BIOCHEMICAL EFFECTS OF MALEIC HYDRAZIDE (1, 2-DIHYDROPYRIQAZINE-3,6-■JluNC) ON RAPHANUS SATIVUS By Lowell Ernest Weller A THESIS Submitted to the School for Advanced Graduate Studies of Michigan State University of Agriculture and Applied Science in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Chemistry 1956 ProQuest Number: 10008656 All rights reserved INFORMATION TO ALL USERS The quality of this reproduction is dependent upon the quality of the copy submitted. In the unlikely event that the author did not send a complete manuscript and there are missing pages, these will be noted. Also, if material had to be removed, a note will indicate the deletion. uest. ProQuest 10008656 Published by ProQuest LLC (2016). Copyright of the Dissertation is held by the Author. All rights reserved. This work is protected against unauthorized copying under Title 17, United States Code Microform Edition © ProQuest LLC. ProQuest LLC. 789 East Eisenhower Parkway P.O. Box 1346 Ann Arbor, Ml 481 06- 1346 ii ACKNOWLEDGMENTS The author wishes to express his sincere appreciation to Professors C. D. Ball and H. M. Sell for their interest and guidance throughout the course of this investigation. The writer is also indebted to Dr, S. H. Wittwer who was instrumental in providing the biological tissue. He is also indebted to the Department of Agricultural Chemistry for the opportunity to pursue graduate study. Grateful acknowledgement is also due Mrs. Audrey Anderson and Mrs. Jean Brehmer for their assistance in the mechanics of the preparation of this manuscript. iii Vita Lowell Ernest Weller candidate for the degree of Doctor of Philosophy Dissertation: Biochemical Effects of Maleic Hydrazide (1,2-Dihydro pyridazine-3>6-dione) on Raphanus sativus Outline of Studies: Major subject: Minor subjects: Biochemistry Organic Chemistry, Plant Physiology and Anatomy Biographical Items: Born, April 17, 1923, Continental, Ohio, Secondary Education, Antwerp High School, Antwerp, Ohio, 1937-U1. Undergraduate Studies, Bowling Green State University, 19iil-l*3, cont, 19li6-U8. Graduate Studies, Michigan State University, 19U8-5>6, M.S. 195>1. Experience: Member United States Army, 19U3-U6. Undergraduate Assistant, Bowling Green State University, I9I46-U 8 . Research Associate in Agricultural Chemistry, Michigan State University, 19U8-5>6. Member of the American Chemical Society, Associate member of the Society of Sigma Xi, iv BIOCHEMICAL EFFECTS OF MALEIC HYDRAZIDE . (1,2-DIIIYDROPYRIDAZINE-3 ,6-DIONE) ON RAPHANUS SATIVUS By Lowell Ernest Weller AN ABSTRACT Submitted to the School for Advanced Graduate Studies, Michigan State University of Agriculture and Applied Science in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Chemistry 1956 Approved ~LO Lowell E. Weller v Many reports have been published concerning the use of maleic 1 hydrazide to prolong the shelf life of a number of plant root crops and storage organs. It is assumed that storage life is prolonged through decreased respiration rates and/or growth inhibition. Dewey and Wittwer (l) noted that a preharvest foliar treatment of radishes (Raphanus sativus) with maleic hydrazide (2^00 ppm or 0.02 M) solution inhibits new root and shoot growth of topped radishes (Radishes clipped near the base of the petioles without removal of the vegetative growing points). The radish plant appeared to be an ideal choice of biological material to study the growth inhibition induced by maleic hydrazide since it is readily cultured under a variety of conditions and the effectiveness of the treatment can be ascertained after only a few days storage. An investigation of the MH-treated and control fleshy radish root samples taken U8 hours after a preharvest foliar treatment with maleic hydrazide and also topped radishes held in storage for five days after the 1*8 hour treatment revealed that there was no difference in beta amylase, phosphorylase and phosphatase activities. Alpha amylase and pectin-methyl-esterase activities could not be detected. Gross chemical analysis of both the fleshy root (1*8 hours after treatment and stored as topped radishes for 7 days) and shoot (U8 hours after treatment) tissues revealed that there were no essential differ­ ences induced by treatment. Both the root and shoot tissues of control Maleic hydrazide is l,2-dihydropyridazine-3,6-dione and is abbreviated MH. Lowell E. Weller vi and treated plants were the same with respect to per cent dry weight, ether extractives, Kjeldahl nitrogen, reducing and non-reducing sugars, and polysaccharides other than starch. Starch in the treated shoot tissue was about double that of the corresponding control tissue. 1 A comparative quantitative examination of the DPNH oxidase system of shoot tissue (U8 hours after treatment) and topped radishes (1*8 hours after treatment, stored 7 days and separated into fleshy root and shoot stub) was conducted. The oxidative system was analyzed quantitatively for the activity of its known constitutive enzymes; that is, (a) diaphorase, (b) DPNH-cytochrome c reductase and (c) cytochrome oxidase (2). The DPNH oxidase activity was the same in both the control and treated root tissue. The activity of the constitutive enzymes was also the same in both types of tissue. The MH-treated shoot tissue exhibited a marked inhibition of the DPNH oxidase system. This inhibition resulted from partial failures in both DPNH-cytochrome c reductase and cytochrome oxidase systems. The DPNH oxidase activity of the treated shoot stub tissue was somewhat lower than that of the corresponding control tissue. The inhibition resulted from a reduced activity of the diaphorase system. The activity of the cytochrome systems was the same for both the control and MH-treated tissue. Muir and Hansch (3) have suggested that maleic hydrazide inhibits growth by reacting with thiol compounds in a manner similar to the DPNH is reduced diphosphopyridine nucleotide. Lowell E. Weller vii addition of thiols to maleic acid. An investigation of the in vitro interaction of thiols with maleic hydrazide indicated that no addition compound resulted from the incubation of maleic hydrazide and thiols at a neutral pH, either in the presence or absence of tissue homogemate. 1. The interaction of maleic acid and a thiol compound was noted. D. H. Dewey and S. H. Wittwer, Proc. Am. Hort. Soc., 57, in press (1956). 2. E. C. Slater, Biochem. J., U6, U8U, h99 (1950). 3. H. M. Muir and C. Hansch, Plant Physiol., 28, 218 (1953). viii TABLE OF CONTENTS PAGE HISTORY OF THE DEVELOPMENT OF THE CONCEPT OF PLANT GROWTH SUBSTANCES..................................... 1 INTRODUCTION . . ............................................ 6 REVIEW OF THE LITERATURE........................ . .......... 10 Effect of maleic hydrazide on plant celldivision ................... and enlargement. 11 Effect of maleic hydrazide on the morphology of plants . . 13 Effect of maleichydrazide on the auxin economy of plants • 14 Effect of maleic hydrazide bn the carbohydrates 15 Effect of maleichydrazide on enzyme systems. . . . . . . . 17 Effect of maleic hydrazide on plantrespiration ........... 18 Toxicity of maleic hydrazide......... .................... 19 STATEMENT OF THE P R O B L E M ................ ................... 23 of plants . EXPERIMENTAL.................................................. 25 Effect of maleic hydrazide on the carbohydrase and phosphatase activity of radishes.. ....................... 26 Preparation of biological material ...................... 26 Analytical m e t h o d s ........................... Results and discussion............ .................. 28 35 Effect of maleic hydrazide on the composition of radishes . 40 Preparation and analysis of radishsamples......... . . 40 Results and discussion................................ 46 Effect of maleic hydrazide on the DPNH oxidase activity of radishes.......................................... • Introduction............................................ 50 50 ix Preparation of DPNH oxidase system .................... 53 Analytical methods .................................... 58 Results and discussion ................................ 70 Maleic hydrazide and thiol compounds...................... 90 Introduction .......................................... 90 Analytical methods .................................... 94 Results and discussion ................................ 95 SUMMARY...................................................... 98 BIBLIOGRAPHY......... 101 X LIST OF TABLES TABLE I, PAGE Effect of Maleic Hydrazide on the Beta Amylase Activity of Tissue Preparations...................... 35 Effect of Maleic Hydrazide on Enzymic Activity of a Commercial Beta Amylase Preparation.................. 36 III. Phosphorylase Activity of Tissue Preparations........ 37 IV. Phosphatase Activity of Tissue Preparations.......... 38 V. The Effect of a Preharvest Foliar Spray of Maleic Hydrazide on the Composition of Radishes.......... 48 The Effect of a Preharvest Foliar Spray of Maleic Hydrazide on the Composition of Radishes.......... 49 Relative Activities of the DPNH Oxidase System and Constitutive Enzymes.............................. 89 Interaction of Maleic Hydrazide and Thiol Compounds. . 96 II. VI. VII. VIII. xi LIST OF FIGURES FIGURE 1. 2. 3. 4. 5* 6. 7. PAGE The DPNH oxidase activity of control and maleic hydrazide treated radish tissues .............. . . 75 The diaphorase activity of control and maleic hydrazide treated radish tissues ...................... 79 The cytochrome oxidase activity of control and maleic hydrazide treated radish tissues ...................... 80 The effect on the absorption at 550 mp. produced by the addition of 0.4 ml. of 10“4 M cytochrome c to the DPNH oxidase system of control and maleic hydrazide treated radish tissues .......................... 82 The effect on the absorption at 340 mp. produced by the addition of 0.4 ml. of 10“4 M cytochrome c to the DPNH oxidase system of control and maleic hydrazide treated radish tissues .......................... 83 The effect on the absorption at 340 mp. produced by the addition of 0.4 ml. of 10”4 M methylene blue to the DPNH oxidase system of control and maleic hydrazide treated radish tissues. 85 The DPNH-cytochrome c reductase activity of control and maleic hydrazide treated radish tissues............ 87 HISTORY OF THE DEVELOPMENT OF THE CONCEPT OF PLANT GROWTH SUBSTANCES Our earliest records show that naturalists of the seventeenth century, in their concern for an understanding of the world about them, brought forth speculation on the nature of the factors underlying the development of plants. It was not until the latter half of the nine­ teenth century, when modern science of plant physiology was really born, that the German botanist Sachs started a detailed study designed towards an understanding of plant development. As a result of his studies he proposed a generalized theory involving "organ-forming sub­ stances" to explain the facts of plant growth and organ production. The original observations which led directly to the first isola­ tion of a plant hormone have been attributed to the British naturalist Charles Darwin. Darwin was primarily concerned with the elucidation of the mechanism of plant responses to external stimuli, for example, unilateral light and gravity. As a result of his research he proposed that the stimuli were perceived by one part of a plant and that some "influence" must be transmitted since the results of the stimuli were expressed in a growth response at some other site of the plant. Boysen-Jensen (1) demonstrated that the "influence" would pass through such a non-living substance as gelatin. Thus, the response of a coleoptile to unilateral light was unimpaired by inserting a gelatin block between a severed tip and the shoot stump. Paal (2) showed that the tip of the coleoptile could influence the growth of the stump 2 independent of any external stimulus. These studies as well as others demonstrated that there exists in the tip of the coleoptile a substance which passes into and down the side of the stump in contact with an asymmetrically replaced severed tip, stimulating the extension of growth of the zone below the tip and giving rise to the curvature observed* It was not until 1926 following the now classical studies of Went, that the isolation of the chemical "messenger” was finally accomplished* The isolation and the technique which arose from it marked the beginn­ ing of the modern era of plant growth substances* Went*s original contribution to the study of plant growth substances was that of a quantitative method for the measurement of the curvature produced by small agar blocks containing the hormone. Since these hormones are active at extremely low concentrations, below the sensitivity of chemical tests, the development and refinement of the biological test ncrtr set the stage for the final isolation of the hormone in pure crystalline form and its subsequent structural elucidation. The chemists, in an attempt to isolate a sufficient quantity of the growth hormones for structural elucidation, chose a more readily it available starting material than coleoptiles. Kogl and Haagen-Smit (3) discovered that human urine was a source of particularly active material* They devised a purification procedure which yielded less than a gram of crystals which was quite active in the Avena test* The chemical constitution of this compound was established and it was subsequently named "auxin A ”. These workers then attempted to obtain "auxin A ” from a plant source, namely corn oil and barley malt. A 3 like series of purifications as used in the case of the urine yielded "auxin A ” and a new compound of similar structure which they called "auxin B". Three years later these workers in conjunction with Erxleben (U) isolated a third active substance from urine called "heteroauxin" (now known as 3-indoleacetic acid). Following these historic years in the development of our know­ ledge of plant hormones, "auxin A and B" were regarded as the natural plant growth substances and "heteroauxin" was probably a product elaborated by microorganisms. From the time of the first report of the isolation of "auxin A and B" many investigations have been directed towards obtaining these auxins from a great variety of plants. This includes efforts by the original investigators to repeat their earlier isolation. To date, no one has been able to obtain either of these auxins from any source. Efforts of the organic chemist to synthesize compounds of the structure assigned to "auxins A and B" have also been unsuccessful. Such evidences as these and others have caused several research workers to "privately express doubt as to the existence of auxins A and B". Even those who are most sympathetic toward the "auxin A and B" existence must admit that there is no rigorous evidence to which they can point. While there is considerable doubt as to the existence of "auxins A and B", there is certainly no doubt as to the existence or importance of 3-indoleacetic acid in the auxin economy of a wide