inhibitor, involved the formation of a Michaelis complex via a transaldimination reaction, (c) Values of the first order rate constant, k2, for disappearance of the 337-nm absorbance (which were the same as those for the slow phase of the biphasic growth of the quinonoid at 508 nm) do not show a hyperbolic de- pendence with increasing ethionine concentration, but are essentially constant, suggesting that they reflect an 153 enzyme conformational change. This result conforms with the previous findings of June gt gt. (193,19U), (d) when the interaction of L-alanine with trypto- phanase was examined under comparable conditions, values of k1 were substantially smaller than those obtained with ethionine, whereas values for k2 were essentially the same as those obtained with ethionine. Since k2 values were comparable for both inhibitors, this presented further sup- port to the authors previous suggestion (l93,l9u) that k2 reflects an enzyme conformational change, (e) Examination of the effect of pH on quinonoid forma- tion revealed that as the pH was increased, kl also in- ' creased, while the relative amplitude of the fast phase of the quinonoid diminished. This supports the proposal that the H20—nm form (B-conformation) is the one poised for reaction, i.e., the active form of the enzyme. In contrast, k2 appeared to be less sensitive to pH. Since the rate of disappearance of the 337-nm form (y-conformatin) corresponded to k2 of the quinonoid, this added further reinforcement to the argument that the y-form is not the active form of the enzyme, and finally (f) Substitution of deuterium at the a-position of the inhibitor L-alanine, affected both the extent and rate of quinonoid formation. Thus, deuterium substitution slowed down the fast phase of the quinonoid growth, but left the slow phase virtually unchanged. This was interpreted as 15“ additional evidence that the slower phase reflected an enzyme conformational change. In order to render further validity to Scheme II (Figure 7.2), which was based on interaction of inhibitors with tryp- tophanase, rate studies with amino acid substrates should be performed and analyzed in terms of the same scheme. The following chapter will address some aspects of these studies; namely, the interaction of the substrate S-benzyl- L-cysteine with tryptophanase at pH 8.0. CHAPTER 8 STOPPED-FLOW STUDIES OF THE REACTION OF S-BENZYL—L-CYSTEINE WITH TRYPTOPHANASE A. Introduction The study of the reaction of S-benzyl-L-cysteine with tryptophanase was undertaken in order to investigate the kinetics of the spectral forms at 337, A20 and 508 nm, and to determine the correlations which may exist between the time progress of these forms with this substrate, to the progress of the same forms observed with the inhibitors, ethionine and L-alanine, as investigated by June gt gt. (197). In particular, the objective was to analyze the fast decay at A20 nm, the slow decay at 337 nm, and their correlation to the biphasic (fast and slow) growth at 508 nm. S-benzyl-L-cysteine (SBC) was chosen for two reasons: First; tryptOphanase is able to catalyze this substrate to form products. Hence, if the time behavior in the progress of the curves at 337, A20 and 508 nm, for this substrate, can be interpreted in the same fashion as was done with the inhibitors, ethionine and L-alanine (197), then this 155 156 should confirm the earlier findings of June gt gt. (193, 194,197), and in particular the scheme (Scheme II, Figure 7.2) presented by June gt gt. (197), for the mechanism of quinonoid formation. Second; changes in the quinonoid band for the reaction of SEC with tryptophanase occur on the stopped-flow time scale, unlike those for SOPC and other substrates, where the rates of formation and decay of the quinonoid are faster than the stopped-flow time scale. June gt gt. (193,19A,l97) also reported on slow ab- sorbance changes at 355, 390 and A20 nm. They attributed (19“) changes at 355 nm to the 6-form of the enzyme (Scheme I, Figure 7.1), whereas those at 390 nm were attributed to changes in the spectrum of free PLP (19A). The slow growth of absorbance at A20 nm, following the fast phase was due to the nonenzymatic formation of a Schiff base between excess free ethionine and PLP (197). B. Materials and Methods B.l. Reagents Pyridoxal 5'-phosphate (PLP); S-benzyl-L-cysteine (SBC); DL-dithiothreitol (DTT); potassium [ethylene diamine tetracetic acid] (KEDTA); and potassium N-(2-hydroxyethyl)- piperazinepropanesulfonic acid (Kepps), were all obtained from Sigma Chemical Company. All other reagents were 157 analytical grade. B.2. Tryptophanase Preparation TryptOphanase from Escherichia coli B/lt7-A was prepared by Iraj Behbahani Nejad as described by Watanabe and Snell (198) including the modification of Suelter et al. (199). Holoenzyme was prepared from stock apoenzyme by incubation in a dialysis buffer which consisted of 25 mM Kepps, 0.1 M KCl, pH 8.0, 1 mM KEDTA, 1 mM DTT, and 5 UM PLP, for one hour at 37°C. The enzyme had a specific activity of AS umole min.l mg-l when assayed with 0.6 mM S-(o-nitrophenyl)- L—cysteine (SOPC) in 50 mM potassium phosphate, pH 8.0, and 50 mM KCl, at 30°C (200). Protein concentration was determined spectrophotometrically by using e(278)=0.795 mL mg_:L cm-l (18A). B.3. Scanning Stopped-Flow Experiments Activated holotryptOphanase (5.0“ mg mL'l) was diluted 1 with the dialysis buffer to a concentration of A.5A mg mL- and pushed against five different concentrations (0.125, 0.25, 0.5, 1 and 2 mM) of SEC in the same buffer at 21.2°C. The final concentrations of the reactants, after mixing, were hence half the initial. All spectra were scanned over the wavelength range 326-A91 nm. Since absorbance over the wavelength range A75-5A0 nm 158 was shown (175,188) to represent only that due to quinonoid, scanning was limited up to 491 nm, due to the intense absorp- tion of that band [emax (508) = 40400 M"1 om‘l (197)] in order to simultaneously monitor changes at 337 nm [cm (337) = 3500 M.1 cm.1 (194)] and 420 nm [Emax (420) = 4600 M"1 cm-l ax (194)]. Analysis of the quinonoid band was performed at 478 nm (the left shoulder of the band); however, in order to avoid confusion in terminology, disscussion in this text shall con- tinue to refer to the implicit 508 nm wavelength. 