THE ROLE or ’DXYGEN'IN PHOTORESPIRATIONi-f. '7 7‘- iii-3'} Thesis for the Degree of Ph. D. MICHIGAN STATE UNIVERSITY GEORGE HUNTLY'LORIMER' ’ 19.72 "“HESll This is to certify that the thesis entitled THE ROLE OF OkYGEN IN PHOTORESPIRATION presented by George Huntly Lorimer has been accepted towards fulfillment of the requirements for Ph. D. degree in Biochemistry 77. 6‘, “747417 Major professor Date March 10, 1972 0-7639 ABSTRACT THE ROLE OF OXYGEN IN PHOTORESPIRATION By George Huntly Lorimer The role of oxygen in photorespiration, in the inhibition of net photosynthetic CO2 fixation, the warburg effect, and in the synthesis and metabolism of glycolate has been investigated. Oxygen consumption occurs during P-glycolate formation in chloroplasts and glycolate oxidation in peroxisomes. Glycolate oxidase (E.C. 1.1.3.1) was purified to homogeneity from.the peroxisomes of spinach leaves. Glycolate oxidase catalysed the oxidation of glycolate to g1yoxy1ate, glyoxylate to oxalate and L-lactate to pyruvate. The KM(02) for each of these three reactions was the same, namely, about 1.9 x 10-4M at 25°, which suggests that oxygen reacts with the same intermediate in each reaction. The energy of activation (Ea) for each substrate with oxygen as the terminal electron acceptor was 11.5 Kcal. It was concluded that the overall reaction is dependent upon the steady state concentration of the reduced flavin enzyme complex. The apoenzyme of glycolate oxidase was prepared by treatment with 1M KBr. The reactivation of the George Huntly Lorimer apoenzyme with flavin mononucleotide (FMN) was followed spectrophotometrically. in 31.19 and j._r_r 11.533 experiments with 1802 dealt with the second site of oxygen consumption in photorespiration, namely that involved with glycolate synthesis. The technique of combined gas chromatography - mass Spectrometry was used. When detached spinach leaves were exposed to an atmosphere of 100% oxygen containing 1802, a single atom of oxygen-18 was rapidly incorporated into the carboxyl group of glycine and serine, the metabolic derivatives of glycolate. This incorporation occurred only in the light as a photorespiratory event. No label was incorporated into the hydroxyl group of serine which is derived from water. Under the same conditions, glycerate and phosphoglycerate did not become labeled. The data was consistent with glycolate formation by a two or a four electron oxidation of a sugar phoSphate from the photosynthetic carbon cycle. A large pool of erythronic (or threonic) acid was also identified but it did not become labeled either 14 with oxygen-18 or with carbon-14 during a 10 min C0 2 photosynthesis - photorespiration experiment. [u-IACJ-ribulose diphOSphate was synthesized for use as a precursor of phosphoglycolate. In the presence of Nb ions and hydrogen peroxide, ribulose diphOSphate was non-enzymatically oxidised to phospho- glycolate and phosphoglyceraldehyde. A previous report George Huntly Lorimer that ribulose diphosphate carboxylase (E.C. 4.1.1.39) catalysed the oxidation of ribulose diphOSphate by molecular oxygen to phosphoglycolate was confirmed by 64C}ribulose diphosPhate. The other product the use of was proven to be phosphoglycerate. A manometric assay to follow oxygen consumption by this reaction was developed. Spinach leaf ribulose diphosPhate carboxylase was purified by two methods to homogeneity as revealed by polyacrylamide disc gel electrophoresis. The first method included (NH4)2304 fractionation, zonal centrifugation in a sucrose density gradient and hydroxylapatite column chroma- tography. The second method included DEAE-cellulose column chromatography and zonal centrifugation. Both purified preparations catalysed the oxidation of ribulose diphosphate. This activity has been named ribulose diphosphate oxygenase. Ribulose diphosphate carboxylase co-purified with ribulose diphosphate oxygenase. Several lines of evidence were marshalled to evaluate the homogeneity of the purified preparation but it has not been unequivocally established that the two activities are associated with one and the same protein. During storage in ammonium sulfate the oxygenase was generally. more stable than the carboxylase. Under these conditions ribulose diphOSphate carboxylase underwent polymerisation. Oxygen uptake by ribulose diph03phate oxygenase was a linear function of time and enzyme concentration. George Huntly Lorimer The oxygen uptake depended upon the presence of ribulose diphosphate, Mg2+, oxygen and enzyme. Enzyme which had been boiled for 2 min was inactive. The activity was stimulated by, but not absolutely dependent upon, the presence of dithiothreitol. This stimulation was not due to the oxidation of dithiothreitol. The pH optimum for oxygen uptake was about 9.3. The stoichi- ometry of oxygen consumption to ribulose diphosphate consumption was one to one. The oxygenase activity was specific for ribulose diphosphate. Ribulose-S-phosphate, ribulose, fructose-6-ph03phate, fructose-1,6-diphosphate and 3-ph03phoglycerate were inactive when tested with 14C the standard manometric assay and/or by the use of labeled substrates. The absence of activity with 3- phosphoglycerate established that the reaction with ribulose diphosphate did not first proceed by carboxy- lation followed by oxidation of the resultant 3-phOSph0- glycerate. Under 100% oxygen the KM for ribulose diphOSphate was about 1.5 x 10-4M. The reaction products of ribulose diphosphate oxygenase were identified by gas chromatography - mass Spectrometry of the trimethylsilyl derivates of phospho- glycolate and phosphoglycerate. Experiments with 1802 and H2180 were performed with the enzyme system. The results were consistent with those obtained.ig;gigg. Oxygenation of ribulose diphoSphate proceeds with the incorporation of a single atom of oxygen, derived from George Huntly Lorimer molecular oxygen, into the carboxyl group of phospho- glycolate. No incorporation occurred into phospho- glycerate. By experiments with H2180, the carboxyl oxygen of phOSphoglycerate was shown to be derived from water. The enzyme did not catalyse the exchange of the carboxyl oxygens of phosphoglycolate or phosphoglycerate with those of the medium. A mechanism consistent with these observations, involving the formation of a ribulose diphosphate peroxide intermediate, was proposed. The overall reaction is 2+ Ribulose-1,5-diphosphate + 02-—————C>2-Phosphog1ycolate + 3-Phosphoglycerate Phosphoglycolate biosynthesis from ribulose diphOSphate and its subsequent oxidative metabolism serves as a biochemical model to explain photoreSpiration. THE ROLE OF OXYGEN IN PHOTORESPIRATION By George Huntly Lorimer A THESIS Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Biochemistry 1972 o’l‘" ACKNOWLEDGMENTS I am grateful to Prof. N. E. Tolbert, who, in addition to supporting me financially, has given me encouragement, advice and considerable latitude at all times. Many of the experiments described in this thesis were performed with the cooperation of Dr. T. John Andrews. It is a pleasure to acknowledge the many fruitful hours of discussion which usually preceded and followed each experiment. The excellent technical assistance of Jack Harten, Sandra wardell and Diana Wied in various aSpects of the work is to be noted. My introduction to the technique of mass spectrometry was made painless by the patience of Dr. Ray Hammond, Dr. Roger Laine and Mark Bieber. I am solely responsible for any errors in the interpretation of the mass spectra. Thanks are also due to Dr. K. Raschke and Dr. P. Wo1k of the MSU - AEC Plant Research Laboratory for the use of their equipment, and to Dr. W; L. Ogren, USDA Soybean Laboratory, Urbana, Illinois, for the gift of purified soybean ribulose diphosphate carboxylase. ii Finally I should like to record the cheerfulness and independence of spirit of my wife, Freia. Her patience and encouragement are especially appreciated. iii TABLE OF CONTENTS LIST OF TABLES. LIST OF FIGURES . LIST OF ABBREVIATIONS INTRODUCTION. LITERATURE REVIEW . The C- 3 Photosynthetic Carbon Reduction Cycle . The C- 4 Dicarboxylic Acid Pathway . The Warburg Effect. Photorespiration. . . The Role of Glycolate in Photorespiration . Glycolate Biosynthesis. . . . . . . Ribulose Diphosphate Carboxylase. . . Oxidation of Carbohydrates and Related Compounds by Oxygen . . . . . Oxygenases. . . Chapter I. STUDIES ON PEROXISOMAL GLYCOLATE OXIDASE. Introduction. Materials . Methods . Protein Determination. Glycolate Oxidase Assays . . Calibration of the Oxygen Electrode. Isolation of Spinach Peroxisomes . Purification of Glycolate Oxidase from Spinach Peroxisomes. . Polyacrylamide Gel Electrophoresis of Glycolate Oxidase. . . . . Determination of the Km for Oxygen . Determination of the Energies of Activation . Preparation and Reactivation of the Apoenzyme of Glycolate Oxidase . iv Page . viii ix xii \lUl-PDJUJ 00 l-' 11 26 34 39 39 4O 40 40 4O 42 42 43 44 44 45 Chapter Page Results and Discussion. . . . . . . . . . . 45 Purification of Glycolate Oxidase from Spinach Peroxisomes. . . . . . 45 Polyacrylamide Gel Electrophoresis . . . 48 Visible Spectrum of Glycolate Oxidase. . 48 Determination of the for Oxygen . . . 51 Determination of the Energies of Activation . . . . 52 Preparation and Reactivation of. the Apoenzyme of Glycolate Oxidase . . . . 57 II. INCORPORATION OF MOLECULAR OXYGEN INTO GLYCINE AND SERINE DURING PHOTORES- PIRATION IN SPINACH LEAVES. . . .1. . . . 69 Introduction. . . . . . . . . . . . . . . . 69 ‘Materials . . . . . . . . . . . . . . . . . 71 Mbthods . . . . . . . . . . . . . . . . . . 7l Photorespiration in [180] Oxygen . . . . 7l Extraction of Glycine and Serine . . . . 72 Extraction of Organic Acids. . . . . . . 73 Preparation of‘ge Si Derivatives18 . . 74 Synthesis of [1 0? Glycine and E 01 Serine . . . . . . 74 Synthesis of [130] Glycerate . . . . . . 74 Isotope Analysis . . . . . . . . . 74 Results . . . . . . . . . . . . . . . . . . 76 Gas Chromatography-Mass Spectrometry of Glycine and Serine . . . 76 Gas ChromatographyeMass Spectrometry of the Organic Acid Fraction. . . . 77 Investigation of Egtraction Procedure. . 88 Incorporation of 0 into Glycine and Serine . . . . . 90 Lack of Incorporation of 180 into Glycerate and other Organic Acids. . . 98 Discussion. . . . . . . . . . . . . . . . . 98 III. ERYTHRONIC ACID - A [1401002 WILD GOOSE CHASE . . . . . . . . . . . . . . . . . . 108 Introduction. . . . . . . . . . . . . . . . 108 Materials . . . . . . . . . . . . . . . . . 110 Chapter Page Methods . . . . . . . . . . . . . . . . . . 110 The [Mojco2 Photosynthesis - Photo- respiration Pulse Chase Method . . . . llO Fractionation of the Aqueous Extract . . 113 Paper Chromatography . . . . . . . . . . 114 Autoradiography. . . . . 115 High Voltage Paper ElectrOphoresis . . . 115 Gas Liquid Chromatography - Mass Spectrometry . . . . . 115 Gas Liquid Chromatography - Radio- activity'Measurements. . . . . . . . 116 Liquid Scintillation Counting. . . . . . 116 Results and Discussion. . . . . . . . . . . 116 IV. A MODEL SYSTEM FOR THE OXIDATION OF RIBULOSE DIPHOSPHATE TO PHOSPHOGLYCOLATE. 123 Introduction. . . . . . . . . . . . . . . . 123 ‘Methods . . . . . . . . . . . . . . . . . . 123 Preparation of [U-14CJ-Ribulose Diphosphate. . . . . . . . 124 Manganese Catalysed Oxidation of Ribulose Diphosphate by Hydrogen Peroxide . . . . . . . 126 Results . . . . . . . . . . . . . . . . . . 127 Identification of Reaction Products. . . 127 V. THE ENZYMATIC OXIDATION OF RIBULOSE DIPHOSPHATE BY MOLECULAR OXYGEN . . . . . 135 Introduction. . . . . . . . . . . . . . . . 135 MBterials . . . . . . . . . . . . . . . . . 136 Mbthods . . . . . . . . . . . . . . . . . . 136 Exploratory Assays . . . . . . . . . . . 136 RuDP Oxygenase Assay . . . . . . . . . . 137 RuDP Cizboxylase Assays. . . . . . . . . 139 (a) C Assay. . . . . . . . . 139 (b) Spectrophotomeizic Assay. . . . . 140 Standardization of [ C] NaHCO3. . . . . 141 Standardization of RuDP. . . . . 142 Protein Determination. . . . . . . . . . 142 vi Chapter Page Polyacrylamide Gel Electrophoresis . . . 142 Analytical Ultracentrifugation . . . 143 Identification of Reaction Products by Mass Spectrometry. . . . . . . . . . . 143 Results . . . . . . . . . . . . . . . . . . 144 Purification of RuDP Carboxylase from Spinach Leaves . . . . 145 Alternative Procedure for the Purifica- tion of RuDP Carboxylase . . . . . 154 Assessment of Purity and Homogeneity . . 162 RuDP Oxygenase Assay . . . . . . . . . . 168 pH Optimmm . . . . . 168 The Stoichiometry of the Reaction. . . . 176 Proof of Reaction Products . . . . . . . 176 Analysis of the Mass Spectra of (Me Si) P- -g1ycolate . . . . . . . 185 Analyéis 3f the Mass Spectra of (Me Si)4 P-glycerate . . . . . . . . . 186 Substgate4 Specificity. . . . . 187 Mechanisrii8 of RuDP Oxygenafg Studied with 18 80] Oxygen and [ OJ Water. . . 189 (a) [ 80] Oxygen . . . . . . . 189 (b) [180] water. . . . . 198 Summary of Other Properties .of RuDP Oxygenase. . . . . . . 200 Activity as a Function of Oxygen Concentration. . . . . . . . 200 Activity as a Function of RuDP Concentration. . . . . . . . . . . . . 201 Stability. . . . . . . . . . . . . 201 Cyanide Inhibition . . . . . 202 Copper Content of RuDP Carboxylase . . . 202 Reaction Intermediate Derived from Oxygen . . . . . . . . . . . . . . . . 203 Discussion. . . . . . . . . . . . . . . . . 203 Mechanism of RuDP Oxygenase. . . . . . . 203 Enzyme Purity. . . 206 The Relationship of RuDP Oxygenase to Photorespiration . . . . . 210 CONCLUDING DISCUSSION . . . . . . . . . . . . . . . 215 BIBLIOGRAPHY. . . . . . . . . . . . . . . . . . . . 218 vii Table 10. 11. LIST OF TABLES Purification of Glycolate Oxidase from Spinach Peroxisomes. . . . . . Retention of 180 During Extraction Procedures . . . . . . . . . Incorporation of 180 into Serine . Incorporation of 180 into Glycine and Serine as a Function of Time . Recovery of Radioactivity During Fractionation. . . . . . . . Reaction Requirements for the Manganese Catalysed Oxidation of Ribulose Diphos- phate by Hydrogen Peroxide . . . . . Purification of RuDP Carboxylase from Spinach Leaves . . . . . Co-purification of RuDP Carboxylase and RuDP Oxygenase . . . . . . . . . Oxygen Uptake by RuDP Oxygenase. Substrate Specifity of RuDP Oxygenase. The Mechanism of RuDP Oxygenase. viii Page 46 89 96 97 114 133 153 161 175 188 191 Figure 10. ll. 12. 13. LIST OF FIGURES The glycolate pathway . The reaction of glucose and fructose with molecular oxygen in alkaline solution . Visible spectrum of glycolate oxidase . Lineweaver-Burk plot for the determination of the K for oxygen of glycolate oxidase for each of the three substrates. . Arrhenius plot for the catalytic activity of glycolate oxidase with oxygen and each of the three substrates. Reactivation of the apoenzyme of glycolate oxidase with FMN. . . . . . . . . . Reactivation of the apoenzyme of glycolate oxidase. First order plot the results. Lineweaver-Burk plot for the determination of the KM for FMN . Double reciprocal plot of the first order rate constants for the reactivation of the apoenzyme of glycolate oxidase by FMN . . . . . . . . . . . . . . . . . Gas chromatography of a silylated extract from spinach leaves containing glycine and serine. . . . . . (ag‘Mass Spectrum.of (Me3Si) Glycine b 'Mass Spectrum of “Me3Si)3 Serine. Gas chromatography of a silylated extract from Spinach leaves containing organic acids . (a)-(c) Mass Spectrum of (MeBSi)2 oxalate, (Me Si) glycerate and Me3Si)2 sucgina e . . . . . . . . . . . ix Page 10 29 50 54 56 59 62 64 67 79 81 83 85 Figure Page 13. (d)-(e) Mass Spectrum of (MeBSi)3 malate and (Me 3Si)4 erythronate. . . . . . . . . 87 14. Comparison of part of the mass Spectrum of authentic (Me Si)3 glycine with that of the same compgund3 extracted from a leaf which had been exposed /t$ 100% oxygen containing 18.1 atoms 7. 80 for 2 min in the light. . . . . . . . . . . . 92 15. Comparison of part of the mass Spectrum of authentic (Me Si)3 serine with that of the same compgund3 extracted from a leaf which had been exposed t1 100% oxygen containing 18.1 atoms % 80 for 2 min in the light. . . . . . . . . . . 94 16. The mechanism of action of transketolase and the proposed mechanism of oxidation of a,B-dihydroxyethylthiamine pyrophos- phate to glycolate and thiamine pyrophosphate . . . . . . . . . . . . . . 106 17. Apparatus for the [1 4CJCOZ photosynthesis- photorespiration' 'pulse- -chase" . . . . . 112 18. The fixation of [IACJCOZ in photosynthesis and 1Ehe subsequent photorespiratory loss . . . . . . . . . . . . . . 118 19. Gas-liquid radiochromatogram of the sily- lated organic acids extraized from a Spinach leaf exposed to [ CJCO2 for 5.5 min followed by 100% oxygen for 1 min . . 121 20. (a) Paper chromatography of the reaction products of them mlHZOZ oxidation of RuDP. . . . (b) Re- -chromatography of peak 11 of the chromatogram shown in Fig. 20a. . . . 129 (c) Paper chromatography of the reaction products of the Mn/H 02 oxidation of RuDP, after treatmen with alkaline phOSphatase . . . . . . . . . . . . . 132 129 21. Zonal centrifugation of RuDP carboxylase after (NH4)2804 fractionation . . . . . . 149 22. Hydroxylapatite column chromatography of RuDP carboxylase. . . . . . 152 Figure Page 23. Co-purification of RuDP carboxylase and RuDP oxygenase by DEAE cellulose column chromatography . . . . . . . . . . 157 24. Co-purification of RuDP carboxylase and RuDP oxygenase by zonal centrifugation. . 160 25. Polyacrylamide gel electrophoresis of purified RuDP carboxylase . . . . . . . . 164 26. Schlieren patterns of purified RuDP car- boxylase obtained in the Spinco Model E ultracentrifuge . . . . . . . . . . . . 167 27. Oxygen consumption by RuDP oxygenase as a function of time. . . . . . . . . . . . . 170 28. Oxygen consumption by RuDP oxygenase as a function of enzyme concentration. . . . . 172 29. The pH optimum.of RuDP oxygenase. . . . . . 174 30. The stoichiometric consumption of oxygen by RuDP oxygenase. . . . . . . . . . . . . . 178 31. (A) Gas liquid chromatographic separation of the silylated reaction products from the RuDP oxygenase reaction. . . 180 (B) The trimethylsilyl derivatives of authentic P-glycolate (III) and P-g1ycerate (IV). . . . . . . 180 32. Mass Spectrum of (MeBSi)3-P- glycolate (70 eV. ). . . 182 33. Mass Spectrum of (Me 3Si)4-P-glycerate (70 eV. ). . . 184 34. Comparison of part of the mass Spectrum of authentic (Me Si)3 -P- -glycolate (a) with that of the sam e compound isolated from an incubigion of RuDP with the enzyme in 82% 0] oxygen (b) . . . . . . . . 