variety of plants. The investigation associated with the isolation and discovery of the unique properties of 3-indoleacetic acid is a nearly typical example of a fundamental investigation which has led to discoveries of great economic importance. h 3-Indoleacetic acid was first obtained as a degradation product of protein as early as 1885 (5)* by Ellinger in 1905 (6). It was first synthesized in Germany However, it remained for Kogl, Haagen-Smit and Erxleben to recognize the unique growth regulating properties of this compound. It was this compound which has served as a model for the chemists who subsequently synthesized a very wide range of com­ pounds of similar structure for use in the control of physiological processes occurring in plants. Much of the early work on growth hormones has been concerned chiefly with the stimulation of growth processes. It was soon recog­ nized, however, that a single substance may both inhibit and stimulate a plant growth pattern; the type of response depending, in part, upon the concentration of growth regulating substance. Thusly, the classi­ fication of growth regulating substances on the basis of stimulators or inhibitors becomes a relative situation depending upon the conditions of its use. It seemed natural that the growth stimulation process would attract the most attention in the early studies. Consequently, many synthetic growth regulating substances structurally related to 3-indole­ acetic acid have been recognized as a result of the research initiated by the discovery of the unique properties of this naturally occurring hormone. However, it was soon realized that under certain conditions growth inhibition could be a desirous effect. One compound which is generally regarded as a growth inhibitor is maleic hydrazide (l,2-dihydropyridazine-2,6-dione). Although this compound is not structurally related to 3-indoleacetic acid, it is one of the compounds whose 5 biological properties were discovered in the search for growth regulat­ ing substances. Literally hundreds of reports have been received with regard to the practical applications of this growth inhibitor. usually the case the more fundamental studies have followed. As is However, very little information is as yet available as to how this compound can bring about growth inhibition. 6 INTRODUCTION 7 INTRODUCTION In 1395, Foersterling (7), during an investigation of the reactions of hydrazine hydrate with phthalic and maleic anhydride, isolated a com­ pound which he called maleic acid hydrazide1 having the following structure: 0 II /°\ H-C N-H II H-C I N-H V II o It was not until nearly fifty years later that any report concerning this compound appeared in the literature# Arndt (8) reported the use of diazomethane in a study of the tautomeric forms of such amides as maleimide, cyclic hydrazides, malonamide, uracil and barbituric acid. tautomerizehalf-way in many cases, particularly if favored by two hydrazide groups onthe ring formation. a He found that dihydrazides same carbon atom or bysix-membered aromatic In later studies Arndt and co-workers (9) report that hydrazinedicarbonyl group -C-NH-NH-C- lying between two atoms can 0 0 1Maleic acid hydrazide is known more commonly now as simply maleic hydrazide. Both of these names are actually misnomers. From the point of systematic organic nomenclature it is 1,2-dihydropyridazine-3,6-dione. Although all three names appear in the literature today, the most common name, maleic hydrazide, will be used exclusively throughout this manu­ script and will be abbreviated MH. 8 have only one —CONH- group tautomerize due to the adjacent positive ■ + + — charges of the OC—NH—NH—CO mesomer which facilitates movement of a + H from nitrogen to oxygen. Maleic hydrazide is slightly acid in character and forms salts readily with alkalies• The free compound is completely water soluble at 0.2$ but is not soluble at the 1.0$ level. insoluble salts from solution. Heavy metals precipitate Both the soluble salts and the rela­ tively insoluble salts function as growth regulants. 0 is reported as in excess of 2^0 The melting point 0 (7), 260 o dec. (9) and 299.^-300 (10). Advantage is taken of the acid property to render maleic hydrazide more soluble and therefore more versatile biologically. tions which are available commercially are: Two common formula­ MH-30, which is a formula­ tion containing 5$% diethanolamine salt of maleic hydrazide (equivalent to 30$ maleic hydrazide) and h2% inert ingredients which include wetting agent and sticker and which is a formulation containing 1*3.3$ sodium salt of maleic hydrazide (equivalent to h0% maleic hydrazide) and £l.7$ inert ingredients. Schoene and Hoffman (11) first recognized the unique biological properties of maleic hydrazide in 19U9* They demonstrated that maleic hydrazide, when applied as a spray, temporarily inhibited stem growth of tomato. Plants showing this response resumed growth from lateral buds about two months after treatment. They also observed that some chlorosis appeared and that root growth was also inhibited in several other species of plants. Since this first report of the unique growth regulating properties of maleic hydrazide, a great many reports have appeared in the literature describing various responses of plants to 9 this compound, usually related in some way to inhibition of growth. The question that still remains unsolved is how does this compound affect the metabolic systems in plants. 10 REVIEW OF THE LITERATURE 11 REVIEW OF THE LITERATURE There are scores of reports in the literature concerned with many varied responses of plants to treatment with maleic hydrazide* A great many of these reports are primarily concerned with the practical appli­ cation of this compound to gain a particular objective without regard as to why or how a certain result came about. There are much fewer reports of a more fundamental nature designed to gain an understanding of how maleic hydrazide alters the biochemical reactions occurring within the organism. This literature review will cover only a selected group of papers falling into the latter category. This then requires a certain amount of selection on the part of the reviewer; certainly the selection of any two reviews might conceivably vary considerably. No attempt will be made to maintain the chronology of reports with respect to the elucidation of our knowledge of the interaction of maleic hydra­ zide and plants. Effect of Maleic Hydrazide on Plant Cell Division and Enlargement Darlington and HcLeish (12) reported that roots of Vicia faba which had been exposed to an aqueous solution containing 0.005 M maleic hydrazide for twenty-four hours showed no mitosis for two days. Lower concentrations did not stop mitosis, but did cause breakage of chromo­ somes at mitosis. Breakage was confined to heterochromatin whereas x-rays are known only to break euchromatin. These authors did not 12 observe this phenomenon with all plant species studied* Deysson and Deysson (13) also investigated the mitosis-inhibiting action of maleic hydrazide and other known mitotic poisons* Using mitotic poisons bear­ ing structural similarities to uracil, namely, maleic hydrazide, barbital and antipyrine, they could not demonstrate that added uracil would reverse the mitosis-inhibiting action of the above agents. There' fore, they concluded that these mitotic poisons did not exert their effect through interference with the metabolism of uracil. Greulach and co-workers (ll*) observed that maleic hydrazide at concentration of 1 to 2000 ppm inhibited mitosis and cell division in onions in proportion to the concentration of maleic hydrazide. Mitosis was resumed after transferring from maleic hydrazide solution with recovery marked at low concentration. At concentrations of 1 ppm many more roots were produced indicating a possible stimulating effect. Additionally (15>) sectioned terminal buds from young beans sprayed with 0.01 M solution of maleic hydrazide showed only a few scattered mitotic figures at one week and none at two weeks following treatment. Apical cells of the treated plants were enlarged and vacuolated. Finally, Gruelach and Haesloop (16) concluded from a study of the effects of maleic hydrazide on internode elongation, cell enlargement and stem anatomy that cell enlargement was only slightly affected. Inhibition of cell division accounted for practically all of the ob­ served growth inhibition. 13 Effect of Maleic Hydrazide on the Morphology of Plants Moore (17) listed the easily visible effects of maleic hydrazide on plants as (a) a temporary suspension of stem elongation from ter­ minal buds or death of terminal buds and adjacent tissues, (b) expansion of lateral buds some time after the terminal bud had been affected, (c) a transient intensification of green in leaves of stunted plants, (d) a localized accumulation of anthocyanins or other non-green pig­ ments, (e) narrowing of leaves on both monocots and dicots, (f) several patterns of leaf chlorosis, (g) an interference with water adsorption, apparently caused by death of root tips, (h) suppression of nodule formation on bush beans, and (i) total, temporary or male sterility. A number of plant species were treated with maleic hydrazide at G.05> to Q*h% as reported by Naylor and davis (Id). They found that maleic hydrazide is remarkably uniform in plant responses from species to species with some loss of sensitivity developing with age. The effect of maleic hydrazide in every species was, (a) cessation of * activity of terminal meristerns, (b) cessation of elongation of internodal region and (c) increase in stem diameter. New leaves were not permanently affected since normal leaves were eventually produced. Histological studies by Rao (19) revealed that a pre-harvest foliar spray of maleic hydrazide induced inhibition of differentiation of tissues in the bud and root primordia of potato tubers and onion bulbs. There was also a retardation of cell division. Hi Effect of Maleic Hydrazide on the Auxin Economy of Plants Growth inhibitors, in general, do not cause lateral buds to develop* However, one agent that does inhibit stem elongation and that also breaks apical dominance is x-ray irradiation, which also causes the destruction of 3-indoleacetic acid or auxin* auxins control apical dominance (20)* Also, it is known that Many reports have been received that maleic hydrazide also breaks apical dominance (U, 11, 17, Id, 21, 22) and it is, therefore, concluded that maleic hydrazide in some way interferes with the normal auxin metabolism* Leopold and Klein (23) reported that maleic hydrazide did not show growth stimulation in standard pea and straight growth tests at low con­ centrations but does inhibit growth in the above tests at dilutions of one part in ten million* 3-Indoleacetic acid completely overcame the maleic hydrazide inhibition* Additionally, there is no reaction in vitro between maleic hydrazide and 3-indoleacetic acid as determined by diffusion studies. Other studies by these same workers (2U) re­ vealed that growth inhibition by high concentrations of auxin can be relieved by the addition of maleic hydrazide. These authors conclude that maleic hydrazide is not a growth regulator since it is not capable of promoting growth in the absence of auxin but it is an antiauxin and acts in opposition to auxin in growth* Other reports (25, 26) have been presented that show that maleic hydrazide accelerates the rate of 3-indoleacetic acid destruction. The antiauxin effect of maleic hydrazide on growth then is ascribed to the accelerated removal of endogenous auxin* 15 Gautheret (27) found that maleic hydrazide at low levels enhances the stimulating effect of 3—indoleacetic acid and at higher levels the two chemicals act antagonistically. A similar direction of response was obtained in root elongation studies (28). Cell proliferations which can be induced by 3-indoleacetic acid can be completely inhibited -4 by maleic hydrazide at 10 _e to 10 M (29). Effect of Maleic Hydrazide on the Carbohydrates of Plants A common response of plants treated with maleic hydrazide is the accumulation of anthocyanin pigments (17, 18 , 30, 31, 32, 33). Such a response is usually interpreted to mean that in some way carbohydrate metabolism has been impaired and that the resulting carbohydrate accumu­ lation results in increased anthocyanin production. Currier and collaborators (3h) working with barley plants and Naylor (35) working with young corn plants both noted that maleic hydrazide treated plants exuded a fluid rich in sucrose. Naylor additionally found that quantitative sugar analysis indicated a tre­ mendous accumulation of sucrose in shoots and roots twenty days after treatment depending upon the concentration of maleic hydrazide. No glucose accumulation occurred as a result of treatment. Probably few other plants have been so widely used in investiga­ tions of the effect of maleic hydrazide on the carbohydrate economy of plants than the sugar beet. Of particular importance in these inves­ tigations was the effect of maleic hydrazide upon the sucrose content. Mikkelsen and collaborators (36) found that foliar sprays of maleic 16 hydrazide produced significantly higher sucrose percentage and did not influence yield. Wittwer and Hansen (37) used a pre-harvest foliar application of maleic hydrazide (2500 ppm) to suppress top growth, root growth and storage breakdown and preserve the sucrose content. In addition to preserving sucrose content, such treatment also increased sugar content* Wittwer and coworkers (38, 39, 1*0) used a pre-harvest foliar spray of maleic hydrazide to extend the storage life of a number of roots and other storage organs. Pre-harvest application of maleic hydrazide to carrots had little effect upon the carbohydrates of organs subsequently held in storage (38). Conflicting reports have been re­ ceived as to the effect of pre-harvest application of this compound to potatoes. Highlands (1|1) and Kennedy and Smith (h2) report that maleic hydrazide had no effect upon the reducing sugar content following storage* Patterson and coworkers (39) showed that reducing sugars accumulated in chemically treated tubers. Although these results seem contradictory, different experimental conditions may account for the seemingly con­ tradictory results* Petersen (1±3) suggested that a block in normal carbohydrate metab­ olism exists which either induces protein use as a respiratory substrate or else inhibits protein formation. This suggestion was prompted by the observation that maleic hydrazide treated tobacco plants contained larger amounts of reducing sugars, reduced soluble nitrogen, and less protein. Phouphas and Goris (hh) found that in vitro cultures of arti­ choke containing maleic hydrazide showed an increased concentration of sucrose and a corresponding decrease in inulin. 