8.4. Data Analygis Data were fitted by the appropriate mathematical func- tion using the nonlinear curvefitting program KINFIT4 (l8). Errors listed are marginal standard deviations, and there- fore include the effects of coupling among the parameters. The absorbance A, for single exponential growth or decay was fitted using the following equation: A = Am — AAe (l) The three adjustable parameters were: Am, the absor- bance at infinite time; AA, the total change in absorbance, i.e., Am - A where AO equals the absorbance at zero time, 0) and kl, the apparent first order rate constant. Biphasic data were analyzed as the sum of two expon- entials according to the following equation: 159 A = Aoo - AA e"klt - AA e-k2t 1 <2) 2 where A00 is the measured absorbance at infinite time; AAl and AA2 are the changes in absorbance due to the fast and slow phases respectively and k1 and k2 are the apparent first-order rate constants for the fast and slow processes. C. Results and Discussion C.1. Absorbance Changes at 337 nm The slow decay (Figure 8.1) observed at 337 nm was fitted to a single exponential (Equation 1), and the results appear in Tables 8.1, and 8.2.A. Total absorbance changes are shown in Table 8.2.8. The rate constant, k2, for the slow decay at 337 nm has an average value of 0.40:0.12 sec-l ably with 0.51i0.l4 sec-l, obtained by June gt gt. (197) with , which compares favor- inhibitor ethionine. This slow decay is interpreted (194, 197) to reflect the enzyme conformational change from the y to the 8- form (Figure 7.1). C.2. Absorbance Changes at 420 nm A fast decay (up to about 200 msec) followed by a slow growth occurred at 420 nm (Figure 8.1). The fast decay 160 439 439 m 1 0 ’t E 006‘ I 2 ‘~. '3 '06 8 .03« 1". ‘I 2: GD g < \ m .001, On I... .... 8 0 2'5 5'0 7’5 160 an MSEC <1 '05 337 nm “a." ”0.0 o 0 ' ° 1 .. ......g... i 420 "m oo.ooz.!::... ...‘000 .. L ' r t r r O or. . 0' 1 H2 3' ‘4 5 6 SECONDS Figure 8.1. Changes in Tryptophanase Absorbance at 337 and 420 nm as a Function of Time After Mixing with S-Benzyl-L-Cysteine in 25 mM Kepps, 0.1 M KCl, pH 8.0, 1 mM KEDTA, 1 mM DTT and 5 uM PLP. After Mixing, Concentrations Were 1 mM S-Benzyl-L-Cysteine and 2.27 mg mL-1 Trypto- phanase. (The rapid decrease in absorbance at 420 nm is expanded in the inset.) 161 Table 8.1. Rate Constants and Amplitudesa for Absorbance Changes at 337, 420 and 478 nmd When 2.27 mg mL‘1 Tryptophanase is Mixed with 1 mM S-Benzyl- L—Cysteine. Wavelength (nm) kl (sec-l) k2 (sec-l) Total A 337 b (slow decay) 0.48 (1) 0.042 420 (fast decay) uu.2 (3) 0.083 A78d (biphasic growth) 42 (4) 0.68 (2) 0.75C aThe data at 337 and 420 nm were fitted with Equation 1. The data at 508 nm were fitted with Equation 2. bNumbers in parentheses are the marginal standard deviation estimates for the last significant figure. - C56% fast phase; 44% slow phase. d5(478 nm)/e(508 nm) = 0.54. Also see the second paragraph of Section (B.3) in this chapter. 162 Table 8.2.a. Rate Constants for 337, 420 and 478 nmd As a Function of S-Benzyl—L-Cysteine (SBC) Con- centrations Determined with Scanning Stopped- Flow Spectrophotometry.a Km and Vmax Values were Determined by KINFIT4 (18) According to the Lineweaver-Burke Method. 478 nmd 337 nm 420 nm (Biphasic Growth) SBC (slow decay) (fast decay) (Fast) (Slow) (mM) k (sec'l) k (secIl) k (sec-l) k (sec-l) 2 1 1 2 0.0625 0.53(3)b 21.3(2) 21(1) 0.125 0.35(3) 28.9(3) 30.1(5) 0.25 O.2l(2) 36.2(3) 38 (2) 0.5 O.42(1) 43.6(4) 42.0(6) O.70(1) 1.0 0.48(1) 44.2(3) 45 (2) 0.47(2) Km (mM) 0.080(9) 0.077(4) vmax(sec"1)C 48(1) 48.4(5) aThe concentrations of the reaction components after mixing were 25 mM Kepps, 0.1 M KCl, pH 8.0, 1 mM KEDTA, 1 mM DTT, 5 uM PLP and 2.27 mg ml.‘1 tryptophanase. All data were fitted to a single exponential. bNumbers in parantheses are the marginal standard deviation estimates for the last significant figure. CThe values of k1 at infinite SBC concentration. d8(478 nm)/e(508 nm) = 0.54. Also see the second paragraph of Section (B.3) in this chapter. 163 Table 8.2.B. Total Absorbance Changes at 337, 420 and 478 nm.a The Noise Level Was Approximately 0.001 A for All Absorbance Measurements. 478 nm (Biphasic Growth) 337 nm 420 nm SBC (mM) (Slow Decay) (Fast Decay) (Fast) (Slow) 0.0625 0.018 0.064 0.205 0.125 0.020 0.073 0.240 0.25 0.038 0.083 0.289 0.5 0.043 0.079 0.39 0.18 1.0 0.043 0.083 0.42 0.33 a e(478 nm)/e(508 nm) = 0.54. Also see the second paragraph of Section (B.3) in this chapter. 164 was fitted to a single exponential (Equation 1), and the values for kl (Table 8.2) exhibited a hyperbolic dependence on SEC concentration, yielding a value of 0.081 mM for the Michaelis constant, and 48.5 sec-1 for kl at infinite SBC concentration. June gt gt (197) reported that the slow growth of ab- sorbance at 420 nm following the fast decay was due to the nonenzymatic formation of a Schiff base between excess free ethionine and PLP, and as such did not reflect directly on the mechanism of tryptophanase catalysis.y C.3. Absorbance Changes at 478 nm (508 nm) Absorbance changes at 478 nm included a biphasic growth (Figure 8.2) followed by decay as substrate was depleted (Figure 8.3). The biphasic growth was initially fitted to the sum of two exponentials; however, due to substrate de- pletion, it was clearly evident that the slow phase of the biphasic growth was strongly coupled to the shutoff region of the reaction. Only the reaction with 1 mM SBC had a sufficiently long slow growth, free from interference by substrate depletion (i.e., decay of the quinonoid) that fitting to the sum of two exponentials was attempted. The values of k1 (for the fast growth), and k2 (for the slow 1 and 0.68:0.02 sec-l, respectively growth) were 42:4 sec- (Table 8.1). Later, the slow growth for the reaction of tryptophanase with 0.5 and 1.0 mM SBC was fitted to a 165 th) . 