195 35. Comparison of part of the mass spectrum of (Me3 Si) -P- -glycerate (a) with that of the3 sam 2 compound isolated from an incu- bagion of RuDP with the enzyme in 82% 0] oxygen (b). . . . . . . . 197 xi ne— iii. ’L’ Ammediol BSTFA DCPIP DEAE DTT EDTA a GPD a KG Me3Si PCMB 3-PGA POPOP PPO RuDP SDS TEAE THFA TPI Tris LIST OF ABBREVIATIONS 2-amino-2-methyl-1,3-propanediol bis(trimethylsi1yl)trifluoroacetamide dichlorophenolindophenol diethylaminoethyl dithiothreitol ethylenediamine tetraacetate a glycerophoSphate dehydrogenase iodoacetamide a-ketoglutarate trimethylsilyl para-chloromercuribenzoate 3-phosphoglycerate phenyl-oxazoly1phenyl-oxazolylphenyl 2,5-diphenyloxazole ribulose-1,5-diph08phate sodium dodecyl sulfate triethylaminoethyl tetrahydrofolic acid triose phosphate isomerase tris(hydroxymethyl)amino methane xii INTRODUCTION In 1920 the late Otto warburg observed that oxygen inhibited net photosynthetic C02 fixation in Chlorella. This phenomenon, now referred to as the warburg effect, has been observed in a wide variety of photosynthetic tissues. The warburg effect can be explained, at least in large part, by photorespiration, the light dependent uptake of O2 and release of C02 that occurs in the photosynthetic tissues of all higher plants. Photorespiration is the physiological mani- festation of the glycolate pathway of metabolism, which has been the subject of extensive investigations by Tolbert, Zelitch, Bassham, Whittingham and their asso- ciates. The glycolate pathway is an important adjunct to the photosynthetic carbon reduction cycle and estimates of the quantity of carbon flowing through this pathway under natural conditions range from 30 to 90% of the total carbon fixed during photosynthesis. The function of the glycolate pathway is at present obscure. Whatever the function, it results in the tqmake of 02 and release of C02, thus decreasing the efficiency of photosynthesis and ultimately of growth. This research reported in this thesis has 2 concentrated on the role of oxygen in photorespir- ation. Chapter I reports some modest and rather diverse observations on one of the enzymes involved in photorespiratory 02 uptake, glycolate oxidase. Chapter II reports the discovery of a second site of oxygen consumption, that involved in the synthesis of glycolate. The succeeding chapters describe the research which arose out of the results reported in Chapter II and also from the report Bowes g£.g1., (1971) that ribulose diphosphate carboxylase catalysed the oxidation of ribulose diphosphate to P-glycolate and P-glycerate. LITERATURE REVIEW The 0-3 Photosynthetic Carbon Reduction Cycle Since its enunciation by Calvin and his associates in the mid-1950's, the C-3 photosynthetic carbon reduction cycle has been the subject of several books (Calvin and Bassham, 1962 and Zelitch, 1971), symposia (Tolbert, 1963; San Pietro 25 $1., 1967; Goodwin, 1967 and Hatch gEH§1., 1970) and reviews (Stiller, 1962; Bassham, 1964; Gibbs, 1967 and Walker and Crofts, 1970). Although a number of nagging questions remain unanswered, there has been no major change in the overall Scheme. The pathway is ubiquitous in all higher plants and algae. It is quantitatively the most important carbon dioxide fixing, gluconeogenic system. The 6-4 Dicarboxylic Acid Pathway The discovery of the C-4 dicarboxylic acid pathway (see review by Hatch and Slack, 1970) was accom- panied by Speculation that this might represent an altogether different pathway of C02 fixation. Although this pathway is undoubtedly important, Subsequent work (ijrkman and Gauhl, 1969; Moss and Rasmussen, 1969; Slack §£H§1., 1969 and Hatch, 1971) has indicated that 3 4 the most probable function of the C-4 dicarboxylic acid pathway is that of a C02 concentrating mechanism. Thus, 002 is initially fixed as malate, oxalacetate or asPartate in the mesophyll cells of C-4 plants. These acids are then transported to the bundle sheath cells where they undergo decarboxylation, the 002 released being efficiently re-fixed by the classic C-3 photo- synthetic carbon reduction cycle. The Warburngffect warburg (1920) first observed that oxygen inhibits photosynthetic C02 fixation in Chlorella. This phenomenon has subsequently been observed in a wide variety of algae and higher plants, (see review by Turner and Brittain, 1962) and isolated chloroplasts (Arnon g£H§1., 1954 and Ellyard and Gibbs, 1969). The investigations of Gaffron (1940), McAlister and Myers (1940), Tamiya and Huzisige (1949) and Turner $5.21., (1956), established a number of general features con- cerning the phenomenon. (a) The inhibition affects not only the net C02 uptake but also the net oxygen output. (b) The magnitude of the inhibition is markedly depend- ent upon the light intensity, the C02 concentration and the oxygen concentration. In general, inhibition is greatest under saturating light intensities, limiting C02 concentrations and high oxygen concentrations. 5 The inhibition can be substantially reduced by saturating concentrations of C02. (c) The inhibition is reversible, thus distinguishing it: ((1) The inhibition is not Simply due to an increase from irreversible photo-oxidation phenomena. in the rate of "dark" (mitochondrial) reSpiration, s ince this process is saturated by about 2% oxygen, whereas the Warburg effect continues to increase between 20 and 100% oxygen. Rh 0 torespiration Photorespiration is defined as the light dependent uptake of oxygen and release of C02 that 0c(liars in the photosynthetic tissues of all higher plants. The subject has been comprehensively reviewed by Jackson and Volk (1970). Measurements of the rate of photorespiration are based upon a variety of indirect methods. Owing to the internal re-cycling of the gases involved, the Se measurements all underestimate the true rate of I3.":101mre3piration. Nevertheless, it is clear that .s ignificant changes in the reSpiratory processes occur up (311 illumination such that under certain conditions the rate of photorespiration may be a significant fraction of the rate of photosynthesis. Higher plants fall into two classes based upon thieir respiratory responses to illumination: 6 (a) C-3 Plants: These plants fix C02 primarily by means of the C-3 photosynthetic carbon reduction cycle. They are capable of high rates of photoreSpiration, whether this is measured by 002 release or by oxygen * uptake. Their C02 compensation point is high, between 40 - 50 ppm. (b) C-4 Plants: These plants possess the C-4 dicar- boxylic acid pathway in addition to the C-3 photosyn- thetic carbon reduction cycle, as outlined in page 1. Due to extremely efficient CO2 fixation, it has proven difficult to demonstrate photoreSpiratory C02 release from these plants. However, Jackson and Volk (1969) have demonstrated unequivocally that increased oxygen uptake does indeed occur in these plants. These plants are further characterized by their low CO2 compensation p0 iSluts, from 1 to 5 ppm. Whatever the method used to measure photo- res‘piration, the results lead to the same conclusion, namely, that photorespiration exceeds that occurring in the dark. A large number of investigations have bQQn conducted in which the effects of light intensity, wa-Velength, C02 concentration, oxygen concentration and temperature have been documented (see review by \ % QThe CO compensation point is the external concentration Q0 C02 it which there is no net uptake or release of 2 7 Jackson and Volk, 1970). From these results it is evident that there exists a very close connection between photosynthesis and photorespiration. In general, the conditions required to demonstrate a maximum Warburg effect are those now known to promote photoreSpiration; i.e., saturating light intensities, limiting C02 concentrations and high oxygen concen- trations. The Warburg effect is therefore in large Part due to photoreSpiration. The substrate pools for photorespiration and dark respiration are clearly different. Goldsworthy (1966) has shown that, following a period of photo- synthesis in [14C] C02, the Specific activity of the C02 subsequently released into C02 free air in the light is 1.5 times that released in the dark. Clearly pho torespiration makes greater use of recently fixed I)l‘_1°t:osynthate than does dark reSpiration. By performing s:L'Ilfllar experiments Zelitch (1966) has shown that the Mediate substrate pools for photorespiration are 3111a 11 and rapidly turned over. '1‘ % Role of Gljcolate in Photorespiration Concurrent with the physiological investigations 0 :IE photorespiration it was recognized that a considerable DQI‘tion of the total carbon fixed during photosynthesis bagsed through the glycolate pathway. Under natural Qghditions, it has been estimated that so - 75% of the 8 total carbon fixed passes through the glycolate pathway (Zelitch, 1959 and Atkins g1; 31., 1971). Furthermore, the results of Wilson and Calvin (1955), Bassham and Kirk (1962), Pritchard £531. (1962) and Tolbert (1963) have clearly established that the physiological con- ditions which enhance glycolate synthesis and metabolism are precisely those under which photorespiration and the Warburg effect become apparent. The close relation- 3 hip between the Warburg effect, photorespiration and glycolate metabolism has been summarized by Gibbs (1969). The elucidation of the glycolate pathway (Fig. 1) has been largely due to the work of Tolbert's grOUp (Tolbert and Cohen, 1953; Richardson and Tolbert, 196 1a; Rabson g; 11;, 1962; Kearney and Tolbert, 1962; ijinez e_t a_1., 1962; Orth gt 31., 1966; Hess and Tolbert, 1966, 1967; Chang and Tolbert, 1970 and Bruin e_t. _a_1_., 19 7O). The finding that many of the enzymes associated with the glycolate pathway are located in the peroxi- Somal (microbody) fraction, distinct from both mito- chondria and chloroplasts (Tolbert, 1971) has added a further dimension to our understanding of photorespiration and the glycolate pathway. The metabolism of glycolate by both higher plants and algae has been comprehensively jbQXriewed on several occasions (Tolbert, 1963; Zelitch, 1964; Hess, 1966; Anderson, 1969; Bruin, 1969; Jackson and Volk, 1970 and Tolbert, 1971). Examination of the glycolate pathway (Fig. 1) Figure l: The Glycolate Pathway. The enzymes involved are: (1) (2) (3) (4) (5) (6) (7) (8) (9) P-glycolate phosphatase Glycolate oxidase Catalase Transaminase Glycine decarboxylase Serine hydroxymethyl transferase Transaminase Hydroxypyruvate reductase Glycerate kinase 10 ? 2105303 CHZOH \ flszNHz co OH COO” 0'2) HIO:HO C00“ COO” (3) THFA+ H20- -——NAD 002+ NH3<————>NADH2 THFA- CHZOH CHZNHZ COOH (6) ADP ATP NAD NADH2 RCNH2 RCOJ/ $5120.03 (IZHZOH cleZOH $H20H C \HQH CHOH 0-0 a fHNHz QQ OH (9) COOH (8) éOOH (7) COOH 11 reveals the most probable sites of photorespiratory oxygen uptake and C02 release. PhotoreSpiratory oxygen uptake is clearly at least in part associated with the oxidation of glycolate to g1yoxy1ate, the re action catalysed by glycolate oxidase. No other oxygen consuming, photorespiratory reactions are known, however. The oxidation of glyoxylate to oxalate, also catalysed by glycolate oxidase (Richardson and Tolbert, 1961b), is probably only of significance in nitrogen deficient plants. In the presence of amino donors glyoxylate undergoes transamination to yield glycine. In turn, glycine is converted to serine by the action of glycine decarboxylase and serine hydroxymethyl transferase (Kisaki and Tolbert, 1969, 1970 and Kisaki gt 2;, 1971). These _i_p_ 33.-3E studies and the in 137.9. studies of C(>ssins and Sinha (1966) and Marker and Whittingham, ( 1966) indicate that the carboxyl group of glycine is the most immediate source of photorespiratory 002 I: e lease. SEESlzzycolate Biosynthesis Benson and Calvin (1950) first observed that g:lycolate was among the most rapidly labeled compounds formed during [14CJC02 photosynthesis by algae. Subse- q-\1ent1y, Schou e_t a_l. (1952) demonstrated that the 12 glycolate so formed was uniformly labeled. However, despite considerable effort, the mechanism of glycolate synthesis has remained unsolved since then. In the intervening years the optimal conditions for glycolate synthesis were defined (Wilson and Calvin, 1955; Bassham and Kirk, 1962; Pritchard e_t EL, 1962 and To leert, 1963). Synthesis of glycolate and of its metabolic derivatives, glycine and serine, was found to be enhanced by saturating light intensities, 1 imiting C02 concentrations and high oxygen concen- trations. In addition, alkaline pH's (9.0 and above) greatly enhanced the formation of glycolate, glycine and serine both in 33.-2’2 (Orth e_t a_l.., 1966) and in cell free chloroplast preparations (Dodd and Bidwell, 1971). The significance of this observation was not understood. The labelling of P-glycolate during [14CJCO2 photosynthesis has been demonstrated (Benson gt a_l_., 1952). Furthermore, Richardson and Tolbert (1961a) Ila-Va described a specific phoSphatase, located in the Q11:Itoroplast, which catalyses the hydrolysis of P- glycolate to glycolate. Thus, in 111.19, glycolate may a): ise from P-glycolate. However, whether it does so Q3 (3‘0 ———«> + H-C-OH H HC-OH Hfi‘OH H20 \ CH 0P0 CH20P03 {55%)CH20P03 2 3 (lsnzopo3 H-f-OH c 0/'\0H $ + 0 0H / ‘$ HC-OH CH20P03 Thus, enolization (1) is followed by the attachment of C02 to the C-2 of RuDP (2) to form the 2-carboxy-3-keto intermediate which subsequently undergoes hydrolytic cleavage between C-2 and C-3 to yield 2 molecules of 3-PGA (3). The evidence to date supports this mechanism. Mullhofer and Rose (1965) demonstrated that carbon-carbon cleavage occurs between the C-2 and C-3 of RuDP. There- fore, carbon dioxide becomes attached to the C-2 of RuDP. Further, by performing the reaction in D20, they demonstrated that the deuterium becomes attached to the carbon that was originally the C-2 of RuDP. This result 23 eliminates the possibility of hydride transfer from the C-3 of RuDP to the developing C-2 of the 3-PGA. The existence of the 2-carboxy-3-keto inter- mediate is inferred from the studies of Wishnick gt _1. (1970). They have reported that 2-carboxy-D-ribitol diphosphate, an analogue of the proposed intermediate, is an extremely effective inhibitor of the carboxylase reaction (Ki about 2 x 10'7M). This inferrence is based on Wolfenden's (1969) proposal that unusually tight binding of an inhibitor might be characteristic of transition state analogues. Other apparent examples of this phenomenon have been reported; e.g., 2-phospho- glycolate for triose phosphate dehydrogenase (Wolfenden, 1969), oxalate for lactic dehydrogenase (Novoa gt 31., 1959) and pyrrole-2-carboxylate for proline racemase (Cardenale and Abeles, 1968). Fiedler gt al., (1967) have studied the carboxy- lase reaction using RuDP labeled with tritium in the C-3 position. In confirmation of the results reported by Mullhofer and Rose (1965) they found that, following reaction with C02, 98% of the radioactivity was associated with the medium. Enzymatic enolization reactions are frequently characterized by isotope exchange with medimm protons. However Fiedler 35 31., (1967) reported that in the absence of 002, RuDP acquired no tritium when incubated in THO with the enzyme. They were unable to rule out the possibility that during enolization the 24 proton was prevented from exchanging by its tight binding to the enzyme. Examples of such tight binding of reaction protons derived from enolized substrates have been found in enzymatic isomerisation reactions (wang 25 51., 1963 and Rose and O'Connell, 1961). While the proposed mechanism would not predict a requirement for CO2 for proton exchange at C-3, it is possible that CO2 might be required to activate the protein in order to carry out the enolization step. The activation phenomena described by Pon £3 31., (1963) would be consistent with this interpretation. Nor would such an interpretation be without precedent. The enolisation of pyruvate catalysed by pyruvic kinase requires the other substrate, ATP. However, analogues of ATP, phosphate and arsenate are equally effective (Rose, 1960). The experiments of Fiedler 25 31., (1967) with RuDP tritiated at C-3 revealed a marked isotope effect. The tritiated substrate reacted at 20% of the rate of the unlabeled substrate, possibly signifying that the enolisation step is rate determining for the overall reaction. 2+ The role of Mg2+ or Mn in the reaction has been studied by Wishnick'g£.§l., (1970). They reported that Mg2+ was not required in the binding of RuDP to the carboxylase. ’They further demonstrated formation 2+ of a dissociable enzyme-Mn complex by measurements of 25 the effect of this species on the longitudinal nuclear magnetic relaxation rate of water protons. They suggested that Mg2+ was involved in the formation or Stabilization of the 2-carboxy-3-keto intermediate. This suggestion was supported by the observation that Mg2+ increased the affinity of the enzyme for the intermediate analogue, 2-carboxy-D-ribitol diphosphate, as determined by gel filtration studies. The interaction of RuDP carboxylase with oxygen was first proposed as early as 1949 by Tamiya and Huzisige, long before that path of carbon in photo- synthesis or the central role of the carboxylase was realized. Based upon their extensive investigations of the physiological conditions required to demonstrate the Warburg effect and upon the effect of cyanide under these conditions, Tamiya and Huzisige (1949) proposed that there was a competition between oxygen and CO2 for the carboxylating enzyme, and further, that cyanide acted upon this enzyme. After examining the kinetics of photosynthesis and photoreSpiration Ogren and Bowes (1970) were lead to the conclusion that the rate limiting step in both processes was controlled by one and the same enzyme. They reported that RuDP carboxylase is inhibited by oxygen and that the inhibition was competitive with respect to C02. Following their own speculation they then demonstrated that oxygen will in fact substitute 26 for C02, the products of the reaction being 2-phospho- glycolate and, they presumed, 3-phosphoglycerate (Bowes gt _a_1_., 1971). Oxidation of Carbohydrates and Related Compounds by Oxygen The oxidation of carbohydrates and related come pounds by oxygen and Species derived from oxygen falls into two broad classes; (a) reactions with molecular oxygen in alkaline solutions and (b) free radical reactions involving the oxygen free radicals, the superoxide free radical, H02-, and the hydroxyl free radical, HO-. Most of the reported research is of a descriptive nature and little attention has been given to the rather complex mechanisms and kinetics. To my knowledge, the oxidation of sugar phOSphates of biological interest by either class of reactions has not been reported. (a) Reactions with molecular oxygen in alkaline solution: In the presence of strong alkali, sugars undergo complex transformations which depend upon the conditions of temperature, alkali concentration and the presence or absence of metal ion catalysts. Nef (1914) observed that in the presence of oxygen, alkaline solutions of glucose and fructose could be oxidised to yield a mixture of monocarboxylic acids. Formic acid and arabonic acid were the principal products but significant quantities of glycolic, glyceric and erythronic acids were also formed. Since glucose and fructose yielded the same oxidation products, it was clear that the hexoses first 27 underwent a Lobry de Bruyn-Alberda van Ekenstein trans- formation to yield the various enediols. Nef (1914) proposed that these were then split by oxygen into acids containing one to five carbon atoms. Thus, oxidation of the 1,2-enediol was thought to produce formic and arabonic acids and so on. This is illustrated in Fig. 2. The relative quantities of the various acids formed is then a reflection of the steady state concen- tration of each of the enediols. During a kinetic analysis of these reactions, Bamford £5 31., (l950a,b) demonstrated the intermediate formation of a peroxide. Thus, they and subsequent investigators (Dubourg and Naffa, 1959 and DeWitt and Kuster, 1971) have represented the reaction of oxygen with the enediol or enolate anion as proceeding by the following‘mechanism: o I $1 $1 $1 Rl-C\OH c a o 02 c=o o-c-oa + - 9 9o o A 0H 6’é'OH R c’0 - - - I - c - OH I R2 2 \OH R2 R2 (1) However, the mechanism may be more complex. Gleason and Barker (1971a,b) have found that the formic acid, produced from ribose-Z-(BH) by the action of oxygen, contains a substantial proportion of the label. They have pr0posed two concurrent mechanisms (i) 28 Figure 2: The reaction of glucose and fructose with molecular oxygen in alkaline solution. 