17 Cytological examination of floral structures treated with high concentration of maleic hydrazide, $.0 to 10%, showed an abundance of starch in the parenchyma cells which remained for a longer period of time than in flowers not treated* The preservation of starch was correlated with retarded flower maturation (1*5). The author suggested that retardation of starch digestion may account in part for lower respiration observed by other investigators* Effect of Maleic Hydrazide on Enzyme Systems Only a few reports have been received with regard to the effect of maleic hydrazide on enzyme systems. Some of the reports which have been received are rather incomplete. Greulach (1*6), studying the effect of maleic hydrazide on starch synthesis and breakdown, reported that this chemical does not block the starch breakdown process. Further, he suggests that it may block either hydrolysis or phosphorolysis but it certainly does not block both pro­ cesses. The observed delayed disappearance of starch from the attached treated leaves could be due to the larger quantity of starch initially present in them or to a slowing of their processes by maleic hydrazide but was more likely due to the lower rates of respiration, assimilation and to interference with translocation in the treated plants owing to maleic hydrazide injury of the sieve tubes. Isenberg and coworkers (1*7) found that maleic hydrazide sprayed upon the foliage of onion affects respiration through partial in­ activation or inhibition of one or more of the dehydrogenases. In a later publication (1*8), they report that succinic dehydrogenase and 18 respiration were stimulated at low maleic hydrazide concentration and inhibited at higher concentrations, Marre (1*9* 50) found that maleic hydrazide and other antiauxins, in concentrations near those that reversibly inhibit growth in vivo, inhibited the activity of a preparation of dehydrogenase in vitro, Concentrations of 3-indoleacetic acid which were inactive in the absence of an inhibitor appear to be able to reverse the inhibiting effect of antiauxin upon the dehydrogenase system. Maleic hydrazide at concentrations of 10 to 10 M stimulates the enzymatic oxidation of 3-indoleacetic acid by indoleacetic acid oxidase with an optimum pH of 5*6 (5l). In contrast to this is the report of Gortner and Kent (52) that pineapple indoleacetic acid oxidase is not inhibited by maleic hydrazide at a concentration of 1.8 x 10 3 M, pH optimum 3«25» Morel and Demetriades (53) have reported that maleic hydrazide inhibits colony growth of culture tissue, and decreases peroxidase activity. Both 3-indoleacetic acid and 2,U-dichlorophenoxyacetic acid enhanced the activity of Jerusalem artichoke tissue with respect to polyphenol oxidase but maleic hydrazide, which inhibits growth, also activates this enzyme. vity and proliferation. There is no correlation between enzymatic acti­ Maleic hydrazide also decreased catalase activity. Effect of Maleic Hydrazide on Plant Respiration The inhibiting effect of maleic hydrazide on the respiration of plant organs has been reported by many investigators (1*7, U6, 5U, 55). 19 Some of these reports have been mentioned earlier. Naylor and Davis (51-t) suggested from the evidence now available from growth and res­ piration studies that maleic hydrazide acts either as a poison or as a biological antagonist competing for the receptor portion of an enzyme(s) concerned in respiration. A phenomenon generally assumed to be associated with growth is respiration although it is now known that the proportion of respiration energy actually utilized in growth, that is, in cell division and increase in size, is small (56, 57)* Though there is no positive correlation between growth and respiration, some respiration is essential for growth. Certainly a partial or com­ plete inhibition of respiration by lack of oxygen or respiratory poison is accompanied in higher plants by cessation of growth. Toxicity of Maleic Hydrazide The development of the use of maleic hydrazide on human foodstuff necessitated the study of the toxicity of this chemical. studies by Tate (58) in this regard will be summarized. The extensive Chronic oral toxicity studies with the sodium salt of maleic hydrazide have been conducted on rats and dogs. Rats have been kept exclusively on a diet containing from 0.5 to 5.0$ active maleic hydrazide as sodium salt (equivalent to 0.6 to 6.0$ sodium salt of MH) two years from weaning and through successive generations. Dogs similarly were fed from 0.5 to 2.0$ active maleic hydrazide as sodium salt (equivalent to 0.6 to 2.14$ sodium salt of MH) for one year. Rats of both sexes grew to normal adult weight on the sodium maleic hydrazide diets and the dogs responded in weight gains equal to or better than controls. 20 The parent, generation of rats were mated, and their progeny were mated through three successive generations to produce through weaning two successive litters in each generation. Sodium maleic hydrazide had no effect on fertility of rats regardless of food level as judged by ratio of pregnancies to matings, ratio of litters born to pregnancies and by size and weight of litters at birth. No deviations from normal were noted in periodic blood and urine examinations of both rats and dogs. Autopsies were performed on rats that died during the two-year test and on those checked for this purpose, to observe any deviations from normal including occurrence and characteristics of any tumorous growth such as cysts, abscesses or neoplasms. No changes were noted which could be correlated in frequency or severity with dosages of the chemical. No evidence of tissue damage related to sodium maleic hydra­ zide was noted in either the rats or dogs at autopsy. Histopathological examination of liver, spleen, kidneys, and bone marrow of dogs fed daily doses of 1.0 g. of sodium salt of maleic hydrazide for one month showed no significant effect in one animal and some increase in destruction of red blood cells in a second animal but there were no changes of any significance in kidney or in bone marrow. The acute toxicity for rats, reported as the LD£G, has been found to be 2,35 g. per kg. for the diethanelaraine salt of maleic hydrazide and 6.95 g. per kg. for the sodium salt of maleic hydrazide. Chronic feeding studies of maleic hydrazide as dietha no la mine salt at a 1.0# active maleic hydrazide level (equivalent to 1.9# of the diethanolamine salt of MH) in the daily diet of rats showed a mortality of 21 out of 2k animals at the end of 11 weeks feeding. Further chronic 21 feeding studies of the diethanolamine fraction proved that the diethanolamine itself was responsible for this mortality and not the active ingredient maleic hydrazide. Since maleic hydrazide as diethanolamine salt has desirable pro­ duction advantages and response characteristics on plants, studies have been initiated to determine the residues of diethanolamine itself when this salt of maleic hydrazide is applied on foliage. A preharvest foliar spray of 3 pounds active maleic hydrazide (l gallon MH-30 contains 2.8 pounds diethanolamine) per acre of potatoes showed 8.7 ppm maleic hydrazide residue in these tubers. No diethanolamine was present by chemical analysis using paper chromatography, indicating that diethanolamine does not translocate into potatoes. Maleic hydrazide does not affect the growth of mice or the growth of testicular tumors in mice. Mice sprayed daily with 0.001 to 0.2$ maleic hydrazide or given drinking water with the same concentration for three weeks gained as rapidly as checks and showed no toxic symptoms. Older white mice with testicular tumors were injected subcutaneously for ten days with 0.1 ml. each of nine concentrations from 0.05$ to the undiluted 30$ maleic hydrazide as diethanolamine salt. Growth of tumors was net inhibited nor did any concentration produce symptoms of toxicity. Embryos of Rana pipiens were placed in 0.1 and 0.0%% maleic hydrazide as diethanolamine salt for U8 hours then transferred to tap water and observed for six weeks (59). ments were not significantly different. Length and body width measure­ No observable effect was noted at 0.05$ maleic hydrazide but at 0.1$ there was retardation of eye development, smaller myotomes and gills and a more sluggish response to 22 %'ight and touch when compared to controls. Circulation was apparently normal. i Eggs of Bufo sp, in the early blastula stage were placed in 0.02# maleic hydrazide for U8 hours then transferred to tap water and observed. Five days later larvae were selected for tail tip amputation to study the rate and amount of regeneration. Maleic hydrazide had no observable effect on embryonic or larval development, reflexes, or amount and rate of tail tip regeneration (59 )* Gastrulae of the salamander Amblyostoma punctatum were placed in 0.5, 1.0 and 2.0# maleic hydrazide for ten days, then transferred to tap water and observed. The posterior third of the tail was clipped from larvae and mitosis from the regenerated tail tip examined. The 0.5# maleic hydrazide was not particularly toxic but higher concentrations were. There were no superficial morphological differences between controls and survivors in size or size of the regenerated tail tips. The mean number of mitotic figures was lower in maleic hydrazide treated larvae but this was considered due to starvation rather than a direct « effect of maleic hydrazide itself (5 9 )• 23 STATEMENT OF THE PROBLEM 2k STATEMENT OF THE PROBLEM It has been well established that maleic hydrazide is a plant growth inhibitor under a variety of conditions. A preharvest foliar application of this chemical will prolong the storage life of a number of economically important roots and storage organs such as onions, potatoes, beets and carrots. An appropriate foliar spray of this chemical, when applied as a preharvest treatment to radish plants, will cause growth inhibition in the roots subsequently stored. Visual observations suggested that storage life was extended as a result of growth inhibition and lowered respiration rates. The immediate problem at hand becomes that of attempting to gain some understanding of why a maleic hydrazide treated root shows inhibition of growth when compared with an untreated root. It is realized that this study will not yield direct information as to the mechanism of the action of maleic hydrazide but should yield some information as to the effect of this chemical in inhibiting growth* It is recognized that the changes in the plant brought about by this compound may be termed a primary effect or more likely an effect many steps removed from a primary one* Data obtained in such a study as this may yield information or suggest an approach to the study of the mechanism of maleic hydrazide inhibition of growth* 25 EXPERIMENTAL 26 EXPERIMENTAL Effect of Maleic Hydrazide on the Carbohydrase and Phosphatase Activity of Radishes Preparation of Biological Material The results of a preliminary investigation by Dewey and Wittwer (60 ) suggested that the radish would be a nearly ideal plant for a biochemical study of growth inhibition induced by maleic hydrazide. They noted that a preharvest foliar spray of a maleic hydrazide solution would inhibit new shoot growth in topped radishes which were stored for short periods of time. Topped radishes were clipped near the base of the petioles without removal of the vegetative growing points. The radish can be cultured readily either in the greenhouse or in the field and develops a fleshy root in a few weeks. Additionally, the effectiveness of the treatment can be determined only a few days following harvest, A commercial variety of radish (Raphanus sativus), Ferry Morse*s Red Comet, was- chosen for this study. The plants were cultured using conventional techniques in plots in the greenhouse. When the roots had developed to a commercially feasible size (about8-10 g.) the plants were ready for treatment. Each plot was dividedinto two equal portions. The foliage of one-half of each plot was treatedwith a 2^00 ppm (0,02 molar) solution of the sodium salt of maleic hydrazide prepared by the proper dilution of MH-UO. The spray was applied with a hand sprayer at 27 the rate of one liter per one hundred square feet. Forty-eight hours after treatment samples from the respective plots were taken for investi­ gation. This time interval was sufficient to cause inhibition of new shoot growth on topped radishes (approximately 1/h inch of shoot portion remaining) as determined by a previous field study (60). Assays to determine whether treatments were effective were conducted as follows. Representative plants were harvested from both treated and control plots and washed. The shoot portion was clipped so that approxi­ mately 1/h inch of shoot remained attached to the fleshy root. These roots were then placed in ventilated polyethylene bags and stored for five days at room temperature or for longer periods of time at lower prevailing greenhouse temperatures. After such a period of storage, the controls exhibited appreciable new shoot growth in contrast to little or no new shoot growth for the treated radishes. Tissue samples. Tissue samples, obtained after being subjected to various experimental conditions, were prepared for investigation according to the procedures described below and referred to in this section with the following designations: Tissue A. Root tissue was obtained forty-eight hours after treat­ ment with a foliar spray of a 0.02 M solution of maleic hydrazide. The tissue was macerated in a Waring Blendor and the resulting macerate lyophilized. 