ENZYMES FOR OXIDATION OF ALPHA-HYDROXYACIDS IN ROOTS 0F GREEN PLANTS Thesis for the Degree of M. S MICHIGAN STATE UNIVERSTTY MAN PING KO 1975 i s 5W... .mlza’f: THEME E ‘ ‘ - ' W J ‘ "J'IF'T “5-9. A. Q. t» :....'. ‘e'gfingb a "‘ }' £12., m:- 8 &‘ ~ BUUK BINDERY INC. LIBRARY BINDERS smuarm. mum; / \L\ L ' ABSTRACT ENZYMES FOR OXIDATION OF ALPHA-HYDROXYACIDS IN ROOTS OF GREEN PLANTS BY Man Ping Ko Homogenates of young tissues of various higher plants were assayed for glycolate oxidase, a peroxisomal enzyme in green leaves of higher plants, and for glycolate dehydrogenase, a functionally analogous enzyme characteristic of certain green algae. In all the leaves examined, a glycolate dehydrogenase of the type found in green algae was not detectable although excess glycolate oxidase was always present. Glycolate dehydrogenase was also not detectable in the roots examined and glycolate oxidase was present at low level in the roots of only certain species. Glycolate oxidase activity in the roots of watercress was found to be not different from that occurring in the leaf except in sub- strate specificity. The enzyme activity in the roots was particulate, capable of reducing the dye 2,6 dichlorophenol-indophenol and molecular oxygen in the presence of L(+)-lactate, glycolate and D(-)-lactate, in order of decreasing rate. The reduction was insensitive to cyanide or other inhibitors of the respiratory electron transport. NAD-Dependent L(+)-lactate dehydrogenase was not found in the leaves, but it occurred widely in the roots. Hydroxypyruvate reductase Man Ping Ko which was present at high level of activity in the leaves was also present in the roots. Both the lactate dehydrogenase and hydroxy- pyruvate reductase from roots of pea seedlings were partially purified by ammonium sulfate fractionation, DEAE cellulose ion exchange chroma- tography and Sephadex G-200 gel filtration chromatography. Some of their properties were examined and compared to those of the same enzymes from leaves or animals. The root lactate dehydrogenase was nearly specific for L(+)- lactate and NAD, but the D(—)-isomer was also reduced at a very slow rate. In the reverse direction this enzyme reduced not only pyruvate but also hydroxypyruvate and glyoxylate in the presence of NADH. . Optimal activity occurred at pH 9.1 in the direction of pyruvate formation and at pH 7.0 in the reverse direction. It resembled closely heart L(+)—lactate dehydrogenase in its molecular weight, electro- phoretic mobility and its response to heat treatment, inhibition by excess substrate and sulfhydryl group inhibitors. Hydroxypyruvate reductase or D(-)-glycerate dehydrogenase from the roots was specific for NADH and hydroxypyruvate and glyoxylate with a rather broad pH optimum which peaked at pH 5.9. In the reverse direction, it oxidized DL-glycerate in the presence of NAD, with optimal activity at pH 9.6. Its molecular weight, inhibition by excess hydroxypyruvate and sulfhydryl inhibitors resembled closely glycerate dehydrogenase from beef liver or leaves. Hydroxypyruvate reductase from roots was not particulate, but was present in the cytosol as it is in the liver tissue. The possible physiological roles of these enzymes of alpha- hydroxy acid oxidation in the roots were discussed. ENZYMES FOR OXIDATION OF ALPHA-HYDROXYACIDS IN ROOTS OF GREEN PLANTS BY Man Ping Ko A THESIS Submitted to Michigan State University in partial fulfillment of the requirements for the degree of MASTER OF SCIENCE Department of Biochemistry 1975 ACKNOWLEDGEMENTS I want to express my sincere thanks to Professor N. E. Tolbert for his guidance, encouragement and support during the course of this work. I would like to thank also Drs. D. P. Delmer and J. L. Fairley for their serving as members in my guidance committee as well as for their helpful suggestions on this manuscript. I thank also the postdoctoral fellows, fellow graduate students and technicians in Dr. Tolbert's laboratory for the many stimulating and helpful discussions in pursuing my research. The financial assistance of the National Science Foundation and the Department of Biochemistry, Michigan State University, is also appreciated. ii TABLE OF CONTENTS INTRODUCTION. . . . . . . . . . . . . . . . . . . . . . . LITERATURE REVIEW . . . . . . . . . . . . . . . . . . . . Enzymes of Glycolate Oxidation in Photosynthetic Tissues of Plants, Algae and in Animals and Other Nonphotosynthetic Organisms. . . . . . . . Enzyme of Lactate Oxidation in Higher Plants . . . Hydroxypyruvate Reductase in Plants and in Animals MATERIALS AND METHODS . . . . . . . . . . . . . . . . . . Plant Materials. . . . . . . . . . . . . . . . . . Chemicals. . . . . . . . . . . . . . . . . . . . . Preparation of Cell Free Extracts of Plant Tissues Preparation of Particulate Fractions . . . . . . . Partial Purification of L(+)-Lactate Dehydrogenase and Hydroxypyruvate Reductase from Pea Roots . . Sucrose Density Gradient Centrifugation. . . . . . Biochemical Assays . . . . . . . . . . . . . . . . MAD-Lactate Dehydrogenase. . . . . . . . . . . . . NAD D-Glycerate Dehydrogenase or Hydroxypyruvate Reductase O O O O O O O O O O O O O O 0 O O O O I Glycolic Acid Oxidase or Dehydrogenase . . . . . . Catalase . . . . . . . . . . . . . . . . . . . . . Cytochrome c Oxidase . . . . . . . . . . . . . . . Estimation of Molecular Weight by Gel Filtration Chromatography . . . . . . . . . . . . . . . . . iii Page 10 10 11 12 14 15 l6 l6 16 17 17 18 Electrophoretic Studies. . . . . . . . . . . . . . . RESULTS . O O O O O O O O O O O O O O O O O O O O O O O C 0 Survey of Plant Tissues for Glycolate Oxidase, Glycolate Dehydrogenase and Lactate Dehydrogenase. . . . . . . . . . . . . . . . . . Enzymes of Alpha-Hydroxyacid Oxidation from Watercress Roots . . . . . . . . . . . . . . . . . Some Properties of the Alpha-Hydroxyacid Oxidase Activity from Watercress Roots Separation of Hydroxypyruvate Reductase and L-Lactate Dehydrogenase from Pea Roots . . . . . . Molecular Weight Estimations . . . . . . . . . . . . pH Optima of Hydroxypyruvate Reductase and Lactate Dehydrogenase from Pea Roots . . . . . . . . . . . Substrate Specificities of L-Lactate Dehydrogenase and Hydroxypyruvate Reductase from Pea Roots . . . Inhibition of the Root Lactate Dehydrogenase and Hydroxypyruvate Reductase. . . . . . . . . . . . . Thermal Stability. . . . . . . . . . . . . . . . . . Kinetic Properties of the Root Lactate Dehydrogenase and Hydroxypyruvate Reductase. . . . . . . . . . . Subcellular Localization of Hydroxypyruvate Reductase in Pea Roots . . . . . . . . . . . . . . Polyacrylamide Gel Disc Electrophoresis. . . . . . . DISCUSSION. 0 O O O O O O I O O O O O O O I O O O O O O . . Alpha-Hydroxyacids Oxidase in Roots of Higher Plants Root Lactate Dehydrogenase . . . . . . . . . . . . . Root Hydroxypyruvate Reductase . . . . . . . . . . . Root Microbodies . . . . . . . . . . . . . . . . . . LIST OF REFERENCES 0 O O 0 O O O O O O O O O O O O O O O O 0 iv Page 19 21 21 27 29 33 39 39 44 50 50 54 54 6O 61 61 63 65 65 7O Table II III IV VI VII VIII IX XI XII XIII LIST OF TABLES A survey of glycolate oxidizing enzymes in crude extracts of leaves of higher plants. . . . . . . . . . Oxidation of glycolate and lactate by extracts of wheat and watercress roots . . . . . . . . . . . . . . Distribution of hydroxypyruvate reductase and lactate dehydrogenase in extracts of tissues of higher plants. Distribution of glycolate oxidase, lactate dehydro— genase and hydroxypyruvate reductase in watercress roots following differential centrifugation. . . . . . Substrate specificity of crude alpha-hydroxyacid oxidase from watercress roots. . . . . . . . . . . . . Partial purification of lactate dehydrogenase from pea roots I O O O O O O O O O O O O O O C I C O C O O 0 Partial purification of hydroxypyruvate reductase from pea roots . . . . . . . . . . . . . . . . . . . . Substrate and coenzyme specificity of hydroxypyruvate reductase and lactate dehydrogenase from pea roots . . Inhibitors of root lactate dehydrogenase and hydroxy- pyruvate reductase . . . . . . . . . . . . . . . . . . Michaelis constants for the root lactate dehydro- genase and hydroxypyruvate reductase . . . . . . . . . Distribution of hydroxypyruvate reductase in sub- cellular fractions of pea root homogenates . . . . . . Comparison of lactate dehydrogenase from pea roots, heart (human) and soybean cotyledon. . . . . . . . . . Comparison of hydroxypyruvate reductase from pea roots, spinach leaves and beef liver . . . . . . . . . Page 23 26 28 3O 34 35 36 49 51 55 57 66 67 Figure LIST OF FIGURES Page Relative stability of glycolate oxidase, lactate dehydrogenase and hydroxypyruvate reductase in an extract of watercress roots. . . . . . . . . . . . . . . 