29 H H W H H W H H MUG O 20 0 2H 0 O C + O H 0 0 H O 0 0 H C CIC +C CICIC + C R R H R A7 \7 ‘7 2 2 2 0 O O H H H H H H H 0 O H H O H O O O O mu” 0 O O 4.6.-. _ 0/ G: _ H2 _m H2 _ _ H2 _ _ \QICICIRI \CHCICIRI CICFCIRI ClClmulClR Ir CICICJCIR H n_o H _ _ e: _ _ _“e H O O O H H 0 e H H H 9H O O O H H— O H 0 e 02 02% H W H 02 0 _ _ _ H H _ _ _ _ H \ ICICICICIC CICICICICIC H _ _ _ _ mu _ H O H H H H H 30 enolization and oxidation and (ii) hydride transfer of the C-2 hydrogen atom to C-1, followed by enolization of the resulting ketone and oxidation of the enediol. of 0-H o I I ll (IS-H T-(II- T-(ll <_-. e———* 6?-’0‘-H ="6' HO-(II-H l R R R One may also conceive a free radical mechanism leading to the formation of the glycosulose peroxide (1) which would be consistent with the tritium labeling experiments of Gleason and Barker. However, Bamford 25 31., (l950a,b) were unable to accelerate the reaction by means of free - radical initiators, thus favoring an ionic mechanism. The mechanism of decomposition of the glycosulose peroxide (1) intermediate to yield the two carboxylic acids is unknown. Conceivably, the peroxide could decompose intramolecularly by a concerted mechanism, perhaps involving a cyclic peroxide (Mechanism A). ”,0 R1 '6) R1 R1 ' C\09 Mechanism A \C -\C {-69 9 I —€I’ I -———(=- + 'O‘C'OH o/c-OH o l I R _ 0’ R2 R2 2 \OH Alternatively the peroxide could decompose by the addition and elimination of a hydroxyl ion (Mechanism.B). 31 R R - C = 0 H°\'1_l \OH Mechanism B 0 a ———e + H0 - O - C - OH R _ C = O 2 \\ R2 OH Since the mechanism of decomposition is of importance in the interpretation of some of the experimental results, a discussion of more completely understood analogous reactions is of interest. Hydrogen peroxide will cleave a-diketones to form carboxylic acids (Bunton, 1961). The first step in this reaction is a nucleophilic attack upon the carbonyl group to form the peroxide intermediate shown below. The structure of this peroxide intermediate is strikingly similar to the proposed peroxide intermediate in the alkaline oxidation of sugars. By using H2180 it has been possible to distinguish between two possible mechanisms of decomposition of the peroxide intermediate (Bunton, 1961). 0 o 180 130 ,180 180 -"-C-R+H180-° R-d-d-Rno R-o-d-R RC 2 2—2-«> . O—OH 18 18 Mechanism 1 18H? 0 ’0 32 18H? 18? 180 18(: 18 Mechanismll R-(II-C-R _I_-120:R-(.( +18/C—R 0_0H OH H 0 Mechanism 1, analogous to reaction pathway A in the oxidation of sugars, proceeds intramolecularly and leads to a 50% enrichment of the carboxyl oxygens of the resultant acids. Mechanism II, involving an attack by water leads to a predicted 75% enrichment. The exper- imental data indicates that Mechanism II is operative. Returning to the alkaline oxidation of sugars, one would therefore predict on the basis of this analogy, that only one atom of oxygen will be incorporated into the resultant carboxylic acids from molecular oxygen; i.e. the decomposition of the glycosulose peroxide involves the addition and elimination of a hydroxide ion. (b) Free Radical Reactions: The reactions of carbohy- drates with hydrogen peroxide have been reviewed by Moody (1964). Owing to secondary reactions and the effects of metal ion free radical catalysts, the nature of these reactions is poorly understood. In the absence of metal ion catalysts, the reaction probably proceeds by an ionic mechanism. 33 H H H I m | _ I c = 0 HO o-c - 0 c I 00H9 ‘Iz——\ H20 0” ‘\o H - I - 0H ---<=' H - I - 0H :3 + R R H o \C/ I 00H9 R CHZOH CHZOH é fnzon z’ ‘\ 9 0 o (I: {'6‘ 9 H0 o-cl: - 0' H2O 00H ,5 + H - I - 0H'---c> ‘H - I - o - H o H \\ a’ R R ? 00H9 R. “““c’ However, in the presence of redox metal ions, it is clear that oxidation occurs by free radical mechanisms. The nature of the reaction depends upon the peroxide concentration and the nature of the metal ion. In the first instance, the oxygen free radical formed depends upon the oxidation number of the initial cation. Thus Xm+ + H202 e x(m+1)+ +1 0H- + OH' n+ a (n-l)+ + - Y + H202 N + H, + Ho2 In addition radicals may arise by secondary processes OH ' + H202 * H 02 + H02 2 H02 + H202 * H20 + 0H + 02 34 Both the hydroxyl free radical, OH. and the superoxide free radical, H02° are powerful oxidizing agents. The diversity and relative quantity of the products formed make it clear that there are several primary and secondary oxidations. Oxygenases The nature of oxygenases, enzymes which catalyse oxygen-fixation reactions, have been reviewed extensively (Hayaishi, 1962; King 95 31., 1964; Mason, 1965; Bloch and Hayaishi, 1966; Guroff gt 91-: (1967); Hayaishi and Nozaki, 1969 and Hamilton, 1969). Several sub-classes of oxygenases may be recognized. (i) Dioxygenases catalyse the incorporation of both atoms of molecular oxygen into a molecule of substrate. 180 4 S18 S + 2 02 (ii) Internal Monoxygenases catalyse the incor- poration of a single atom.of oxygen into the substrate, the other atom of oxygen being reduced to water by electrons derived from the substrate itself. 18 1 18 18 8H2 + 02 s o + H2 0 (iii) External Monoxygenases catalyse the incorporation of a single atom of oxygen into the substrate, the other atom of oxygen being reduced to water by electrons derived from an additional electron donor. 35 18 18 18 S +' AH2 + 02 r S 0 +' A + H2 0 (iv) d-Ketogluterate - dependent Oxygenases catalyse the incorporation of one atom of oxygen into the substrate concomitantly decarboxylating a-keto- glutarate (d-KG) to succinate with the incorporation of the second atom of oxygen into the substrate (Holme g_t_ 31. , 1971). 180 1 818 S + dKG + 2 0 + CO + 2 180 succinate - With the notable exception of lipoxygenase, oxygenases have been found to contain at least one additional component besides the protein moiety. The dioxygenases generally contain either ferrous or ferric iron as the sole cofactor, although in the case of tryptophan dioxygenase the iron is complexed in the form of a heme. The monoxygenases are a more diverse class with respect to the nature of the cofactor(s). Some such as lysine monoxygenase and imidazoleacetate monoxygenase appear not to contain any metal components but instead contain organic cofactors, in this case flavin adenine nucleotide. Others, such as dopamine B-hydroxylase contain copper and no organic cofactor. More complicated electron transfer systems exist such as the microsomal cytochrome P-450 hydroxylating system. Studies on the mechanism of oxygenase action have concentrated on elucidating the nature of "active 36 oxygen" and the mechanism by which it is formed. Recent evidence (Hirata and Hayaishi, 1971 and Strobel and Coon, 1971) has implicated the superoxide radical in the 'mechanism.of tryptophan dioxygenase and the hydroxylation reactions catalysed by cytochrome P—450. Both enzymes contain heme iron. The work of Knowles 35 a1. (1969) and Massey 35 31. (1969) has established that reduced flavins are also capable of generating superoxide radicals upon interaction with molecular oxygen. In addition a number of biologically active reducing agents generate radicals in the presence of oxygen, e.g. various sulphydryl reagents (Sosnovsky and Zaret, 1970), reduced ferredoxin (Misra and Fridovich, 1971) and reduced quinones (Misra and Fridovich, 1972). There is little question then that superoxide radicals are produced in biological systems. Indeed, an enzyme, superoxide dismutase (erythrocuprein) has recently been described which catalyses the dismutation of superoxide (MCCord and Fridovich, 1969). - - + .02 + .02 +2H——c>02 +H202 However, the concept of superoxide involvement in oxygenase reactions is not new, (see Hayaishi, 1962), and schemes similar to that proposed by Mason (1965) for metapyrocatechase have been suggested for other oxygenases (Bloch and Hayaishi, 1966). 37 OH\\ OH 0- OH I I\_0 H 0\ SIFe 2 + I , H Fez-L— UO-O / H OH OH Despite the attractiveness of these proposals and the experimental observations, the ability to inhibit a given oxygenase reaction with superoxide dismutase (Hirata and Hayaishi, 1971 and Strobel and Coon, 1971) does not constitute unequivocal evidence that the super- oxide radical is the reactive form of oxygen. Rather a cautious interpretation is required since secondary reactions (see p.33) could generate other radical species from superoxide. A cursory glance at a listing of oxygenases reveals that for the most part the substrates are either aromatic or highly reduced. "In general, oxygen-rich cmmpounds, such as carbohydrates, are not favorable substrates for oxygenases" (Hayaishi and Nozaki, 1969). In this reSpect rat kidney inositol oxygenase is exceptional. This enzyme catalyses the oxygenation of myo-inositol to D-glucuronate, with the incorporation of one atom of oxygen (Charalampous, 1960). This is an internal monoxygenase but the details of the mechanism 38 remain to be elucidated. Work to date has not dis- tinguished between a dehydrogenation-oxygenation and a dehydration-oxygenation mechanism (Crandall, 1964). CHAPTER I STUDIES ON PEROXISOMAL GLYCOLATE OXIDASE Introduction The role of glycolate oxidase in photorespiratory oxygen uptake is now well established. However, although the enzyme has been studied with DCPIP as the electron acceptor by several investigators, little is known about the role of the natural electron acceptor, oxygen, in the reaction in its native state. Previous work in this laboratory (Baker and Tolbert, 1967) indicated that precipitation of the enzyme with ammonium sulfate in the course of purification of the enzyme altered its kinetic and Spectral properties. Accordingly, in order to study the enzyme in a state that more probably reflects it in zi!g_activity, the enzyme was purified by first isolating the peroxisomes. The subsequent purification procedures avoided recourse to ammonium sulfate precipitation. This chapter reports this method of purifying glycolate oxidase, the reactivity of the enzyme towards oxygen and the preparation and reactivation of the apoenzyme. 39 40 Materials Spinach (Spinacia oleracea L.) was purchased from local markets or cultivated in a growth chamber. During the day period of 11 hours the light intensity was about 3,000 foot candles and the temperature was 21°. During the subsequent 13 hours of darkness, the temperature was 15°. TEAE-cellulose was obtained from Serva Company and Sephadex-G25 (medium) from Pharmacia. Catalase was obtained from Worthington Biochemical Corp. Methods Protein Determination Protein was determined by the method of Lowry .gt‘al., (1951), using bovine serum albumin as a standard. Glycolate Oxidase Assays (1) For routine purposes glycolate oxidase was assayed anaerobically by 2,6-dichlorophenolindophenol (DCPIP) reduction as previously described (Tolbert gt.§1., 1968), except that cyanide was omitted. The DCPIP concentration in this reaction is not saturating. Thus, when an estimate of the maximum reaction rate was required, the assay was performed at several DCPIP concentrations, a Lineweaver-Burk plot constructed and the maximum.reaction rate determined by extrapolation. 41 (2) The enzyme was also assayed by use of the oxygen electrode. The electrode was calibrated as described below. The complete reaction mixture contained the following components (in umoles, unless otherwise stated): Tris (Cl) buffer, pH 8.5, 240; flavin mono- nucleotide (FMN), 0.10; sodium glycolate, 25 or lithium L-lactate, 31 or sodium g1yoxy1ate, 84; 500 units cata- lase and glycolate oxidase in a total volume of 3.2 ml. The reaction mixture was flushed with 100% oxygen for 5 min prior to initiating the reaction with substrate. The temperature was as indicated in the text. The reaction was linear until about 35% of the oxygen had been consumed. The reaction was also proportional to enzyme concentration. One unit of enzyme activity is defined as the amount of enzyme required to consume one umole of oxygen or reduce one pmole of DCPIP per min at 25°. Calibration of the Oxygen Electrode The method was based upon that described by Goldstein (1968) in which catalase was used to release stoichiometric amounts of oxygen from.sodium.perborate. Ten to fifty pl aliquots of freshly prepared 0.033M sodium perborate were added to 3.2 ml of 0.05M sodium phosphate, pH 7.0, containing 500 units catalase and the pen deflection recorded. The pen response was a linear function of the quantity of perborate added. 42 The response of the oxygen electrode to temper- ature was not a linear function. As the temperature increased, the pen response EE£.§E increased in an approximately Nernstian manner. However, this was in part offset by the decrease in the solubility of oxygen. Thus it was necessary to calibrate the electrode at the temperature at which it was to be used. Isolation of Spinach Peroxisomes Spinach peroxisomes were isolated by isopycnic sucrose density centrifugation in the 54 m1 tubes of an SW 25.2 rotor as previously described (Tolbert gt 31., 1968) or in the B-29 or B-30 zonal rotor (Donaldson, 1971). Purification of Glycolate Oxidase from.Spinach Peroxisomes The subsequent procedures, all performed at 4°, were dependent upon the volume of the solution containing the peroxisomes. When the peroxisomes were isolated in the SW 25.2 rotor, Method A was used. When the perox- isomes were isolated in the zonal centrifuge, Method B was used. Method A: The peroxisomes were broken by dilution with 3 volumes of 0.02M glycylglycine, pH 7.5, so that the final concentration of sucrose was 0.5M. The diluted solution was centrifuged at 144,000g for 60 min to remove membraneous material. From 5 to 10% of the activity was routinely found in the precipitate. The supernatant solution was dialysed against 50 volumes 43 0.005M Tris-Cl, pH 8.7 for 6 hours. The dialysate was reduced in volume to 2.9 ml by vacuum dialysis overnight. The concentrated solution was centrifuged at 14,000g for 20 min to remove the slight precipitate which had formed. The supernatant solution was applied to a 1 cm (i.d.) x 10 cm column of TEAE-cellulose equilibrated with 0.005M Tris-Cl, pH 8.7, and the enzyme which did not stick to the column, was washed through the column with this buffer. Method B: The peroxisomes were broken by dilution with 3 volumes of 0.005M Tris-Cl, pH 8.7. The diluted solution was centrifuged at 144,000g for 60 min. The supernatant solution was applied to a 2.5 cm (i.d.) x 20 cm column of TEAE-cellulose, equilibrated with 0.005M Tris-Cl, pH 8.7, and the enzyme eluted with this buffer. The eluate containing the enzyme was concentrated by ultrafiltration using a PM 30 membrane (Amicon Corp.). Polyacrylamide Gel Electrophoresis of Glycolate Oxidase Polyacrylamide gel electrophoresis of the TEAE- cellulose purified glycolate oxidase was performed at pH 9.3 as described by Davies (1964) and at pH 7.0 as described by Williams and Reisfeld (1964). In both cases the gel concentration was 5.5%. Twenty to thirty pg of protein were applied. Following electrophoresis the gels were stained for protein with 0.5% (w/V) Amido-Schwartz in 7.5% (v/v) acetic acid for 2 hours and 44 destained in 7.5% (v/v) acetic acid. Other gels were stained for glycolate oxidase activity by incubation at 250 in the dark in a reaction solution containing 4.0 m1 0.1M Tris-Cl, pH 8.5, 0.25 ml 2M sodium glycolate, 0.01 mg phenazine methosulfate and 1.8 mg nitroblue tetra- zolium. A control gel was incubated in the same reaction solution less glycolate. Determination of the Km_for Oxygen The Km for oxygen for each of the three reactions catalysed by glycolate oxidase (glycolate to g1yoxy1ate, g1yoxy1ate to oxalate and L-lactate to pyruvate) was determined at 250 by analyses of the reaction progress curves rather than by measurement of initial reaction rates under different oxygen concentrations. Calcu- lations showed that the organic acid concentrations were saturating throughout the course of the reactions. Increasing the concentration of the organic acids did not alter the progress of the reaction. Determination of the Energies of Activation The energies of activation for each of the three reactions catalysed by glycolate oxidase were determined by measuring the initial rate of oxygen consumption at various temperatures. Correction was made for the effect of temperature upon the pen response. 