0 Of} . ' w ' e. 0.4a . - ' U .‘ LIJ o z o , <1 I; 3: o $0.4“. g 0.3. O .. m o 00 C) e a ‘ <02 (12 . . r . . 0 25 50 75 100 MILLISECONDS 0'0 V T I T I C) I (Z ‘4 6 SECONDS Figure 8.2. Biphasic Changes in Tryptophanase Absorbance at 478 nm as a Function of Time After Mixing with S-Benzyl-L-Cysteine in 25 mM Kepps, 0.1 M KCl, pH 8.0, 1 mM KEDTA, 1 mM DTT and 5 uM PLP. After Mixing Concentrations Were 1 mM S-Benzyl-L-Cysteine and 2.27 mg mL"1 Trypto- phanase. (The data for the first 110 msec are expanded in the inset.) 166 A.pHococHSU on» no soapgoo oammzofin ozp mzocm pomcfl 059V .ommcmLQOpdzLB Hugs we wm.m paw ocfiopmzolqlahwcomlm ZE m.o opoz mcoflumpucoocoo meaxfiz Loum< .mqm z: m can BBQ SE H .o .m.m opzwfim mazoumm 0%. . ow . om . om r m. . m o 0 00000000000000. .0 C C . C . mozooum . . . p m p w p m L o 0.... — o .njo .r O .0 oo _.o .. V or MW ~.o W .. .o O S . 8 . nu 0 00 o H .. so V 8 o N . V 3 o_mw .Ir a: . 3 Ed 3 .md T $00 090000. $6 1 Rd 167 single exponential, and the corresponding values for k2 are shown in Table 8.2. The fast growth was fitted to a single exponential, and the kl values (Table 8.2) exhibited a hyperbolic de- pendence on SEC concentration yielding a value of 0.077 1 mM for the Michaelis constant, and 48.4 sec- for kl at infinite SBC concentration. C.4. Absorbance Changes at 360 nm A slow decay occurred at 360 nm with all SBC concentra- tions. Only scanning spectra were available at this wave- length, and it was not possible to reliably analyze the data due to the low total absorbance changes at this wave- length. June gt gt. (194) attributed changes at this wave- length to the 0-form of the enzyme (Scheme I; Figure 7.1). C.5. Absorbance Changes at 397 nm A fast decay (about 200 msec) followed by a slow decay occurred at 397 nm. Both phases had small total absorbance changes. The fast decay is due to the 420-nm form, since 397 nm is a shoulder of this band (188). The slow decay reflects changes in the spectrum of free PLP (194), and as such, it is not directly involved in the mechanism of tryptophanase catalysis. 168 C.6. Correlations Between the Fast Phases at 420 and 478 nm (508 nm) The results obtained for kl, along with the correspond— ing Km and Vmax values (Table 8.2) at 420 and 478 nm, clearly show the correspondence between the fast decay at 420 nm, and the simultaneous fast growth of the quinonoid at 478 nm (insets in Figures 8.1 and 8.2). This confirms the findings of June gt gt. (197) who reported that the 420-nm band (B- conformation) is the active form of the enzyme, i.e., the form directly preceding the quinonoid, and not the 327— nm band (y-conformation) as previously reported by Marino and Snell (184) and Snell (175). C.7. Correlations Between the Slow Phases at 337 and 478 nm (508 nm) The rate of disappearance of the 337-nm absorbance is the same as the rate of the slow phase of quinonoid forma- tion at 478 nm, and is essentially unaffected by SBC con- centration (Table 8.2). The average rate constant for the slow decay at 337 nm obtained with SBC, 0.40:0.12 sec-l, compares favorably with 0.51:0.14 sec-l , obtained by June gt gt. (197) with the inhibitor ethionine, and agrees closely with the rate constant for the conversion of the 337-nm form, conformation y, to the 420-nm form, conforma- tion 8, following a rapid decrease in pH (194). Thus the 169 337-nm form apparently does not form quinonoid directly but is first converted to the 420 nm form before the a-proton of an amino acid substrate, or inhibitor (197), is removed to form quinonoid. D. Preliminary Principal Component Analysis of the Reaction of S-Benzyl-L—Cysteine with Tryptophanase In this section, an attempt will be made to describe, in brief, some highlights of the principal component analysis of rapid scanning wavelength kinetic experiments. Later, some aspects of the method will be used in the analysis of the reaction of S-benzyl-L-cysteine (SBC) with tryptOphanase. Details of the method are discussed in the doctoral disser- tation of R. Cochran (17); see also, Cochran and Horne (201, 202) and Cochran gt gt. (203). D.1. Introduction A typical scanning stopped-flow kinetics experiment generates an absorbance-wavelength-time surface. This surface contains information about the static spectrum and concentration-time profile of each absorber in the reaction. The ultimate goal of such experiments is to identify one or more kinetic mechanisms that satisfactorily explain the concentration-time profiles of each of these absorbers. However, one must first extract these 170 concentration profiles from the experiments. Moreover, one needs to be able to extract the static spectra and con- centration profiles before proposing a specific kinetic mechanism, since a kinetic mechanism presupposes that the concentration profiles are already known. Prior to analysis of such kinetic experiments, the number of absorbers which occur in a reaction is not generally known. Although one usually knows which of the chemicals initially mixed are absorbers, and although one sometimes knows which products are absorbers, one usually does not know how many light absorbing transient intermediates occur in the reaction. In a scanning stopped-flow experiment, absorbance is measured at t wavelength channels across a selected spectral region that is rapidly and repeatedly scanned as the re— action progresses. If the scanning rate is fast compared to the fastest absorbance change in the reaction, the resulting M consecutive spectra may