32 Separation of lactate dehydrogenase and hydroxy— pyruvate reductase from homogenates of pea roots by DEAE-cellulose chromatography . . . . . . . . . . . . 38 Elution profiles of lactate dehydrogenase and hydroxy- pyruvate reductase from Sephadex G-200 column. . . . . . 41 Molecular weights of lactate dehydrogenase and hydroxypyruvate reductase from pea roots . . . . . . . . 43 pH Curves for the reduction of pyruvate, hydroxy- pyruvate and glyoxylate by lactate dehydrogenase and hydroxypyruvate reductase from pea roots . . . . . . 46 Effect of pH upon the oxidation of L—lactate by lactate dehydrogenase and the oxidation of DL- glycerate by hydroxypyruvate reductase from pea roots. . 48 Heat stability of L-lactate dehydrogenase and hydroxypyruvate reductase from pea roots . . . . . . . . 53 Distribution of microbody and mitochondrial marker enzymes and hydroxypyruvate reductase from pea roots in a continuous linear sucrose density gradient. . . . . 59 vi DCPIP EDTA FMN NAD(P) NAD(P)H PCMB Tricine LIST OF ABBREVIATIONS 2,6 Dichlorophenol-indophenol, Sodium Salt Ethylenediamine Tetra-acetic Acid, Disodium Salt Flavin Mononucleotide (Riboflavin Phosphate), Sodium Salt B-Nicotinamide Adenine Dinucleotide (Phosphate), Disodium Salt B-Nicotinamide Adenine Dinucleotide (Phosphate), Reduced Form, Disodium Salt p-Chloromercuribenzoic Acid, Sodium Salt N—tris-(Hydroxymethyl)-methyl Glycine vii INTRODUCTION Interest in the metabolism of the alpha-hydroxy acids, such as glycolic acid and lactic acid, in both plants and animals is ages old and a great number of reports have appeared concerning these problems. Whereas many reports have been concerned with glycolate metabolism in photosynthetic tissues of plants, and with lactate metabolism in animals, very little attention has been devoted to the study of either acid in the nonphotosynthetic tissues of the green plants and in par- ticular to the roots. Since virtually nothing is known about enzymes which oxidize alpha-hydroxyacids in the roots, the goal of the research was to survey for them and then to elucidate some of their properties. These exploratory experiments have dealt only with a comparison between properties of the enzymes from roots with those found in leaves or in animals. Three enzymes have been studied: glycolate oxidase, lactate dehydrogenase and hydroxypyruvate reductase or glycerate dehydrogenase. A fourth enzyme, a-glycerophosphate dehydrogenase, has also been investigated in roots and in the leaves, although it is not an alpha- hydroxyacid oxidizing enzyme. As part of my research program, this latter enzyme was characterized from leaves of soybean and spinach and these results are being published as a separate manuscript (12). LITERATURE REVIEW Enzymes of Glycolate Oxidation in Photosynthetic Tissues of Plants, Algae and in Animals and Other Nonphotosynthetic Organisms Glycolate oxidase has been extensively studied in green leaves and, recently, glycolate dehydrogenase has been shown to be a similar but quite distinct enzyme in the algae. Glycolate oxidase in the leaves (103) and glycolate dehydrogenase in the algae (76,105) have been extensively reviewed, along with other enzymes related to glycolate metabolism. Both enzymes seem important because of their role in photo- respiration, in which half of the carbon fixed in photosynthesis is lost during glycolate biosynthesis and metabolism (104). Glycolate oxidase, the first enzyme of the glycolate pathway in the peroxisomes of the leaves, was first described by Claggett, Tolbert and Burris (14) and is present in all green leaf tissues. The enzyme is a. flavoprotein, (49,126), with a molecular weight of 270,000 (29). It is not coupled to oxidative phosphorylation (124). The activity of the enzyme in etiolated leaves is markedly increased upon exposure of the tissue to the light (59,107), but its development does not parallel chloroplast development (27,107). The enzyme also oxidizes at a slower rate other short chain L(+)-alpha-hydroxymonocarboxylic acids to the corresponding keto acids (29), and it is therefore an alpha—hydroxyacids oxidase. 3 In contrast to glycolate oxidase, the enzyme that oxidizes glycolate in most unicellular green algae does not couple to oxygen as electron acceptor (15,81). In fact, the algal enzyme is not an oxidase but a dehydrogenase which can use DCPIP as electron acceptor but the natural electron acceptor is unknown (81). Glycolate dehydrogenase has been found to be of general occurrence in algae (76,105). It was in an unidentified fraction, but there are reports recently that it is in the mitochondrial (16,94) and the microbody (33) fractions. The failure to utilize oxygen as the electron acceptor during glycolate oxidation and the difference in intracellular loca- tion are not the only differences between the algal enzyme and the oxidase in higher plants. Glycolate oxidase will oxidize lactate as an alternative substrate at 1/3 the rate of glycolate but it has absolute stereospecificity for L(+)-lactate (14,126). The enzyme is insensitive to cyanide or sulfhydryl inhibitors and it is activated by the addition of FMN (29). In contrast, the algal glycolate dehy— drogenase oxidizes D(—)-lactate as an alternative substrate; it utilizes L(+)-lactate only a little or not at all; it is inhibited by cyanide and sulfhydryl binding reagents; and it is not affected by FMN (35,81). There may also be another enzyme for glycolate metabolism in marine diatoms (84), which might be distinct from either the leaf oxidase or the algal dehydrogenase. The diatom enzyme oxidizes L(+)- lactate faster than glycolate and D(-)-lactate. It is in the mito- chondria (85) and its oxidation of L(+)-lactate or glycolate is coupled to oxygen uptake, and it is therefore an oxidase. The coupling to oxygen is, however, inhibited by cyanide and other electron transport 4 inhibitors. An enzyme of similar nature may also exist in roots of rice seedlings and in developing rice seeds (108). The enzyme in rice roots also oxidizes L(+)-lactate faster than glycolate and D(-)-lactate and the oxidation is coupled to oxygen, but its sensitivity to electron transport inhibitors has not been tested. Enzymes of glycolate oxidation also exist in nonphotosynthetic tissues. Enzymes for catalyzing the oxidation of glycolate are present in fungi (2,28,97), bacteria (56,68), and mammalian liver and kidney (60,74,114). Glycolate oxidase from mammalian liver and kidney is similar to that in the leaf peroxisomes in intracellular localiza- tion, prosthetic group and catalytic properties (60,74). In the non- photosynthetic tissues of the plant, i.e., the roots, glycolic acid oxidase originally was reported absent or found in very low levels of activity (52,106), and it was presumed not to exist. However, glycolic acid and glyoxylic acids were found in the roots and glyoxylic acid was suggested to be the hydrolysis product from allantoin and allantoic acids (58), which constituted more than half of the total nitrogen in some roots (78,79). Mothes and Wagner have extensively reviewed the work done on the root glycolate oxidase (80), and concluded that the activity of the enzyme was a function of state of development. Thus etiolated leaf and roots, when allowed to turn green in light had the active enzyme, while roots kept in the dark did not have the enzyme. The formation of the enzyme in the roots in light was not due to sub- strate activation or induction because adaptive formation was not observed when glycolic acid was given to the roots even if light was present, when light was not applied too long. They gave examples of roots not capable of