45 Preparation and Reactivation of the Apoenzyme of Glycolate Oxidase The method of Massey and Curti (1966) was used ‘with minor modifications. To a solution of glycolate oxidase, purified from spinach peroxisomes to the stage before TEAE-cellulose chromatography and containing about 2 units, was added an equal volume of 2M KBr in 0.1M sodium pyrophOSphate, 3 x 10-3 ‘M EDTA, pH 8.5. The solution was allowed to stand on ice for two hours before desalting on a l x 15 cm column of Sephadex G25. The apoenzyme was assayed by DCPIP reduction in the absence of FMN with concentration of enzyme sufficient to detect 1% holoenzyme. The reactivation of the apoenzyme was followed anaerobically by DCPIP reduction as described except that the reaction was initiated by the addition of FMN. The concentration of FMN was determined spectrophotometri- cally at 450 nm using a value for the molar extinction coefficient of 1.22 x 104 cm-lM-1 (Gibson gt.al., 1962). Results and Discussion Purification of Glycolate Oxidase from Spinach Peroxisomes The purification of glycolate oxidase from spinach peroxisomes by the two methods is presented in Table I. The biggest problem concerning both methods and one which was never satisfactorily overcome is the lability of the enzyme eSpecially in dilute solution. 46 um saga mmH081 om mo Gowumasmcoo amwmxo mo dump ESEmeE pmmmao muuxm am no: cowumummmud mweyo .wwmamwoson mvsuo msu cw wuw>auom mehncm Hmuou man does woman mm3 pama» N ways .cofiuosomu mHmoQ mo mumu Enawxme Umumaommuuxm we“ do woman xua>fluowuom owmwomam maww>wuom HouOH . u m .aomcwam wx N.m Scum coaumwsmwuucmu Hmcow an woumHomH mmEomeonmm .m oomhmz m.o om.o~ o.mN qm.o MH¢DAM m.uom camaommm kww>wuom HmuoH samuoum .5833 we N; 88m «.3 3m 3 emuflofl mmaoflxoumm .< 85m: .mmZOMHXQmmm mo .m madman 50 Ec com 0mm com 03 03 o5 can own omm P 0.0 mod m 0.0 a 0.0 . mos mod 51 position of the two peaks is in agreement with the Spectrum recorded at pH 8.3 by Frigerio and Harbury (1958) but differs from that recorded at pH 7.4 by Zelitch and Ochoa (1953) who found peaks at 450 nm and about 365 nm. Free FMN has peaks at 370 nm and 450 nm at pH 7.0 (Hemmerlich £5 31., 1964). However, as the pH of the free FMN is raised from.pH 9 to pH 12, the absorption at 375 nm decreases and a new peak develops at 335 nm. These changes have been attributed to the ionization of the 3-imino position in the isoalloxazine ring (Hemmerlich gt 31., 1964). Thus the differences in the spectra at pH 8.7 and pH 7.4 can be attributed to the effect of pH upon the FMN moiety. Subsequent to the termination of this study similar spectral properties were reported for the pig liver enzyme by Schuman and Massey (1971). Their study was considerably more comprehensive than that undertaken here. Interestingly the pig liver enzyme shows an even more pronounced shoulder at 420 nm. Investigations by Schuman and Massey (1971) have shown that the absorption at 420 nm is due to the presence of another chromophore besides FMN. The chemical nature of this chromophore remains to be identified. Determination of the K for Oxygen The Km of glycolate oxidase for oxygen with each of the three substrates was determined as described in 52 the Methods section. Lineweaver-Burk plots (Fig. 4) indicate that the Km (02) for each of the three reactions is the same, namely about 1.9 x 10~4M oxygen at 25°. Air (20% oxygen) saturated water contains 2.6 x 10-4M oxygen at 250. Regular Michaelis-Menten plots (not shown) indicate that the enzyme is effectively saturated with oxygen at a concentration of about 8.0 x lO-AM. This would correspond to an oxygen concentration in the gas phase of about 60%. This result is regarded as physio- logically significant since in 2339 measurements of photorespiration indicate that the overall process is not even saturated by 100% oxygen. Further discussion of the physiological significance of this finding is postponed until Chapter V. The original purpose of these experiments was to gain an insight into the mechanism of action of glycolate oxidase. The fact that the Km for oxygen is the same with each of the three substrates suggests that oxygen reacts with an intermediate which is either the same in each of the three reactions or so struc- turally similar that small differences in reactivity with oxygen cannot be detected. Determination of the Energies of Activation The Arrhenius energy of activation (Ea) was determined for each of the three reactions with oxygen as the terminal electron acceptor. The results (Fig. 5) 53 NW8 omN> .omN um No 28 om.o mcwmuaoo Hmum3 pmumusumm Had :Hmuoua w8\aw8\mMH051 on H No as mH.o u 0mm as an oompaxo mumaoomaw w: w .ammmm mtouuooam amwmxo . H8 «.0 awe poEfimcou nowhxo mmH051 mm oommmuaxm mm > .Aowhllov oumamxomaw new Ax xv oumuomHuA .AQIIIIQV mumHoowaw .mmumuumnnm omnnu men mo some now mmmtho mumaooaaw mo umwhxo Mom omNM men mo cofiumcfiahmump oru How uon Musmnum>mmBoGHA .q muswwm 54 me Figure 5. 55 Arrhenius plot for the catalytic activity of glycolate oxidase with oxygen and each of the three substrates, glycolate 0—-——0, L-lactate A A and glyoxylate 0-—-—O. Slopes represent the lines of best fit. The calculated values for E are glycolate (11,500 cal.), L-lactate (1; ,200 cal.) and glyoxylate (12,400 cal. 10 56 A o A.‘ o A . o o .0033 .0031. .0035 .0036 .0037 VT (°KI 57 show that the values for Ea for each of the three reactions are close to being the same. Since the partial reaction common to all three overall reactions is the oxidation of the reduced flavin, this result suggests that the rate limiting step in each of the three reactions is the same, namely, the oxidation of the reduced flavin. However, the overall reaction rates for the oxidation of the three substrates are quite different. In order to be consistent with the results reported here it is necessary to conclude that the overall rate of reaction is dependent upon the steady- state concentration of the reduced flavin-enzyme complex. However, lacking further data, it does not seem profitable to speculate further about the steps involved in the mechanism. Preparation and Reactivation of the Apoenzyme of Glycolate Oxidase The apoenzyme of glycolate oxidase was prepared as described in the Methods section. The apoenzyme was completely inactive in the absence of FMN (Fig. 6). The recovery of active reconstituted holoenzyme was rather variable, ranging between 35 and 65%. The reason for this variability was not investigated. When the enzyme was dialysed against bromide for 12 hours as described by Massey and Curti (1966) in order to remove FMN only 10% of the activity was recovered. The reactivation of the apoenzyme with FMN at 250 58 .chuoua w: ma .2 10H x om.~ u Coaumuucmocoo zzm .omm n H "22m nuwa mmmpwxo mumaoozaw mo mshuamoam mnu mo cowum>auommm .o unawam 59 zzu.’ mo 009” 60 did not take place immediately (Fig. 6). The order in which the reactants were added did not alter the progress of the reactivation, indicating that glycolate has no role in the reactivation process. Kinetic analysis of the reactivation process indicated that it was a first order event (Fig. 7). A family of curves similar to that obtained in Fig. 6 were obtained by varying the FMN concentration. Two kinetic constants may be obtained from such data. Firstly, the Km for FMN can be calcu- lated from the final steady state reaction rate (Ve) under different concentrations of FMN. A Lineweaver- Burk plot of the data obtained in this manner gives a value for the Km for FMN of 4 x 10-6M (Fig. 8). This agrees with the value of 3 x 10-6M previously determined by Zelitch and Ochoa (1953). Analysis of the rate constants obtained at different FMN concentrations revealed that the reactivation process was not a simple one-step process. The simplest model for the reacti- vation would be of the type k E+F1~m.—i1—>*E-FMN k-1 * where E - FMN represents a catalytically active binary complex. In this case kact = k+1 [FMN] + k_1 (Strom 3531., 1971) and a plot of kact versus [FMN] should yield a straight line with a slope of k+1 and an intercept on the ordinate 61 Hosam m£w> map ma .aoauontmu mHmon mo dump ummcwa m> paw u .msau and um Gowuosvmn mHmun mo camp .0 ouswwm cw :3oem muasmmu mau mo uoaa nopuo umufim .ommpwxo mumaooxaw mo 08%Namoam mnu mo cowum>wuommm .m muswwm 62 k: 0.435 min" minutes [I 00’me '0 F. (aA/M - I.) ¢ 63 How .hmmmm cOHuoDva mHmUa anoummam wnmvcmum .zzm EM msu mo GOHumaHEMmumU msu Mom HOHQ xusmuum>wm3mcHA .m mustm 64 o _Zzu=\oo_. m ‘ d d 22... :92 x 3 uzv. 65 axis of k_1. However, the plot of kact versus [FMN] was hyperbolic, indicating that the activation process is not adequately described by the above model. A two step mechanism involving the rapid for- mation of an E-FMN complex followed by a slow conversion of this complex to a catalytically active state may be considered. k k * E + FMN a—Ll» E - FMN +2 E - FMN k k -l -2 In the case where k+1k+2 + k+1k_2 >> k_1k_2 it can be shown (Strom £5 31., 1971) that 1 ___ k-1 + k+2 , 1 + 1 Fact k+1W+2+k-2) [FMN] k+2+E-2 Here a double reciprocal plot of l/kact versus l/EFMNJ should yield a straight line. Fig. 9 shows that such an analysis of the data does yield a linear relationship. The following relationships were thus obtained: k+ 6 l 2 + k_2 = 2.0 min”1 and k.+1 = 2.4 x 10 M- /k_1l+k.+2 This kinetic analysis is consistent with the concept that the simple combination of the apoenzyme of glycolate oxidase with FMN is not sufficient for catalytic activity. This binary complex must then undergo an additional reaction (possibly a conformational transition) in order to become catalytically active. It is interesting that Massey and Curti (1966) have reported a similar phenomenon for the reactivation of the 66 .mzzmg mamum> AZZM an mmmvon mumHoo%Hw Ho 08%Ncmoam msu mo aOHum>Huommu mnu Scum vmchHnoV mucmumcoo mumu “mono umuHm msu mo HOHQ HmoonHomu mHndon .m muszm 67 T2 _zzaxme w "Q ‘N >l/L st (UM) 68 zapoenzyme of D-amino acid oxidase by FAD. CHAPTER II INCORPORATION OF MDLECULAR.OXYGEN INTO GLYCINE AND SERINE DURING PHOTORESPIRATION IN SPINACH LEAVES Introduction The inhibition of net photosynthetic CO2 fixation by oxygen, often referred to as the Warburg oxygen effect (warburg, 1920), has been observed in a wide variety of algae and higher plants (Turner and Brittain, 1962) and isolated chloroplasts (Ellyard and Gibbs, 1969). It has become evident that this phen- omenon is due to photoreSpiration; that is, the light dependent uptake of oxygen and release of C02 which is thought to be associated with the glycolate pathway of metabolism (Jackson and Volk, 1970). Photorespir- ation is especially evident under conditions of high light intensity, limiting CO2 and high oxygen concen- trations. In these circumstances a large part of the total carbon fixed during photosynthesis flows through the glycolate pathway (Tolbert, 1963). Recently, many of the enzymes of the glycolate pathway have been located in the peroxisomal (microbody) fraction, as distinct from both chloroplasts and mitochondria (Tolbert, 1971). The origin of phosphoglycolate and 69 70 glycolate is one of the most interesting problems in photosynthetic carbon metabolism and one which is of fundamental importance in photorespiration. Glycolate may arise from phosphoglycolate by the action of a Specific phosphatase, which is located in the chloro- plast (Richardson and Tolbert, 1961), but whether it does so exclusively is not clear. Two radically different mechanisms have been proposed to account for the formation of uniformly labelled glycolate during photosynthetic [IACJCO2 fixation. One, proposed by Tanner g5 31., (1960), Stiller (1962), and Zelitch (1965) suggests that glycolate arises by means of a hitherto undiscovered reductive condensation of two 14C labelling experi- molecules of C02. However, the ments of Hess and Tolbert (1966) and Coombs and Whittingham (1966) tend to discount this possibility. The other mechanism, proposed in various forms, suggests that glycolate, phosphoglycolate or both are formed as the result of the oxidation of one or more intermediates of the photosynthetic carbon cycle (Wilson and Calvin, 1955; Bassham and Kirk, 1962; Tolbert, 1963; Coombs and Whittingham, 1966; and Gibbs, 1969). If the latter hypothesis is correct the nature of the oxidant is of considerable interest. To investigate the possibility that the oxidant is molecular oxygen, or is derived from it, experiments were performed in which detached spinach leaves were allowed to photorespire in an atmOSphere 71 of [180] oxygen. This chapter concerns the incorporation 18 of [ 0] into the products of the glycolate pathway, glycine and serine. Materials Spinach was grown as previously described (Chapter I, p. 40). Younger leaves, weighing about 2 gm and about 10 cm long, were harvested from mature plants immediately before use. [180] Oxygen (93.5 atoms Z) and [18 OJHZO 93 atoms Z) were obtained from Miles Laboratories Inc., Elkhart, Indiana. Silylating reagents were obtained from Regis Chemical Co., Chicago, Illinois. Solvents were redistilled where necessary. Methods Photorespiration in [18010xygen A Spinach leaf was placed in a stoppered 2.2 cm (i.d.) x 10 cm test tube and the volume of the gas Space determined by measuring the volume of water required to fill the tube completely. This volume was 31.0 t 0.5 ml. The water was removed, except for the last 1 ml which covered the bottom of the petiole, and the leaf was allowed to photosynthesize in a stream of humidified air at 250 while being illuminated from two Sides with white light of about 4,000 foot candles. The 72 air entered and left the tube by means of syringe needles passing through the Stopper. After about 40 min the system was flushed with 50 ml of 100% oxygen, followed immediately by an injection of 6 ml of isotopic oxygen. At various times after introduction of the isotope, the leaf was killed by filling the tube with boiling 90% (v/v) ethanol. Extraction of glycine and serine. After the leaf had remained in boiling 90% (v/v) ethanol for at least 3 min, the solution was decanted and the leaf further extracted for 3 min each with boiling 50% (v/v) ethanol and boiling absolute methanol. The extracts were combined and evaporated to dryness. The residue was dissolved in a minimum volume of chloroform~methanol-0.2'M formic acid (25:60:15 by vol.). The chloroform phase was removed and washed twice with 0.2 M formic acid. The aqueous phases were combined and washed twice with chloroform. Sufficient Dowex 50, H+ form was added to the aqueous phase to render the supernatant solution colorless. This solution was removed and the resin was washed with 0.2 M formic acid. Glycine and serine were eluted from the resin with lJM NHhOH and the eluate was evaporated to dryness. The residue was dissolved in a minimum volume of water and applied as a streak to a 24 cm x 50 cm Sheet of Whatman 3MM chromatography paper. Glycine and 73 serine markers were applied on both Sides of the main Streak. Chromatography was performed for 17 hr at 250 with n-butanol-propionic acid-water (10:5:7 by vol.) as solvent. The chromatogram.was dried and thin strips were cut from both sides and from the middle. These strips were Sprayed with ninhydrin solution (0.2% (w/v) in water-saturated n-butanol) and heated at 900 until color development was adequate. The area of the chromatogram containing both glycine and serine was located by comparison with the markers. Glycine and serine have very similar Rf values in this system. This area of the chromatogram was cut out and extracted with water. The resultant solution was evaporated to dryness in preparation for silylation. Extraction of organic acids. The supernatant solution plus washings from the Dowex 50 step were combined and evaporated to dryness. The residue was dissolved in 8 m1 of 0.05 M NH40H, the pH of the resultant solution being about 9.4. This solution was incubated with approximately 3 units of alkaline phOSphatase (Sigma Type VII) at 250 for at least 30 min and then applied to a 0.7 cm (i.d.) x 6 cm column of Dowex 1-acetate. The column was washed with water and the organic acids were eluted with 42M acetic acid. This fraction was evaporated to dryness and the residue used to prepare the Mb3Si derivatives. 74 Preparation of Me3$i derivatives. The dried residues obtained as described above were suspended in 50 ul of acetonitrile and 50 ul of bis(trimethylsi1yl)trifluoroacetamide containing 1Z (v/v) trimethylchlorosilane under dry conditions and heated at 1500 for 15 min to facilitate the Silylation reaction. Standards containing glycine, serine, gly- cerate, malate and erythronate (1 ug/ul) were Similarly prepared. Samples containing glycine were allowed to stand at room temperature for at least 48 hr before analysis to ensure the formation of (Me3Si)3 glycine (Bergstom 25 31., 1970). 180] [18 Synthesis of [ Glycine and 0] Serine A mixture of glycylglycine (1.0 mg) and seryl- glycine (1.1 mg) was dissolved in 190 pl of 6 N HCl and 10 ul of [IBOJHZO (93 atoms Z). This solution was heated for 20 hr at 110° in a sealed vial. [180] Glycerate Synthesis of Calcium DL-glycerate (3.5 mg) was dissolved in 90 ul of 2N HCl and 10 pl of [1801H20 (93 atoms z). This solution was heated at 900 for 17 hr in a sealed vial. Isot0pe analysis Aliquots (l - 4 ul) of the above silylated samples were analyzed using an LKB-9000 combined gas chromatograph - 75 mass Spectograph equipped with a 1.4 m x 3 mm (i.d.) silanized glass column packed with 3Z (w/v) SE-30 on silanized Supelcoport (100-120 mesh, Supelco Inc., Bellefonte, Pa.). The column temperature was 1000 for glycine and serine and programmed at a rate of 50/min from 500 to 2000 for the organic acids. The flow rate of the heliwm carrier gas was 30 ml/min. The temperature of the ion source was 2900 and the ionizing voltage 70 eV. Isotope incorporation was measured by the procedure of Thorpe and Sweeley (1967). The normal isotopic abundance in fragment ions was determined experimentally from.the mass spectrum of the compound in question, rather than from probability theory based on empirical formula. This value (F) was determined by the ratio (P-+ 2) observed / P observed, where P and (P‘+ 2) are the intensities of the ions at m/e = P and m/e = (P‘+ 2) in the standards. In samples containing 180 the following correction for normal isotopic abundance was made (P'+ 2)corrected = (P + 2)observed - F x Pobserved Finally, a 18 _ (P +2)corrected o A 0 — P +'(P + 2) x 1004 observed corrected For most determinations the intensities of the various ions were determined by direct measurement of peak heights either from oscillographic recordings or from normalized bargraphs (Sweeley 25 31., 1970). When more accurate data were required for detection of any 76 possible incorporation into the hydroxyl group of serine, the accelerating voltage alternator unit (multiple ion analyzer) was used. This technique involves the con- tinuous recording, at constant magnetic field, of the intensities of two ions formed by electron impact on the gas chromatographic effluent. The ions were care- fully focused by manual setting of the magnetic field strength (for the ion at the lower m/e value) and by decreasing the accelerating voltage (for the ion at the higher m/e value) (Sweeley EE.