0° until used. Such tissue preparations were stored at temperatures below Replicate samples were prepared in this manner from olants obtained from two separate plantings. Tissue B. Root tissue was obtained forty-eight hours following the treatment described above. The topped roots were stored in 28 ventilated polyethylene bags for 96 hours at room temperature, at which time the effectiveness of the treatment was evident. After removal of the remaining shoot stub, the tissue was frozen and an acetone powder prepared as follows. The frozen tissue was macerated with two weight equivalents of cold acetone in a pre-cooled Waring Blendor at 0°. The slurry was filtered and the residue again macerated with one weight equivalent of acetone, re-filtered and washed with cold acetone. The residue was then placed in a vacuum desiccator over calcium chloride and stored in the cold. Following the removal of the last traces of solvent, the powders were stored at a temperature below 0°. Tissue C. Root tissue was obtained forty-eight hours after a foliar treatment with an 0.02 M solution of maleic hydrazide. The topped radishes were stored in ventilated polyethylene bags for 96 hours at room temperature. The remainder of the shoot portion was removed after storage and the root tissue placed in polyethylene containers, immediately frozen and stored in the frozen state until used. For comparative purposes, control tissue preparations, corresponding to the preparations Tissue A, B and C, were also prepared in the same manner from similarplants of the same planting but were not treated with maleic hydrazide. Analytical Methods Phosphorylase. Enzyme solutions for investigation were prepared from both control and MH-treated tissue as follows: 1 g. of Tissue A was extracted with 25 ml. of water for 1 hr. at 25 1 g. of Tissue A was extracted with 25 ml. of succinate buffer pH6.5 for 1 hr. at 25°; ; 29 1 g. of Tissue A was extracted with 25 ml. of acetate buffer pH 5.0 for 1 hr. at 25°; 100 mg. of Tissue B was extracted with 20 ml. of water for 1 hr. at 50 g. Tissue C was extracted with 50 ml. of water for 1 hr. at 25°. After the extraction period had elapsed the samples were centrifuged, filtered and the enzyme activity was determined. Preparation of reagents. Cori ester solution. inorganic phosphate was prepared as follows. A solution free from One gram of the dipotassium salt of glucose-1-phosphate was dissolved in 10 ml. of water. Approximately 25 mg. of calcium oxide was added, the solution thoroughly mixed and allowed to stand for 10 minutes. After filtering normal sulfuric acid was added to make neutral to a phenol red internal indicator. The solution was diluted to 1±0 ml. and 60 ml. of succinate buffer was added. This solution required refrigeration for storage. Succinate buffer. A mixture of 59 g. of succinic acid and 81 g. of sodium carbonate was cautiously dissolved in distilled water and boiled to remove carbon dioxide. one liter and mixed. Starch amylose. The solution was cooled, diluted to The pH should be 6.U. One hundred milliliters of succinate buffer was added to 800 ml. of water, heated to boiling and removed from heat source. Ten grams of Merck’s Lintner soluble starch suspended in 100 ml. of water was added with stirring. and centrifuged clear. The mixture was cooled to room temperature The clear solution of amylose was decanted off, preserved with toluene and stored at room temperature, 30 Determination of phosphorylase activity (6l). flasks were placed in a water bath at 25°. Two 50 ml. volumetric To each flask was added 1 ml. of tissue extract and 1 ml. of 1.% starch amylose previously o brought to 25 . One milliliter of Cori ester in succinate buffer, pH 6.U, was added to one tube and mixed. After 30 minutes, 5 ml. of 6.66% molybdate solution was added to both flasks. the enzyme. This inactivated Now 1 ml. of Cori ester was added to the flask to which none was added initially. This became the blank. Both flasks were diluted to about 30 m l ., 5 ml. of 7.5 N sulfuric acid was added and mixed. Five milliliters of freshly prepared h% ferrous sulfate was added and the solution diluted to the 50 ml., mixed and the optical density determined with a photoelectric colorimeter using a red filter (630 mp) (62). The optical density readings were translated into pg. of phosphorus by reference to a previously prepared standard curve. The phosphorylase unit is expressed as pg. of phosphorus set free from Cori ester at 25° at pH 6.U in 60 minutes by the phosphorylase enzyme in 1 ml. of tissue extract. Phosphatase. Enzymic solutions for investigation were prepared from control and MH-treated tissues as described: 1 g. Tissue A was extracted with 50 ml. of water for 1 hr. at 38°; 200 mg. Tissue B was extracted with U0 ml. of water for 1/2 hr. at 50 g. Tissue C was extracted with 50 ml. water for 1 hr. at 25 . The samples were then centrifuged, filtered and an aliquot of the filtrate used for the determination. 31 Determination of phosphatase activity (63). Five milliliters of the tissue extract was added to 10 ml. of water and equilibrated at 38°. The substrate, 10 ml* of $ -glycerophosphate at pH 5*8, previously warmed to the same temperature, was added. Three milliliter samples were taken at appropriate times thereafter and delivered immediately into 5 ml. of 6*66% molybdate. The phosphorus determination was con­ ducted as outlined in the previous section under phosphorylase. A phosphatase unit is expressed as the mg. of phosphorus set free from ^-glycerophosphate at 38°, pH 5*8 in 60 minutes by the phosphatase enzyme in 1 ml. of tissue extract. Alpha amylase* An enzyme extract was prepared from control and MH- treated tissue by extracting 50 g. of Tissue C with 50 ml. of water for .o 1 hour at 25 . Preparation of reagents. 1. A stock iodine solution was made up to contain 11 g. of iodine and 22 g. of potassium iodide in 500 ml. of water. 2. Iodine solution A. To 15 ml. of the stock solution, 8 g. of potassium iodide were added and diluted to 500 ml. with water. 3. Iodine solution B. To 2 ml. of the stock solution, 20 g. of potassium iodide were added and diluted to 500 ml. with water. U. The dextrin solution contained 0.6 g. of Merck*s reagent dextrin made up to 1000 ml. with water. 5. A buffer of pH h*k-hS was prepared with 120 ml. of glacial acetic acid, I61j. g. of anhydrous sodium acetate diluted to a volume of 1000 ml. 32 6. A beta amylase solution was prepared using a commercial pre­ paration of beta amylase so that 5 ml. of solution contained 100 mg* of beta amylase, 7* A buffered alpha araylodextrin solution was prepared from a suspension of 10 g, of Merck*s Lintner soluble starch, 25 ml, of acetic acid-sodium acetate buffer, 5 ml* of beta amylase solution. This solution was brought to 500 ml, with water and then saturated with toluene, 8, A color standard was prepared by pipetting 5 ml, of iodine solution A into a comparator tube and adding 1 ml, of dextrin, solution. This standard should be prepared just prior to using and should not be used longer than four hours. Determination of alpha amylase activity (61;). Twenty milliliters of buffered alpha amylodextrin was placed in a 50 ml, flask containing o 5 ml, of water and the mixture placed in a water bath at 25 • After temperature equilibration, 5 ml. of the extract to be tested was added. At appropriate time intervals 1ml. of the reaction mixture was added to 5 nil* of iodine solution B in a comparator tube, then shaken and compared with the standard* Alpha amylase units are the number of mg, of soluble starch dextrinized by the alpha amylase 0 at 25 in 1 ml, of tissue extract in one hour . . and pH u,!i# Beta amylase. "Extractions of control and MH-treated tissues were con­ ducted as follows: o 1 g. of Tissue A was extracted with 50 ml. of water for 1 hr. at 25 ; 33 £00 mg, of Tissue B was extracted with 50 ml, of water for 1 hr, o at 2£ . Centrifugation of the filtrate yielded a solution which was used for the determination. Preparation of reagents, 1. A buffer of pH l;.U-l4.£ was prepared using 3 ml, of glacial acetic acid, lj,l g, of anhydrous sodium acetate and made up to 1 1 . with water, 2. A N/20 alkaline ferricyanide solution was prepared using l6.£ g, of potassium ferricyanide, 22 g. of anhydrous sodium carbonate and diluting to one liter. This solution was stored in a dark container. 3. An acetic acid reagent containing 200 ml. of glacial acetic acid, 70 g. of potassium chloride, 20 g. of zinc sulfate heptahydrate made up to one liter with water. It. A buffered starch solution consisting of 10 g. of Merck’s Lintner soluble starch, 2£ ml, of acetate buffer pH It.U was brought to £00 ml, with water and saturated with toluene. Determination of beta amylase activity (66, 6?). To 20 ml. of the buffered starch solution in a 12£ ml. Erlenmeyer flask was added sufficient water so that on subsequent addition of the enzyme solution the total o volume was 30 ml. When the flasks and contpnts had come to 2£ , the enzyme solution was added and the hydrolysis allowed to proceed for fifteen minutes or longer. At the end of this time the reaction was stopped by the addition of 20 ml. of one percent sulfuric acid. 3k To a 2 ml. aliquot of the resulting solution was added 10 ml, of N/20 ferricyanide reagent and the tube was immersed in a boiling water bath. After 20 minutes the tube was cooled in cold running water and the contents poured immediately into a 125 ml. flask. The tube was then rinsed with 25 ml. of acetic acid reagent and the rinsings were added to the flask. One ml. of fifty percent potassium iodide solution was added and the contents of the flask were thoroughly mixed. Titration was then carried out with N/20 sodium thiosulfate to the complete disappearance of the blue color of the starch indicator. Beta amylase unit is designated as the number of mg. of soluble starch converted to maltose by the beta amylase of 1 ml. of tissue extract in one hour at 25° and pH In Ii. Pectin-methylesterase. of water at 38 o One gram of Tissue A. was extracted with 30 ml. for one hour, centrifuged, filtered and the resulting solution used for the determination below. Substrate. A 0*5$ citrus pectin solution in 0.1 M sodium chloride was prepared by adding five grams of citrus pectin slowly with stirring to approximately 600 ml. of boiling 0.1 M sodium chloride. was cooled rapidly and diluted to one liter. The solution A drop of toluene was added and the solution stored in the refrigerator until used. Determination of pectin-methylesterase activity (65). equipped with a stirring device was immersed in a 25 bath. o A 250 ml. beaker constant temperature Fifty milliliters of substrate was added and allowed to temperature equilibrate. The pH of the solution was adjusted to 7.0 (glass electrode) by the addition of 0.05 N sodium hydroxide. At zero time 5 ml. of enzyme 35 solution was added and the pH again adjusted to 7.0. The pH was maintained near f by the periodic addition of 0.05 N sodium hydroxide* The amount of alkali added over a definite period of time gives a criterion of enzymatic activity. A unit of pectin-methylesterase is defined as the mg. of methoxyl liberated from pectin by the enzyme contained in 1 ml. of tissue extract in 60 minutes at 25°. Results and Discussion Beta amylase: The results of the investigation of the beta amylase activity of the respective tissue preparations are summarized in Table I. TABLE I EFFECT OF MALEIC HYDRAZIDE ON THE BETA AMYLASE ACTIVITY OF TISSUE PREPARATIONS Beta Amylase Activity* Extraction Solvent Water (3 ml. * 60 mg. Tissue A) Reaction Time (min.) , 15 Water (5 ml. * 100 mg. Tissue A) 20 Water (5 ml. *= 50 mg. Tissue B) 20 Control MH-Treated 59.6 59.6 1*6.5 1*6.2 16.0 17.1 16.0 16.0 58.6 58.0 1*6.5 1*5.8 17.1 18.8 18.8 17.U ^Results expressed as mg. soluble starch converted to maltose by the beta amylase in 1.0 ml. of tissue extract in one hour at 25 and pH l*.l*. Upon inspection of the results in Table I it becomes immediately apparent that the beta amylase activity is essentially the same in both 36 the control and treated tissue preparations. Additionally^ the effect of added maleic hydrazide on the enzymatic activity of a commercial preparation of beta amylase (Nutritional Biochemicals Corporation) of unknown purity was studied* In addition to yielding information as to the effect of maleic hydrazide on the activity of beta amylase in vitro, such a study also yielded a check upon the experimental procedures used in estimating the activity of the enzyme* The results of such a study are given in Table II, TABLE II EFFECT OF MALEIC HYDRAZIDE ON ENZYMIC ACTIVITY OF A COMMERCIAL BETA AMYLASE PREPARATION Maleic Hydrazide (Molarity) Beta Amylase Activity* 36.1 36.1 1 x 10-6 33.9 3lx.2 1 x 10-4 33.7 33.9 1 x 10-3 35-6 35.2 Q (control) Expressed as mg* of soluble starch converteg to maltose by 1,0 mg, of beta amylase preparation in one hour at 25 and pH lulu Here again, it is evident that under the conditions used in this investigation that at concentration of 1CT3 to 10-5 M* maleic hydrazide had no observable effect on beta amylase activity. Alpha amylase: The results of the study of the alpha amylase activity of the tissue preparations can be stated summarily as follows. None of 37 the extracts from any of tissue preparations A, B or C exhibited any detectable alpha amylase activity over periods as long as 90 minutes. Various procedural changes were made but none of the changes resulted in a measurable activity. As a further check on the validity of the experimental procedure a commercial preparation of alpha amylase (Nutritional Biochemicals Corporation) of unknown purity was assayed. Additionally, the in vitro effect of maleic hydrazide on the activity of the commercial alpha amylase was studied. Under the conditions of this investigation, concentrations of maleic hydrazide up to 10~3 M. were without any detectable effect on the activity of alpha amylase. Phosphorylase: Typical data obtained in the study of the phosphorylase activity of the various tissues are given in Table III. TABLE III PH0SPH0RYLASF ACTIVITY OF TISSUE PREPARATIONS Phosphorylase Activity Extraction Solvent Reaction Time (min.) Water (1 ml. * 20 mg. Tissue A) 30 Water (l ml. 33 5 mg. Tissue B) 20 Water (1 ml. * 1 g. Tissue C) 20 Control 87 (blank 83) 89 79 83 (blank 81*) 80 86 (blank 83) 8k MH-Treated 85 (blank 83) 88 82 83 (blank 83) 80 93 (blank 87) 95 ^Results expressed as pg. of inorganic phosphorus liberated from Cori ester by the phosphorylase enzyme in 1 ml. of tissue extract in one hour at 25° and pH 6.1*. 36 None of the various tissue extracts examined exhibited any appreciable amount of phosphorylase activity. The variations in pg. of phosphorus expressed in Table III fall within the limits of experi­ mental error of the procedure which amounts to approximately +1* pg. Various extraction procedures using acetate and succinate buffers were used in an effort to obtain a more active enzyme preparation. The Cori ester solution was increased from 1% to 3% and the starch concentration increased from 1% to %% in the experimental procedure without any appreciable effect. Increasing the volume of tissue extract increased the "blank" phosphorus reading so that it was beyond the upper limit of the method used to determine phosphorus. However, using larger amounts of tissue extract did not give appreciably greater phosphorylase activity. Phosphatase; Typical data obtained in the study of the phosphatase activity of the tissue preparations are given in Table IV, TABLE IV PHOSPHATASE ACTIVITY OF TISSUE PREPARATIONS Extraction Solvent Water (5 ml. = 100 mg. Tissue A) Viater (5 ml* = 25 mg. Tissue B) Water (5 ml. * 5 g. Tissue C) Time (rain,) 0 (blank) 15 30 30 0 (blank) 15 30 0 (blank) 15 30 mg. of Phosphorus * Control .TIH-Treated 0.260 1.53 1.19 1.29 0.365 o.!U±o 0.360 0.565 1.720 1.170 0.2l*2 1.1*7 1.27 1.30 0.380 0.1*1*0 0.360 0.555 i.5Uo 1.21* * Results are expressed as mg. of inorganic phosphorus liberated from ^-glycerophosphate by the action of the phosphatase enzyme in 1 ml. of the tissue extract in one hour at 33 and pH 5*3. 