greening and therefore devoid of the enzyme, but 5 they also cited examples of roots that turned very green in the light but still had no enzyme activity. It is therefore suggested that not only the stage of development but also the intrinsic genetic capacity to produce the enzyme was involved. Mitsui et a1. (77) reported the existence of glycolate oxidase in roots of paddy field rice plants, upland rice plants and barn millet: the enzyme activity was low at the beginning of germination, but gradually increased after germination and then decreased again. This has been confirmed by Tolbert et al. (108), who also studied the enzyme in greater detail and suggested that the enzyme was a lactate oxidase rather than a glycolate oxidase on the basis that it oxidized L(+)-1actate faster than glycolate. Glycolate oxidase has also been reported in the microbody fractions of the roots of wheat (19), carrots, castor bean, potato tubers and castor bean endosperm (44). The enzyme from the castor bean endosperm has been studied and found to be somewhat different from the one in the leaves, being inhibited by cyanide (98). The property of glycolate oxidase in other nonphotosynthetic tissue has not been examined except for the rice roots. Since Tolbert et al. (108) found that the proper— ties of the oxidase in rice roots were different from that in the leaf, the enzyme from the roots of some other plants was examined in this thesis. Enzyme of Lactate Oxidation in Higher Plants In sharp contrast to the extensive work on NAD-dependent L(+)‘ lactate dehydrogenase in animals, reports about the purified enzyme from plant sources have been almost nil. This is because alcohol is presumed to be the traditional end product of anaerobic respiration 6 in plants, while lactate is the product of anaerobic respiration in animal tissues. Thus alcohol dehydrogenase has been found to be widely distributed in plants and has been isolated, purified and studied (18,23,66,83). However, it has been known for a long time that both alcohol dehydrogenase and lactate dehydrogenase played a significant role in the anaerobic period of germination in which pyruvate was metabolized either to ethanol (11,64,70) or to lactate (4,6,17,93). Leblova et a1. (65) found that lactate dehydrogenase was present in the seeds; its activity decreased during the first 12 hours of germination and then rose during the later stage of germina- tion; the change in enzyme activity and lactate concentration was reciprocal. Others have found that detached rhododendron leaves and buckwheat seedlings (7,25) accumulated lactic acid during anoxia and that it rapidly disappeared upon returning the tissue to air. In attempts to explain these phenomena, lactate dehydrogenase has been isolated and partially purified. The enzyme was isolated by King from soybean cotyledon (50), and was found to be similar to the lactate dehydrogenase of muscle in mammals in many of its properties. Davies et a1. (21) purified the enzyme from potato tuber and found it to be regulated by ATP at low pH but not at high pH. Isoenzymes of lactate dehydrogenase in meristem tissues of roots have also been reported (37), but this was not confirmed (50). Evidence that lactate dehydrogenase is widely distributed in plants seems numerous. It was detected in a particulate fraction from rhizomes of Equesitum (3), in extracts from potato tubers, squash fruit and turnip roots (95). The nature of these activities has not been studied. NAD—Dependent D(—)-1actate dehydrogenase, which has 7 been found in bacteria (22,99), in slide mold (30), in fungi (31,87), in algae (36,117), and in certain invertebrates (67,92), has not been reported in higher plants, nor has the NAB-independent D(-)-lactate dehydrogenase which was found in algae (33,69,117), bacterial membrane (34,46,48) and even in mammals (10,112,113). Lactate dehydrogenase from roots has not been isolated or studied, although it has been reported to be active in the meristem zone of the root tips (51). Hydroxypyruvate Reductase in Plants and in Animals Stafford et a1. (95) were the first to report on this enzyme from parsley leaves by an assay involving the reduction of hydroxypyruvate to glycerate, as a NAD-linked D-glycerate dehydrogenase. At about the same time, Zelitch reported on a NADH glyoxylic reductase in spinach leaves (122) which he later crystallized from tobacco leaves (123). Later work of Holzer and Holldorf (42) and Landahn (61) indicated that NAD glycerate dehydrogenase and NADH glyoxylate reductase were the same enzyme. Further work on the enzyme by Tolbert and his associates showed that in leaves it was located in the peroxisome (111), where it functioned chiefly as hydroxypyruvate reductase in the glycolate pathway to provide one of the sources of glycerate for gluconeogenesis and synthesis of sucrose (40,45,88). A similar enzyme has been reported in livers of animals (120) and in bacteria (13,53,62). Roseblum et a1. (89) and Sugimoto et a1. (96) have purified the enzyme from beef liver, and Kohn and his associates (54,55) carried out extensive studies on the properties of hydroxypyruvate reductase from spinach leaves and indicated that these enzymes were very similar. 8 The enzymes from different sources, although differing in molecular weights, are composed of two equal subunits, and use either hydroxy- pyruvate or glyoxylate as substrate but have a greater actiVity with hydroxypyruvate. In addition, the activities of these enzymes are similarly modified by anions and are sensitive to sulfhydryl binding reagents. The enzymes from beef liver and beef spinal cord are also similar and inhibited by nucleotides (ATP) and other glycolytic inter- mediates (91). In animals as well as in plants, hydroxypyruvate reductase is involved in serine metabolism (8,86,115). In the leaves, this occurs whenever there is a low level of glycine formation from glycolate (40). Hydroxypyruvate reductase is found in the cytosol of liver (74), but it exists in the peroxisomes of the leaves (111) and in a microbody- 1ike (16) or mitochondria-like (94) fraction of the algae. The enzyme has not been studied in the nonphotosynthetic tissues of the plant, although it was reported to be in the roots (95). Its subcellular location in the roots has not been reported. MATERIALS AND METHODS Plant Materials Seeds of sorghum (Sorghum bicolor L. var. Texas 610), corn (Zea mays L.), peas (Pisum sativum L. var. Adelman), broad bean (Vicia faba L.) were sterilized with 0.5% sodium hypochlorite solution for 20 minutes, rinsed with distilled water several times and then soaked in distilled water for approximately 8 hours; they were germinated in moist sterile vermiculite in wooden flats at 30° and grown in the greenhouse for 7-10 days. Seeds of wheat (Triticum vulgare L. var. Eva), barley (Horedium valgare L. var. Liberty), and oat (Avena sativa cv. Victory) were sterilized in a similar manner and then germinated in the dark in moist filter papers and grown in the greenhouse in half strength Hagland's solution (41) for 10 days at 30°. Watercress (Nasturtium officinale R. Br.) from the local market was grown in full strength Hoagland's solution in the greenhouse at 30° for 8-10 days. During these periods, the Hoagland's solution was changed once every two days and the roots were washed with tap water twice. With this treatment, algal or microbial growth was kept at minimum and no significant growth of any microorganism was observed during these short growth periods. 