§l-: 1966). Results Gas-Chromatography - Mass Spectrometry of Glycine and serine Owing to the small metabolic pool Sizes of both phOSphoglycolate and glycolate, it was necessary to investigate photorespiratory incorporation of [180] by analyzing the metabolic products of glycolate, glycine and serine. Both of these compounds accumulate in relatively large pools. In addition, the MeBSi deriv- atives of glycolate, oxalate and pyruvate have nearly identical gas chromatographic retention times, emerging with the tail of the solvent front, such that a clean mass Spectrum of (iMe3Si)2 glycolate could not be obtained. The MeBSi derivatives of glycine and serine were formed, rather than the more commonly used 77 N-trifluoroacetyl-n-butyl esters, since the latter esterification would result in the loss of at least half of any 180 incorporated into the carboxyl group. (Mie3Si)3 glycine and (Me3Si)3 serine were readily separated by gas chromatography from each other and from several other compounds present in the sample (Fig. 10). The mass Spectra of authentic samples of (MeBSi)3 glycine and (MeBSi)3 serine were Similar to those previously reported (Vandenheuvel and Cohen, 1970; Bergstrom, 35 31., 1970). The only ion in the mass Spectrum of (MeBSi)3 glycine (Fig. lla) suitable for measuring 180 incorporation was that at m/e 276, which results from the loss of a methyl group from the parent compound and thus contains both carboxyl oxygen atoms. The spectrum of ('Me3Si)3 serine (Fig. 11b) contains prominent ions at m/e 204 and m/e 218. The ion at m/e 204, which may be regarded aS'Me3Si-0-CH2 CH.= + NH - SiMe3 contains the hydroxyl oxygen only, while that at m/e 218, MeBSi - +NH = CH - COO - SiMe3, contains only the carboxyl oxygen atoms. Gas Chromatography - Mass Spectrometry_of the Organic Acid5FractIOn The MeBSi derivatives of the organic acid fraction were prepared as described in the Methods section. The peaks in the gas chromatogram (Fig. 12) were identified by their mass Spectra (Figs. 13a,b,c,d,e). Comparison of authentic standards with regard Figure 10. 78 Gas chromatography of a silylated extract from spinach leaves containing glycine and serineo The column temperature was maintained at 100 . The tracing Shown is the response of the total ion current detector. Mass spectra Showed that peak I was (Me Si) glycine and peak II was (Me Si)3 rin This tracing was obtained far a 3leaf exposed to lOOZ oxygen for 5 min in the light. Smaller peaks for glycine and serine were observed for Shorter exposure times or when exposure occurred in the dark. RESPONSE 79 1 «.JW‘AJLIIJ *II 6 TIME(MIN) 10 80 A.>o ONV mcHumm MAHmmmzv Ho Eduuumam mmmz .Anv A.>m ORV mcHoxHu MAHmmmZV mo Esuuoomm mam: .AmV HH mustm 81 ON 2E gum e_mN CNN 09 0.1 03 om 1 TON 60m m3 1 mx ma 1? 1 19.2310, so v8 ms 18 “.6 .mm 2:2 1 Amozgmnxznwux m m 18 2E m1 cm as: mm .3256 19.6 w m OOH £6 owN CNN 09 0.1 02 cm H #174. h L TPFl. bl L _ 1 mvN _ 19.. 2K .1 ml {X 10:. ms 1 192310, so 18 0 .8 2:2 1 Angm/ b loo ZINI v I _ no.2 mN Am£v_m\ NP NZ _ 91.0 NA .w V 09 AllSNBlNI HAHN-138 ALISNBINI BAllV'IEH 82 .Amumcomunu m mqfiHmmmSV HOV oumcounwxum 1VAHmmmZV u > .wumHme m mAHm 62v 1 >H .mumcmoosm AHmmmzv 1 HHH .mumumo%Hw AHm 62v 1 HH .mummeo AHmmmSv n H “mBOHHom mm mxmmm mDoHHm> mnu pmHmHucmpH muumaonuooam mmme uamsvomnsm .mHSumumaEmu GEDHoo Anunuv .Houomumv ucmuuso aoH Houou mo masommmu A v .mpHom OHmeHo wchHmucoo mm>mmH nomcHam Scum uowuuxm vmumH%HHm m mo asamuwoumaouno mow .NH oquHm 83 .22.: mzz. on ON or - p — _ — 8 1 _ x ........................... . l \\\\\ M GNP] VA \\\ 8 \ N \\\\\ m 1 .5 H 2:1 . fi\\\‘ 8N] =1 SSNOdSBU 84 Figure 13 (a) Mass spectrum of (MeBSi)2 oxalate (70 eV.) (b) Mass Spectrum of (MeBSi)3 glycerate (70 eV.) (c) Mass Spectrum of (MeBSi)2 succinate (70 eV.) RELATIVE INTENSITY RELATIVE INTENSITY RELATIVE INTENSITY 85 100 20 (Me3Si)2— OXALATE 0‘ /O-Si(Me3) 807 MW 234 117 147 (I: ‘ 50_ 73 0’ \O-Si(Me3) 1 20- 219 1— 1 I 1 1.111111 1 [I r T T Y T T I T 1 r r'r I T so 100 140 180 220 m/e 100 25 73 (W39 )3 -GLYCERATE CH2- 0—51(Me3) 80“ H-C-O-Si(Me3) — MW 322 (I: sod I x _ . — 147 O O 51(Me3) 40... X4 _. 2 20 189 292 307 A Ti! lJf F‘Lt I rlLr TL r r fiL] I rLI LIL r I I r I r T 32‘2 I so 100 140 180 220 260 300 340 mm 100 11 (Me3Si)3— SUCCINATE 80‘ Mw 252 O‘C,O—Si(Me3) 7 so— 147 I — 73 ('le 40“ CH2 x4 >— 2 1': 20- 117 0’ \O-SIIMS3) I M 262 ~— 11 I- L .1 [L P r I r T I I I I T r Y I I I I f so 100 140 180 220 260 86 Figure 13 (Continued) (d) Mass Spectrum of (Me3Si)3 malate (70 eV.) (e) Mass Spectrum of (Me3Si)4 erythronate (70 eV.) 87 ON 08 omm om” oi ooz om ONN low 9m MON mom 13.23101qu mx E 13 . AmoszIOIWII low 13.23101“qu vmv >22 18 Amazemnob/o me22 1 mo .m o w 18 A 5:. to meNonumu map mo macaw cowhxo anon mo ownmsoxm muoHQEoo Mom pouovaum umnu mH mHmmauamumm aH man> msav .mom cam mom m\E um mGOH osu mo mHthmam kn vmchuno mmsHm>o .dmN paw NmN m\8 um mcoH map mo mHm%Hmcm %n vmckuno mmSHm>n .Ucon mpHHQSQSSH mo mHthoHpms ha hHoHom muaooo cOHumuomuoocH umsu wcHabmmw .omH mo GOHumHomHooaH Mom USHOHumHQ mmonu mum mHmmsucmuma aH mmsHm> mgfim .mucmEmusmmmE muoa Ho mounu mo mwmum>m mnu mH mDHm> 50mm oow.o n¢m.o on.q mo.N COHuomuuxm Hmum< eAm.eHV oem.e sem.o «Ame.ev mw.e «AH0.HV mH.~ soauomuuxm museum mumumo%Hw mcHumm maHomHo AN mmHoEv ucmuaoo omH mHmamm mMMDQmuomm ZOHHUHuom0Hpmu mnu Eoum waHUHoomu m mumuu Home: mnu .EmmwoumEouso mmw onu mm oomnu HSSOH may .mumaoua hum eAHm 02v HH> new mumams AHmmmzv H> .mumsHoosm NAam 02v > .mumuoome aHmmmzv >H .Amoabo sou oBu mchucoo AHamQOHaV CBOch: HHH .mumHoowa AHmmmZV mHnmnoum HH .oummeo NaHmmmzv H ”m30HHom mm mum mxmmm mDOHHm> N mnH .CHE H Mom mezxo NOOH up ©03OHHom SHE m.m Mom oumo g on pmmomxw mmmH nomchm m Eonm pmuomuuxo mpHom QHCNWHO pmumeHHm man mo EmuwoumEousooHomu UHDvHHnmmu .mH seawae 121 VII VI 2 j 20 MINUTES IIWCPN BSNOdSBB 80133130 122 It was therefore concluded that neither erythro- 14C nate not its phOSphate ester became labeled with under these experimental conditions. Even if erythronate were an artifact formed during the extraction and fractionation procedures, and there is no evidence that eliminates this possibility, the failure to detect any 140 in erythronate precludes the possibility that this compound, or its phOSphate ester or the compound from which it is artifactually formed, have a direct role in photosynthetic-photoreSpiratory carbon metabolism. Further investigation of the metabolism of erythronic acid did not therefore seem warranted. CHAPTER IV A MODEL SYSTEM FOR THE OXIDATION OF RIBULOSE DIPHOSPHATE TO PHOSPHOGLYCOLATE Introduction The results of the ipqgigp [180] oxygen exper- iments described in Chapter II indicated that glycolate biosynthesis involved the incorporation of one atom of oxygen, derived directly or indirectly from.molecular oxygen. It was not possible to determine which form of oxygen was the reactive Species. However, Since the oxygen atom of the hydroxyl free radical, OH“, exchanges with water (Kasanowsky pp 31., 1956), the reactive Species might be the superoxide radical H02: or hydrogen peroxide. None of the oxygen atoms of H02- or H202 exchanges with those of water (Dole pp al., 1952; Cahill and Traube, 1952). Therefore, in addition to molecular oxygen pp; pg, these Species must be considered as potential candidates. The possibility that the reactive Species was Singlet oxygen could not be excluded either. Indeed the apparent requirement of light for incorporation of 180 could be interpreted as evidence that an "activated" Species of oxygen was the reactive Species. The assumption that a 4-electron oxidation of a 123 124 ketose sugar by molecular oxygen would proceed by a concerted mechanism with the incorporation of an atom of oxygen into each of carboxyl groups formed, lead to the conclusion that glycolate formation involved a 2-electron oxidation to yield phosphoglycolate and/or glycolate and an aldose sugar phosphate. In view of the reactivity of hydrogen peroxide and the free radicals derived from it towards carbo- hydrates (See review by Moody, 1964) it was decided to investigate the reactions of hydrogen peroxide with intermediates of the C-3 photosynthetic carbon reduction cycle. Initially it was planned first to investigate the non-enzymatic reactions and then proceed with enzymatic Studies. However, before this could be completed, the experiments described in the next chapter became imperative. This chapter then describes some limited Studies on manganese catalyzed oxidation of ribulose diphOSphate by hydrogen peroxide. Methods Preparation of_LU - 14C] - Ribulose DiphOSphate Uniformly labeled [14C] ribulose diphosphate was synthesized by the method of Wishnick and Lane (1969) with the following modifications. Uniformly labeled [14C] glucose-6-phosphate (35 uCi/umole) was used rather than glucose thus dispensing with the hexokinase Step. 125 Secondly the pyruvic kinase-ATP regenerating system was omitted. Thirdly, Since the preparations were conducted on a much smaller scale, the RuDP was separated from the other reaction components by paper chromatography (Whatman 3MM) in butanol : propionic acid : water (10:5:7). Unlabeled RuDP was run on both sides of the sample streak. Following chromatography, the RuDP markers were treated by the method of Bandurski and Axelrod (1951). A sample strip was scanned for radioactivity. Generally there were two peaks, the major one co-chromatographing with RuDP. The minor peak was probably sugar mono- phosphates. The major peak was eluted from the paper with.water and lyophilized. The residue was redissolved in 1 ml of water and pH adjusted to neutrality by the addition of Tris base. To ensure radiochemical purity, a sample was subjected to high voltage paper electrophoresis at pH 3.4 as described by Bieleski and Young (1963). Electrophoresis was conducted for 1.5 hours at 1800 Volts, 25 mA. [14C] glucose-6-phosphate and unlabeled RuDP were included as markers. Following electrophoresis, strips were Stained for RuDP and scanned for radioactivity. More than 90Z of the radioactivity co-electrophoresed with RuDP. The remainder co-electrophoresed with glucose-6-phosphate. Radiochemical purity was further indicated by the chromatographic behaviour of reaction controls in several solvent systems. The specific 126 1 activity of the [U- 4C] RuDP was assumed to be the 14 same as the [U- C] glucose-6-phosphate from which it was made. Manganese Catalyzed Oxidation of Ribulose Diphosphate by Hydrogen Peroxide All reactions were performed at 250. The basic reaction consisted of the following components in a total volume of 250 pl: 10 mM bicine-NaOH, pH 8.4, 10 mM MnClZ, 30 mM hydrogen peroxide and 0.2 mM [U-IAC] RuDP (2.3 uCi/ umole). The reaction was generally allowed to proceed for 30 min or 1 hour before team- ination by the addition of 75 units of catalase. A control omitting hydrogen peroxide was run concurrently. The reaction products were identified by paper chromatography in several solvent systems. No one solvent was capable of resolving the products and reactants. Other details are given in the figure legends. After it became apparent that 3-phosphoglyceral- dehyde was a reaction product it was possible to follow the reaction spectrophotometrically by coupling the production of 3-phosphoglyceraldehyde to NADH2 oxidation by the use of triose phosphate isomerase and a-glycero- phosphate dehydrogenase. The spectrophotometric assay contained the following components in a total volume of 250 pl: 10 mM bicine-NaOH, pH 8.4, 10 mM MnClz, 30 mM H202, 0.15 mM NADHZ, 0.8 mM RuDP, 10 units triose 127 phosphate isomerase and 2.7 units of a-glycerophOSphate dehydrogenase. Results Identification of Reaction Products A two electron oxidation of RuDP with carbon- carbon cleavage occurring between C-2 and C-3 can conceivably yield 2 sets of reaction products: i.e., 2-phosphog1ycolate and 3-phosphoglyceraldehyde and/or 2-phosphoglycolaldehyde and 3-phosphoglycerate. It was not possible to separate all these compounds from one another in any one chromatographic system. Following the reaction of [U-14C] RuDP with hydrogen peroxide, an aliquot was subjected to paper chromatography in n-butanol : propionic acid : water (10:5:7 by volume) for 24 hours. The 2 and 3 carbon phoSphate esters listed above were included as markers. After chromatography the location of the compounds was determined either by use of the phOSphate Spray reagent of Bandurski and Axelrod (1951) in the case of the markers and by scanning for radioactivity. The resultant chromatogram (Fig. 20a) revealed 2 peaks of radioactivity, the faster moving one having a pronounced shoulder. Peak I corresponded in position to ribulose diphOSphate. A decision on the composition of peak II could not be made since the markers all ran in this region. However, the control, [U-14C] RuDP incubated without hydrogen 128 .mpxnmpHmHoomeum >H pom op%amvauoome 1m HHH .oumumounnm HH .mumHoo%Hwnm H mum3 muwxuma onH .musoz qH mwB hndmuwoumEouno Ho SOHumuao 65H .AmSDHo> >9 MHNV Hmum3uHoom£m mmSVEoumxm uco>Hom 0:9 .MON oHSme :H SBOLm Emuwoumaouno wnu mo HH xmmm Ho %£mmuwoumaouaonmm .oumumowaum Hum mp%nmpHmHmo%kum .mp%nmon uHoowaum .oumHoomeam nuH3 posdmuwoumaousonoo HH xmom .mnsm omuomoucb ou mpcommmuuoo H xmmm .muson «N mm? knmmuwou ImEouno mo GOHumudp may .AmESHo> hp NumuoHv Houm3upHum HMOHaoumuHocmusn mm3 Emumxm ucm>Hom maH .mnnm Ho SOHumono o m\oz mnu Ho muosooum COHuomoH can we Nnmwuwoumaouao Hommm .bom seawae .oON mudem 129 e_mto Eat Eu ON m... 0.. d d I HH an com 130 peroxide, showed only one peak, that due to RuDP. Thus, on this alone it can be concluded that at least one of the two sets of reaction products has been formed. The area of the chromatogram corresponding to peak II was cut out and the radioactivity eluted with water. The solution was reduced in volume and re- chromatographed for 14 hours in phenol : water (7:3 by volume). Markers were included and the resultant chromatogram stained and scanned as before. The chromat- ogram (Fig. 20b) shomx12 peaks of radioactivity which best corresponded with 2-phosphoglycolate and 3-phospho- glyceraldehyde. Although the other 2 markers overlapped the ratio of radioactivity in the 2 peaks favors assign- ment to 2-phOSphoglycolate and 3-phOSphoglyceraldehyde. In order to confirm that the acid product was 2-phOSphoglycolate and not 3-phosphoglycerate, an aliquot of the original reaction solution was treated with alkaline phosphatase and chromatographed for 17 hours [14C] glycolate and in phenol : water as before. glycerate markers were run.alongside. Two radioactive peaks were obtained, the faster moving (Peak III, Fig. 20c) corresponding in position to ribulose. This peak probably also contained glyceraldehyde. The slower moving peak (Peak II) co-chromatographed with glycolate. No radioactivity was found to co-chromatograph with glycerate (Fig. 20c). Further proof that the reaction products were 131 .m .ouwumume mu Hp Ho GOHuHmoa Home may on mvcommmuuoo HHH xmom .mumHoo Hw mo w nuHB ponmmuwouoEounouoo HH xmmm .AmomsmpHmumowa mchummo anmnoua vcmv mafia nmuommnan cu mucoamouuoo H xmom .muno: NH mmB nmmuwoumEOHLQ Ho SOHumuap one .AoEDHo> %n MHNV umumsuHocmna mm3_Eoum%m ucm>Hom mSH m umnamone mcHmeHm suHB udeummuu Hmuwm .mosm Ho SOHumono o m\qz mnu Ho muoopoua COHuommu mnu mo SnamuwoumEouno Hmamm .UON oustm 132 oq 5.9.5 Eat Eu om ON 9 d 53 cm 133 2-phosphoglycolate and 3-phosphoglyceraldehyde was obtained from the Spectrophotometric assay. The require- ments for the reactions are given in Table 6. The results indicate that the reaction requires RuDP, MhClz and hydrogen peroxide. Equimolar concentrations of magnesium do not substitute for manganese. The oxidant is clearly not oxygen Since catalase completely inhibited the reaction. Furthermore the requirement for both coupling enzymes for the oxidation of NADH2 demonstrates that the product of the RuDP oxidation is indeed 3- phosphoglyceraldehyde. TABLE 6.--REACTION REQUIREMENTS FOR THE MANGANESE CATALYSED OXIDATION OF RIBULOSE DIPHOSPHATE BY HYDROGEN PEROXIDE Reaction P-glyceraldehyde Assay n moles NADH2 oxidised / min. Complete 1.15 - RuDP 0.12 - M’nCl2 0.32 - M'nCl2 + 0.01‘MIMgCl2 0.32 - TPI, or - 0 CPD 0 Complete + 152 units catalase 0 The complete reaction is described in the Methods section. 134 The requirement for manganese ions in the hydrogen peroxide dependent oxidation of RuDP strongly suggests that the oxidation proceeds by a free radical mechanism. A plausible mechanism is shown below. However, apart from the nature of the reaction products, there is no experimental evidence to support this mechanism or, for that matter, to exclude other mechanisms. 2 CH20P03 CHZOPO3 CHZOPOB (IIHZOPO3 f=0 H20 |=0 0H° f=0 fOH C \ H‘f'OH o H-f—O o __D H_?