be regarded as instan- taneous, and the absorbance-wavelength-time surface may be represented by a PXN matrix 3, the actual experimental surface. Cochran and Horne (201,202) developed the method of statistically weighted principal component analysis to determine the minimum number of absorbers required to explain each scanning experiment, and how many of these change concentration during a reaction. The authors describe 171 two kinds of principal component analysis (PCA) useful for kinetic experiments. Each requires only the matrix 5 from a scanning stopped-flow experiment, and each gives a lower bound estimate of the number of solutes in the reac- tion with a Spectrophotometric response. Second moment matrix principal component analysis, called M analysis, gives a lower bounds estimate that is sensitive to the linear dependence of the concentrations of the detectable species. Sample covariance matrix principal component analysis, called § analysis, gives an estimate that is sensitive to the linear dependence of the time rates of the concentrations. These two estimates are not necessarily the same, and the authors show that application of both analyses enables discrimination between mechanistic alter- natives that would be indistinguishable with either analysis alone. The authors first develop rules for the application of M and S analyses to errorless data. With this simplifica- tion, the M and S estimates of the number of solutes are the ranks of respectively, the second moment matrix, M, and the sample covariance matrix, g. Later, they discuss the effects of random measurement errors, and in particular, the case in which the variance of the measurement errors, as a function of wavelength, is not constant throughout the experiment. The goal is to infer the essential ranks of the two matrices; i.e., the ranks that would be obtained if 172 measurement errors could be eliminated. The details of utilizing a general error variance model to develop a weighting scheme, are described in R. N. Cochran's doctoral dissertation (l7). Cochran and Horne (201) used the var- iance model to weight the absorbance matrix A, before per- forming the principal component calculations. The applicability of PCA requires only that the response at each wavelength channel be a linear function of the con- centrations of each detectable species, i.e., that Beer's Law holds. The starting point for weighted M and S analyses is the properly weighted (201) experimental absorbance matrix, 5w, obtained from a scanning stopped-flow experiment. Based on A M analysis reconstructs an absolute absorbance —W’ matrix, AP. The essential rank m corresponds to the minimum number of linearly independent absorbers (eigenvectors) re- quired to interpret the experiment, and is determined by finding the lowest value of r for which the reconstructed absorbance matrix fir fits the experimental matrix A to within its random errors. S analysis forms the experimentally weighted difference matrix (gw - §;) [formed by subtracting the average of all the weighted N spectra, 5;: from each spectrum in AW] to reconstruct a difference absorbance matrix, (Ar - E). The essential rank 8 corresponds to the minimum number of linearly independent absorbers (eigen- vectors) whose concentrations must have changed during the 173 experiment, and is determined by finding the lowest value of r for which (3r - E) fits the experimental matrix (t - E) to within its random errors. Two criteria were used by Cochran et al. (203) to de- termine, via M analysis, how well the experimental surfaces were fit by reconstructed surfaces: (1) visual comparison A of plots of the reconstructed absolute surface, A —r’ and the experimental surface, 5, and (2) examination of the weighted residual surfaces, i.e., plots of the residuals (A — t). The residuals should be random when plotted versus time and wavelength. The same protocol was followed with S analysis. D.2. Data Analygis In the present work, weighted M analysis was used to analyze the reaction of S-benzyl-L-cysteine with trypto- phanase. Reconstructed absolute surfaces were plotted for A A3 through A which correspond respectively to r=3 through 7, 7 eigenvectors (absorbers). Residual surfaces, (A? - g) were also plotted for the same r values. Careful visual comparison of the reconstructed absolute surfaces revealed that A6 was the closest match to the experimental surface to within its random errors. Figure 8.4 shows the experi- mental surface g for the reaction of 0.125 mM SBC with tryptophanase. Figure 8.5 shows the reconstructed surface A 56. For comparison, 55’ shown in Figure 8.6 did not 174 “,37%9 -- , /// Z‘.‘ ’ ’l ’v / ‘~ o:.:._ ’I/I RESOLVICCRBBFSETBPEH 2.27flG/HL TRYPTOPHRNHSE 0 0.1251101 S-BENZYL-CYSTEINE Figure 8.4. l) ’6 9 ., 9 7‘ , Experimental Surface, A, for the Reaction of 0.1%5 mM S-Benzyl-L-Cysteine with 2.27 mg mL‘ TryptOphanase. BDNVBHOSSV 175 RESOLVSCCRBHFSEBEVHBSSURF o 0 .125 M! S-BENZYL-CYS TE [NE . Figure 8.5. Reconstructed Absolute Surface, 35, for the Re- action of 0.125 mM S—Benzyl-L-Cysteine with 2.27 mg mL“1 Tryptophanase. 176 -- -“-“‘ ~~¢§~ \ 'l e \} 1;,” ‘ ‘ ‘9 ” mus.“ -. w ° \- :. {‘{9’ 9; vunm\\ .‘\“~‘:f‘§ \;35§S¥ngyo \ - I0 ‘SEQ’I’J‘AS 30%{x ’ RES'YLV3CCRBHFSESE‘IRBSSURF . 