10 Chemicals DEAE-Cellulose (Whatman DE-52, microgranular) was obtained from Whatman and Sephadex G-200 from Pharmacia. TEMED (N,N,N',N' tetra methylenediamine), MBA (N,N'-methylenebisacrylamide) and acrylamide were from Canalco. Glycolic acid and sucrose were the products of Mann Research Laboratory. All enzymes and chemicals were from Sigma Chemical Company, J. T. Baker Chemical Company, or Mallinckrodt Chemicals. Chemicals used were of reagent or analytical grade and were used without further purification. Preparation of Cell Free Extracts of Plant Tissues The plant materials were rinsed several times with distilled water, blotted dry and separated into leaves and roots with razor blades or scissors. Each type of tissue was weighed to the nearest gram, minced in cold grinding medium and homogenized. The grinding medium and homogenizing procedures varied according to the type of plant material, but generally, two volumes of grinding buffer was used for each gram of tissue and the grinding medium used was 0.05 M potassium phosphate buffer at pH 7.5 with 2% (w/v) polyvinylpoly— pyrrolidone added during the grinding. The plant tissues were ground in a chilled mortar with a pestle for several minutes with or without sand. The preparations were passed through six layers of cheesecloth and centrifuged at 500 x g for 10 minutes. The supernatants were assayed for enzymes or as a starting point for subsequent enzyme iso- lation. All work was carried out at 0-5°. 11 Preparation of Particulate Fractions For localization of enzymes in particulate fractions, either method A or method B was used. In method A, the plant tissue was minced with scissors in l-2 volumes (w/v) of grinding medium which consisted of 0.4 M sucrose and 20 mM glycylglycine. The slurry was then ground gently with a mortar and pestle and passed through 4 layers of cheesecloth. After centrifuging at 270 x g for 10 min to remove whole cells and debris, the homogenate was decanted and centrifuged for 25 min at 37,000 x g. This produced a supernatant and a pellet, which was gently resuspended in grinding medium. The pellet, which contained the microbodies and mitochondria among other organelles, was the crude particulate fraction for enzyme assays. In all cases, catalase and cytochrome c oxidase was assayed to check for the breakage of the particles. Method B was used only for the differential centrifugation of pea root homogenates. The tissue was homogenized in a chilled mortar and pestle with one volume by weight of grinding medium (0.5 M sucrose and 0.02 M glycylglycine at pH 7.5). The resulting slurry was squeezed through 8 layers of cheesecloth, and the pH (approximately 7) was readjusted to 7.5 with potassium hydroxide. The sap was then centri- fuged at 270 x g for 20 min, and the resulting pellet was resuspended in the grinding medium. The sap was further centrifuged at 6000 x g for 20 minutes. This pellet was also resuspended in grinding buffer. A "mitochondrial" fraction was prepared by centrifugation of the sap at 37,000 x g for 20 min and resuspension of the pellet in the grinding medium. The supernatant fluid after the last centrifugation was designated the "supernatant" fraction. In each case, catalase and 12 cytochrome c oxidase were assayed for the localization of the par- ticular organelles in each fraction. Partial Purification of L(+) Lactate Dehydrogenase and Hydroxypyruvate Reductase from Pea Roots Peas (Pisum sativum L. var. Adelman) were soaked for 4 to 5 hours in distilled water and germinated in moist sterile vermiculite for 7 days in the dark at 30°. The pea seedlings were then washed ten times with tap water to free the roots from vermiculite and then rinsed twice with distilled water. The roots were separated from the plants with scissors and then cut into small pieces with a sharp razor blade. The minced roots were ground in a Waring Blendor at top speed for 60 seconds with equal parts (w/v) of a buffer containing 0.05 M potassium phosphate at pH 7.5, 1 mM mercaptoethanol and 2% polyvinylpolypyrroli- done. The brei was passed through 6 layers of cheesecloth and centri- fuged at 37,000 x g for 20 min. The supernatant was referred to as the "crude extract." L(+)-lactate dehydrogenase and hydroxypyruvate reductase were partially purified from this crude extract by the following steps. The results of each stage of purification procedures are summarized in Table VI for lactate dehydrogenase and in Table VII for hydroxypyruvate reductase (glycerate dehydrogenase). (a) Ammonium sulfate fractionation Cold saturated (NH4)280 solution at pH 7.0 was added to the 4 crude extract to make the final solution 30% saturated with ammonium sulfate. The proteins in the solution were allowed to precipitate for 30 minutes in the cold room (4°). The precipitate was removed by centrifugation at 10,500 x g for 30 minutes and discarded. The 13 supernatant was made 60% saturated with respect to ammonium sulfate by adding cold saturated ammonium sulfate. The protein, precipitated between 30-60% ammonium sulfate solution, contained more than 95% of the total activities of both L(+)-lactate dehydrogenase and hydroxy- pyruvate reductase. The precipitate was collected by centrifugation and resuspended in a small volume of 50 mM potassium phosphate buffer at pH 7.0 and 1 mM mercaptoethanol. The solution was desalted in a Sephadex G-25 column (2.5 cm x 35 cm) before being applied to a DEAE- cellulose column. (b) DEAR-cellulose treatment Approximately 450 mg of the desalted protein was added to a 2.5 cm x 30 cm DEAE-cellulose (Whatman, DE-52) column previously equilibrated with several bed volumes of 5 mM potassium phosphate at pH 7.0 and 1 mM mercaptoethanol. The column was then washed with 1 bed volume of the equilibrating buffer. The proteins were then eluted at a flow rate of 0.5 ml per minute with 3 bed volumes of a linear gradient of 0 to 0.5 M potassium chloride in 5 mM potassium phosphate at pH 7.0 and 1 mM mercaptoethanol. Six-milliliter fractions were collected with a Gilson automatic fraction collector. The chloride content of the fractions was determined with a Barnstead Purity meter. By this procedure, the L(+)-lactate dehydrogenase and hydroxy- pyruvate reductase were eluted separately from the column (Figure 2), with the hydroxypyruvate reductase eluting first. The fractions which showed an increase in specific activity over the previous stage were combined. The pooled proteins were concentrated by adding saturated ammonium sulfate solution until 70% saturated. These precipitated 14 proteins could be stored in the pellet form in the freezer at -4° for several weeks without loss of enzyme activity. (c) Gel filtration chromatography in Sephadex G-200 The concentrated protein with lactate dehydrogenase activity from the previous step was dissolved in 4 ml of 50 mM potassium phosphate at pH 7.0 and 1 mM mercaptoethanol and applied to a Sephadex G-200 column (1.6 cm x 90 cm) previously equilibrated with the same buffer. Proteins were eluted with 300 ml (2 bed volumes) of the same buffer at a flow rate of 4 ml per hour. Fractions of 3.3 ml were collected and those with lactate dehydrogenase activities were pooled and concentrated by 67% (NH SO4 precipitation. Hydroxypyruvate 4)2 reductase was partially purified in the same manner except that the buffer used was at pH 6.5. Both proteins could be kept as ammonium sulfate suspension or as a frozen pellet at -20° for a long period. The lactate dehydrogenase seemed to be more labile in solution than hydroxypyruvate reductase. In potassium phosphate buffer at pH 7.0 with 10% glycerol, lactate dehydrogenase lost 85% of its activity after 2 days at 0-4°. Hydroxypyruvate reductase, on the other hand, could be kept in 10% glycerol in 50 mM phosphate buffer with 1 mM mercaptoethanol over a period of one week at 0-4° without significant loss in activity. Sucrose Density Gradient Centrifugation The isolation method for microbodies followed that described by Huang and Beevers (44) with little modification. The grinding medium contained 0.4 M sucrose, 1 mM EDTA, 10 mM KCl (potassium chloride), 1 mM MgCl (magnesium chloride), 10 mM dithiothreitol, 0.15 M Tricine 2 15 buffer adjusted to pH 7.5 and 0.5% by weight of polyvinylpyrrolidone added during the grinding. The pea roots were ground gently in medium with mortar' and pestle after being first chopped to small pieces with razor blades. The homogenate was passed through 8 layers of cheese— cloth and centrifuged at 270 x g for 10 min. Ten milliliters of the supernatant was layered on a 40-ml linear gradient composed of 30-60% sucrose over a cushion of 5 ml 60% sucrose. All sucrose solutions were expressed as percentage by weight and made up in 0.1 M Tricine buffer at pH 7.5. After centrifugation for 4 hours at 21,000 rpm in a Beckman L2-6SB ultracentrifuge with Spinco rotor SW 25.2, the gradients were collected in 1.5-ml fractions using a density gradient fractionator. All steps were carried out at 4°. The fractions were assayed for catalase, cytochrome c oxidase and hydroxypyruvate reductase. The density of the fraction was determined in the Bausch and Lomb refractometer. Biochemical Assays All assays were run at 25° and initial rates were recorded. The spectrophotometric assays were done on a Gilford 24008 recording spectrophotometer. In all cases, the enzyme assays were tested to check if activity was dependent on protein concentration over the range used. Oxygen uptake was measured with a Rank Brothers oxygen electrode. Protein was determined by the method of Lowry et a1. (71) using bovine serum albumin as standard. All enzyme activities were expressed as nmoles (substrates converted) per minute. 