§0£0_H M OH H-(li-OH H-(ll—OH H-(ll-OH H ,0 CHAPTER.V THE ENZYMATIC OXIDATION OF RIBULOSE DIPHOSPHATE BY MOLECULAR.OXYGEN Introduction The results of the ipgyiyp 1802 experiments described in Chapter II indicated that glycolate bio- synthesis involved the incorporation into the carboxyl group of one atom of oxygen, derived directly or indirectly from molecular oxygen. In a brief commun- ication Bowes pp a1., (1971), reported that a purified preparation of soybean ribulose diphosphate carboxylase catalysed the oxidation of ribulose diphosphate by molecular oxygen to yield P-glycolate and, they assumed, P-glycerate. This chapter confirms and considerably extends that initial observation with a purified prep- aration of Spinach ribulose diphosphate carboxylase. The reaction to be considered is: Ribulose diphosphate + 02 ~ P-glycolate + P-glycerate The reaction was catalysed by purified preparations of RuDP carboxylase, but the oxidative reaction is hereafter referred to as RuDP oxygenase. The experiments described in this chapter were 135 136 part of a cooperative project with Dr. T. John Andrews. Materials Spinach was grown as previously described in Chapter I. DEAE-cellulose (0.66 meq.g-1), hydroxylapatite (Bio-Gel HT) and Dowex - A950W -XZ (200-400 mesh) were obtained from Bio-Rad Laboratories. Sephadex G-25, medium, was a product of Pharmacia. All commercial preparations of enzymes were obtained from Sigma Chemical Co. The barium salt of 3-PGA was obtained from Boehringer-Mannheim Corp. and converted to the Tris salt by treatment with Dowex 50+ form and adjustment to neutrality with Tris base. 2-P-glycolate was from General Biochemicals. Silylating reagents were obtained from Regis Chemical Co. All other chemicals were from Sigma Chemical Co. [14 Corp., [14C] fructose diphOSphate from New England Nuclear C] NaHCO3 was obtained from Amersham Searle Corp., and [14C] glucose-6-phosphate and [lacJ-fructose- 6-phOSphate from Calatomic. [14C]-ribulose-diphosphate was prepared (Chapter IV). [180] oxygen (93.5 atoms Z) and [180] water (20.6 atoms Z) were obtained from Miles Laboratories. Methods Exploratory Assays In developing an assay, it was decided to concentrate 137 upon methods which would reflect oxygenation and which could not possibly be confused with carboxylation. Methods based on the disappearance of RuDP or appearance of P-glycerate were bypassed, since carboxylation would clearly interfere with either. Attempts were made to develop assays for glycolate which could be formed from P-glycolate by the action of alkaline phosphatase. They were discarded due to lack of sensitivity, inter- ference by ribulose and/or glycerate or non-Stoichio- metric conversion. For example, ribulose interfered with the colorometric Calkin's test for glycolate and with the phenylhydrazone assay of glyoxylate. Attempts were then made to assay oxygen consumption. For economic reasons (RuDP costs $1.50 mg-l) reaction volumes had to be kept to a minimum. When a one ml solution was equil- ibrated with lOOZ oxygen and oxygen consumption measured with a standard Clark oxygen electrode, back diffusion of oxygen into the electrode chamber resulted in an unacceptably high blank rate. RuDP Oxygenase Assay A.manometric method was used in most of this work, even though this method has several disadvantages. Single side arm warburg flasks with a total volume of about 3‘ml were used. The Gibson Constant Pressure Respirometer was run at 160 shakes min-1. During initial investigations the temperature was 30°. Subsequently it 138 was found that by Operating at 25°, a temperature nearer to ambient, the large pressure fluctuations associated with tipping in the contents of the Side arm could be minimized. The apparatus was equipped with Tygon tubing which was permeable to oxygen and thus it was not ideally suited for use with 100Z oxygen. However, by flushing the apparatus with lOOZ oxygen for two to three hours prior to use, the blank rate due to leakage of oxygen was substantially reduced. The reason for this effect is not known. The standard reaction mixture contained the following components in a total volume of 1.0 ml: in the flasks, 100 umoles ammediol-Cl, pH 9.3, 10 umoles M3012, l umole EDTA, 0.4 umoles dithiothreitol and enzyme (commonly 1 to 2 mg protein) to give a rate of oxygen consumption of between 2 and 8 ul per min, and in the Side arm 2 umoles RuDP. With the side arm.vent open, the system was flushed with humidified lOOZ oxygen for 3 minutes. The oxygen supply was turned off, the vent closed and the system.allowed to equilibrate for a further 9 min. The reaction was then initiated by tipping in the RuDP. Readings were taken at 90 sec intervals. A water blank or alraction blank (less enzyme or with boiled enzyme) was run concurrently. The values reported are corrected to standard temperature and pressure. 139 RuDP Carboxylase Assays 14 (a) following components in a total volume of 250 pl: 25 C Assay: The reaction contained the umoles Tris-Cl, pH 7.80, 2.5 umoles‘MgClz, 0.015 umoles EDTA, 1.25 umoles dithiothreitol (freshly prepared), 12.5 umoles [14C] NaHC03 (0.72 uc/umole), 0.125 umoles RuDP and sufficient enzyme to give rates less than about 16,000 dpm per min Of enzyme reaction. The enzyme was preincubated at the reaction temperature of 300 for 10 min before initiating the reaction with RuDP. The reaction was terminated after 1 min by the addition of 0.5 ml 2N HCl. The solution was then quantitatively transferred to a glass scintillation vial and evaporated to dryness in an oven at 95°. To ensure complete removal of the unreacted 14C02, an additional 0.5 m1 2N HCl was then added and the samples redried. After cooling, 1.0 ml water was added followed by 14 ml of the scintillation cocktail (Chapter III). The cross channel ratio method of counting was used. The rate of reaction was propor- tional to enzyme concentration. However, the progress of the reaction was not consistently linear but appeared to depend upon factors related to the state of purity of the preparation and the previous treatment of the enzyme. This may be related to the activation phenomena reported by Pon pp a1., (1963). In general, activity declined after a Short time. Since this was not the primary purpose of this study, it was not extensively 140 investigated. In order to avoid this non-linearity, the reaction was terminated after 1 min. (b) Spectrophotometric Assay: The reaction contained the following components in a volume of 250 ul: 25 umoles Tris-Cl, pH 7.80, 2.5 umoles MgClZ, 0.015 umoles EDTA, 1.25 umoles dithiothreitol (freshly prepared), 12.5 umoles NaHCO 0.125 umoles NADH, 2.5 umoles ATP, 3, 10 ul of coupling enzyme containing 1.07 units yeast P-glycerate phosphokinase, 1.17 units rabbit muscle P-glyceraldehyde dehydrogenase, 1.00 unit yeast triose phosphate isomerase and 0.90 unit rabbit muscle a-glycer- ophosphate dehydrogenase, and sufficient carboxylase to give a reaction rate not exceeding 2.0 nmoleS/min. The temperature was 25°. The enzyme was preincubated for 10 min before initiating the reaction by addition of RuDP. A reaction blank less carboxylase was routinely included. The rate of reaction was proportional to enzyme concentration. A rapid non-linear rate lasting about 2 minutes was followed by a rate which was linear for the succeeding 10 min. The rapid non-linear rate was also Observed in the reaction blank. The rate was determined from the linear portion of the reaction. A mixture of the coupling enzymes for the carboxy- lase assay was prepared in the following manner. Fifty ul yeast P-glycerate phosPhokinase containing 160 units, 125 ul rabbit muscle P-glyceraldehyde dehydrogenase containing 175 units, 15 ul yeast triose-P-isomerase 141 containing 150 units, and 100 pl rabbit muscle a-glycero- phOSphate dehydrogenase containing 134 units were added to 710 pl 0.05 M Tris-Cl, pH 7.7, 0.002 M EDTA and 0.005 M DTT. (NH45S04, which inhibits the carboxylase reaction, was removed by desalting on a 0.7 x 15 cm column of Sephadex G-25 equilibrated with 0.05 M Tris- Cl, pH 7.7, 0.002 M EDTA and 0.005 M DTT. The enzymes were recovered in 1.5 ml and Stored under N2 at 4°. Under these conditions the coupling system was active for at least 2 weeks. Standardization of [14C] NaHC03 A 0.35 ml aliquot of [14C] Na2C03 (ostensibly 58 uCi/umole) was added to 1.65 ml 0.303 M NaHCOB. To a scintillation vial was added 0.1 ml l‘M Hymamine hydroxide in methanol and 1 ul of the diluted [14C] NaHCO3. This was then followed by 0.9 ml water and 14 ml of scintillation cocktail (Chapter III). Quadruplicate samples were prepared. A dummy vial containing no added 140 was prepared Since phOSphorescence generated by the scintillant in alkaline solution caused spurious counts. The sample vials were counted after the activity in the dummy vial had declined to normal background levels. They were recounted 2 or 3 hours later to ensure that they too had reached a constant count rate. 142 Standardization of RuDP Stock solutions of RuDP were standardized by the 14C carboxylase assay and also by the Spectrophoto- metric carboxylase assay. Both methods involved the use of excess amounts of carboxylase. The RuDP from Sigma was found to be 85Z pure by weight. The RuDP synthe- sised by Paulsen and Lane (1966) was 63Z pure by weight. Spectrophotometric analysis of the RuDP revealed some UV absorbing material, with a peak at 260 nm. This was most probably adenine nucleotides, which are used in the enzymatic synthesis of RuDP. On a molar basis it amounted to no more than lZ contamination. Protein Determination Protein was determined spectrophotometrically at A280 and A260 against appropriate buffer blanks. The formula suggested by Layne (1957) was used. Protein (mg/ml) = 1.55 A280 - 0.76 A26() Polyacrylamide Gel Electrophoresis Fifty to 100 ul aliquots of the purified preparation were subjected to polyacrylamide gel electrophoresis by the method of Davis (1964) on 5.5Z or 6Z gel as indicated. The samples were applied as a solution containing lOZ glycerol. Electrophoresis was performed at 5 ma/tube for a period of time corresponding to twice the time taken for the tracker dye to run through. The gels were stained for a minimum of 3 hours 143 in 0.5Z (v/v) Amido Schwartz in 7.5Z (v/v) acetic acid and destained in 7.5Z (v/v) acetic acid. Analytical Ultracentrifugation Ten m1 of the uniformly suspended (NH4)2SO4 precipitated enzyme was centrifuged at 27,000 g for 15 minutes. The precipitate was dissolved with 1.0 ml of 0.025 M glycylglycine, pH 7.7, 0.10 M KCl and 0.001 M DTT, and the resultant solution desalted on a 0.7 x 15 cm column of Sephadex G-25 equilibrated with the same buffer. Analytical ultracentrifugation was conducted in a Spinco MOdel E Analytical Ultracentrifuge at 3.90 and a protein concentration of 11.6 mg/ml. After a Speed of 50,740 rpm.was obtained, photographs were taken at 4 min intervals. The phase angle was 70°. Identification of Reaction Products by Mass Spectrometry Following the reaction, the solution was trans- ferred to a test tube and frozen until use. After thawing the tube plus contents were held in a boiling water bath for about 40 sec to precipitate the protein. (It was not necessary to remove the coagulated protein Since most of it adhered to the test tube and what little was carried over was trapped at the top of the ion exchange column.) After cooling, the solution was applied to a 0.5 x 4 cm column of Dowex 50 H+ form, in order to remove the buffer and Mgz+. The column was washed with 1.5 ml water and the eluate evaporated to dryness. To 144 ensure anhydrous conditions 0.1 ml of absolute ethanol was added and re-evaporated to dryness. Forty ul BSTFA containing lZ (v/v) trimethylchlorosilane were added to the dried residue and heated at 1100 for 10 min to facilitate the silylation reaction. Standards containing P-glycolate and P-glycerate (2.5 umoleS/ml) were similarly prepared. Aliquots (0.5 to 1 ul) of the above silylated sample were analyzed using an LKB-9000 combined gas chromatograph - mass Spectrometer, equipped with a 1.4 m x 3 mm (i.d.) silanized glass column packed with 3Z (w/v) SE-30 on silanized Supelcoport (100-200 mesh, Supelco Inc., Bellefonte, Pa.). The column temperature was 1500 and the flash heater temperature was 170°. The flow rate of the helium carrier gas was 30 cc/min. The temperature of the ion source was 2900 and the ionizing voltage 70 eV. Results The report of Bowes‘gp.al., (1971) that a purified preparation of soybean RuDP carboxylase catalysed the formation of P-glycolate from RuDP was confirmed. Details of these experiments are in manuscript (Andrews, Lorimer and Tolbert). A sample of soybean RuDP carboxylase, purified by the method of Paulsen and Lane (1966), was kindly furnished by Dr. W; L. Ogren, USDA Soybean Laboratory, Urbana, Ill. [14CJ- RuDP prepared 145 as described in Chapter IV was incubated with this enzyme in the presence of lOOZ oxygen. Treatment Of the products with alkaline phOSphatase was followed by paper chromatography with n-pentanol saturated with 5 M formic acid. Besides unreacted [14C1' ribulose, 2 additional radioactive products were formed which co- chromatographed with marker [14C] glycolate and [14C] glycerate. No glycolate formation occurred under 100Z N2. In addition the reaction appeared to occur much more rapidly at pH 9.0 than at pH 7.8. Purification of RuDP Carboxylase from Spinach Leaves All operations were performed at 0 to 4°C. At each stage of the purification a 1.0 ml aliquot was removed and applied to a 0.7 x 15 cm column of Sephadex G-25 (medium) equilibrated with 0.025 M Tris-Cl, pH 8.0 and 0.001 M DTT, in order to remove phenolic material and (NHA)ZSO4 which interfere with the protein and carboxylase assays reSpectively. Prior to use, the pH of the saturated (NHL4)2804 was adjusted with NHAOH so that a 1:5 dilution gave a pH of 7.3. Initial Enzyme Extraction: About 100 g (fresh weight) washed, deribbed Spinach leaves were homogenized for 45 sec in a Waring Blender with 250 ml 0.1 M Tris-Cl, pH 8.0, 0.002 M EDTA and 0.05 M mercaptoethanol. The brei was filtered through 2 layers of cheesecloth and one layer of "Mirah cloth" and the filtrate centrifuged 146 at 18,000 g for 30 min. The clear yellow supernatant solution constituted the initial extract. (NH4)ZSO4 Fractionation: Sufficient saturated (NH4)2804 was added to the initial extract to give 30Z saturation. After standing for 30 min the suspension was centrifuged at 18,000 g for 10 min and the precip- itate discarded. The supernatant solution was brought to 50Z saturation by the addition of saturated (NH4)ZSO4. After standing for 30 min the suspension was centrifuged at 18,000 g for 30 min. The enzyme was stored as the precipitate overnight, and redissolved with about 15 m1 0.1'M Tris-Cl, pH 8.0, 0.002 M EDTA and 0.05 M mercapto- ethanol. Insoluble material was removed by centrifu- gation at 18,000 g for 20 min. This supernatant solution constituted the (NH4)2304 fraction. Zonal Centrifugation: The density of the (NH£)2804 fraction was adjusted by dilution with buffer so that it was just less than that of 12.5Z (w/w) sucrose. The B-30 zonal rotor (International Equipment Co.) was used. With the rotor Spinning at about 2,500 rpm, 60 ml of buffer were pumped into the rotor via the rim line at a rate of about 10 ml/min. The enzyme solution was then pumped into the rotor, followed by a 520 ml linear (by volume) gradient from 12.5Z to 30Z (w/w) sucrose in 0.05 M potassium phOSphate pH 7.60, 0.1 mM EDTA, 0.001'M DTT. This displaced the enzyme solution inwards towards the core. The gradient was then developed at 50,000 rpm for 147 5 hours. After decelerating to about 2,500 rpm the gradient was displaced out of the rotor via the rim line by pumping distilled water into the core. 25 ml fractions were collected. Alternative methods of loading and unloading the rotor have also been attempted without significantly decreasing the half-band width of the carboxylase. The results of the zonal centrifugation step are Shown in Fig. 21. RuDP carboxylase constitutes such a large proportion of the soluble protein of the leaf, that it is not always necessary to assay for carboxylase activity. Experience Showed that the largest peak of protein invariably contained the carboxylase. Thus, the fractions indicated in Fig. 21 were pooled solely on the basis of absorbance. The enzyme was precipitated by the addition of 2 volumes of saturated (NH4)ZSO4 to bring the solution to 66Z saturation. The suspension was centrifuged at 18,000 g for 20 min and the precipitate dissolved with 15 ml 0.005 M potassium phosphate, pH 7.60, containing 1 mM DTT. The resultant solution was desalted on a 2.3 x 2.5 cm column of Biogel P-6 equilibrated with the same buffer. The desalted enzyme was designated the zonal centrifuge fraction. Hydroxylapatite Column Chromatography: The zonal centrifuge fraction was applied to a 1.6 x 11 cm column of hydroxylapatite, equilibrated with 0.005 M potassium phosphate, pH 7.60 containing 1 mM DTT. The column was then washed with 40 ml of this buffer followed Figure 21. 148 Zonal Centrifugation of RuDP Carboxylase after (NH4 ) $04 Fractionation. The fractions indicated ngre pooled on the basis of the absorbance at 280 nm. Previous experiments indicated that this peak contained the carboxylase activity. 280 149 J l RIM 100 300 VOLUME IMLSI 560 CORE 150 by a 120 ml linear potassium phosphate gradient from 0.005 M to 0.050 M. The flow rate was about 15 ml/hr and 10 ml fractions were collected. The results are Shown in Fig. 22. Fractions containing more than 5 umoleS/min of carboxylase activity were pooled. An aliquot was removed for assay and the remainder precip- itated by the addition of 2 volumes of saturated (NH4)ZSO4. EDTA and mercaptoethanol were then added to produce final concentrations of 0.1 and 5.0 mM respectively. The suspension was Stored at 0 to 4°. Before use, an aliquot was withdrawn and centrifuged at 10,000 g for 10 min. The precipitate was dissolved with 1.0 ml 0.25 M glycylglycine at pH 8.6, 0.01 M EDTA and 0.01 M DTT and desalted on a 0.7 x 15 cm column of Sephadex G-25 equilibrated with the same buffer. A summary of two such purifications is shown in Table 7. The specific activity of the carboxylase was marginally less than the value of 1.4 umoles min-1mg-1 reported by Lane's group (Paulsen and Lane, 1966 and Wishnick and Lane, 1971). Note that in order for a comparison of the Specific activities to be made, the values reported in Table 7 should be multiplied by 1.92 to correct for differences in protein determination. The corrected specific activities were 1.25 umoles min-1 mg-1 for preparation A and 1.39 umoles min-1mg-1 for preparation B. The specific activity of the carboxy- lase was constant (within experimental error) across the Figure 22. 151 Hydroxylapatite Column Chromatography of RuDP Carboxylase. A----A RuDP Carboxylase activity (umoles min‘lml‘l). H A280 A————i Absorbance ration, Azgo/A26o 0----0 Specific activity1 of _RuDP carboxy- lase (umoles min-1mg 152 $258.5: 1 p p 2 1| ._z\z_z\mm._oz; 150 III) VOLUME (M LS) 153 .e.e me um mumeemoee z mmo.o an umSOHHom e.H me um mumnmmonm EsHmmmHoa z moo.o nuHB uSOHpmuw doom m >9 UOQOHo>Oo GESHOU¥¥ .uGOHome mumoamona HoocHH m Sn OOQOHo>Op GESHOO¥ Hm Ne.o See o.~ New «smeaememHexoeuem Se me.o new H.H can coHemwsuHeeemu HmsoN no umGOHuomHm ee mm.o oem e.o Nee eOmNA mzv Non-mm OOH ¢¢.o mom m.o omNH uomuuxm HMHuHaH m SOHumumamum em me.o mHH o.