0.125 M S-BENZYL-CYSTEINE . Figure 8.6. Reconstructed Absolute Surface, 35, for the Reaction of 0.125 mM S-Benzyl-L-Cysteine with 2.27 mg mL-1 Tryptophanase. ' 177 reproduce the experimental surface. Inspection of the residual surfaces provided better visual comparison of the fit, and for (A6 — 5) shown in Figure 8.7, the ran- domness is clearly evident. Figure 8.8 shows the residual surface for r=5, which clearly shows non-randomized residuals. The same results were obtained in the M analysis of the re- action of 0.0625, 0.25, 0.5 and 1.0 mM SBC with trypto- phanase. Hence, M analysis indicates that a minimum of six ab- sorbers, with a spectrophotometric response, are present in the reaction of SBC with tryptophanase, since a minimum of six eigenvectors, (r=6), are required to reconstruct the surfaces to within the random errors of the experimental surfaces. Three of these absorbers, namely, the 337, 420 and 508- nm forms have already been analyzed in this study. The fourth absorber is the 360-nm form (o-conformation). The fifth is the nonenzymatic PLP absorber with a maximum at 397 nm, and the sixth is the independent slow growth at 420 nm (following the fast phase), which is probably due to the nonenzymatic formation of a Schiff base between excess free SBC and PLP. As described earlier, application of M analysis yields the minimum number of linearly independent absorbers re- quired to reconstruct the experimental surface. However, according to the results obtained earlier (Section B), Figure 8.7 178 ’V?) >.- H \_ \_ Rec srt th id 131‘ ce,(A6-A) for0 the Re ctio on of 0.125 mMC S- Be nzyl -LS Cye stein with 2. 27 mg mL- '1 Tr rypto phan (A ’ 4‘ #- a Figure 8.8. econ OHJW O 179 structed Residual Surface, (A - A), r he Reaction of 0.125 mM S-Benzyl-L- ysteine with 2.27 mg mL-1 Tryptophanase. 180 concentration changes of some of these absorbers should not be linearly independent in time. In particular, the coupling of the fast decay at 420 nm to the fast growth of the quinonoid at 508 nm, and the slow decay at 337 nm to the slow growth of the quinonoid. S analysis must be per- formed in order to verify these findings. CHAPTER 9 SUGGESTIONS FOR FUTURE WORK A. AMP-Aminohydrolase Four modes of action are suggested. The first in- volves a systematic study of product (IMP) effects on the catalytic reaction, which may yield information helpful in clarifying the role of product on the enzyme activation. The second involves studies on the activation of enzyme by (H+) ion. This would constitute a follow up of the work done by Campbell and Suelter (147), in which they re- ported on the activation of AMP-aminohydrolase (from rab- bit skeletal muscle) by (H+). Stopped-flow studies on the uptake and release of (H+) may provide information on the conformational changes in the enzyme following interaction of K+. The third suggestion would be to conduct sedimen- tation velocity studies on K+-free AMP-aminohydrolase, as a function of temperature, to verify the conclusions made about the dissociation of the tetrameric enzyme at low temperatures. The fourth should involve stOpped-flow studies on the effects of ADP, ATP and GDP, on the catalytic reaction of the enzyme, as a function of pH. This should 181 182 help clarify the present ambiguities as to the effect of these nucleotides on the regulatory role of AMP-amino- hydrolase at, and below physiological pH levels. B. Tryptophanase Since the equilibrium concentrations of the spectral forms of tryptophanase are pH dependent (193,194), the re- action of S-benzyl-L-cysteine with tryptophanase should be performed as a function of pH. One important point that could be resolved from such an experiment is whether the d-form can also form a Schiff base directly with S-benzyl- L-cysteine, which would then form the quinonoid. The other would be to study the pH dependence of the quinonoid formed between this substrate, and to compare such results to those obtained with the inhibitor ethionine (197). Fixed wavelength pushes should be done with S-benzyl— L-cysteine at 360 nm in order to reliably analyze the mechanistic role of the 6-form. Finally, complete principal component analysis of the reaction of S-benzyl-L-cysteine with tryptophanase is suggested. APPENDICES Fiji; APPENDIX A Uncomplicated "Clean" Reaction and Its Independence of Wavelength For an uncomplicated "clean" reaction, where only the reactant and product absorbances change with time, the ab— solute absorbance (At) at any time (t), and at any wave- length is: At = (eB> + (eS)(S) + (ep>(b)> (E0), where (SO) is the initial substrate concentration, and (E0) the initial enzyme con- centration, then 183 184 (so) = (S) + (P) Substitution of (2) into (1) yields: At = (b)[(eB)(B) + (eE> + (Ae>

+ (63>(so>1 where The absorbance at infinity (Am) is: 00 A = (b)[(€B)(B) + (EE)(E) + (EP)(SO)] The absorbance at zero time (A0) is: A0 = (b)[(eB)(B) + (EE)(E) + (6P)(SO)] Subtraction of (6) from (1) yields: A - A = (Ae)(b)(P) t 0 Subtraction of (6) from (5) yields: Am - A0 = (Ae)(b)(SO) (2) (3) (4) (5) (6) (7) (8) 185 Dividing (7) by (8) yields: t 0 (P) A - A (SO) Rearranging equation (9) yields: (9) (10) Equation (10) is independent of the wavelength (A). APPENDIX B MODEL A Kl kl Step 1 E1 + 3.1—"Bis FTEa 2 Km kp Step 2 Ea + S V__—"" EaS —->Ea + P where (Ei) is the inactive enzyme; (S), the substrate; (Ea), the active enzyme, (P) the product; capital (K)'s, are equilibrium constants and small (k)'s are rate con- stants. The initial enzyme concentration (E0) is: E = Ei + E S + Ea + EaS (l) 0 i The initial substrate concentration (SO) is: S = S + E 0 s + Ea + 2EaS + P (3) i From the equilibria in Steps (1) and (2) above: 186 187 (Ei)(S) EiS = —— (3) K1 and (E )(S) EaS = _;i____ (4) Km Substituting (3) and (4) into (1), and solving for (E1) yields: E —E(l+S/K) E.