16 NAB-Lactate Dehydrogenase This enzyme was assayed by following changes in absorbance at 340 nm (26). For the determination of pyruvate reduction, the 1—ml reaction mixture contained 0.1 M potassium phosphate buffer at pH 7.0, 0.01% Triton X—100, 0.12 mM NADH, appropriate amount of enzyme extract and 10 mM potassium pyruvate. For measurement of the reverse reaction, that involved the oxidation of either D- or L—lactate, the l-ml mixture contained 0.1 M NaOH-glycine buffer at pH 9.2, 0.01% Triton x-100, 3 mM NAD, enzyme and 20 mM D- or L-lactate (lithium salt). In both cases the endogenous rate was monitored for 2 to 3 minutes and the reaction was initiated by the addition of substrates. Since pyruvate reduction lies far to the right, it was used routinely for detection of lactate dehydrogenase. NAD D-Glycerate Dehydrogenase or Hydroxypyruvate Reductase This enzyme was assayed the same as the lactate dehydrogenase described above except that the substrate was replaced with equimolar of Li—hydroxypyruvate or DL-glycerate (calcium salt) and the pH of the buffers were changed to 6.2 or 9.2, respectively. Glycolic Acid Oxidase or Dehydrogenase This enzyme was assayed by following the anaerobic reduction of 2,6 dichlorophenol-indolephenol (DCPIP) at 600 nm (109). The reaction mixture of 2.5 ml contained 0.08 M sodium pyro— phosphate at pH 8.5, 0.12 mM DCPIP, 0.01% Triton X-100, 0.1 mM FMN and either 8 mM glycolic acid (neutralized with NaOH) or 20 mM D— or L-lactate (lithium salt). The reaction was initiated by addition of l7 substrate after reading the endogenous rate for 5 minutes. To test for sensitivity of the enzyme to O or cyanide, assays were run 2 aerobically or in the presence of 2 mM potassium cyanide, respectively. Changes in 0.0. were converted to nmoles of dye using the extinction coefficient of 21.9 cmZ/umoles for DCPIP (1). The oxidase was also assayed by measuring the disappearance of 02 with a Rank oxygen electrode at 10 mvolts. The reaction mixture of 2.5 ml contained the complete mixture of the DCPIP assay minus the dye (DCPIP). Oxygen concentration was calculated from its solubility from air into water. All solutions for the assay were air saturated except the enzyme. At 25°, the reaction mixture held 685 nmoles oxygen (38). Percentage changes in oxygen concentration recorded in 5 minutes were converted to nmoles oxygen/min. Catalase This enzyme was used as the peroxisome marker enzyme and was assayed spectrophotometrically at 25° by following the disappearance of H202 at 240 nm (72). One to one hundred microliters of extract was added to a 3-ml mixture containing 1.25 x 10’2 M H202 and 1/15 M sodium phosphate at pH 7.0. The disappearance of H202 was calculated by using the extinction coefficient of 0.036 cmz/umole. Cytochrome c Oxidase Cytochrome c oxidase, the mitochondrial marker, was assayed by following the oxidation of reduced cytochrome c at 550 nm (110). The enzyme was placed in the bottom of the l-ml quartz cuvette and 20 ul of 1% digitonin was added. After mixing and a l min incubation period, 0.9 ml of 0.05 M potassium phosphate buffer at pH 7.0 was added. The l8 reaction was then initiated by the addition of 100 ul of solution containing 10 mg/ml cytochrome c reduced with dithionite. A sample without the enzyme was used in order to measure the rate of nonenzymatic oxidation of reduced cytochrome c. The extinction coefficient of 21.0 cmZ/umole for cytochrome c was used to calculate the amount of cytochrome c oxidized (73). Estimation of Molecular Weight by Gel Filtration Chromatography Gel permeation chromatography was used to estimate the molecular weightsHw ucmHm mowcmNmTZE N msHm TNua>Huo¢ m>wumem «mucmHm Hmnmfln m0 mw>mwa mo muuwuuxw wpsuo ca mwfixucw mcfluflvflxo mumaoomam mo >m>usm m .H wanna .mucmam v0 a; .wpmaoowam £uw3 >uH>Huom ou m>flumHmu >ua>fiuom mo mucsoEm ucmoHMHcmHmcH no on mmumoflpcfl :0: .wocwmnm uflmnu ca mmfiufl>fluom on w>flumawu mum mmsHm> opp .No Ho mpflcmxo mo mocwmoum wzu CH m>mmmm mom .mmmmm mHmOQ may CH mHmOQ use wumHoowam £ua3 wmmmm msu mo mammn on» so new ammwm mpouuomao mo map CH No new oumaoomam Lufl3 Somme mnu mo mammn may no w mm Ummmwumxo mum moHuH>Huom m>apmHmm i 24 O OH HO O OO OOH Ha OO OOH names O O O O OO OOH O Om OOH mmmuoumumz O mm O O OHN OOH m OOH OOH «.essmuom O mm me O OOH OOH N OR OOH umo O mm om O NO OOH a OO OOH mama O O O O OOH OHH Om NO OOH .Tcuoo O O O O OO OHH OH OO OOH smaumm mumma OHmoo mumuomalo mumuomalq mumHoo>H0 mumuomalo mumuomHIA mDMHOOSHU mumuomalo mumuomqu mpmHoowaw unmam No no LAO mafia mOflcmxo :5 m mafia mufl>fiuom O>Humflmm Iomscflucoov H magma 25 in the C4 corn plant should be reexamined in view of the possibility suggested above that the alpha-hydroxyacid oxidase in C plants may 4 be modified. A final concentration of 2 mM cyanide stimulated enzymatic activity in all cases by both the DCPIP and the O electrode assays. 2 This has been observed since the discovery of glycolate oxidase and attributed to removal of the aldehyde or keto acid product which normally severely inhibits the utilization of the hydroxyacids. In the roots of the same plants, the situation was entirely different. Except in the roots of two plants (wheat and watercress), glycolate oxidation was not detected by the same assay procedures. Even in the roots of watercress and wheat the levels of enzyme activities were very low, at least 50 times lower than the level found in the leaves (Table II). The alpha—hydroxyacid oxidase system in the crude extracts of roots of wheat and watercress oxidized L-lactate faster than glycolate, and also oxidized D—lactate to some extent. In these two plants at least, the system in the roots seemed to be modified from that in the leaves. Whereas the activity in the roots of watercress was stimulated by 2 mM potassium cyanide and inhibited by O in the DCPIP assay, very much similar to the system 2 in the leaves, the activity in the roots of wheat was inhibited by 2 mM cyanide and also by 02 in a similar assay. Since the alpha- hydroxyacid oxidase activities in the crude extracts were very low, they had been concentrated from both root sources to confirm these observations by differential centrifugation. To look for other enzymes of alpha-hydroxyacid oxidation, NAD— 1actate dehydrogenase was assayed by using pyruvate as the substrate, 26 .xmmmm mumHoowflm UHQOHmmcm wnu cfi umnu mo w mm pmmwoumxw mum mwfiufl>fluo¢ .cfimuoum mE\CHE\mHmOD wmaoec m.o mmz uomuuxm umwn3 now new Camuoum mE\:HE\mHmOQ mmaosc m.a mMB uomuuxo mmmuoumumx Low mufl>wuom camaommm .pocuws mHmOQ UHQoummcm on» an ©w>8mm¢ x. o o o 0 me mm mm boa ooa ummnz o o o o mmH HHH Om 50H OOH mmwuoumuwz mumuomHna mumuomqu mumaooxaw wumuomHlo wumuomHuq mumaooxaw mumuomalo oumuomalq oumHoowao ucmam Amov Mad moan mpflcmmo SE m msHm Nufl>fluo¢ m>wumem amuoou mmwuoumum3 tam umwn: mo muomuuxw SQ mumuomH can mumHoomam mo coflumvflxo .HH magma 27 and hydroxypyruvate reductase or glycerate dehydrogenase was also assayed by using hydroxypyruvate as the substrate (Table III). Hydroxypyruvate reductase was extremely active in the leaves of C 3 plants, but only about 1/10 as active in the C plants, corn and 4 sorghum, as has been found previously (109). Lactate dehydrogenase was not present in the leaves and the active peroxisomal hydroxy- pyruvate reductase does not use pyruvate as a substrate. Some lactate dehydrogenase was present in the roots of the C plants. 