~ mHH emueumemHexoeeem ae mm.o ace m.H Gem :OHumwseHeusmu Hesou I» mowumSOHu omnm em Ne.o omm w.o can ow A mzv Non-om OOH om.o men e.o mHHH someuxm HmHuHsH < COHumemmHm cHououm H H-ws -ses -sHe oe~\ow~ we pHOHw mm 081 meoEJ OHumm GHOuoum mm>¢MH mUxo mnam .1111. .Ommexonumu mmnm To .owN£mmuwoumEouoo aEsHoo mmOHDHHOO m3 mmmamw>xo manm pom mmMH>xonHmo mnam Ho SOHHMOHHHHDQIOU .mm meste 157 1.111,.le 20 $310er g c- 1 1 d1 1 oo' .o‘ e' N' d d d d 1.114 1le z0:) $310er I I I I I I I I I I J l I 1 10 st 00 N 90 110 130 FRACTION N UMBER 70 158 than that of 12.5Z (w/w) sucrose. Buffer was added, when necessary to reduce the density of the protein solution. With the B-30 zonal rotor Spinning at about 2,500 rpm, a 500 ml linear (by volume) sucrose density gradient from 12.5Z to 30Z (w/w) sucrose in 0.025 M glycylglycine pH 7.6, 0.001 M EDTA, 0.001 M DTT, was pumped into the rotor via the rim line at a rate of about 10 ml/min. Sufficient 35Z (w/w) sucrose was then pumped into the rotor until the 12.5Z (w/w) sucrose emerged from the core. The protein solution (about 30 ml) was carefully layered over the 12.5Z sucrose, a further 50 ml overlay of buffer layered over the protein solution, and these were then pumped into the rotor. The gradient was centrifuged for 4.5 to 5 hours at 50,000 rpma After decelerating to about 2,500 rpm the gradient was diSplaced out of the rotor via the core line by pumping 35Z (w/w) sucrose into the rotor at the rim. 25 ml fractions were collected. The results are Shown in Fig. 24. Fractions 8 through 14 were pooled and 1.5 volumes saturated (NH4)ZSO4 were added. The resultant suSpension was stored at 4°. Before use, the enzyme was desalted as previously described. A summary Of the co-purification of RuDP carboxylase and RuDP oxygenase is given in Table 8. RuDP carboxylase and RuDP oxygenase purified together as if they were the same protein. However, the ratio of carboxylase activity to oxygenase activity did 159 illllllllll .OmmH>xonHmo Anna o....o .ommamw>xo mmnm .1111. .owNn Omchw>xo mesa pom mmmH>xonHmo mesa mo :OHDMOHHHHDQIOU .qN OHDme 160 on l1.11111 ZO $310M O 531 3 .§ 1:01,.le ‘200 5310er -|1 I I .4 g I ” Io xxx-”T ,H’II" -1 a ”we“ '- < ' -7\T I 1 : \ q g I \ I I I - .8. f 1. "CORE" A280 4- VOLUME IM LSI 161 coaumw q~.o on mm Hq mom oo.~ mom udmwuuamo HmfiON mm.o no mm mm dad mm.H oom mmOHSHHwo m¢mm N¢.o ooH mNH OOH on om.o oo¢.~ pomuuxm HmHuHaH o N Mmmcmwbno N N lolk AwEv mmmahxonumo meHM awexmmaoan wwmww aflaxmmaoa1 cw~< :Hmuoum mmwcwwxxo mnnm mmmamxonnmo mnam Mm3 pmhmamua mm3 Acamuohm w: oov m8%ncm .ommazxonumo masm commando mo mammposdouuomam How mpwewamuommaom hog flog mug mpg flag .mN musmflm 164 TL 3; flow 7: .‘rrlhu‘ . T3 165 60% saturated (NH4)2804, the preparation was run in the Model E Analytical Ultracentrifuge. A.minor contaminant was revealed (Fig. 26a). Electrophoresis of the same enzyme preparation on the following day also revealed the presence of additional bands which had not been present immediately after preparation (Fig. 25c). These results were taken as evidence that the enzyme was polymerizing. Such behaviour had been observed before by Kleinkopf gt 31., (1970), for the purified barley enzyme, and by Ridley $3.31., (1967) for the purified Spinach enzyme. To confirm that polymerization was indeed continuing, gel electrophoresis and analytical ultracentrifugation were again performed on the same enzyme preparation after 25 days storage in (NHh)2304. Electrophoresis revealed additional bands which were much more intense than previously (Fig. 25d). Analytical ultracentrifugation performed under exactly the same conditions as before showed that the contaminant had increased in concentration and that in addition an even faster sedimenting peak was now apparent (Fig. 26b). The calculated sedimentation coefficients ($20, buffer) for the three species are 18.6, 26.2 and 34.1. Polyacryla- mide gel electrophoresis of a different preparation of enzyme, stored in the same manner for 49 days, revealed evidence of further polymerization (Fig. 25e). Thus, apart from this known aggregation phenomenon, the prepar- ation appeared to be homogeneous by these commonly Figure 26. 166 Schlieren patterns of purified RuDP car- boxylase obtained in the Spinco MOdel E ultracentrifuge. Both photographs were taken 24 min after reaching a speed 8f 50,740 rpm. The temperatuge was 3.9 and the phase angle was 70 ._ The protein concentration was 11.6 mg ml in both cases, and the buffer was 0.025 M glycylglycine, pH 7.7, 0.1MIKC1 and 0.001M DTT. The upper figure was obtained after storing the enzyme as a precipitate in (NHfi) SO for 11 days and the lower figure w s 3btained after 25 days of storage in (NH4)2804. 167 168 used criteria. RuDP Oxygenase Assay With the manometric assay described in the IMBthods section, the rate of oxygen uptake was linear until about 50% of the RuDP had been consumed (Fig. 27). The rate was also proportional to enzyme concentration (Fig. 28). The requirements for the reaction are listed in Table 9. The oxidation of RuDP was clearly enzymatic since no oxygen uptake occured with boiled enzyme. Like the carboxylase, the oxygenase reaction was dependent upon Mg2+ and was stimulated by DTT. The oxygen uptake in the presence of DTT was not due to the oxidation of DTT, since the rate of oxygen uptake by controls without RuDP or enzyme was the same whether or not they contained DTT. The reaction also depended upon oxygen, there being no gas uptake under 100% nitrogen. This result confirmed the [14C] data in which no P-glycolate was formed under nitrogen. Lastly, of course, the reaction depended upon RuDP. pH thimum The pH optimum.for the oxygenase reaction is shown in Fig. 29. While these measurements are complicated by buffer effects, the pH optimum appears to be about 9.3. Details concerning the pH optimum are in manuscript (Andrews, Lorimer and Tolbert). 169 Figure 27. Oxygen consumption by RuDP oxygenase as a function of time. The reaction was initiated by the addition of RuDP. D _acomplete reaction -—-—'reaction blank, less enzyme or RuDP 170 _ h p 0 ms. 2 omznmzoo zmeca .1 1.0 _ 10 MINUTES 171 Figure 28. Oxygen consumption by RuDP oxygenase as a function of enzyme concentration. 172 .15 - 0 1 72.2 No mmzo — 5 ml 1.2 mg ENZYME 0.5 173 Figure 29. The pH Optimum for RuDP Oxygenase D—Cj in 0.1 M ammediol-Cl buffer. l—. in 0.1 M glycine-KOH buffer. 174 40,. 0 0 3 2 FIGS PIC_E WG—0pbc — 0 1 pH 100 175 TABLE 9.--OXYGEN UPTAKE BY RuDP OXYGENASE. THE REACTION iMIXTURE WAS AS DESCRIBED IN THE METHOD SECTION LESS THE COMPONENTS INDICATED. Contents 02 uptake nmoles/min/mg protein Complete 70 - DTT 38 - MgC12 - RuDP - enzyme boiled enzyme (2 min., 100°) OOOOO - 02, + N2 176 The Stoichiometry of the Reaction When an accurately determined quantity (2 moles) of RuDP, standardized as described in the Methods section, was incubated with the enzyme under 100% oxygen, and the reaction allowed to run to completion, a total of 2 umoles of oxygen was consumed " “I indicating that oxygen and RuDP were consumed with a stoichiometry of 1 : 1 (Figure 30). One may conclude, additionally, that under these conditions there is l ‘ \. little or no carboxylation of RuDP. Proof of Reaction Products The reaction products were identified by com- bined gas liquid chromatography - mass spectrometry. Two major compounds (peak I and II in Fig. 31) were present which co-chromatographed with authentic standards of the trimethylsilyl derivatives of P-glycolate (peak III) and P-glycerate (peak IV). Mass spectra (Fig. 32 and 33) confirmed that the products were indeed P- glycolate and P-glycerate, i.e. the mass spectra obtained from peak I and II were identical to those obtained from peaks III and IV reSpectively. The RuDP used for these studies did not contain any detectable P- glycolate or P-glycerate. On one occasion, the small peak preceding peak I (Fig. 31a) was sufficiently large to merit identification. Its mass Spectrum was con- sistent with it being the trimethylsilyl derivative of Figure 30. 177 The stoichiometric consumption of oxygen by RuDP oxygenase. The reaction was initiated by the addition of 2 umoles RuDP. 178 1 mxxo mmnoz; MINUTES Figure 31. 179 (A) Gas liquid chromatographic separation of the silylated reaction products from the RuDP oxygenase reaction. (B) The tri- methylsilyl derivatives of authentic P- glycolate (III) and P-glycerate (IV). Mass spectrometry confirmed that peaks I and III were (Me Si) .- P-glycolate (Figure 32) and Seakg II and IV were (Me3Si)4 P-glycerate (Figure 33). 180 11 JA’L-- u L. 80I 20... mmzommwm 2mm 60'. mi. 15 10 MINUTES 181 A.>o ORV mumaoo%awumumaflmmmzv mo Efiuuommm mmmz .Nm munwfim 182 l w“ «5.2 onx own hmm 0\Cg own omN 0¢N con .05 _ mum mam Amozzmno/oso w Agmaoinoouf m . Amuzgm om“ 0N— NS th >22 330050155923 om mm. TON low Tom OOH ALISNBLNI BAIIV'IBH 183 A.>m QC mumuoomawumuqfimmozv mo 53.30on 3% .mm musmflm F 184 ES cm? 9.} 00¢ owm owm e_mm 3N own own e_Nfi 0.0 L n h p n b p h — p h L h b F — b P h b b h b p F P r b h b L b F b + J. J. J]. 44 1.1. J41 I... 1.. 4 1:114 ‘1. «<4 144 Xv.2\ 4&1 :4 41 J A _ _ _ m m :m omv Em Am§xmoxoxo Rm 5 AngVWIOl— BM _ vnv >>S_ .mozxmloiuononmr _ .v m «mozxmro mmeoboi am 2): ms Iom oou m" ALISNBLN I HAHN-lab! 185 the ethyl ester of P-glycolate. This ester was pre- sumably formed during the drying of the sample with absolute ethanol. At this stage, following ion exchange chromatography on Dowex 50 H+, the solution was acidic, which would catalyse esterification. This peak was absent from the standards which were not treated with Dowex 50 H+ form or ethanol. Analysis of the Mass Spectra of (Me Si Q P-glycolate The mass spectrum of (Me3Si)3 P-glycolate contains the molecular ion at m/e 372 (Fig. 32). The most intense ion occurs at m/e 357 (M-15), an ion which arises by the loss of a methyl group from the molecular ion. The ion at m/e 328 (M-44) most probably arises by a rearrangement involving the loss of the carboxyl group to give an ion with the structure, CH2 = +0 - P - (OMeBSi)3. The assignment of structure was confirmed by the absence of 18O in this ion in samples of (MeBSi)3-P-glycolate containing 180. The ions at m/e 315 and m/e 299 are common to the trimethylsilyl deriv- atives of phOSphate esters (Zinbo and Sherman, 1970) and are related to one another by the metastable ion at m/e 284. The ion at m/e 315 most probably arises by a rearrangement of the molecular ion to give the ion with the structure, (Me3SiO)2 - P(OH) = +0Me3Si, and a metastable ion at m/e 268 supports this contention. It should be noted that the ability of trimethylsilyl 186 groups to undergo migration in the mass spectrometer is well documented (DeJongh gt 31,, 1969 and McCloskey 3511., 1968). The ion at m/e 299, which has the struc- ture (MeBSiO)2 - P(O) - 0+ = SiMez, clearly arises from two sources. Firstly it may arise by rearrangement from the ion at m/e 357, and a metastable at m/e 250 supports this transition. Secondly, it may arise from the ion at m/e 315 by the loss of -CH3 plus H and a metastable at 284 supports this transition. Inter- estingly, in samples containing 180 in the carboxyl group, the ions at m/e 315 and 299 were also found to contain 180. Thus, the rearrangement leading to the formation of the ions at m/e 315 and 299 involves a transfer of -0Me3Si to the phosphorus atom rather than simply the transfer of #Me3Si to one of the available oxygen atoms of the phosphate group. Other phosphate ions are located at m/e 227 and 211 with a metastable ion at m/e 196 relating these ions. Analysis of the'Mass Spectra of (Me Si , P-glycerate The (Me3Si)4-P-glycerate Spectrum contained no molecular ion (Fig. 33). The ion at m/e 459 is due to the loss of a methyl group from the molecular ion. The prominent ion at m/e 357 most probably corresponds to the structure (MeBSiO)2-P-(0)-OCH2-CH = +O-Mie3Si, i.e. the loss of the silylated carboxyl group from the molecular ion, and the metastable ion at m/e 269 supports 187 this interpretation. The structure of the ion at m/e 387 is most perplexing. This ion is common in the mass spectra of the MeBSi derivatives of sugar phosphates (Zinbo and Sherman, 1970). On the basis of deuterium labeling, they assigned the structure Me3Si-+0 = P- (OMe3Si)3 to this ion. In the case of the sugar phos- phates this ion is thought to be derived from that m/e 459 and a metastable at m/e 326 supports this contention. The spectrum of (Me3Si)4 P-glycerate also contains an ion at m/e 459 and a metastable ion at m/e 326. However, in this case the ion at m/e 459 contains a total of 11 methyl groups, while the structure proposed by Zinbo and Sherman (1970) for the ion at m/e 387 contains a total of 12 methyl groups, an obvious discrepancy. In samples containing 180 in the carboxyl group the ion at m/e 387 was found not to be labeled. A possible Structure consistent with this observation is [MeBSiO-CHZ-O-PmMeBSi) (OSiMe2)2]+. The spectrum also contains the familiar phosphate ions at m/e 315 and 299 (and the related metastable at m/e 284) and at m/e 227 and 211 (and the related metastable at m/e 196). Substrate Spec ifity A number of substrates, other than RuDP, were tested for activity by the standard oxygenase assay (Table 10). Fructose diphosphate and fructose-6- phosphate gave a very marginal rate above that of the .ommahxonumo mnnm ammn%0m mo cowumumamum onu nuw3 Cowuommu kn pmsflanmumnk 188 o m mnoaxm mumaamonmuo mmouUSHm o N mnoaxm mumnamonmwu omouosum u H muoaxm mumumomawonamosmum «o u muoaxm omoannam - H Tons 32a8fi$ $333 *2: 2: Tons 323233 8332 uoahom floumaoomawg cum N .IJluo mumaoomamumflowq oxmuan o mumm m>wumamm AZV .ocoo mummumndm mm- 75« - t (I) Z LU '— Z 50- . Lu 2 ’2 _l UJ a: 25- 1 440 1.50 1.60 1.1.0 1.530 1.60 m/e 198 derivatives of authentic samples of P-glycolate and P-glycerate are compared with similar spectra for P-glycolate and P-glycerate obtained after incubation of RuDP with the oxygenase under an atmosphere con- taining 82% 1802. In the case of the P-glycolate spectra, a comparison of the relative intensities of the ions at m/e 357, 359 and 361 clearly shows that one, and only one, atom of 180 has been incorporated. The apparent increase in the relative insensity of the ion at m/e 361 is due to the combination of one atom of oxygen-l8 29 with the naturally occurring isotopes of silicon, Si 30 (natural abundance 4.7%) and Si (natural abundance 3.1%). It is not due to the incorporation of two atoms of oxygen-l8. Additionally, a comparison of the ions at m/e 328 and 330, ions lacking the carboxyl group, 180. These results therefore prove shows the absence of that one atom of 18O was incorporated into the carboxyl group of P-glycolate. For the P-glycerate Spectra, a comparison of the relative intensities of the ions at m/e 459 and 461 clearly shows that no oxygen, derived from molecular oxygen, has been incorporated. By default then, the carboxyl oxygen of P-glycerate must be derived from water. In order to confirm this, the oxygenase assay was run in the presence of H2180. [180]‘water: (b) In order to perform these experiments, the 199 reaction was reduced in volume and run in 4 x 1 cm vials equipped with serum rubber caps. The components were: 20 ul 0.25 M AmmedhflrC1 pH 9.3, 1 ul 1.0 M MgClz, 2 ul 0.01 M DTT, 2 M1 0.05 M EDTA, 20 01 enzyme (20.7 mg/ml) and 60 01 [ISOJHZO (20.63 atoms z). The vial was flushed with 100% oxygen and the reaction initiated by the in- jection of 15 pl 0.025 M RuDP. The final [1803H20 content was thus 10.3 atoms %. The reaction was allowed to proceed at 250 for 60 min with shaking, and terminated by freezing. The Me3Si-derivatives of P-glycolate and P-glycerate were prepared as in experiment 2 above. The results (Table 11) confirmed that the carboxyl oxygen of P-glycerate was derived from.water. However, oxygen-l8 was also incorporated into the carboxyl group of P- glycolate. The labeling of the P-glycolate carboxyl group is to be expected since the keto oxygen of RuDP most probably exchanges with the medium via the hydrated form: 0H20P03 cnzopo3 011201303 18 H20 18 =0 fla an- O-C-OH —-—4——-¢=' C- 0 HC-OH HC-OH HC-OH The absence of incorporation of 18 0, from.molecular oxygen, into P-glycerate during the RuDP oxygenase reaction might have been due to an enzyme catalysed exchange of the carboxyl oxygens with the medium. To ensure that the enzyme did not catalyse any exchange of the carboxyl oxygens of the products with those of the 200 medium, the enzyme was incubated with P-glycolate and P-glycerate in the presence of [180]H20 under 100% oxygen. The reaction was performed in 4 x 1 cm vials equipped with serum rubber caps, and consisted of the following components: 20 ul 0.25 M.Ammediol-Cl pH 9.3, 1 01 1.0 MIMgClz, 2 ul 0.01 M DTT, 2 ul 0.05 M EDTA, 10 ml 0.05 M P-glycerate, 10 ul 0.05 M P-glycolate and 65 ulIIBOJHZO (20.63 atoms %). The vial was flushed with 100% oxygen and the reaction initiated by the injection of 20 ul enzyme (20.7 mg/ml). The final [180] H20 content was thus 10.3 atoms %. The reaction was allowed to proceed at 25° for 60 min with shaking and terminated by freezing. The MeBSi-derivatives were prepared as in eXperiment 2 above. The results (Table 11) indicate that the enzyme did not catalyse the exchange of the carboxyl oxygens of either phosphate ester with the medium. Summary of Other Properties of RuDP Oxygenase The experiments described in this section were performed by Dr. John Andrews. Like all experiments described in this chapter they were the outcome of joint experimental planning. They are included here for the sake of completeness. Activity as a Function of Oxygen Concentration At both 250 and at 30°, oxygenase activity was a linear function of the oxygen concentration in the gas phase from 0 to 100% oxygen. This indicates that 201 the enzyme was not saturated with 100% oxygen in the gas phase. The physiological significance of this result will be discussed later. Activity as a Function of RuDP Concentration Owing to the insensitivity of the oxygenase assay, attempts to determine the KM for RuDP have proven difficult to perform. However, a Lineweaver-Burk plot of the best data indicates a KM for RuDP under 100% oxygen of about 0.2 mM. This must be regarded as an operational K value, since the other substrate, oxygen, 'M was present in sub-saturating concentrations. Stability Inactivation experiments have not yielded consistently reproducible results. Upon storage in (NH4)2804, one preparation showed a constant level of both oxygenase and carboxylase activity for about 3 weeks. Then both activities declined precipitously, suggesting that they were one and the same protein. However, in other preparations there has generally been a slow loss of carboxylase activity which has not been paralleled by equivalent losses in oxygenase activity. The oxygenase activity in general seems to be more stable. This result suggests that the two activities are associated with different proteins. 