=O a m (5) l (l + S/Kl) Define (E), as the total concentration of active engyme; then E = E + E S (6) Substituting (4) into (6), and solving for (Ea) yields: E = E (7) (l + S/Km) Substitution of (7) into (5) yields: 188 (E0 — E) E. = (8) (l + S/Kl) Substitution of (8) into (3) yields: (SHEO - E) EiS = (9) (S + Kl) Substitution of (8) into (4) yields E s = ——————(E)(S) (10) S + Km Subtraction of (1) from (2), using (8) and (9) for the values of (El) and (EiS), respectively; and finally solving for (S) yields: K (E - E) S = SO - P - E0 = i§l§§l + l O (11) (S+Km) (S + Kl) Since, 88 = kp (Bag) (12) and and 189 then substitution of (10) into (12) yields: P kp(E)o Solving for (E6) yields: (3)020)T (E') = 0 k, at large (t), (E5) (1- -k’t e R? t=t t=0 = e-k't ) + (E0)0 e'k' (B)(EO)T t —(B+k_ )dt (21) (22) (23) (24) (25) 199 let (B)(EO)T (E6) = -———————— , for large (t) (26) k7 Substitution of (26) into (24) yields: -k't -k't v = t _ 9 (E0) (E0)m(l e ) + (E0)Oe (27) (E0)0 is the concentration of (E6) at zero time; i.e., prior to the addition of activator. Thus (E0)0 is assumed to be close to zero, since at zero time, (E') the active enzyme concentration, prior to the addition of activators is itself very small. Therefore, k"3) (28) v = v _ ' (E0) (E0)m(l e Substitution of (28) into (4) yields the final working equation: -k't dP kp(Eb)m(l-e ) _E = (29) (1 + Km/S) REFERENCES 10. 11. 12. 13. 14. 15. REFERENCES Roughton, F. J. W., Proc. R. Soc., 8115, 475 (1934). Chance, B., J. Franklin Inst., 229, 455, 613, 637 (1940). Gibson, 0., J. Physiol., 117, 49P (1952). Sturtevant, J. M., Rapid Mixing and Sampling Tech- niques in Biochemistry, Editors: Chance, B., Eisen- hardt, R. H., Gibson, Q. H., and Lonberg-Hohn, K. K., Academic Press, New York (1964), p. 89. Dye, J. L., Acc. Chem. Res., t, 306 (1968). Dye, J. L., and Feldman, L. H., Rev. Sci. Instrum., 11, 154 (1966). Dewald, R. R., Dye, J. L., Eigen, M., and deMaeyer, L., J. Chem. Phys., 39, 2388 (1963). Feldman, L. H., Dewald, R. R., and Dye, J. L., in "Solvated Electron, Adv. Chem. Ser., 5Q, 163 (1964). Ho, G.-H., Dye, J. L., and Suelter, C. H., J. Biochem. and Biophys. Methods, 4, 287 (1981). DeBacker, M. G., and Dye, J. L., J. Phy. Chem., 15, 3092 (1971). Minnich, E. R., Long, L. D., Ceraso, J. M., and Dye, J. L., J. Am. Chem. Soc., 9_, 1061 (1973). Hansen, E. M., Ph.D. Dissertation, Michigan State Uni- versity, East Lansing, Michigan (1970). Coolen, R. B., Papadakis, N., Avery, J., Enke, C. G., and Dye, J. L., Anal. Chem., 31, 1649 (1975). Ho, G.-H., Ph.D. Dissertation, Michigan State Uni- versity, East Lansing, Michigan (1976). Papadakis, N., Ph.D. Dissertation, Michigan State University, East Lansing, Michigan (1974). 200 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 201 Coolen, R. B., Ph.D. Dissertation, Michigan State University, East Lansing, Michigan (1974). Cochran, R. N., Ph.D. Dissertation, Michigan State University, East Lansing, Michigan (1977). Dye, J. L., and Nicely, V., J. Chem. Educ., 48, 443 (1971). Hill, R., Proc. R. Soc., B100, 419 (1925). Monod, J., Wyman, J., and Changeux, J.-P., J. Mol. Biol., 12, 88 (1965). Adair, 0. 3., J. Biol. Chem., 63, 529 (1925). Adair, G. 8., Proc. Roy. Soc. London Ser., A109, 292 (1925). Koshland, D. E., Nemethy, G., and Filmer, D., Bio- chemistry, 5, 365 (1966). Eigen, M., Nobel Symposium, 5, 333 (1967). Hammes, G. G., and Wu, C.-W., Science, 172, 1205 (1971)- Herzfield, J., and Stanley, H. E., J. Mol. Biol., 5g, 231 (1974). _ Suelter, C. H., Kovacs, A. L., and Antonini, E., FEBS Letters, g, 65 (1968). Ellis, K. J.,Kuntz,K., and Sturtevant, J. M., J. Biol. Chem., 246, 6631 (1971). Frieden, C., J. Biol. Chem., 235, 5788 (1970). Ainslie, Jr., G. R., Schill, J. P., and Meet, x. E., J. Biol. Chem., 247, 7088 (1972). Frieden, 0., Ann. Rev. Biochem., 38, 471 (1979). Pierce, T. H., Ph.D. Dissertation, Michigan State University, East Lansing, Michigan (1981). Cukier, R. I., Levine, H. B., Shuler, K. E., J. Comput. Phys., 55, l (1978). Blum, J. J., Arch. Biochem. Biophys., 5, 486 (1955). -— Aust, A. E., Ph.D. Dissertation, Michigan State Uni— versity, East Lansing, Michigan (1975). 36. 37. 38. 39. 40. 41. 42. A3. 44. 45. 46. 47. 48. A9. 50. 51. 52. 53. 202 Badwey, J. A., and Westhead, E. W., J. Biol. Chem., 251, 5600 (1976). Spivey, H. 0., Flory, W., Peczon, B. D., Chandler, J. P., and Koeppe, R. E., Biochem. J., 41, 119 (1974). Rodriguez, H. B., and Halloway, M. R., Biochem. Soc. Trans., 5, 83 (1976). Kirschner, K., Gallego, E., Schuster, I., and Good- all, D., J. Mol. Biol., 55, 29 (1971). Shill, J. P., and Neet, K. E., Biochem. J., 123, 283 (1971). Hatfield, G. W., Ray, Jr., W. J., and Umbarger, H. E., J. Biol. Chem., 245, 1748 (1970). Mankovitz, R., and Segal, H. L., Biochemistry, 8, 3757 (1969). Vagelos, P. R., Alberts, A. W., and Martin, D. B. J. Biol. Chem., 238, 533 (1963). Rabin, B. R., Biochem. J., 102, 22c (1967) Whitehead, E., Prog. in Biophys. and Mol. Biol., gt, 321 (1970). Schmidt, G., Hoppe-Seyler's Z. Physiol. Chem., 179, 243 (1928). Schmidt, G., Hoppe—Seyler's Z. Physiol. Chem., 208, 185 (1932). Schmidt, G., Hoppe-Seyler's Z. Physiol. Chem., 219, 191 (1933). Conway, E. J., and Cooke, R., Biochem. J., _5, 479 (1939). Zielke, C. L., and Suelter, C. H., The Enzymes, 5, ed. Boyer, 3rd, ed., Academic Press, New York (1971). Brams, J., J. Am. Chem. Soc., 8t, 4997 (1959). Atkinson, M. R., and Murray, A. W., Biochem. J., 104, 100 (1967). Burger, R., and Lowenstein, J. M., J. Biol. Chem., 242, 5281 (1967). 54. 55. 56. 57. 58. 59. 60. 61. 62. 63. 64. 65. 66. 67. 68. 69. 70. 71. Chapman, A. G., 203 £38, 8309 (1973). Lee, Y., J. Suelter, C. C. L., Fed Kalckar, H. Biol. Chem. H. Proc., 25, 756 (1966). , Byrnes, 3 E. w., and Atkinson, 227, 0. E., Berry, A. 987 (1957). J. Biol. Chem., , and Winely, M., J. Biol. Chem., 167, 461 (1947). Byrnes, E. W., Su, J.-C., Li, 2, 536 (1966). Wolfenden, Setlow, B. Coffee, C. J., and Kofke, W. A., J. Biol. R. 3 3 and Suelter, C. H. Res. Commun., 55, 422 (1965). c.—c, and Ting, c. Tomozawa, chemistry, 1, 3965 (1968). Y. 3 , Biochem. Biophys. C., Biochemistry, and Bamman, B., Bio- Burger, R., and Lowenstein, J. M., J. Biol. Chem., 241, 1244 (1966). 359, 6653 (1975). Chem., Lee, Y., The Enzymes, 3, Academic Press, New York (1959)- Ashby, B., 8728 (1978 Setlow, B. and Frieden, ). 3 C ' 3 and Lowenstein, £33. 