3 Whether hydroxypyruvate reductase is present in the roots could not be resolved by this one assay on the crude homogenate, since lactate dehydrogenase can catalyze the reduction of both pyruvate and hydroxy- pyruvate (75). In the root extracts of two plants (i.e., corn and sorghum), neither pyruvate nor hydroxypyruvate reduction was observed, possibly due to inactivation of the enzyme during the grinding procedure. Enzymes of Alpha—Hydroxyacid Oxidation from Watercress Roots Since extracts of the roots of watercress showed activities for glycolate oxidase, hydroxypyruvate reductase, and lactate dehydrogenase, it was possible that there were one to three enzymes catalyzing these reactions and further experiments were conducted to test these possibilities. It was possible to separate the NAD requiring enzyme activities from that of glycolate oxidase by dif- ferential centrifugation. By method A described in the Materials and Methods, about half of the glycolate oxidase activity, measured in terms of DCPIP reduction with either glycolate or L—lactate as sub- strates, remained particulate, while 89% of the activity for 28 Table III. Distribution of hydroxypyruvate reductase and lactate dehydrogenase in extracts of tissues of higher plants Hydroxypyruvate Lactate Plant Tissue reductase dehydrogenase Broad bean Leaves 280 0 Roots 26 13 Corn Leaves 57 0 Roots 0 0 Peas Leaves 297 0 Roots ll 3 Sorghum Leaves 35 0 Roots 0 0 Watercress Leaves 136 0 Roots 104 35 Wheat Leaves 550 0 Roots 25 15 29 hydroxypyruvate reduction and 86% of the activity for pyruvate reduc- tion were in the supernatant (Table IV). ‘The results indicated that lactate was being oxidized by at least two different enzymes, an alpha-hydroxyacid oxidase and a NAD-dehydrogenase. Ammonium sulfate fractionation also indicated that there were different enzymes catalyzing these reactions. Most of the glycolate oxidase was lost during the fractionation, probably because of its particulate nature, but 16% of the hydroxypyruvate reduction and 50% of the pyruvate reduction were recovered in the 40% (NH SO4 pellet 4)2 of the root homogenate. It was also possible to distinguish the glycolate oxidase from the NAD-linked dehydrogenases by their rela- tive stabilities in root homogenates prepared by grinding in 50 mM potassium phosphate buffer at pH 7.5. In the homogenate, after centrifugation at 500 x g for 10 minutes to remove cell debris, glycolate oxidase activity, measured with L-lactate as substrate, remained stable over a period of at least several hours, while that of NADH pyruvate reductase and hydroxypyruvate reductase declined slowly (Figure 1). This experiment was performed with watercress roots that had been frozen for two months so this stability pattern for the enzymes may not be exactly the same with fresh tissues. Some Properties of the Alpha-Hydroxyacid Oxidase Activity from Watercress Roots The alpha-hydroxyacid oxidase activity from watercress roots differed from that of glycolate oxidase of the leaves in preferen- tially utilizing L-lactate, and in this respect was similar to that in the rice roots (108). The particulate fraction (37,000 x g pellet) of the root homogenates was used in further experiments. 3O mm m OOH umHHOO O x OOO.OO OO Om mmm ucmumcummsm O x OO0.00 N m 00H m oov ucmumcummnm m x OON o m mmmamumo m O OO OOHHOO O x OOO.OO OOOOOO mm mm vaH ucmumcnomsm m x ooo.hm mum>su>m noutwsmp OOH Om OOOH unnumcummsm O x Ohm .moaz wumuomH m ma OmH umaamm m x OOO.Om wum>su>m mmmuoscmu mm OOH mmom usmumcuom5m m x ooo.nm Imxouomn mum>nu>m OOH HO OOHO unnumcummsm O x OON .mo4z usxouuxm mO O.~ Om HOHHOO O x OOO.OO mm O.H HO unnumcummsm O x OOO.OO OHOoo OOH O.H on ucmumcquSO m x Ohm .mumuUMHIH ommcfixo wannaxonvhn mm O.H NH umHHmm O x OOO.OO unemHa OO m.O NH ucmumcuwmsm O x OOO.OO OHOoo OOH m.O mm ucmumcummsm O x Ohm .mumHooOHO a cflwuoum mE\:wE\moHosc sws\mmaosc cofluomum mwumuumnzm moexncm coausnfluumwa muH>Huom oamaommm >uw>wuom Hmuoa cofiummsuHuucmo Hafiucmuwmmao mcw3oHHom muoou mmmnoumumz ca wmmuoswwu mue>su>mhxou0hn can wmmcwmoupmnoo mumuona .ommcon oumaoomam mo coausnfiuumwn .>H OHQMB 31 .HHVIAuv ommuoswwu mum>summ>xouwmnlmOOZ Ocm .HnTIAVV wmmuoswmu wum>5uxmumO¢z .HnYIAUV wmmvflxo mumHoomam "av um mmmuoum mafiusp mHm>HODCH ucmsvmeSO we use Houmu mEHuv mawumfiowaefl poxmmwm mmz usmumcummsm m x 00m one .mouscHE OH you O x com um GOODMHHucmo smnu use .ocovflaouuwmxaom nchH>xaom an maficflmucoo m.h mm um Hemmsn mumnmmonm Edammmuom z mO.O CH vcsoum mums muoou one .muoou mmmuoumums mo uomuuxm cm ca mmmuoswwu mum>su>m umxouvmn Ocm mmmsmmoum>£ow mumuomH .wmmwfixo mumHooxHO mo auHHHnmum m>Humamm .H musmflm 32 D, In D/d 0/ O H newsman 952.. E 25... ON _ fl. /O/26>3;9ontoaz :ch0 22°36 «OOHOOOOm a I / D 1 0/D 3389.356 233... O/O/O\ O to mmov Iowbuo )0 °/. 00. 33 The addition of L-lactate resulted in the immediate utilization of molecular oxygen or the reduction of the artificial electron acceptor, DCPIP. The rates of reactions in both cases were slow. Oxygen con- sumption and dye reduction were not inhibited by 0.5 mM cyanide, 2 ug/ml antimycin A, or 10-6 M rotenone. The reactions were not inhibited by the sulfhydryl binding reagents, 10‘5 M p-chloromercuri- benzoic acid and 10-3 M N-ethylmaleimide. Besides activity with L- lactate, glycolate and some activity with D-lactate, the fraction catalyzed the oxidation of other alpha-hydroxyacids such as DL- alpha-hydroxyisocaprate and glyoxylate (Table V). Assays in which equal amounts of glycolate and lactate were added resulted in activity almost intermediate between that observed for the individual substrates, which suggested that a single enzyme was involved. Separation of Hydroxypyruvate Reductase and L-Lactate Dehydrogenase from Pea Roots Although homogenates of the roots of pea seedlings had no alpha- hydroxyacid oxidase activity, they readily catalyzed the reduction of pyruvate, hydroxypyruvate and glyoxylate. Since peas were easy to grow, the roots of pea seedlings were chosen to further study these dehydrogenase or reductase activities. Partial purification by methods described in the Materials and Methods is shown in Table VI for the L-lactate dehydrogenase activity and in Table VII for the hydroxypyruvate reductase activity with hydroxypyruvate as substrate. Upon passage of the protein fraction from 30-60% saturated (NH 2SO 4) 4 precipitation through the DEAE-cellulose column, the enzymes were well separated into two protein fractions (Figure 2). One protein, designated hydroxypyruvate reductase since it did not utilize lactate, 34 Table V. Substrate specificity of crude alpha-hydroxyacid oxidase from watercress roots Rates of DCPIP reduction and oxygen consumption were expressed as the percentage of the rates with glycolate. Two enzyme preparations were used: one had specific activity of 1.3 nmoles DCPIP/min/mg protein,* the other 4.6 nmoles Oz/min/mg protein.** Final concentra- tion of each substrate was 8 mM. Electron Acceptors Substrates DCPIP % Oxygen % Glycolate 100* 100** L-lactate 176 129 D-lactate 27 71 Glyoxylate 91 -- DL—glycerate 0 -- DL-alpha-hydroxy- 82 -- isocaproate Glycolate (8 mM) plus 131 -- L-lactate (8 mM) 35 Table VI. Partial purification of lactate dehydrogenase from pea roots Total Specific Activity Activity Purifi- Protein nmoles NADH/ nmoles NADH/min/ cation Recovery Fraction (mg) min mg protein (fold) (%) Crude 1201 3.86 3 1 -- 30—60% 468 7.38 16 4.4 100 (NH4)ZSO4 DEAE-52 153 6.67 44 13.6 89 Sephadex 18 4.93 274 85.5 67 G-200 36 Table VII. Partial purification of hydroxypyruvate reductase from pea roots Total Specific Activity Activity Purifi— Protein nmoles NADH/ nmoles NADH/min/ cation Recovery Fraction (mg) min mg protein (fold) (%) Crude 1201 38.9 32 1 100 30-60% 468 31.4 77 2 80 (NH4)ZSO4 DEAR-52 59 11.1 189 6 28 Sephadex 8.6 10.7 1240 40 27 G-ZOO 37 .mmumuumgm mm HIV 089553 Home . H Din: mumHHonmHm . HOIOO mumafimm Ixxouown mchs .mwonuwz cam OHMHuwumz may CH OmQHHommc mm mmmuospwu wum>su>m>xoucms use mmmcwm louvmnmv mumuomH How memmmm OOHm use A O E: owm um cHououm Mom UmuouHGOE 0Hw3 mGOHuomum ucmsHmmm .OmuomHHoo mums HE O mo mcoHuowum .2 m.OIH.O Eoum mcHocmu Hollow OOHHOHLU Edemmuom mo ucmemuO HEIOOO m nqu Mao: Mom HE Om um wmude mm3 chuoum man can HocmnumoummonEum 2E H mchHmucoo ummmsn mumnmmonm EsHmmmuom SE m suH3 Owgmm3 mmz HEo Om x 60 m.m .lemav cEsHoo one .