39 Pectin-methylesterase: This pectin enzyme was chosen for study ov^r other possible pectin enzymes because of its simplicity in assaying. No evidence of any pectin-methylesterase activity could be obtained from the tissue preparations investigated initially. Because of the inability to demonstrate activity in the early experiments this enzyme system was not studied in other preparations. However, the validity of the experimental procedure was checked using a tissue preparation from another plant source whose pectin-methylesterase activity had been previously demonstrated. Admittedly, the study of the carbohydrase enzymes outlined above has not been extensive. However, it should be borne in mind that throughout the investigations the main objective was an attempt to find an enzyme system(s) which may have been affected by chemical treatment and in turn could be correlated with growth inhibition. Having demonstrated such a phenomenal would then justify a more exhaustive study of the enzyme system(s) involved. However, the results of the investigation are sufficient to justify the following conclusion. Under the conditions described in this section a preharvest foliar application of maleic hydrazide was without affect on the carbohydrase and phosphatase activity of the radish roots. Additionally, maleic hydrazide did not affect an in vitro system of either alpha or beta amylase. It should be noted that tissue samples were obtained from radishes which had been subjected to two different sets of experimental conditions. In the one instance samples were taken hours after chemical treatment and immediately preserved for investigation. There was no criterion available in this case to indicate whether maleic hydrazide had affected the radishes in any way. In the second case the radishes were harvested 1*0 li8 hours after treatment* topped and held in storage for a period of time• Such tissue exhibited visible evidence of some endogenous difference since the maleic hydrazide treated radishes did not show any appreciable new shoot growth at the shoot stub whereas the untreated radishes did. The changes in biochemical and physiological conditions brought about by this chemical treatment and expressed as growth inhibition may also have existed in the radish at harvest and prior to storage. In the absence of any criterion of difference this remains a moot question. It seemed desirable at this stage of the investigation to concen­ trate further studies on tissue which had been previously treated with maleic hydrazide and stored for a sufficient period of time so that there was some evidence of an effect of the chemical treatment. Effect of Maleic Hydrazide on the Composition of Radishes Preparation and Analysis of Radish Samples Replicate samples containing 25 radish plants averaging from Q to 10 g. per root were collected at appropriate times using care to obtain as nearly equivalent replicates as possible with respect to total fresh weight. Shoot samples were collected 1*8 hours following a foliar spray of a solution containing 2500 ppm (0.02 molar) maleic hydrazide. Root samples were collected also 1*8 hours after treatment but were stored in ventilated polyethylene bags for 168 hours with approximately l/h inch of the shoot portion remaining intact during the storage period. shoot fragment was removed prior to drying. This The fresh weight of the shoot and root portions was obtained separately for each replicate and hi then the samples were dried in a forced air oven at 60° for twenty-four hours • He—weighing the samples yielded the dry weight and from the data the percent moisture was calculated. The samples were ground in an intermediate Wiley mill to pass 60 mesh and used for the analysis to be described below (68). % moisture = fresh weight (lQQ) dry weight Bther extract. Exactly 5.0000 g.of ground tissue was weighed into an extraction thimble and extracted with anhydrous ether in a Soxhlet extractor for sixteen hours. During the extraction the extractor averaged 20 cycles per hour. Following the completion of the extraction, the ether was removed on a steam bath and the residue remaining dried in a vacuum oven over phosphorus pentoxide at 60 o for six hours. The flask and contents were weighed and, knowing the tare weight of the flask, the per cent ether extract was calculated. % ether extract = g. ether extract (100) , 5*0000 g. weight of sample Alcoholic extraction for sugar analysis. The residue remaining in the thimbles after the ether extraction was freed of ether, replaced in the Soxhlet extractor and extracted for sixteen hours with 80$ ethanol. During the alcoholic extraction the extractors averaged fifteen cycles per hour. This extract was used for the sugar determinations described below. Sugars. A 100 ml. aliquot of the 80$ alcoholic extract was transferred to a casserole and evaporated to about 50 ml. on a steam bath. During U2 the evaporation small portions of water were added and the evaporation continued until the odor of alcohol was no longer detectable* time did the volume of liquid fall below 50 ml. water was added and the mass heated to 80°. At no Fifty milliliters of The solution was transferred quantitatively to a 200 ml* volumetric and diluted nearly to volume with water. Three milliliters of saturated neutral lead acetate was added and the flask shaken vigorously and permitted to stand for fifteen minutes. The supernatant liquid was then tested for complete precipitation, diluted to volume and thoroughly mixed. The resulting solution was then filtered into a dry beaker containing one gram of dry sodium oxalate. After shaking and allowing the precipitate to settle, the solution was filtered through a dry paper into a dry flask. Direct reducing sugars. A 50 ml. aliquot of the filtrate and 25 ml. each of Fehling's solution A (Dissolve 3U*639 g* of copper sulfate pentahydrate in water and dilute to 500 ml.) and B (Dissolve 173 g. of sodium potassium tartrate and 50 g. of sodium hydroxide in water and dilute to 500 ml.) were transferred to a 1*00 ml. beaker. The beaker was covered with a watch glass and heated on a Precision electric heater which had been pre-adjusted so that under these conditions boiling would begin after four minutes of heating and continue to boil for an additional two minutes. solution was filtered through a tared Selas crucible. The hot The cuprous oxide remaining in the beaker was transferred Quantitatively with the aid of 80° water to the crucible. The contents of the crucible were then o o thoroughly washed with 80 water, dried at 105 in an oven and the crucible and contents re-weighed to ascertain the amount of cuprous oxide present (aliquot l/8). ii3 Calculate the milligrams of glucose from Munson Walker table* % reducing sugar » g. glucose (8 ) (100 )________ 5.0000 g. dry wt. of sample Non-reducing sugars * A 50 ml. aliquot of the above clarified aqueous sugar solution was transferred to a I4OO ml. beaker. Five milliliters of 12 N. hydrochloric acid was added to the beaker and then covered with a watch glass. Hydrolysis was allowed to proceed for twelve hours with a o laboratory temperature in excess of 25 . After the hydrolysis period had elapsed the solution was neutralized with about 20 ml. of 3 N. sodium carbonate to make the solution neutral to methyl orange. The contents of the beaker were then transferred to a 100 ml. volumetric flask and diluted to volume. The amount of reducing sugar present in a 50 ml. aliquot was determined as described earlier, (aliquot l/l6) Calculate the milligrams of invert sugar from Munson Walker table. (g. of invert % non-reducing * (sugar after — sugar (inversion 5.0000 g. Starch. l/2 g. of invert) sugar before ) (0 .95 ) (16 ) (100 ) inversion ) dry wt. of sample The dried insoluble residue remaining after the 80$ alcoholic extraction was transferred to a 300 ml. flask* Sixty milliliters of waterwere added in 10 ml. portions and the mixture stirred to assure complete wetting of the mass as well as complete dispersion. The mixture was then heated to boiling in a water bath for one hour with occasional shaking. After cooling, 100 mg. of malt diastase was added to the flask followed by two milliliters of toluene and the mass incubated for 2ii hours at 38°. In order to prevent the dispersed material from settling, kh the flask was shaken from time to time and any residue on the sides of the flask was washed down with water. After incubation, the en­ zymes were inactivated by heating in a boiling water bath for fifteen minutes. The solution was diluted to a volume of about 90 ml. and 160 ml. of 95% ethanol were added in 2$ ml. portions with thorough shaking between additions. This procedure precipitated gums, pectins and other interferring polysaccharides. After the solution had at­ tained room temperature, it was filtered into a $00 mi^ volumetric flask and washed to volume with 60% ethanol (by volume). A 100 ml. aliquot was freed of alcohol by evaporation on a steam bath accord­ ing to procedure as described under the sugar section above. The volume was adjusted to approximately 7$ ml. and $ ml, of 12 N, hydro­ chloric acid was added and the hydrolysis carried out in a water bath at 80 o for three and one-half hours. After cooling, one milli­ liter of a 10% phosphotungstic acid in 1% hydrochloric acid solution was added and the solution mixed and permitted to stand for fifteen minutes. filtered. The solution was then diluted to a volume of 100 ml. and Fifty milliliters of the filtrate was neutralized with about 10 ml. of 3 N. sodium carbonate to a methyl orange endpoint. The resulting solution was diluted to a volume of 100 ml. and a $0 ml. aliquot used to determine the glucose present according to the procedure described earlier, (aliquot 1/20) Calculate the milligrams of glucose from Munson Walker table. % starch * g* glucose (20) (100) (0.90) 5.000b g. dry wt. of sample-- U£ Polysaccharides. The insoluble residue from the starch determ­ ination was transferred to a 300 ml* wide mouthed flask and 100 ml* of water was added* Ten milliliters of 12 N. hydrochloric acid was added and the flask heated in a boiling water bath for two and one-half hours with intermittent shaking* After cooling, one milliliter of 10$ phosphotungstic acid in \% hydrochloric acid was added andthe contents thoroughly to stand for fifteen minutes* mixed and permitted The resulting solution was then diluted to £00 ml. in a volumetric flask. (Three milliliters of water in excess of £00 ml* was added to allow for the pres­ ence of the insoluble material present). A 100 ml. aliquot was removed and neutralized to a methyl orange endpoint with about 8 ml* of 3 N. sodium carbonate. The solution was made to volume in a 200 ml. volumetric flask and the reducing sugar determined in a £0 ml* aliquot, (aliquot 1/20) Calculate the milligrams of glucose from Munson Walker table. % polysaccharides* g. of glucose (20) (0.9) (100) £.0000 g* dry wt. of sample Nitrogen. The semi-micro Kjeldahl method was used for the determination of nitrogen* Thii’ty to forty milligrams of sample tissue was digested in a Kjeldahl flask with 3 ml. of concentrated sulfuric acid and approximately 100 mg. of a catalyst consisting of one part potassium sulfate and three parts copper sulfate. Digestion was continued for h6 eight hours* The samples were distilled into excess standard acid and the excess acid determined by titration with standard base using a methyl purple indicator. % nitrogen « (ml. of O.QQ97 N. HCl) (Q.Qll;) (100) g. weight of sample Results and Discussion The results of the analysis for gross nutritive constituents of radish root and shoot tissues are given summarily in Tables V and VI. These tables are self explanatory and require only a few comments* In the instance of the root analysis (Table V), the chemical treatment has had little effect upon the plant as revealed by these analyses. The combined reducing and non-reducing sugars in the treated tissues were about one per cent higher than that in the corresponding control tissue. There was some difference in starch content, treated samples being somewhat less, but on an absolute basis this difference was small and probably not significant. The data on the analysis of the shoot tissue (Table VI) reveal that maleic hydrazide treatment has had no large effect upon the gross constituents. It should be noted that the starch content of the treated tissue was double that of the corresponding controls. From a nutritional aspect it is of interest to note here that root and shoot samples were also analyzed which were obtained 216 hours k7 OUTLINE OF THE METHOD OF CARBOHYDRATE ANALYSIS Dry plant tissue ground to pass 60 mesh (£*0000 g*) Extract 16 hrs, in a Soxhlet extractor with anhydrous ether Residue Ether extract Extract 16 hrs. in a Soxhlet extractor with 80$ ethanol Residue Reducing sugars Disperse in water Incubate 2h hrs, at 38° with malt diastase Residue Acid hydrolysis 2 1/2 hrs. at 100° Residue Acid hydrolysis Non-reducing sugars Acid hydrolysis Starch Acid hydrolyzable polysaccharides hS 43 bD US • P t 8 0) TJ 03 P ■ 8 •rH P 05 to P c tf bD P 10 bD G o ■3 (S bD P CO •G 0H TG O 03 P 1 C o 2£ P0J o o ■>8s*CJ iR H a P P 0} O p rH os bo p : o p n} P 03 snJ s , p o P 03 O S *H E P Q) p 43 c0 TO P EO h k9 & 2ID T5 > (1) P < 00 ) on •8 •H THE EFFECT OF A PREHARVEST FOLIAR MALEIC HYDRAZIDE ON THE COMPOSITION SPRAY (F OF RADISHES CO £ & O iH to rH 0) ( od p •H r~i CM ft 0) Pi 3 vO Ov VO O OO • * vO ON co vO a Ov U \ • p j ON on 9 UN 0 CO r- vO PT • rH • p t • U \ OO c*c-- P t vO U N • rH • CM CO vO • {A un U N CM UN • UN O « CM r-~ Ov on ON UN PT * O UN • ON CM Pi • O CM CM • ON CM PJ • vO Ov O Pf • rH vO • ON 00 P t M3 • ON P t UN • vO CM O • P t CM CO • UN 0vO O P— O • ON UN UN * vO CM rH • Pt P t e• UN CM vO • O rH O • ON rH r• CO P t • UN Os • ON vO ON CO * O CO Ov vO CO Pf UN • O • vO• p xz bD *rt a 6t f (D ,0 *H P rH O XJ 50 following treatment. This period corresponds to the period of U8 hours after treatment plus the 168 hours of storage used previously for the samples analyzed. These treated root tissues had less reducing sugar, more, non-reducing sugar and slightly more than one per cent more total reducing and non-reducing sugar than the controls. The ether extrac­ tives of the treated samples were also about one per cent less than the corresponding control. Per cent dry weight of the treated samples was approximately one per cent higher. Analysis of the shoot tissue (216 hours after treatment) showed that the treated tissue had a higher dry weight and lower nitrogen content than the controls • The reducing sugars of the treated samples were double that of the untreated samples. The control tissue gave no reducing sugar but the treated tissues did contain less than one per cent. Additionally, mineral analyses were obtained on samples equivalent to those used for gross constituent analysis. These results, obtained spectrographically, did not reveal any marked difference in mineral content as a result of maleic hydrazide treatment. Effect on DPNH ^ ~ of Maleic Hydrazide -- — —p - the ■- TiI Oxidase Activity of Radishes Introduction The almost complete inhibition of cellular respiration by agents which interfere with the normal functioning of cytochrome oxidase or of other cytochrome components of the respiratory chain indicated the obli­ gatory role of these enzymes in the production of biosynthetically use­ ful energy. Although this is only one of a number of systems involved 51 in the transfer of electrons to molecular oxygen it is generally be­ lieved to attribute substantially to the sum of the total electrons transferred in biological systems. It has long been known for many years that DPN is necessary for the oxidation of a number of intermediary metabolites by tissue* The 1 DPN is reduced by the metabolite, activated by a specific dehydrogenase, 1 to DPNH* There are two well known ways whereby the DPNH may be oxidized to reform DPN, (a) by the anaerobic reaction with a substrate (S): S ♦ DPNH — ^ DPN 4 SH2 which requires a single enzyme, SH3 dehydrogenase and (b) by the aerobic reaction: H+ + DPNH + 1/2 Oj, — > DPN + H20 which requires a complex of enzymes which may be called DPNH oxidase system, A less well known system involved in the possible oxidation of DPNH is: DPNH + fumarate — > DPN + succinate. This reaction was demonstrated indirectly by Dewan and Green (69) who found an oxido-reduction system linking # -hydroxybutyrate, which is a typical DPN-requiring dehydrogenase and succinic dehydrogenase, which does not require DPN* It has been shown by Slater (70) that this is a slow reaction and it is doubtful whether it is of great quantitative importance in vivo. XDPN and DPNH are abbreviations used for the oxidized and reduced forms respectively of diphosphopyridine nucleotide. 