607 (1967). Matsuo, I, Ohkido, M., Biochem. Cutaneous Epidermal Differ., Proc. (1976); Edited by Seiji, M., and Bern- U.S. Semin. stein, I. Prus, E., A. 3 Univ. Park Press, J. Biol. Chem., J. M. Kizaki, H. Purzycha-Preis, J. Zydowo, M., Acta Biochem. Pol., 51, 241 (1980). , Wozniak, M., , J. Biol. 253 3 Chem., , and Sakurada, T., Jpn Baltimore, Md. and (1977). Murakami, K., J. Biochem. (Tokyo), 55, 1331 (1979). Niven, D. F., J. Gen. Microbiol., 100, 5 (1977). Collins, P. A., and Knowles, C. J., Nathans, G. R., Chang, D., and Deuel, T. F., Methods Enzymol., 5t (Purine Pyrimidine Nucleotide Metab.), 497 (1978) Barsacchi, and Raggi, R. A. 3 3 Ranieri-Raggi Biochem. J. 3 , M-, 182, —— Bergamini, 361 (1979)- C. 3 72. 73. 74. 75. 76. 77. 78. 79. 80. 81. 82. 83. 84. 85. 86. 87. 88. 89. 204 Lowenstein, J. M., Physiol. Rev., 5g, 382 (1972). Ogasawara, N., Goto, H., Yamada, Y., Biochim., Bio- phys. Acta, 524, 442 (1978). Shiraki, H., Ogawa, H., Matsuda, Y., and Nakagawa, H., Biochim. Biophys. Acta, 566, 345 (1979). Shiraki, H., Ogawa, H., Matsuda, Y., and Nakagawa, H., Biochim. Biophys. Acta, 566, 335 (1979). Ashby, B., and Frieden, C., J. Biol. Chem., 252, 1869 (1977). Koretz, J., and Frieden, C., Proc. Natl. Acad. Sci., U.S.A., 11, 7186 (1980). Embden, G., and Wassermeyer, H., Z. Physiol. Chem., ttg, 226 (1928). Parnas, J. K., Biochem. Z., 206, 16 (1929). Parnas, J. K., Mozolowski, W., and Lewinski, W., Biochem. Z., 188, 15 (1927). Pennington, R. J., Nature, 192, 884 (1961). Suelter, C. R., Thompson, D., Oakley, G., Pearce, M., Husic, H. D., and Brody, M. 8., Biochem. Med., gt, 352 (1979). Young, R. B., McConnell, D. G., Suelter, C. H., Philip, T. A., Muscle Nerve, 5, 117 (1981). Husic, H. D., and Suelter, C. H., Biochem. Biophys. Res., Commun., 55, 228 (1980). Brody, M. S., Diss. Abstr. Int. B (1978). Raggi, A., Ronca-Testoni, S., and Ronca, G., Biochim Biophys. Acta, 178, 619 (1969). Raggi, A., Bergamini, C., and Ronca, G., FEBS Lett., 28. 19 (1975). Rahim, Z. R. A., Lutaya, G., and Griffiths, J. R., Biochem. J., 184, 173 (1979). Sabina, R. L., Swain, J. L., Patten, B. M., Ashizawa, T., O'Brien, W. E., and Holmes, E. W., J. Clin. Invest. 55, 1419 (1980). 90. 91. 92. 93. 9A. 95. 96. 97. 98. 99. 100. 101. 102. 103. 104. 105. 106. 107. 205 Rajendra, W., Chetty, C. S., Indira, K., and Swami, K. 8., Arch. Int. Physiol. Biochim., 88, 379 (1980). Rajendra, W., Indira, K., and Swami, K. 8., Indian J. Exp. Biol., t5, 1487 (1980). Lagerkvist, U., J. Biol. Chem., 233, 138 (1958). Abrams, R., and Bentley, M., Arch. Biochem. Biophys., 12, 91 (1959)- Davey, C. L., Arch. Biochem. Biophys., 5, 296 (1961). Lieberman, I., J. Biol. Chem., 223, 327 (1956). Tornheim, K., and Lowenstein, J. M., J. Biol. Chem., 249, 3241 (1974). Lehninger, A. L., Biochemistry; 2nd. ed., Worth Publishers, Inc., N.Y. (1977), p. 733. Kelley, W. N., Rosenbloom, F. M., Henderson, J. F., and Seegmiller, J. B., Proc. Natl. Acad. Sci. U.S., 51, 1735 (1967). Dagley, S., and Nicholson, D. E., An Introduction to Metabolic Pathways, Wiley, New York (1970). Greenberg, D. M. (ed.), Metabolic Pathways, 5, Academic Press, New York (1970). Henderson, J. F., and Paterson, A. R. P., Nucleotide Metabolism, Academic Press, New York (1972). Mahler, H. R., and Cordes, E. H., Biological Chem- istry (2nd ed.), Harper and Row, New York (1971). Hers, H. G., and Van den Berghe, G., Lancet, t, 585 (1979). Ranieri-Raggi, M., Bergamini, C., and Raggi, A., Ital. J. Biochem., 55, 238 (1980). Ronca-Testoni, S., Raggi, A., and Ronca, G., Biochim Biophys. Acta, 198, 101 (1970). Raggi, A., and Raineri—Raggi, M., Biochim. Biophys. Acta, 566, 353 (1979). Wheeler, T. J., and Lowenstein, J. M., J. Biol. Chem., 254, 8994 (1979). 108. 109. 110. 111. 112. 113. 114. 115. 116. 117. 118. 119. 120. 121. 122. 123. 124. 206 Ronca, G., Raggi, A., and Ronca—Testoni, S., Ital. J. Biochem., at, 305 (1972). Cohen, P., Control of Enzyme Activity, ed., Chapman and Hall, London (1976). Parnas, J. K., and Lewinsky, W., Biochem. Z., 276, 398 (1935)- Coffee, C. J., and Solano, C., J. Biol. Chem., 252, 1606 (1977). Chung, L., and Bridger, W. A., FEBS Lett., 55, 338 (1976). Tornheim, K., and Lowenstein, J. M., J. Biol. Chem., 2A7, 162 (1972). Mockrin, S. C., Byers, L. D., and Koshland, D. E., Jr., Biochemistry, t5, 5428 (1975). Smiley, K. L., Berry, A. J., and Suelter, C. H., J. Biol. Chem., 242, 2502 (1967). Zielke, C. L., and Suelter, C. H., J. Biol. Chem., 246, 2179 (1971). Boosman, A., and Chilson, 0. P., J. Biol. Chem., 251, 1847 (1976). Ranieri-Raggi, M., and Raggi, A., Biochem. J., 189, 367 (1980)- Stankiewicz, A., Spychala, J., Skladanowski, A., and Zydowo, M., Comp. Biochem. Physiol. B, 62B, 363 (1979). O'Driscoll, D., Roring, N. P., and Ross, C. A., Biochem. Soc. Trans., 5, 928 (1976). Ogasawara, N., Goto, H., Yamada, Y., and Yoshino, M., Biochem. Biophys. Res. Commun., 15, 671 (1977). Shiraki, H., Ogawa, H., Matsuda, Y., and Nakagawa, H., Biochim. et Biophysica Acta, 566, 335 (1979). Yoshino, M., Kawamura, Y., and Ogasawara, N., J. Biochem., 55, 299 (1976). Skladanowski, A., Kaletha, K., and Zydowo, M., Int. J. Biochem., t5, 865 (1981). 125. 126. 127. 128. 129. 130. 131. 132. 133. 134. 135. 136. 137. 138. 139. 140. 141. 207 Stankiewicz, A., and Spychala, Y., Comp. Biochem. Physiol. B, 698, 5 (1981). Weber, G., in: Molecular Biophysics, eds. B. Pullman and M. Weissbluth (Academic Press, New York) p. 369 (1965). Kurganov, B. I., Dorozhko, A. I., Kagan, Z. S., and Yakovlev, V. A., J. Theor. Biol., 55, 247 (1976). Fersht, A., Enzyme Structure and Mechanism, Freeman, New York (1977). Raggi, A., Ranieri, M., Taponeco, G., Ronca-Testoni, S., Ronca, G., and Rossi, C. A., FEBS Lett., t5, 101 (1970). Ogasawara, N., Goto, H., and Yamada, Y., Adv. Exp. Med. Biol., 1228, 169 (1980). [Purine Metabolism. Man. - 3:Biochem., Immunol. Cancer Res.]