>£mmumoumEOH£o mmoHsHHOUIMOmO an muoou mwm mo mwumcwmoeon Eoum mmmuosnmu mum>su>m>xouwmc cam mmmcomouwmsmw mumuomH mo COHumHmmmm .m musmHm 40-4 -20 38 O “now n; It»! go uououuaauoo N am 093 (o aouoqmsqv 500 ' ( '-|‘” )( |L 2 50" ugu: x salowu) Panmxo HGVN Tube Number Figure 2 39 was eluted at 0—0.05 M potassium chloride while the other protein, designated L-lactate dehydrogenase because it used L-lactate as well as the other two substrates, was eluted at 0.1-0.2 M potassium chloride. Each of these proteins was further purified 40- to 100— fold by being eluted through a Sephadex G-200 column separately (Figure 3). These partially purified enzymes (Tables VI and VII), though still not homogeneous, were free from each other (as shown by gel electrophoresis) and were used for further characterization studies in all of the subsequent sections. Molecular Weight Estimations The molecular weights of the lactate dehydrogenase and hydroxy- pyruvate reductase were estimated by using Sephadex G-200 molecular exclusion chromatography. Graphical estimation of the molecular weights of root lactate dehydrogenase and hydroxypyruvate reductase was obtained from a standard curve established by plotting the logarithms of the molecular weights of a series of protein standards versus Kav as described in Materials and Methods (Figure 4). The molecular weight of lactate dehydrogenase was found to be 160,000 daltons and that of hydroxypyruvate reductase was 68,000 daltons. pH Optima of Hydroxypyruvate Reductase and Lactate Dehydrogenase from Pea Roots Variation of enzymatic activities with pH was investigated with 0.1 M phosphate/pyrophosphate buffer over a pH range from 4.5—8.5 and with 0.1 M glycine-NaOH buffer over the range from 8.6—10.6. The pH values were adjusted with either NaOH or HCl. 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H _ fi — H Ono-360 whflrodJ hoom MP<>Dm>d>XOmO>I hoom dun. 562:4 832m 053m dwa mmE. ODOmm .232 £83.30 0 2.25630 ..0 00 DO To AD» 44 electrode attached to a pH meter and the results are shown in Figure 5. Lactate dehydrogenase had a sharp pH optimum around pH 7.0 for all the substrates, pyruvate, hydroxypyruvate and glyoxylate. Hydroxy- pyruvate reductase had a broad pH optimum around 5.9. Both enzymes were the most active by far with hydroxypyruvate and both enzymes were able to reduce glyoxylate but at a slower rate. The rate of the reverse reaction at alkaline pH was also determined by the standard assays as described in the Materials and Methods, and the results are shown in Figure 6. The pH optimum of L(+)-lactate oxidation by lactic dehydrogenase was 9.6, and the optimum for glycerate oxidation by hydroxypyruvate reductase was at pH 9.1. SubstrateySpecificities of L-Lactate Dehydrogenase and Hydroxypyruvate Reductase from Pea Roots The lactate dehydrogenase from pea roots catalyzed NADH—dependent conversion of pyruvate presumably to L(+)-lactate because it oxidized L(+)-lactate at a much greater rate than the D(-)-isomer. Therefore, it is probably similar to L(+)-lactate dehydrogenase from other tissues, which also reduces hydroxypyruvate and glyoxylate. Hydroxy- pyruvate reductase, on the other hand, did not use pyruvate and was active only with hydroxypyruvate and glyoxylate. These results are summarized in Table VIII. Both enzymes were specific for NADH and could not utilize NADPH. Both enzymes were able to catalyze the reverse reaction at high pH at a much slower rate than the forward reaction. Lactate dehydrogenase was able to oxidize glyoxylate as well as to reduce it, similar to the lactate dehydrogenase found in animals (24). The stereoisomerism of the substrates that can be 45 .mmmuosvmu mum>su>m>xouchn How mcHH >>mm£ m can mmmcmmouwhsmv mpmuomH How wcHH cHau m ha Owuocwv mH HAVIAVO mum>5a>m>xoupmg muse :Ulnuv mumH>xo>Hm . A Iv mumsbumm mo coHuoHHHOmm .HE H mo mOBHo> Hmuou m cH mummmsn wumnmmonm0u>m\mumnmmonm EsHmmmuom z H.O one OONIO xmomnmmm Eoum mmsxucm OOHMHHSQ wHHMHuHmm opp mo ucsoem OHOHHQOHQQM .28 NH.O mamz .25 OH mmumuquSm "Omchucoo mmusuxHE coHumndosH .muoou mom Bonn mmmuoswmu wum>dnmm>xoucwn Ocm mmmcmmouvmnmo oumUUMH SQ mumH>xo>Hm Ocm mum>5u>m>xouO>£ .wum>su>m mo coHuoscmH wnu Mom mm>nso mm .m OHOOHO 46 (ugamd bm/quI/HQVN salomu) OSOUODOJpAqOO ammo-I o O O (0 q» N O l l l “.3 ‘ID 3 O qIf) c N “In ‘50 232 “.2. ‘2 "3 ‘CD 0 «.— g: ;£’ :3 :13 _IO Sm ‘0 'U ,5 I L I l \ m V O O O O O O o (D d‘ N (uIaIOId bwfquI/HQVN smoum) asomnpaa anAnMdAxmpAH Figure 5 47 .wHw>Hpommmmu .wmmuospmu mum>5u>m>xoucmn 0cm mmmcmmoupanmn mumuOMH Mom HnVIAvv mumumohHmIqa use H—UIAuv wumuomHIH mumz poms mmumuumndm .HE H mo mEdHo> Hmuou O OH Hmmmdp OOHomeImomz 2 H.O Ocm OONIU xmcmnmmm Eoum mosxncm OmeHusm mHHmHuHmm onu mo unsoem ouMHumoummm .25 m afiz .28 ON wumuumndm "UOCHmucoo musuxHE coHumndocH .muoou mom Eoum mmmuoswmu oum>5u>m>xonoxn 2n mumuwoaHmIHO mo coHumono map can mmmcomoucxnmw oumuomH an mumuOMHIH mo coHumcon ms» com: mm mo uommmm .O wusmHm 48 9-6 M In Ion; go 7. so osoIonpou oIoAnIAdAxoapm o 9 In 0 l l o/ / O -0 /o _ O / 2 2% :o I." >0 a: at: "ID 205 fig '2. m I O I a. o 8 0:: e— HQ 0 D 6% \ In Ol— 41: >~ f. O D\ c? l l I‘- 0 O o O In I-6 Ho '10 Ion; I0 7. so asouaboIIpAqao OIDIOM Figure 6 49 Table VIII. Substrate and coenzyme specificity of hydroxypyruvate reductase and lactate dehydrogenase from pea roots Each incubation mixture contained 4-10 ug protein (the fractions eluted from the Sephadex G-200 column in Table VII for hydroxypyruvate reductase and in Table VI for lactate dehydrogenase), 0.12 mM NADH or NADPH, 10 mM substrate, and buffer plus water in a total volume of 1 ml in a l-ml quartz cuvette with pathlength of 1 cm. The buffers used were 50 mM potassium phosphate at pH 7.0 for lactate dehydrogenase and at pH 6.2 for hydroxypyruvate reductase. The substrate-dependent reduction of NAD or NADP by the enzyme was also monitored in l-ml reaction mixtures containing 0.1 M NaOH-glycine at pH 9.5, 7.5 mM NAD or NADP, 20 mM substrate, 4-10 Hg protein, and water. Lactate Hydroxypyruvate dehydrogenase reductase Substrates nmoles/min/mg protein Pyruvate plus NADH 104 0 Pyruvate plus NADPH 0 0 Hydroxypyruvate plus NADH 243 676 Hydroxypyruvate plus NADPH 0 0 Glyoxylate plus NADH 126 250 Glyoxylate plus NADPH 0 0 Alpha-ketobutyrate plus NADH 60 0 Alpha-ketoglutarate plus NADH 10 0 L-lactate plus NAD 46 0 L-lactate plus NADP 0 0 D-lactate plus NAD 12 0 D-lactate plus NADP 0 0 Phenylpyruvate plus NADH 0 O DL-glycerate plus NAD 0 11 DL-glycerate plus NADP 0 O Glyoxylate plus NAD 10 50 utilized by hydroxypyruvate reductase has not been investigated, but all previous work on the enzyme from other sources indicated that it was specific for the D-isomers (55,95,120). Inhibition of the Root Lactate Dehydro— genase and Hydroxypyruvate Reductase p—Chloromercuribenzoate (PCMB) completely inhibited hydroxy- pyruvate reductase at 0.1 mM final concentration, but it had little effect on lactate dehydrogenase (Table IX). The approximate 20% inhibition of lactate dehydrogenase by PCMB was not prevented by addition of 5 mM dithiothreitol, cysteine, mercaptoethanol, or glutathione, which by themselves had no effect on the enzyme. Both enzymes were inhibited by 10 mM oxamate, 10 mM oxalate and 10 mM o-phenanthroline, though to a different extent. It was surprising that the lactate dehydrogenase was completely inhibited by 0- phenanthroline, since no metal requirement is known for this ++ + +++ ++ ++ + , enzyme. Cu , Zn , Mg , Fe , Mn at 1 mM did not affect the activities of either enzyme. Thermal Stability Aliquots of the partially purified enzymes were incubated at various temperatures in a water bath for 20 minutes. After centri- fugation to remove precipitated proteins, the supernatants were assayed for enzyme activities at 25°. The activity of hydroxypyruvate reductase remained constant up to 40° (Figure 7), and half the activity was lost in 20 min at a temperature of about 55°. Lactate dehydro- genase was stable for 20 minutes up to a temperature of 76°. This is a particularly high temperature stability for a plant enzyme, 51 Table IX. Inhibitors of root lactate dehydrogenase and hydroxypyruvate reductase The two enzymes were assayed by measuring the rate of oxidation of NADH in the presence of hydroxypyruvate after prior incubation at 25° for 5 minutes with the inhibitory compounds. Relative Rate of Reaction* Added compound Hydroxypyruvate reductase Lactate dehydrogenase None 100 100 PCMB 0.0125 mM 88 100 PCMB 0.05 mM 68 89 PCMB 0.1 mM 0 81 PCMB 0.25 mM 0 68 Dithiothreitol 5 mM 100 100 Dithiothreitol (5 mM) 100 81 plus PCMB (0.1 mM) Potassium cyanide 10 mM 100 100 Sodium arsenite 10 mM 94 88 Oxamate 10 mM 22 55 Oxalate 10 mM 9 82 EDTA 10 mM 105 100 o-phenanthroline 10 mM 55 0 Iodoacetamide 2 mM 89 67 Ethylmaleimide 1 mM 34 95 * 100% corresponded to 84 nmoles NADH/min/mg protein for hydroxy- pyruvate reductase and 87 nmoles NADH/min/mg protein for lactate dehydrogenase. 