52 Another system which has been described recently which may also be involved in the oxidation of DPNH is a system functioning with the aid of ascorbic acid, or more likely an oxidation product of ascorbic acid. Such a system was first observed in green peas by Mathews (71) and more recently found in cucumbers by Beevers (72). As yet very little is known of this system and its importance in the oxidation of DPNH re­ mains to be evaluated. Slater, (70, 73, 7h) on the basis of his experiments, suggested that the DPNH oxidase system consisted of the following components, the arrows denoting the pathway taken by the hydrogen atoms or electrons on their way from substrate to oxygen. Cytochrome b — > succinate DPNH— ^ Diaphorase— > Factor— ?> Cytochrome c— Cytochrome a— ^ Cytochrome a3 — X>2 Methylene blue -- > 02 The addition of methylene blue enables the reaction to by-pass all the x components except diaphorase. The factor is the BAL-sensitive (Slater factor or antimycin sensitive factor) component of the system and is probably the same as the factor previously found necessary for the oxidation of succinate. This proposed scheme does not, of course, exclude the possibility that further components, not yet identified, may also be required. It is known that the reactions between succinate and cytochrome b is easily reversible. If the reaction between cytochrome b and the factor is also reversible to a certain extent, the slow reduction of XRAL is a common abbreviation for British Anti-Lewsite (2,3-dimercaptopropanol)• 53 cytochrome b by DPNH and the anaerobic oxidation of DPNH by fumarate are understandable since in the absence of oxygen and the presence of fumarate the reaction procepds as follows: ■DPNH-- > Diaphorase— > Factor— > Cytochrome b— > fumarate. The system designated as the DPNH oxidase system then can be re­ solved into the following enzyme components: a) diaphorase, a flavoprotein catalyzing the direct oxidation of DPNH. b) DPNH-cytochrome c reductase, which mediates the oxidation between DPNH and cytochrome. c) cytochrome c oxidase, which facilitates the oxidation of reduced cytochrome c. Although there is some question as to what entities constitute DPNH-cytochrome c reductase, these entities affect the enzymatic re­ moval of hydrogen from DPNH with the transfer of electrons to cyto­ chrome c. Thus, in keeping with the general system as defined by Slater and outlined above, DPNH-cytochrome c reductase will be con­ sidered in this thesis to include that activity >of diaphorase together with the essential elements which accomplish the oxidation of DPNH and the reduction of cytochrome c. Preparation of DPNH Oxidase System The radish plants, which were used as a source of biological material in this investigation, were treated with a 2!?00 ppm solution of maleic hydrazide (0.02 M) applied as foliar spray. taken 2*8 hours after the treatment described above. Samples were Shoot samples were 5U immediately frozen and stored until used* Topped radish roots were stored for 168 hours at greenhouse temperature. Following storage, the fleshy roots and shoot stubs were separated and preserved by freezing. Samples of both treated and untreated radishes were pre­ pared in a similar manner* Tissue extracts for the study of the respiratory enzymes were prepared by a method similar to that used by Keilin and Hartree (75) for the extraction of cytochrome c oxidase* Essentially this pro­ cedure had been used by Slater and others for the preparation of DPNH oxidase system in animal tissues* The procedure consisted of extraction with phosphate buffer and precipitation of the enzymes by acidification with acetic acid. Schneider and Hogeboom (?6) have observed that the majority of cytochrome oxidase activity in animal tissue was found in the mito­ chondrial cell fraction. These investigators (77, 78) have also shown that the presence of DPNH-cytochrome c reductase is not restricted to any definite cell structure* The greatest amount of this enzyme was found in the mitochondrial and submicroscopic particles; some being present in the nucleus. DPNH Since the individual enzyme components of the oxidase system were found in various sedimentable and non- sedimentable fractions, only a general tissue preparation was studied and no attempt was made to analyze cell fractions which might be ob­ tained by differential centrifugation methods. It was realized that using an extraction procedure originally designed for animal tissue might involve some problems when plant tissue was used. Of particular concern here would be such problems 55 as buffer concentration, type of buffer and hydrogen ion concentration. Whatever influence these and other factors might have, the modified procedure of Keilin and Hartree did, indeed, yield an active DPNH oxidase system when applied to plant tissue. Whether the conditions used yielded the most active preparation possible is not known. If absolute enzyme activities were the prime purpose of the investigation this problem would be of upmost importance. However, when enzymatic activity of the preparations obtained under like conditions are com­ pared, then this problem is minimized. The extraction procedure varied depending upon the kind of tissue used. The particular conditions associated with special plant organs were arrived at generally by the trial and error methods. Extraction of root tissue; Thirty grams of frozen tissue was macerated with 90 ml. of cold 0.02 M phosphate buffer pH 7*3 for four minutes in a pre-cooled blendor. The resulting slurry was strained through four thicknesses of cheese cloth and the solution centrifuged in the cold at 2000 rpm for five minutes. The centrifugate was collected and the residue re-extracted with U5 ml. of cold 0.02 M phosphate buffer. Centrifugation was repeated and the second centrifugate added to the original centrifugate and refrigerated if necessary to bring the temperature to 0-k°* The pH was then adjusted to 5.5 by the addition of cold 0.1 N acetic acid and the solution permitted to stand for one hour at about 0°. Centrifugation for five minutes at 2000 rpm yielded a residue and a centrifugate. The centrifugate was held at U° in the refrigerator while the residue was dispersed in 90 ml. of cold 0.02 M 56 phosphate buffer and the acid precipitation repeated. The resulting residue was homogenized with Ii5 ml. of cold 0.1 W phosphate buffer pH 7*3* This solution was stored at I). (Solution A). The combined centri­ fugates from the acid precipitations were held at U° for 12 hours and centrifuged again. The resulting centrifugate was discarded and the residue homogenized with Solution A. necessary, at li°. This solution was stored, when All enzymatic studies using this and all other preparations were conducted within U8 hours of preparation. During the preparation of the solution of enzymes great care was exercised to keep the extraction temperature below 10° at all times, (l ml. of extract was equivalent to 2/3 g. fresh tissue). Extraction of shoot tissue; The enzymatic preparation was obtained from the leaf tissue in essentially the same way as described for the root tissue. However, in the case of the leaves, the presence of large amounts of chlorophyll was a disturbing factor. changes in the extraction procedure. This necessitated some minor Since the procedure was nearly the same as outlined above only the changes will be described in this section. Fifteen grams of frozen leaf tissue was macerated with 90 ml. of phosphate buffer. This solution was then permitted to stand for one hour at h° prior to centrifugation. All centrifugations were conducted for ten minute intervals at a speed of 2000 rpm. The final preoaration was not completely free of chlorophyll but since blank solutions of the enzyme preparation were used consistently the presence of chlorophyll was not a prohibitive factor. fresh tissue). (l ml. of extract was equivalent to 1/3 g. 57 OUTLINE OF PROCEDURE FOR PREPARING DPNH-OXIDASE SYSTEM Frozen tissue Homogenize li min. with cold 0.02 M phosphate buffer pH 7*3 Centrifuge in cold at 2QQQ rpm. RESIDUE 1 *! CENTRIFUGATE 1 Homogenize with 1/2 volume of original buffer. Centrifuge at 2000 rpm. Residue (discard) CE'NTRIFUGa TE 2 (Combine Centrifugates 1 and 2) Adjust pH to^5*5 with cold 0.1 N acetic ac^d. Let stand 1 hr. at 0 , Centrifuge in cold at 2000 rpm. CENTRIFUGATE Ac 3 RESIDUE 2 Disperse in original volume of cold phosphate buffer. Centrifuge at 2000 rpm.____ CENTRIFUGATE h Ac Residue (discard) Adjust pH to^5.5 with cold 0.1 N acetic acid. Let stand 1 hr. at 0 . Centrifuge at 2QQQ rpm. I RESIDUE 3 CENTRIFUGATE 3 (Combine centrifugates 3 and It) Disperse in 1/2 original volume of 0.1 M phosphate buffer pH 7 *3• I o Let stand 12 hrs. at h • Centrifuge at 2000 rpm. RFSID uA: SOLUTION A Homogenate with SOLUTION A Store at u DPNH OXIDa SE PREPARATION Centrifugate (discard) _ 58 Extraction of Hshoot stub*1 tissue: Again, the extraction procedure was essentially the same as used for the root tissue extraction. variations from the outlined procedure will be mentioned. Only Seven and one-half grams of frozen wshoot stub*1 was maceratpd with U5 ml. of cold 0.02 M phosphate buffer pH 7.3. From this point on the procedure was exactly as described above except that the volumes used were one-half of that described in the section on root extraction. was contaminated only slightly with chlorophyll. The final preparation (1 ml. of extract was equivalent to 1/3 g. fresh tissue). Analytical Methods The evaluation of the activity of the total DPNH oxidase system and of its component parts were conducted spectrophotometrically. The procedures used were dependent upon the spectral characteristics of both oxidized and reduced diphosphopyridine nucleotide and oxidized and reduced cytochrome c. The spectra of these compounds are as follows (79): 20 DPN 15 10 DPNH 0 260 300 3U0 380 A - my-. h20 59 These spectra are typical of pyridine nucleotides in the oxidized and reduced form. Thus the oxidation of DPNH can be readily followed by noting the decrease in optical density at 3^iD mp. f. -Reduced cytochrome c 12 10 aX M Oxidized _ cytochrome c— , v 600 Reduced cytochrome c exhibits an absorption maximum at 550 mp. which is not true of oxidized cytochrome c. Thusly the oxidation or reduction of cytochrome c can be followed by observing increases or decreases in optical density at 550 mp. Preparation of special reagents. Reduced diphosphopyridine nucleotide: Reduced DPN was prepared by a modification of a procedure described previously by LePage (80). To a 16 x 150 mm. test tube was added 5 mg. of DPN (Sigma Cozymase - ”90" from yeast) dissolved in 0.6 ml. of water and 1.0 ml. of freshly prepared 1.0$ sodium bicarbonate. Two milliliters of a freshly prepared 3.0$ solution of sodium hydrosulfite in 1.0$ 60 sodium bicarbonate were prepared in a small test tube with cautious shaking to avoid aeration (60 mg. of sodium hydrosulfite dissolved in 1.0$ sodium bicarbonate and made to a 2.0 ml. volume). An 0.1*0 ml. aliquot of sodium hydrosulfite solution was added to the tube containing the DPN sample, mixed gently and allowed to stand twenty minutes at room tenperature without further agitation. To the sample was added 18.0 ml. of a 1.0$ sodium bicarbonate - 1.0$ sodium carbonate (v/v). The mixture was oxygenated for five minutes to remove the excess hydrosulfite. The DPNH solution was neutralized to pH 7-3 by the dropwise addition of an 0.5 M potassium dihydrogen phosphate before use in the enzymatic studies. The nucleotide solution could be stored under refrigeration before neutralization without any appreciable oxidation; however, at pHTs near neutrality, the solution oxidizes slowly even at li°. Such a solution assayed approximately 10 4 molar using the molar extinction coefficient of DPNH at 3^0 mp. of 6.22 x 106 sq. cm. per mole (8l). Oxidized cytochrome c: A 10 - 4 molar solution of cytochrome c was prepared by dissolving 16.5 mg. of cytochrome c (Sigma cytochrome c, based on a molecular weight of 16,500) in 0.005 N hydrochloric acid and diluting to a volume of 10 ml. Such solutions were prepared just prior to using. Reduced cytochrome c: A 10~4 molar solution of reduced cytochrome c was prepared by dissolving 16.5 mg. of cytochrome c (Sigma cytochrome c, based on a molecular weight of 16,500) in 0.10 M phosphate buffer pH 7.3 and diluted to nearly 10 ml. with additional buffer. Reduction was accomplished by adding an amount of sodium hydrosulfite sufficient to give a concentration that is 0.001 M. The preparation 61 was then aerated by shaking for 3 to 5 minutes to oxidize the excess reducing agent. Auto—oxidation was minimized by keeping the phosphate buffered cytochrome c solution at h° (82). Determination of total DPNH oxidase activity. The total system contained the following components added to the absorption cell in the order listed: 0.5 ml. of enzyme solution, 1.5 ml. of glass distilled water, 1.0 ml. of DPNH solution. The blank contained 1.0 ml. of glass distilled water in place of the DPNH solution. Upon addition of the DPNH solution the contents of the cell were mixed and the decrease in optical density at 3h0 mp was obtained. Determination of diaphorase activity (83). The components of the system employed for the measurement of diaphorase activity consisted of 0.5 ml. enzyme solution, 0.3 ml. of 2 x 10 3 M potassium cyanide, 0.1 ml. of 5.3 x 10”4 methylene blue, 1.1 ml. of glass distilled water and 1.0 ml. of DPNH solution. The blank contained the same components except for water in place of DPNH solution. The oxidation of DPNH was followed by observing the optical density decrease at 3-U0 mp. Determination of DPNH - cytochrome c reductase (77). The reaction mixture —g (volume 3.00 ml.) contained 0.5 ml. enzyme solution, 0.3ml. of 2 x 10 M potassium cyanide, 0.8 ml. distilled water, 1.0 ml. of DPNH solution, and O.li ml. of oxidized cytochrome c. except DPNH. The blank contained all components The reaction was followed by recording the rate of increase in optical density at 550 mp. 62 Det.erminat.iQn of cytochrome c oxidase. spectrophotometrically at 25 Schneider (810, Cytochrome oxidase was determined by the general procedure of Hogeboom and The reaction mixture (volume 3.00 ml.) contained 0.5 ml. of enzyme solution, 1.0 ml. of 0.1 M phosphate buffer pH 7.3> 0.8 ml. distilled water, 0.3 ml* of 1* x 10 3 M aluminum chloride and added last 0.1* ml. of 10 4 M reduced cytochrome c. all components except cytochrome solution. The blank solution contained The reaction was followed for six minutes at one minute intervals by noting the decline in optical density at 5^0 mp. Measurement of enzyme activities. All spectroohotometric studies were carried out using a Beckman DU spectrophotometer equipped with a temper­ ature control device so that the cells and contents could be maintained at 25°> the temperature at which all determinations were made. Matched cells of one centimeter light path were used exclusively throughout. All solutions used were temperature equilibrated prior to use. All zero time optical density readings were obtained within 15 seconds following the addition of the last component of the system and tnorough mixing. Optical density readings, obtained at one minute intervals for six minutes, are given on the immediately succeeding pages. Duplicate read­ ings were obtained on enzymatic preparations from two replicate tissue samples. These optical density readings are summarized graphically in Figures 1 through 7 • 63 DPNH-OXIDASE ACTIVITY Extinction x IQ3 at 380 mp.