. Sammons, D. W., Diss. Abstr. Int. B (1977). Sammons, D. W., and Chilson, 0. P., Arch. Biochem. Biophys., 191, 561 (1978). Hampton, A., Sasaki, T., Perini, F., Slotin, L. A., and Kappler, F., J. Med. Chem., t5, 1029 (1976). Lazarus, R. A., Benkovic, P. A., Benkovic, S. J., Arch. Biochem. Biophys., 197, 218 (1979). Dudycz, L., and Shugar, D., FEBS Lett., 107, 365 (1979). Stankiewicz, A., Spychala, J., Makarewicz, W., Comp. Biochem. Physiol. B., 66B, 529 (1980). Doree, M., and Terrine, C., Compt. Rend. Acad. Sci. Ser. D, 275, 1503 (1972). Zielke, C. L., and Suelter, C. H., J. Biol. Chem., 246, 1313 (1971). Ashman, L. K., and Atwell, J. L., Biochim. Biophys. Acta, 258, 618 (1972). Setlow, B., and Lowenstein, J. M., J. Biol. Chem., 243, 6216 (1968). Citri, N., Adv. Enzymol., 31, 397 (1973). 142. 143. 144. 145. 146. 147. 148. -l49. 150. 151. 152. 153. 154. 155. 156. 157. 158. 159. 208 Nachod, F. C., and Wood, J., J. Am. Chem. Soc., 51, 629 (1945). Pauling, L., The Nature of the Chemical Bond, Cor- nell University Press, Ithaca, New York (1960), p. 514. Suelter, C. H., Science, 168, 789 (1970). Suelter, C. H., Metal Ions in Biological Systems, 5, High M-lecular Complexes. Editor - Helmut Sigel; Marcel Dekker, Inc., New York(l974). Smiley, K. L., Jr., and Suelter, C. H., J. Biol. Chem., 242, 1980 (1967). Campbell, J. C. W., and Suelter, C. H., Biochemistry, t5, 4836 (1977). Lee, Y.-P., J. Biol. Chem., 527, 999 (1957). Nikiforuk, G., and Colowick, S. P., J. Biol. Chem., 219, 119 (1956). Kaletha, K., Stankiewicz, A., Makarewicz, W., and Zydowo, M., Int. J. Biochem., Z, 67 (1976). Henry, H., Chilson, 0. P., Comp. Biochem. Physiol., .32, 301 (1969). Kizer, D. E., Cox, B., Lovig, C. A., and DeEstrugo, S. F., J. Biol. Chem., 238, 3048 (1963). Lee, Y.-P., and Wang, M. H., J. Biol. Chem., 243, 2260 (1968). Askari, A., Mol. Pharmacol., g, 518 (1966). Setlow, B., and Lowenstein, J. M., J. Biol. Chem., 243, 3409 (1968). Turner, D. H., and Turner, J. F., Biochem. J., 15, 143 (1961). Makarewicz, W., Comp. Biochem. Physiol., 52, 1 (1969). Nara, S., Siekagaku, 3g, 204 (1960); CA 55, 5815 (1964)- Tomozowa, Y., and Wolfenden, R., Biochemistry, 5, 3400 (1970). 160. 161. 162. 163. 164. 165. 166. 167. 168. 169. 170. 171. 172. 173. 174. 175. 176. 177. 209 Skladanowski, A., Kaletha, K., and Zydowo, M., Int. J. Biochem., 0, 177 (1979). Melander, W. R., Arch. Biochem. and Biophys., 64, 90 (1974)- Zielke, C. L., Ph.D. Dissertation, Michigan State University, East Lansing, Michigan (1970). Askari, A., Nature, 202, 185 (1964). Brown, W. E. L., and Hill, A. V., Proc. Roy. Soc. (London), Ser. B., 55, 297 (1922-1923). Zielke, C. L., and Suelter, C. H., Fed. Proc., 55, 728 (1969). Kuhne, w., Ber., 206 (1875). Nencki, M., Ber., 336 (1875). Hopkins, F. G., and Cole, W. S., J. Physiol., 55, 451 (1903). HOpkins, F. G., and Cole, W. S., J. Physiol., 51, 418 (1901). Happold, F. C., and Hoyle, L., Biochem. J., 55, 1218 (1935). Happold, F. C., and Struyvenberg, A., Biochem. J., 5&. 379 (1954). Wood, W. A., Gunsalus, I. C., and Umbreit, W. W., J. Biol. Chem., 170, 313 (1947). Happold, F. C., Adv. Enzymol. Relat. Subj. Biochem., 19. 51 (1950). Wada, H., Tryptophan Metabolism Vol. 1, Sekai, Hoken Tsushinsha, Ltd., Osaka, Japan, 77-92 (1964). Snell, E. E., Adv. Enzymol. Relat. Areas Mol. Biol., 3;, 287 (1975). Newton, W. A., and Snell, E. B., Proc. Natl. Acad. Sci. U.S.A., 55, 1431 (1962). Yanofsky, C., and Crawford, I. P., Proc. Nat. Acad. Sci. U.S.A., 55, 1016 (1959). 210 178. Newton, W. A., and Snell, E. E., J. Bact., 55, 355 (1965). 179. Newton, W. A., and Snell, E. E., Proc. Natl. Acad. Sci. U.S.A., 5t, 382 (1964). 180. Kagamiyama, H., Wada, H., Matsubara, H., and Snell, E. E., J. Biol. Chem., 2 7, 1571 (1972). Y., 181. Newton, W. A., Morino, and Snell, E. E., J. Biol. Chem., 240, 1211 (1965). 182. Morino, Y., and Snell, E. E., J. Biol. Chem., 242, 5591 (1967). 183. Hogberg-Raibaud, A., Raibaud, 0., and Goldberg, M. E., J. Biol. Chem., 250, 3352 (1975). 184. Morino, Y., and Snell, E. E., J. Biol. Chem., 242, 2800 (1967). 185. Snell, E. E., and DiMari, S. J., in The Enzymes, Vol 2, 3rd ed., P. D. Boyer, Ed., Academic Press, New York, p- 335 (1970)- 186. Kagamiyama, H., Morino, Y., and Snell, E. E., J. Biol. Chem., 245, 2819 (1970). 187. Raibaud, 0., and Goldberg, M. E., J. Biol. Chem., 258: 3451 (1973)- 188. Suelter, C. H., and Snell, E. E., J. Biol. Chem., 252, 1852 (1977)- 189. Heinert, D., and Martell, A. E., J. Am. Chem. Soc. 93, 3257 (1962). 190. Johnson, R. J., and Metzler, D. E., Methods in Enzym- ology, 18A, 443 (1970). 191. Davis, L., and Metzler, D. E., in The Enzymes (Boyer, P. D., ed.) Vol. 7, pp. 33-74 (1972). 192. Metzler, C. M., Cahill, A., and Metzler, D. E., J. Am. Chem. Soc., 102, 6075 (1980). 193. June, D. S., Kennedy, B., Pierce, T. H., Elias, S. V., Halaka, F., Behbahani-Nejad, I., El-Bayoumi, A., Suelter, C. H., and Dye, J. L., J. Am. Chem. Soc., 191. 2218 (1979). 194. June, D. S., Suelter, C. H., and Dye, J. L., Bio- chemistry, _5, 2707 (1981). 195. 196. 197. 198. 199. 200. 201. 202. 203. 204. 205. 211 June, D. S., Ph.D. Dissertation, Michigan State Uni- versity, East Lansing, Michigan (1979). Hillebrand, G. G., Dye, J. L., and Suelter, C. H., Biochemistry, t5, 1751 (1979). June, D. S., Suelter, C. H., and Dye, J. L., Bio- chemistry, 55, 2714 (1981). Watanabe, T., and Snell, E. E., J. Biochem. (Tokyo), 93, 733 (1977)- Suelter, C. H., Wang, J., and Snell, E. E., Anal. Biochem., 15, 221 (1977). Suelter, C. H., Wang, J., and Snell, E. E., FEBS Lett., 55, 230 (1976). Cochran, R. N., and Horne, F. H., Anal. Chem., 55, 846 (1977). Cochran, R. N., and Horne, F. H., J. Phys. Chem., 55, 2561 (1980). Cochran, R. N., Horne, F. H., Dye, J. L., Ceraso, J. and Suelter, C. H., J. Phys. Chem., 55, 2567 (1980). Ivanov, V. I., and Karpeisky, M. Y., Adv. Enzymol. Relat. Areas Mol. Biol., 55, 21 (1969). Coetzee, W. F., and Pollard, E. C., Photochem. Photobiol., 22, 29 (1975). , .1 flf-‘N-‘Q‘W T MAJ 'Molaq padwels 319p aql 43139 anJniaJ s; nooq 31 pafiueqa aq [11M SBNId 'pJoan Jnox NOJ} inoaoaqa siqi aAomaJ 01 dOJp xooq u; aaeld 3S1V1831VN 9N1N80138 -—a-—r SEIHV‘HQI‘I {18W