52 .omm um umsu HO O we Oommoumxo mum mmHuH>Huo¢ .AnVIAvv ommuosvou mum>5n>m>xoupmn How oum>sn>m>xounwn new mamz Sufi: use HIV mmmcmOoncmnoc oumuomH Mom oum>5u>m cam :92 5.33 OOHpH>Huom .Hom Omxmmmm mum3 mucmumcuwmsm onu .mCHmuonm OmumuHmHomHm on» m>oEmH ou COHHMO:MHHucmo Mmumd .mouscHE On How sump Hopes m CH mmHSOOHomEou msoHHm> um OoquSUCH oumz moESNcm OOHMHHDQ mHHmHuHom 059 .muoou mom ECHO ommuuscou mum>nu2®>xowv>c 0cm ommsomouvhaop oumuomHIH mo muHHHQmum ummm .O ousmHm 53 OOHOHO cchaaocTota *0 053.360... cm 00 ow com 0 OiflH O - q I o omopoavom 23333203.. 32.305»: OO 2°83 C) C) «I (3 ¢ C) (D C) a) 00. .92 amIDJadwai Io Ioul I0 °/. so KIIAIIov 54 which are generally inactivated above 35-45°. In this respect the lactate dehydrogenase from roots is similar to the enzyme in cotyledons which also have a high thermal stability (50). Kinetic Properties of the Root Lactate Dehy— drogenase and Hydroxypyruvate Reductase Effect of substrate concentration on the initial rate was determined at pH 7.0 for lactate dehydrogenase and at pH 6.2 for hydroxypyruvate reductase in the presence of a constant amount of enzyme. Both enzymes showed normal Michaelis-Menten kinetics and the Michaelis constants for the substrates and coenzyme are shown in Table X. Both enzymes were inhibited to some extent by high con- centration of substrates and NADH. Pyruvate concentration greater than 10 mM and NADH concentration greater than 0.125 mM were inhibi- tory for root lactate dehydrogenase. Hydroxypyruvate concentration greater than 8 mM was inhibitory for hydroxypyruvate reductase. Hydroxypyruvate reductase from pea roots utilizes hydroxypyruvate at a K.m of 4.8 x 10-4 M but the K.m for glycerate at 2.4 x 10.2 M is so high that the dehydrogenase reaction seems hardly possible in vivo. The same concern is noted for the Km (lactate) of 5.3 x 10-2 M for lactate dehydrogenase. It is presumed in the liver that lactate oxidation occurs by this enzyme because of immense amounts of enzyme and a high concentration of lactate, but neither of these factors is present in the roots of plants to favor the dehydrogenase reactions. Subcellular Localization of Hydroxye pyruvate Reductase in Pea Roots Lactate dehydrogenase is considered to be a soluble or cytoplasmic enzyme from work with animal tissues (74). However, hydroxypyruvate 55 HVIOH x m.v oumuUMHIH 28 ON Odz m.m ommcomonpwnmo oumuomq NIOH x m.m Qsu>m>xOHO>£ 28 OH madz O.O mmmcomouvwnow oumuomq HHIOH x v.m $042 28 NH.O AIHHV oum>su>mmxouwmm O.O mmmcomoucxnoo oumuomq vIOH x O.m mOsuxm O.h ommcmmonvasmn mumuomq NIOH x v.m omz 28 m AIMOO oumuoowHOIqo N.O ommuoswou oum>sn>m>xoucxm OIOH x O.O mum>su>msxouo>n zs OH moaz N.O mmmuoswmu mum>numm>xouosm mIOH x O.H mafiz 28 NH.O AImzv oumHhxoxHO m.O mmMOUSUOH mum>=u>mmxouomm HVIOH x m.v mosu>m>xou©>2 N.O mmmuosvou mum>zu>m>xoucmm HHMHOEV 8M oumupmndm unnumcou oumuumnsm OHQMHHM> 2m moshucm ommuoswou oum>su>mhxou©>n use mmmcomouwmnov oumuomH uoou onu 80m mucmumcoo mHHomnon .x OHQMB 56 reductase was found in the peroxisomes in the leaves (111), although it has not been found in the peroxisomes of the rat liver (74). Therefore, the subcellular distribution of hydroxypyruvate reductase in pea roots was examined. In an experiment by differential centri- fugation in 0.5 M sucrose, as described in method B, 40% of the catalase and 88% of the cytochrome c oxidase was found in the par- ticulate fraction but only 9% of the hydroxypyruvate reductase was in the same fraction and the rest was in the supernatant. The specific activity of hydroxypyruvate reductase in this fraction was the same as that in the crude extract, while its specific activity in the supernatant increased due to particle removal by centrifuga- tion (Table XI). In another experiment, the root homogenate was centrifuged over a continuous linear sucrose density gradient as described in Materials and Methods. The distribution of specific marker enzymes for organelles as well as for the hydroxypyruvate reductase was determined (Figure 8). The peroxisomal marker enzyme, catalase, and the mitochondrial marker enzyme, cytochrome c oxidase, overlapped at an equilibrium density of about 1.21 g/cm3, while hydroxypyruvate reductase was found only at the low density part of the gradient, i.e., the supernatant. Although the gradient failed to separate the organelles in the root homogenate, it was good enough to show that the hydroxypyruvate reductase was present in the cytosol of the cell. It is possible that one of the isoenzymes of hydroxypyruvate reductase is present in the microbodies of the root in too low a level for detection. This possibility is not likely, since in the next section it is shown that in the disc electrophoretic patterns of the iso- enzymes of the root hydroxypyruvate reductase and leaf hydroxypyruvate 57 .>HH>Huom OCOCOOOEOC Hmuou HO O we .1 OOH OOH mHH «>Ho>ooou Hmuoe ucmumcuomsm OO OH. O O HO OO O x OOO.R O OH OH HO OH O OOHHOO O x OOO.Hm O Om OO OON OO OOH OOHHOC O x OOOO H OH O.O Om O OO OOHHOC O x OR. OOH Hm OOH Om OOH OO 8:8 HOV CHopoum Hwy CHououm Hwy CHmuonm COHuomHm CoHuCnHHumHO OE\CH8\mmH08C COHHCQHHumHO O8\CH8\mOH08C CoHuanHumHa 08\CH8\mmH081 ommuoswom ouO>Cummeouomm ommOon o oEOHCoQMNO mmmHmumO .m Oosuo8 CH OonHHomoo mm UmEHOmnom mOB omouosm 2 m.O CH CoHumOCMHHquo HOHHCOHOMMHO mouMCmOOEOC uoou mom mo mCoHuomuw HOHCHHoondm CH ommuoCUmu oum>suwm>xOH©>C mo CoHuCQHHumHO .Hx OHQOB 58 .m80 Hmm 8O CH >HHmcm© mmouosm .uCOHOOCO H8 Com ouCCHE Com powwommmmHO NON: OOHOEJ CH ommHmuOO .uCOHOmHO H8 Com ouCCHE Mom OONHOon o meounoouxo OOHOEC CH ommOon o o8oucoou>0 Iii} .quHOOHO H8 Com musCHE Com OONHOHxO 2O4z mOHOEC CH mmmuoswmu wum>58>m>xouvwm .quCHE Com oumuumnsm mo oHOEC H mo OOCOHmwmmmmHo mcu OCHNHHOHOU 08>NCo mo UCCOEO onu mm OOCHMOU mH HHCC oCO .ommcon o o8ounoou>o an OHHOCOCUOUHE OCO mmOHmumo >n OOHOOHOCH mH HoxHOE HOEOOHxOCmm .uCOHOOHO wuHmCmo mmouosm HOOCHH mCOCCHuCoo m CH muoou mom 808m mmmuosuou oum>su>m>xouomn OCO mthNCo meuOE HOHCUCOCoouHE UCO AOOQOHOHE mo CoHuCCHHumHO .m oHCOHm (99” X u"5) KIIsuao asomns I9 C.“ — 9 T '3' T’ :1 ( Iu-I x Suun) asopIxo oawmqooutg ‘2 | 9. In 00 I T | fl“. 0 .4 I D\\\\\\\\\‘ 1 C) 'l \ / ,—""’3]$u\i8 / / \I ./ j _/ ./ / ° 0 fl / N “susmsxouosm z OIOH x O.H NO NIOH x O.H z OIOH x O.H OOOHOxosHO om 2 mIOH x N.H II 2 NIOH x v.m oumnoomHmlo 8 ”mosHm> M Houm>snmm>xoupmcv mm mom Omuuomou uOC mom CoHuHQHCCH mpmuumndm O0.00 OO82\OOO<2 Oo<2\mmosuwmhxouwmm .Nv mum>suhmxxouvwm mum>smm®>xouvxm muHoHMHoomm mumuumndm mm OOO.OOIOOO.mO OO OOO.~O mm OOO.eO OOO.OO OOOHmz umHsomHoz .mom H0>HH moon .mom Neon uoom OOHuHomonm Ho>HH moon UCO mobmmH CUMCHmm .muoou mom 80mm ommuodvmu mum>5u>m>xOHO>2 mo COmHHOQEOO .HHHx mHQmB 68 vb HOmoumo HHH m80meoumm H0mou>o CoHuOooH HOHCHHmomHuCH Hmzum m0 COHu OO H21 OHO OO mv.mm H2: my mm H28 H.Ov OOH ImuuCooCoov COHuHQHCCH w "meow on OuH>HuHmcmm Osm.o .Omm.o OOH.O mmfiOquomH mo OO OOH.O mm .OOH.O .OOH.O .OOH.O .NOH.O OOHOHHHnos oHuwuoemouuomHm mm H mm vlm m OEHOM OHOOESNCoomH mo COQECZ O0.00 mo> NNH.Nv mom OOCHEOxo HOC mCOHCm 2Q CoHum>Huo< Om m.m m.m O.m COprUHxO oumuoo>H0 O0.00 O.OIO.O HHH O.OIO.O HOOOHQO m.m CoHuonmu wum>nu>m>xonvam "MEHHQO mm .mmm Ho>HH moom .mom mmoq uoom mOHuummoum HomscHucooO HHHx mHnOe 69 and function remain to be determined. 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