* in. _______Control____________ Replicate 1 Replicate 2 MH-Treated Replicate 2 teplicate 1 0 612 638 562 560 590 595 518 525 1 603 630 553 550 577 589 5o8 516 2 593 ,619 581 539 565 578 897 503 3 580 608 530 530 555 567 887 898 8 569 592 517 5i5 588 552 875 883 5 556 579 5o5 507 538 538 868 877 6 582 568 898 1*95 528 526 858 863 0 538 5o5 708 68.0 887 582 587 585 1 528 1*95 702 633 883 538 585 583 2 522 890 698 629 880 536 583 581 3 518 886 692 621 837 533 581 578 8 512 881 688 615 833 531 577 578 5 508 hlS 682 609 829 527 575 570 6 502 bn 673 603 826 525 873 568 0 500 510 899 5oo 880 883 873 868 1 895 505 893 898 876 881 875 867 2 891 502 887 890 873 880 872 865 3 887 898 883 886 872 878 871 863 8 888 898 879 882 870 877 868 861 881 891 876 879 869 875 867 860 5 6 878 888 873 875 868 873 865 858 61* DIAPHORASE ACTIVITY Extinction x IQ3 at 3l*Q m**. in* Control Replicate 1 Replicate 2 _______ ivIH— Treated________ Replicate 1 Replicate 2 0 557 568 563 520 561 560 1*15 1*91 1 539 51*5 51*3 1*98 539 536 388 1*71 2 512 528 511 1*85 5io 512 359 1*1*6 3 1*91 510 1*93 1*72 1*97 1*90 339 1*32 1* 1*71 U93 hi9 1*65 1*79 1*82 329 1*18 5 1*57 1*80 hi3 1*1*9 1*70 1*72 325 1*02 6 1*1*7 1*67 1*62 1*25 1*62 1*60 310 387 0 620 615 661 61*5 612 621 61*2 638 1 571 565 616 60l* 570 576 581* 591* 2 529 52i* 577 560 530 537 557 55o 3 1*96 h90 51*6 528 1*98 508 528 520 I* h lh 1*81 527 507 1*78 1*89 506 505 5 1*59 510 1*91 1*60 1*72 1*98 1*93 6 1*1*8 1*57 502 1*82 1*51 1*63 1*92 1*87 0 573 576 580 593 579 571* 560 561* 1 562 569 57U 583 576 571 557 560 2 552 558 562 577 570 566 552 556 3 51*6 551 555 572 565 559 51*6 552 1* 510* 5U9 551 561* 562 558 51*3 51i? 5 539 51*3 51*1* 557 555 552 538 5)*2 6 535 535 537 51*9 51*8 51*1* 531* 536 65 CYTOCHROME C OXIDASE ACTIVITY Extinction x 103 at 550 mu. in. _______Control____________ Replicate 1 Replicate 2 MH-Treated________ Replicate 1 Replicate 2 0 260 266 215 257 271 269 28a 265 1 2a8 256 203 2aa 261 258 27a 251 2 236 2Uh 191 23a 2a7 2aa 258 238 3 226 232 175 222 233 231 2a5 226 U 213 218 162 210 220 220 233 216 5 203 206 153 202 208 210 221 206 6 19a 196 ia a 190 198 201 211 19a 0 2a5 2a 9 301 303 2a7 257 253 265 1 235 2ao 292 29a 2a3 251 2a6 260 2 227 232 283 286 236 2a6 2ai 253 3 222 227 279 281 231 2a2 236 250 a 216 221 27a 27a 226 238 232 2a6 5 211 216 266 270 222 235 226 2ao 6 206 212 260 263 218 232 222 237 o 288 280 288 270 271 268 2a8 27a 1 285 277 285 267 268 265 2A6 272 2 281 27a 282 26a 265 261 2a5 269 3 278 270 280 260 263 258 2a3 266 a 276 268 277 258 261 256 2a 2 26a 27a 265 275 256 257 258 239 262 5 6 271 262 273 253 256 253 236 260 66 EFFECT OF ADDED CYTOCHROME C ON ABSORPTION AT 55O mp. Time min. _____________ Fxtinction x 103 at 990 mp. Control Replicate1 Replicate 2 MH-Treated_____ Replicate 1 Replicate 2 Root: 0 b 7 0 3 9 3 2 0 1 16 21 III lb 19 12 11 10 2 28 30 28 29 2b 33 21 19 3 39 b2 10 bb 36 b3 33 32 b 92 99 *1 96 b9 93 bb b7 9 66 70 63 69 99 69 92 97 6 77 8b 72 81 68 7b 63 69 0 b 3 b3 30 2 3 1 0 1 15 13 9b bo 9 10 9 8 2 26 23 67 92 16 16 18 19 3 36 33 77 61 22 22 26 22 b bb b2 87 68 27 26 3b 28 5 9i 50 99 76 31 32 bO 33 6 £8 97 100 83 39 38 b7 38 0 0 2 -9 0 -2 1 -8 b 1 b 6 -9 b 2 9 -b 8 2 8 10 -2 7 9 9 0 11 3 12 13 0 10 8 12 3 19 b lb 16 b 13 12 19 9 18 9 17 20 7 16 19 18 8 20 6 20 22 10 18 17 20 11 22 Shoot: Shoot Stub: 67 EFFECT OF ADDED CYTOCHROME C ON DPNH OXIDASE ACTIVITY Time min. ______ E xtinction x 103 a t 3bO mp.. Control R eplicate 2 Replicate 1 MH-Treated Replicate 2 Replicate 1 Root: 0 571 566 $08 512 591 570 $02 $10 1 $62 556 1*97 502 $81* $62 l*9l* 503 2 51*9 5U5 1*8$ m 571 552 1*8$ U9U 3 538 537 h7h 176 557 538 1*77 1*83 1* 530 $21* U63 1*61* $1*9 530 1*71 1*77 5 521 517 1*51* i*$6 $1*0 $21 1*6$ U6k 6 $08 5o5 1*1*3 hh3 531 512 1*1*7 U52 0 39k 39k 63k 6k0 1*10 1*11* 598 $28 1 383 381* 62$ 630 i*o$ 1*08 589 $20 2 375 376 617 621 l*oo 1*03 $81* 5lU 3 368 370 610 61$ 395 398 579 $10 1* 361 366 602 610 390 39U 57 U $06 5 356 360 597 60$ 387 390 568 5oi 6 353 358 590 $98 385 386 $61* 1*98 0 530 528 532 $2l* $10 $08 1*7$ 1*70 1 525 522 528 $20 $o6 $01* 1*71 1*66 2 518 515 522 $17 $02 h99 1*68 1*63 3 512 511 520 5io h99 U96 1*6$ 1*61 1* $08 $08 517 $08 h97 h93 1*62 1*59 5 503 $02 513 $05 h93 1*90 1*60 1*57 6 k99 1*98 $10 $01 k91 1*88 1*58 1*55 Shoot: Shoot Stub: 68 EFFECT OF ADDED METHYLENE BLUE ON DPNH OXIDASE ACTIVITY E xtinction x 10a a t 3i|0 mp.. Time min» Control R eplicate 2 Replicate 1 MH-Treated Replicate 1 Replicate 2 Root: 0 563 560 511 510 588 552 518 520 1 556 58o 892 890 519 525 890 896 2 526 5i8 871 871 898 500 869 873 3 508 5oi 858 851 876 888 852 853 h 891 888 837 835 859 871 838 880 * 873 870 822 826 886 862 815 820 6 858 857 811 509 880 853 809 810 0 879 875 626 680 585 895 598 612 1 859 850 611 623 533 882 588 603 2 839 829 598 606 521 878 580 598 3 1(20 801 575 582 512 865 578 588 h 806 393 559 568 502 857 565 573 $ 392 381 580 888 892 851 558 567 6 381 369 526 530 883 886 583 563 0 863 857 855 887 882 850 850 81(2 1 853 888 888 839 838 8I16 885 838 2 888 881 881 832 832 838 881 832 3 880 835 837 828 828 833 837 829 k 833 829 830 822 823 828 831 825 828 822 828 817 819 822 828 820 i 6 820 818 819 810 812 818 822 815 Shoot:•• Shoot Stub: % ■ 69 DPNH-CYTOCHROKE C REDUCTASE ACTIVITY Extinction x IQ3 at 550 in, _______Control_________________ MH-Treated___ Replicate 1 Replicate 2 Replicate 1 Replicate 2 0 5 7 17S 200 2 h 167 152 1 17 18 182 209 12 17 180 162 2 26 27 197 220 2b 2b 191 171 3 36 39 205 233 3h 3h 198 182 b b5 hi 21b 2l;2 hi hh 20b 192 £ 53 56 222 2b9 52 51 215 200 6 60 65 227 256 60 62 223 207 0 0 2 0 3 1 1 -1 0 1 10 12 9 13 8 10 7 7 2 18 18 16 22 lb lb 13 lb 3 28 26 2b 29 18 20 19 20 b 36 32 29 36 2b 2b 25 2b 5 bo 35 35 b2 28 30 30 30 6 5o bb bl b8 32 32 3b 3b o 5 b -I 1 -1 3 -b 2 l 7 6 2 3 1 5 -1 b 2 10 9 b 6 3 8 2 7 12 12 6 8 5 10 b 10 3 b lb 15 8 10 7 12 6 12 15 17 10 12 8 13 7 13 5 6 17 20 12 15 10 15 9 15 70 Results and Discussion Cells are capable of oxidizing, with considerable ease, a great many types of metabolites many of which are quite stable in air. The enzymes that participate in these energy-yielding reactions are generally called ^respiratory1* enzymes. The sequence of reaction going from sub­ strate to oxygen may be illustrated diagramatically as follows, the arrows indicating the direction of transfer of hydrogen or electrons (79). Substrate —> pyridine nucleotides— > Flavoproteins — > Cytochromes -> — > Oxygen Not only are cells capable of producing energy through the "respiratory" enzymes, but they also possess highly organized systems which operate to trap the energy that is released and subsequently make the energy available to the cells for energy requiring responses. The energy is trapped by the action of multienzyme systems. A multienzyme system may be defined as an organization of enzymes which catalyzes an orderly sequence of reactions to bring about the con­ version of a substrate to the desired product. Multienzyme systems must be approached with some skepticism, although they have proven of some value in revealing details as to how certain metabolic processes occur. When intact tissue is homogenized, a number of factors influence the resulting enzymatic activity observed and hence the nature of the reaction sequence in such preparations. Some of these more obvious factors are: (a) dilution effects resulting from the suspending medium used during homogenization, (b) inhibiting effects related to the liberation of sub- 71 stances through cell fragmentation, and (c) destructive effects such as autolytic processes which may modify the structure of the enzyme proteins and alter their enzymatic activities. Additionally, the instability of the enzymes, substrates and co-factors, the actual concentration of each of these components, the mutual affinities of substrates and co-factors for the enzymes, pH, temperature and salt effect, the introduction of spatial separation of enzyme from substrate and co-factors and the possible influence of hormonal control on enzyme systems must be considered. The only well established effective means presently known for the coupling of energy production with energy utilization by a multienzyme system is the esterification of inorganic phosphate into the adenylate system to form adenosine triphosphate. Adenosine triphosphate ma y b e readily transferred to energy-requiring systems. The esterification of inorganic phosphate can take place at two levels: level. (a) the substrate level and (b) the electron-oxygen transfer The substrate level phorphorylation has been studied in some detail and is characterized by the formation of adenosine triphosphate in absence of oxygen by reactions catalyzed by water soluble enzymes. The electron-oxygen transfer system involves the passage of electrons from the substrate through several electron carrier systems to the final acceptor, oxygen. Relatively large amounts of utilizable energy become available and are trapped also in "high energy” phosphate bonds\ 1The terra "high energy" has been applied to such bonds to indicate that they release a relatively large amount of energy when they are broken and not that there is a strong bonding energy between the phosphate group and the group to which it is attached. 72 depicts the sequence for the transfer of electrons to oxygen as: Substrate * 2H— > Pyridine nucleotides (2e+H+)— > Flavin system (2e+H )— > Cytochrome system (e)— >oxygen. At each level of the electron-oxygen transport system significant quantities of utilizable energy are theoretically available* In the oxidation of DPMh a theoretical value of four ’’high energy" phosphates is possible. Lehninger (86) has determined experimentally that in the oxidation of DPNH by a mitochondrial preparation, three "high energy" phosphates are formed for the transfer of the two electrons from DPNH to oxygen. These results would indicate a 75 per cent trapping efficiency. The electron-oxygen transport system is characterized by its instability, oxygen uptake and existence in the mitochondria. Respiration, that is the oxidation of metabolites with the subsequent production of biologically useful energy, is affected by multienzyme systems residing within the organism. aerobically or anaerobically. Respiration may occur either Although both types of respiration do occur, certainly in higher plants anaerobic respiration can not take place at the exclusion of aerobic respiration, at least for very long periods of time. Aerobic respiration is associated with the electron-oxygen transport system mentioned earlier and is a vital process for higher plants. The cytochrome system is generally believed to play a predominant role in aerobic respiration. However, It is true that the extent of the participation of the cytochrome system is not known. A large body of evidence has been accumulated which indicates that the cytochrome system is associated with a large part of the aerobic respiration. 73 One of the phenomena generally assumed to be associated with growth is respiration. However, it is now known that the proportion of res­ piratory energy actually utilized in growth, that is, in cell division and increase in size, is small (56, 57). Although there is no direct correlation between growth and respiration, some respiration is un­ questionably essential for growth. Surely a partial or complete inhibition of respiration through lack of oxygen or respiratory poisons is accompanied in plants by the cessation of growth. Conversely, if a substance such as maleic hydrazide can bring about cessation or inhibition of growth, it seems reasonable then to suspect that the respiratory system(s) may have been affected by chemical treatment. As mentioned earlier, the cytochrome system functions in the electronoxygen transport system which in turn functions to produce biosynthetically useful energy. It seems logical then to expect that inhibition of growth may be reflected in a lowered respiration rate which in turn means less biologically useful energy available. Conversely, a lowered production of biologically useful energy through inhibition of the enzymes in the electron-oxygen transport system could be reflected in cessation of growth or growth inhibition. Even the inhibition of an enzyme system functioning to produce substrate(s) for the electron-oxygen transport system may be detected as a lowered activity of the enzyme systems involved directly in the electron-oxygen transport system. A study of the electron-oxygen transport system might be approached in at least two ways. A quantitative measurement of the gaseous exchange of respiring tissues would reveal the net result of the action of the "respiratory" enzymes. A second, and a better approach to such a study 7h would be an investigation of the DPNH oxidase system. Such a study would yield not only a measure of the oxygen uptake through the cytochrome system but also an index of the individual activities of the other enzymes which form the component parts of the DPNH oxidase complex. DPNH oxidase activity. Preliminary results were obtained from a study of tissue homogenates by a colorimetric procedure devised originally to measure dehydrogenase activity. In this assay tissue homogenates were incubated for several hours (up to 20 hours) in a buffered system in the presence of a substrate and a hydrogen acceptor. The method of assay depends upon the ability of DPNH to reduce the colorless 2,3,5-triphenyltetrazolium-chloride to its colored reduction product (87 ). This reaction