202 Cyanide Inhibition The oxygenase activity was inhibited about 50% by 1 x 10-4M cyanide. This inhibition appeared to be dependent upon the presence of RuDP. When the enzyme was incubated with different concentrations of cyanide between 0.5 and 2.0 x 10-4M and the reaction initiated by the addition of RuDP, there was a lag before inhi- bition was observed. This behaviour was strikingly similar to the lag in the cyanide inhibition of the carboxylase as reported by Wishnick and Lane (1969). When the enzyme was incubated for 1 hour at 40 with 5 x 10-3M KCN and then the cyanide removed by Sephadex, the oxygenase activity was fully restored. This has also been reported to be true for the carboxylase activity. Copper Content of RuDP Carboxylase Wishnick'gt‘al., (1969) have reported the presence of l g atom of c0pper per mole of enzyme. However, Atomic Absorption analyses of two purified enzyme preparations failed to confirm this report, but revealed the presence of only 0.1 g atom.of copper per mole of enzyme. This discrepancy cannot be attributed to the small differences in the specific activity of the various preparations. Furthermore addition of copper sulphate to a concentration 20 times that of the carboxylase did not stimulate the rate of oxygen uptake. 203 Reaction Intermediates derived from Oxygen Addition of catalase or erythrocuprein (super- oxide dismutase) did not effect the RuDP oxygenase reaction. These results suggest that neither hydrogen peroxide nor the superoxide radical are involved in the oxygenase reaction, although negative results do not unequivocally eliminate the possibility of their involvement. Discussion The term RuDP oxygenase is new. This activity is best described by the rules in Enzyme Nomenclature by the International Union of Biochemistry as ribulose- l,5-diphosphate:oxygen oxidoreductase (E.C. l.l3.l.-). Enzymes in the group 1.13 catalyse reactions "acting on single donors with incorporation of oxygen (oxygenases)". In this group there are no enzymes acting upon sugar phosphates. With the exception of myo-inositol oxygenase and lipoxygenase. ‘Most other enzymes in this group act upon aromatic substrates. Mechanism of RuDP Oxygenase The results of the 180 experiments (Table 11) are consistent with the following mechanism for RuDP oxygenase activity. 204 CH2-0P03 (IIHZ-OPO3 9 CH20P03 (IIH20P03 c=0 C-OH 18o-o-c-OH c H?-OH-————C> -OH ~=> C=0 180 0- OH- H -OH HC-OH HC-OH + CH 0P0 CH 0P0 H OPO 0H 0 2 3 2 3 2 3 \4/ c "Activated 1802"? HC-OH 1 . 18 02 cnzopo3 The oxidative cleavage is depicted as occurring between C-2 and C-3 of the enediol structure of RuDP. Although the results do not eliminate the possibility that the cleavage occurs between C-3 and C-4, this is considered less likely. Specifically labeled [IACJ-RuDP would be needed to clarify this point. The reaction is depicted as proceeding via a peroxide intermediate formed by attack of oxygen on the C-2 of RuDP. The results do not indicate how this intermediate peroxide is formed. Recently a great deal of attention has been devoted to determining the reactive species of oxygen in oxygenase reactions and Hirata and Hayaishi, (1971) and Strobel and Coon, (1971) have reported evidence for the involve- ment of the superoxide radical in the reactions of tryptOphas dioxygenase and the microsomal cytochrome P-450 hydroxylation system respectively. Preliminary evidence indicated that RuDP oxygenase was not inhibited by superoxide dismutase, but this negative result does 205 not unequivocally eliminate the possible involvement of the superoxide radical any more than a positive result would unequivocally establish that the reactive Species of oxygen was the superoxide radical. Prelim- inary evidence also indicated that the purified carboxy- lase did not contain the eXpected 1 g atom of copper per mole of enzyme reported by Wishnick gt al., (1969). This was disappointing since the association of ferrous or copper ions with oxygenases is widespread. There are no reports of other metal ions in purified RuDP car- boxylase. Since the above points remain to be clarified, the mechanism is for the meantime depicted as involving some form of activated oxygen. The mechanism of decomposition of the peroxide intermediate clearly involves the attack and release of hydroxyl ion rather than an intramolecular concerted ‘mechanism, since the latter would result in the incor- poration of molecular oxygen into P-glycerate, which was not observed. The mechanism of decomposition of the peroxide thus appears to be similar to the mechanism of decomposition of the peroxide intermediate formed during the reaction of hydrogen peroxide with a-diketones studied by Bunton (1961) and discussed in the Literature Review. 206 Enzyme Purity An intriguing question raised by these results is whether RuDP oxygenase and RuDP carboxylase are the same or different proteins. The simplest conceptual model would assign the carboxylative and oxygenative activities to different proteins. The consistent differences in percentage yield can be interpreted to support this model. So, too, does the fact that upon storage in (NH4)2304 the carboxylase activity declines more rapidly than does the oxygenase activity. Different pH optima may be interpreted as evidence for two independent proteins. Although evidence for homogeneity was obtained by gel electrophoresis and analytical ultracentrifugation, these results must be interpreted with caution. The carboxylase has an extremely low catalytic activity. Taking the value of 1300 moles RuDP carboxylated per hr per mole of enzyme at 300 reported by Paulsen and Lane (1966) and assuming 8 catalytic sites, calculations reveal that each catalytic site operates at a rate of only about 3 carboxylations per second. Given such low activity and the fact that the oxygenase assay required 1 to 2 mg of protein, it is quite conceivable that a contaminant, undetectable by the techniques used, could well account for the observed oxygenase activity. The second model would assign the carboxylative and oxygenative activities to the same protein and there 207 are several variants of this model. This model is supported by an ostensibly pur enzyme and an apparently similar substrate Specificity. One such variant of this model would be the existence of two independent catalytic Sites, one for carboxylation, the other for oxygenation. Another variant would be the existence of a common catalytic site for both carboxylation and oxygen- ation and that this site is capable of undergoing subtle transformations such that in one state it is capable of catalysing carboxylation while in the other it is capable of catalysing oxygenation. For example, carboxylation might require that the sulphydryl groups, known to be involved at the active site, be in the reduced state, while oxygenation might require that the same sulphydryl groups be in the oxidised disulphide state. The dif- ferences in the pH optima for carboxylation and oxygenation can be rationalized in such a manner. For example, carboxylation might require an amino group to be pro- tonated while oxygenation may require the same group to be unprotonated. The failure to separate the two activities is consistent with this one protein hypothesis. A consistent decrease in the carboxylase to oxygenase activity ratio that is observed during the purification might simply be due to the removal of an activator of the carboxylase and/or an inhibitor of the oxygenase. If this is the case then these effectors must be of high molecular weight since the initial extract was 208 always subjected to gel filtration on Sephadex G-25 before assay. There is precedent for such an effector of the carboxylase. Wildner and Griddle (1969) have described what is termed the "light activating factor" which they isolated from mutant tomato leaves. This factor has a molecular weight of about 6000 and stimu- lated purified tomato leaf carboxylase activity by about 2.7 fold. Alternatively, the decrease in the carboxylase to oxygenase activity ratio might be due to transformation of the catalytic site from a form suited to carboxylation to one suited to oxygenation. The differences in the rates of loss of activity observed upon storage in (NH4)ZSO4 can be rationalized in a similar manner. The observation by Ogren and Bowes (1970) that the carboxylase is inhibited by oxygen and that the inhibition is competitive with respect to C02, suggests that the carboxylase and the oxygenase are one and the same protein. Another item of circumstantial evidence concerns the nature of the cyanide inhibition. The pronounced lag in the inhibition of the oxygenase reaction is strikingly similar to that observed for the carboxylation reaction by Wishnick and Lane (1969). These authors demonstrated the formation of an inactive ternary complex of enzyme, RuDP and cyanide. While a more detailed study of the cyanide inhibition of the oxygenase reaction has not been conducted, owing to the insensitive nature of the assay, the general features of 209 the inhibition are in agreement with those reported for the carboxylase. One final piece of evidence favoring one protein for both carboxylase and oxygenase activity is of a physiological nature. It is the remarkable constancy of the CO2 compensation point within a Species of plants. The CO2 compensation point is the atmospheric concentra- tion of CO2 at which there is no net gain or loss of CO2 during photosynthesis. This is clearly a reflection of the relative carboxylase and oxygenase activities. A low carboxylase to oxygenase ratio would result in a higher CO2 compensation point and vice versa. If, on one hand, the carboxylase and the oxygenase activities were associated with different proteins, one might expect to find varietal differences in the CO2 comp- ensation point, depending on the relative amounts of each protein. If, on the other hand, the two activities are associated with one and the same protein, then the relative activity of each is tightly coupled or fixed, and one would not expect to find varietal differences in the 002 compensation point. ‘Moss gt 31., (1969) have measured the C02 compensation point at 250 in 100 var- ieties of wheat. The value they found was 52 t 2 ppm C02. Similar measurements on 44 genotypes of soybean by Cannell.gt‘al., (1969) gave a constant value of 73't 0.9 ppm C02. While such evidence may be quite fortuitous, it is consistent with the concept that the 210 carboxylase and the oxygenase are one and the same protein. The question whether there are one or two proteins does not invalidate the results obtained here which clearly demonstrate the enzymatic oxidation of RuDP by molecular oxygen, an activity described here as RuDP oxygenase. The Relationship of RuDP Oxygenase to Photorespiration Both the _i_n_ 17.912 and in v_it_r_'2 [180] oxygen results are consistent with the view that the oxygen- ation of RuDP is the primary photorespiratory event in precisely the same manner that the _i_r_1_ Lilo and E 12532 [14CJCO2 results are consistent with the view that the carboxylation of RuDP is the primary photosynthetic event. The warburg effect, photoreSpiration and glycolate synthesis are most evident under conditions of limiting CO2 concentration and high oxygen concen- tration. Furthermore, these processes may, in large part, be overcome by increasing the concentration of C02. It is clear that 32,3132 the concentrations of CO2 and oxygen must govern the relative activities of the carboxylase and the oxygenase. It is proposed that the properties of RuDP oxygenase are consistent with its function as the primary oxidation reaction in photores- piration. It was previously established (Chapter I) that the other oxygen consuming reaction of 211 photorespiration, glycolate oxidase, is effectively saturated under an atmOSphere of about 60% oxygen. Yet photorespiration is known not to be saturated even under atmospheres of 100% oxygen. It was therefore particu- larly interesting that the oxygenase activity was a linear function of the oxygen concentration and that it was not saturated under atmospheres of 100% oxygen. The high pH optimum of the oxygenase reaction is also of physiological interest, since [IACJCO2 fixation studies in gigg with algae (Orth gt al., 1966) and with cell free chloroplast preparations (Dodd and Bidwell, 1971) have demonstrated that the percentage of the total carbon fixed into the intermediates of the photoreSpiratory pathway, glycolate, glycine and serine, increases dramatically at pH values in excess of 9.0. Dodd and Bidwell (1971) eliminated the possibility that this effect might be due to limiting CO2 concentrations at the higher pH values. This response to pH has lacked any enzymatic explanation. This explanation, based on the response of the oxygenase to pH, is attractive. Photorespiration, the warburg effect and glyco- late biosynthesis are known to be stimulated by high light intensity, limiting C02 concentrations and high oxygen concentrations. These conditions would be expected to lead to RuDP accumulation were it not for the activity of RuDP oxygenase. The rather high pH optimum of the oxygenase is consistent with the phenomenon Jagendorf 212 and Neumann (1965) and others have reported that, when unbuffered chloroplasts are illuminated, there is a rapid rise in the pH of the medium. The broken chloroplasts used in such studies are not intact nor are they capable of fixing C02. A rise in the pH of the medium is interpreted as being due to the tranSport of protons into the space enclosed by the lamellar membrane. Thus, in 3312, illumination will result in the stroma becoming more alkaline. This pH shift, of itself, would be eXpected to lead to conditions favoring oxygenation of RuDP with a concomitant decrease in the carboxylation of RuDP. But additionally a shift to more alkaline pH's will effectively shift the equilibrium between C02 and bicarbonate towards bicarbonate. Since CO2 is the substrate for the carboxylase, this pH Shift will further limit the extent of carboxylation. In addition, illumination will clearly result in an increase in the oxygen concentration in the chloroplast. Therefore, the selective requirement of high light intensity for photoreSpiration and glycolate synthesis can be explained firstly by a light dependent pH shift from conditions favoring carboxylation of RuDP to conditions favoring oxygenation of RuDP. This effect is reinforced by the light dependent increase in oxygen concentration in the chloroplast. Whether or not the pH ever rises as high as 9.3 iflfil£ZQ is unknown, but any shift in this direction will favor oxygenation. Indeed the control 213 of photosynthesis and photorespiration might ultimately centre around subtle shifts in the pH of the chloroplast. While the above discussion appears to make teleological sense if nothing else, there is a major discrepancy in the fact that the pH optimum of P-glyco- late phosphatase, also a chloroplast enzyme, is about 6.3 (Richardson and Tolbert, 1961) with very little activity above pH 8.0. Although evidence is lacking, the suggestion that P-glycolate phosphatase is a membrane bound enzyme involved in the tranSport of glycolate out of the chloroplast, is attractive. If this were the case, the pH optimum of the solubilized enzyme may be artifactual. Although the activity of RuDP carboxylase in crude extracts is insufficient to account for the-in 2222 rates of photosynthetic CO2 fixation, few would challenge the pivotal role of the carboxylase. Whether the activity of RuDP oxygenase in crude extracts is sufficient to account for the observed rates of glycolate synthesis and photorespiration iguyigg is difficult to answer meaningfully. The optimum oxygenase conditions of 100% oxygen and pH 9.3 probably do not exist 33 gigg. The activity of RuDP oxygenase in crude extracts is about 25% that of RuDP carboxylase when both are assayed under optimum.conditions. However, if the activities were determined at an intermediary pH, say 8.6, under natural atmospheric conditions of 0.03% 002 and 21% 214 oxygen, oxygenation would exceed carboxylation. This is perhaps an unfair comparison since the KM for 002 for the purified carboxylase is known to be anomolously high (see Literature Review for a discussion of this feature of the carboxylase). The results in Fig. 18 indicate that under 100% oxygen a spinach leaf is capable of photorespiring at rates approaching those of photo- synthesis. Under no circumstances, however, has the rate of oxygenation exceeded that of carboxylation when the assays are performed under their reSpectively optimal conditions. CONCLUDING DISCUSSION The inhibition of net photosynthetic C02 fixation by molecular oxygen, the Warburg effect, can now be explained in terms of the action of RuDP oxygenase which results in the formation of P-glycolate and P-glycerate. Not only does this reaction reduce the pool of RuDP available for carboxylation and thus the rate of C02 fixation but it also leads to the formation of P-glycolate, the subsequent metabolism of which results in the loss of C02. Thus, the oxygenation of RuDP both inhibits C02 fixation and stimulates 002 release. As the glycolate pathway is now conceived, photorespiration involves the uptake of oxygen at two sites, firstly in the synthesis of P-glycolate and secondly in the oxidation of glycolate by glycolate oxidase. The release of C02 occurs at another site, namely glycine decarboxylase. Since the conversion of 2 moles of RuDP to 2 glycolates and then to one serine involves the consumption of 3 moles of oxygen and the release of 1 mole of C02, the photorespiratory quotient is theoretically 3 : 1. This contrasts with the reSpir- atory quotient for dark reSpiration of 1 : 1. 215 216 In order to release one mole of C02, tWO moles of RuDP must first be oxygenated. Therefore at the C02 compensation point one can predict that the rate of RuDP oxygenation is twice the rate of RuDP carboxylation. Furthermore one would also predict that at the 002 compensation point the rate of oxygen uptake is three times the rate of CO2 uptake. Bulley and Tregunna (1970) have attempted to measure the rates of C02 exchange in soybean leaves at the CO2 compensation point. In 21% oxygen, the CO2 exchange rates were determined to be 61 and 87 umoles dm'zbr‘1 at light 4 ergs cm-zsec-l. intensities of 6.5 x lo4 and 28 x 10 At comparable light intensities, the rate of oxygen exchange of soybean leaves at tha CO2 compensation point in 19% oxygen has been measured by Mulchi gt $1., (1971). They reported values ranging from 300 to 350 umoles dm-zhr-l. ‘While these 2 sets of values are not strictly comparable, it is interesting to note that the rates of oxygen exchange exceed those of C02 exchange by 4 to 5 fold. The values for the rates of CO2 exchange are under- estimates owing to the internal recycling of photo- respiratory C02. The requirement of light for photorespiration and for the incorporation of molecular oxygen into the carboxyl groups of glycine and serine is indirect and dependent upon the formation of RuDP, in the same sense that the C02 fixation reactions of photosynthesis are 217 truely dark reactions. 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