is nonspecific and the results obtained therefrom were erratic and in­ conclusive. Hence, a more definitive procedure was adopted for the assay of DPNH oxidase activity of maleic hydrazide treated radishes and comparative controls. In this assay, the enzyme preparation was obtained from an acid-treated phosphate buffer extract of the respective tissues. The method of following the oxidation of DPNH was the spectrophotometric method of Warburg and collaborators based upon the change in spectral characteristics of oxidized and reduced DPN. The results of the measure­ ment of DPNH oxidase activity are summarized in Figure 1, showing the activity of the total system unfortified by additional amounts of cytochrome c or any other co—factors • It should be noted from Figure 1 that in the root tissue the rate of DPNH oxidation is the same for both types of tissue. However, in both the shoot and shoot stub the treated tissues exhibit an activity of about one-half of the respective controls. Shoot o CM CM OJ rH O o rH CO *t1w 0 tissues Shoot Stub -p CM o V\ o CM iH UN -P O CM CM Figure 1. The DPNH oxidase activity of control and inaleic hydrazide treated radish - A E 340 my., values represent the decrease in optical density x 1000. Root 75 E-t 76 Before investigating the individual components of the DPNH oxidase system it seemed desirable to establish the enzymatic nature of the oxidation of DPNH. Singer and Kearney (88) have reported that cytochrome c may be reduced non-enzymatically by pyridine nucleotides, a reaction catalyzed by various flavins. Since heated enzyme preparations (90 o 100 for 5 minutes) failed to change the optical density at 3U0 mp. over periods in excess of six minutes, it was assumed that the oxidation observed was indeed enzymatic. It should be mentioned that the DPNH oxidase activity of all of the tissue preparations was considerably lower than might be anticipated when compared with similar preparations obtained from animal tissue. However, in the studies with animal tissue vital organs have usually been used. Such tissue might be expected to exhibit higher orders of enzyme activity in general. A comparison of plant and animal tissue on such a basis may be unjustified. A lower oxidative response could conceivably be peculiar to a system containing sodium dithionite reduced substrate. Slater (70) observed that DPNH prepared by the nonenzymatic modified method of Ohlmeyer (89) was not satisfactory for certain phases of his oxidative study. Pre­ sumably this was due to interference of the products of oxidation of the reducing agent. However, an equally plausible suggestion might be that the lower oxidative level was due to impurities in the DPN used or the result of interaction of reducing agent and such impurities as might have occurred in the DPN. In this study cnemically reduced DPN (cozymase - 90) was used exclusively waicn when assayed spectrophotometrically was determined to be more than 90 per cent pure. 77 Diaphorase activity. In investigating the DPNH oxidase system for factors essentially associated with the saccess or failure of the system in the various tissues, quantitative determinations of diaphorase activity were performed. The experimental system designed to measure this activity contained cyanide to inhibit the cytochrome oxidase portion of the respiratory chain and methylene blue to act as a hydrogen acceptor to by-pass the cytochrome system. It has been established that methylene blue is capable of oxidizing reduced diaphorase. Cyanide was used at a final concentration of 2 x 10“4 M, which concentration has been shown by Lockhart and Potter (90) to be sufficient to block completely the activity of cytochrome oxidase. These workers also showed that at concentrations of 10-2 M cyanide the reduction of cytochrome c itself was inhibited and concluded that even 10“3 M cyanide has some depressant action on the reduction of cytochrome c. That 10 M cyanide does not inhibit diaphorase was reported by Adler et al. (91) and later observed also by Lockhart and Potter (90). that DPN reacts with cyanide (92). Meyerhof and co-workers have reported Colowick and associates (93) not only confirmed the conclusion that DPN is capable of forming a complex with cyanide but also presented evidence that DPNH is unaffected by cyanide. Therefore, the site of action of cyanide in the DPNH oxidase system seems to be the terminal part of the oxidative sequence rather than the inter­ action with the components of the initial diaphorase system. In the diaphorase determination, it was noted that in the systems con­ taining both methylene blue and cyanide, the inherent rate of autoxidation of the substrate was accelerated. This effect was also noted by Slater (70) who stated that the reason for this reaction was not known. In the 78 data reported here the diaphorase activities are corrected for this autoxidation by use of a "blank" determination. It should be noted in Figure 2 that the diaphorase activity of the treated tissue preparations is essentially the same as the respective controls except for the treated shoot stub tissue* These data then suggest that the lowered DPNH oxidase response of the treated shoot is not associated primarily with limited diaphorase activity but rather point to a failure in the cytochrome components* Lower diaphorase activity of the treated shoot stub suggests that lowered DPNH oxidase activity may be associated with some inhibition of the diaphorase system. Cytochrome oxidase. Direct measurement of the cytochrome oxidase by the spectrophotometric method of Hogeboom and Schneider (8b) confirmed the suggestion of a deficiency in the cytochrome system in the case of the shoot (Figure 3)* However, the level of cytochrome oxidase activity remained the same for both the control and treated root, as might have been expected since the DPNH oxidase was the same. The cytochrome oxidase activity of the control and treated shoot stub tissues was essentially the same. That the metabolism is now generally accepted* of DPNH normally requires cytochrome oxidase Slater (70) has demonstrated that 5 x 10 M potassium cyanide produced a 96.9 per cent inhibition of the DPNH oxidase system. The fact that the oxidation of cytochrome c was indeed enzymatic was confirmed by the fact that no change in absorption at S*?Q mji* occurred in the oresence at of 2 x IQ"4 M cyanide. a final concentration of 1.3 x 10 M. Cytochrome c was used This concentration of cytochrome c 79 M1 I 0 ) o X5 rH E-t X * o • cr\ o rH w •o c m rH Shoot Stub 0 CO H X CO O •H -P • -P rH -H XI £ CO O CM o o~\ a O CM o rH -P CO O -P 0 C •H Shoot a CM m a ■H rH CD T> • TJ cO O M I —I CD T) C O CD nj -H -P -p co a a ODTJ o P -H •p q q CO -H 0) >5, 'O o CD •H CO N g cO JJ 3 © q *h q T) K U >: CO d) X tflT! -P O O CD •h &x: Q> -P rH a -P S ^ ?» C 0) T)OW C rH CD CO 5h X P. rH CD O CM P Jh -P C*H W q O o cd 0 • rH H CO Cm g > O O O CM CM CM rH rH O co CH • >» • 4. P O E O >fl C •H CD H O 5 M' rH * a; q i (0 o co o q o O CD CD J0 P 0 fX i—1 Root CO -P H X .V rr\ TJ CD CM x:H E CD CD C x: CM U 0 bO ‘H CP -p CD s a Shoot Shoot Stub rH O o 8 o C\J & O -G o o o r*\ CM o o •rim ° " a v x r\ o o -p ^ E* V * I (0 0) la G CM 5>0 mp. in a system where cytochrome oxidase is not inhibited. Cytochrome c was used at a final concentration of 1.3 x lCf5 M. Both types of root preparation show a parallel reduction of cytochrome c with the oxidation of DPNH (Figure 2i). In the case of the shoot preparations, where the DPNH oxidase of the treated preparation was about one-half that of the corresponding control, a lower rate of reduction of cyto­ chrome c in the treated preparation suggests a lowered cytochrome c reductase activity. In the instance of the shoot stub, also, the treated preparation exhibited a lower DPNH oxidase activity but the rate of reduction of cytochrome c is essentially the same, indicating that the cytochrome c reductase system in both preparations are at about equal levels. For all tissue preparations, evidence of cytochrome oxidase was not apparent over the 6 minute assay period. Cytochrome c. A relatively adequate amount of endogenous cytochrome c must be assumed in the root tissues since added cytochrome c did not change the rate of oxidation of DPNH appreciably (Figure 5). However, a high cytochrome c concentration in the tissue extracts, prepared in the manner described in the experimental section, was not expected. The fractionation procedure was designed to remove most soluble tissue metabolites. The addition of cytochrome c, 1.3 x 10“° M, did accelerate Hoot Shoot Stub iH o CM CM r-i o CO r—i •rim ° " a v + XA -P O CM o -p tA CM iH iH O CM o CM Figure ii. The effect on the absorption at $$0 my., produced by the addition of O.i* ml. of 10-4 M cytochrome c to the DPNH oxidase system of control and maleic hydrazide treated radish tissues + A K 5f0 mu. represent the increase in optical density x 1000* Shoot 82 a3 o $ m -p o o m CM CM iH vr\ o CM Q Or\ c 8 O o CM O O 5 d 73 p •d rH T5 <0 o CM O UN rH m t> P p O G •H Shoot E CM Q CO rH o CM iH •H CO d d O p rH •H p P S E CX o P Tf G *o c3 •H d O rH d P O (0 "d G c0 o P d G C G a o O O d • -d P. cm E o a p O E p PJ d ro P to p R G cd 00 a (n c d d o 00 G •H to Ph P TS d *H G & X O O to CO d p @ P 23 P iH P-. CO d O *> P p d • P H G P g cO rH P o O Root CM o O p d d feaOS Cm P cm rH ' CHa OH I H00C-CH-CH2-S-CH OH \ / \ / • 0I K o c C Later Hopkins and collaborators (103, lOlt) showed th a t maleic acid also reacts w ith the sulfhydryl groups of proteins and could thus in h ib it 92 the action of succinic dehydrogenase. Although dehydrogenases in general contain sulfhydryl groups (79) other dehydrogenases investi­ gated were not inhibited by the disappearance of free sulfhydryl groups. This was interpreted to mean that although a free sulfhydryl group was essential to succinic dehydrogenase, this is not necessarily true for other dehydrogenases. Muir and Hansch (10J.) further suggested that undoubtedly a study of other stable organic compounds with double bonds activated by groups such as amides, esters, ketones and nitriles would lead to the discovery of many compounds quite toxic to plant cells. Naylor and Davis (£ii) reported that maleic hydrazide had a de­ leterious effect on root tip respiration and suggested that it would seem entirely possible that it exerts its influence on growth by in­ hibiting respiration. Just what portion of the oxidative metabolism of the plant is affected was not indicated but they speculated that a dehydrogenase was in some way prevented from functioning normally. The speculative nature of this suggestion should be emphasized. Isenberg, Odland, Popp and Jensen (U7) reported that a number of dehydrogenases were inhibited in tissue obtained from maleic hydrazide treated plants. However, it should be pointed out that these workers used the 2,3,5-triphenyltetrazolium chloride as a means of measuring dehydrogenase activity. It has been shown by Brodie and Gots (105) that the transfer of hydrogen from enzymatically reduced DPN to triphenyltetrazo1ium chloride is mediated by diaphorase; thusly, inhibition of only diaphorase could appear as dehydrogenase inhibition. The long incubation period used by Isenberg and coworkers also leads to some un­ certainties as to the results. 93 In a later publication, Isenberg and coworkers (J.i8) presented more convincing evidence that maleic hydrazide treatment did suppress respi— ration and depress succinic dehydrogenase activity. In fact, these investigators reported that low concentration enhances both respiration and succinic dehydrogenase activity while just the reverse is true at higher maleic hydrazide concentrations. Marr4 (50) reported that maleic hydrazide inhibits dehydrogenase activity in the presence of various substrates although the results were erratic* In another report (h9) this same author indicated that maleic hydrazide in concentrations near those that reversibly inhibit growth in vivo have inhibited the activity of a preparation of dehydro­ genase in vitro* Indoleacetic acid appeared to be able to reverse the inhibiting effect of maleic hydrazide upon the dehydrogenase system. It is true that the suggestion of Muir and Hansch (101) that maleic hydrazide reacts in the same manner as lactones and maleic acid in adding sulfhydryl compounds at the double bond appears inviting. However, the similarity in structures may be misleading with respect to the ease of addition at the double bond. In view of the published reports reviewed above concerning the possible inhibition of dehydro­ genase activity by maleic hydrazide, it seemed desirable to ascertain if compounds carrying a free sulfhydryl would indeed add to the double bond of maleic hydrazide. The absence of any data in regard to this problem made this area of investigation seem necessary. 9U Analytical Methods A technique similar to that of Morgan and Friedmann (102) was employed for the study of the interaction of maleic hydrazide with thiol compounds. A stock solution of maleic hydrazide was prepared by dissolving lb.Ol g. of maleic hydrazide in 300 ml. of water, ad­ justing the pH to 7 and finally making to a volume of 500 ml. (U0 ml. is equivalent to 0.01 mole of maleic hydrazide). One-hundreth mole of the sulfhydryl containing compound was added to U0 ml. of stock solution. The pH was again adjusted to 7 and the resulting solution diluted to 5>0 ml. A tissue homogenate was prepared by macerating radish shoot tissue with two weight equivalents of water. The boiled tissue homogenate was heated in a boiling water bath for 10 minutes. In the studies involving the use of the tissue homogenates, 0.01 moles of both mercaptoacetic acid and maleic hydrazide and 10 g. of tissue homogenate were contained in a volume of 50 ml. and incubated at a pH of 7• A. 1.0 ml. aliquot was titrated at zero time and at various o time intervals after incubation at 37 • Titrations were carried out by adding a 1.0 ml. aliquot to 5.0 ml. of 0.01 W sulfuric acid. Twenty milliliters of 0.02 N iodine was added and the excess iodine back titrated with a standard sodium thiosulfate solution using starch as the indicator. Blank solutions were prepared which contained only maleic hydrazide, thiol compound or homogenate in amounts equivalent to that used when in combination. Maleic acid was used in place of maleic hydrazide under like conditions as a check on the experimental procedure. 95 Results and Discussion The data in Table VIII clearly demonstrate that thiols do not add readily to the double bond of maleic hydrazide under the experimental conditions used. In the case of mercaptoacetic acid there is a small decrease in titratable thiol groups following incubation, however, this decrease can be accounted for in the blank. This probably arises from the slow oxidation of the thiol compound during long incubation. In marked contrast are the results with maleic acid which is known to add thiols at the double bond (102). In this instance after only four hours of incubation approximately 60 per cent of the thiol groups can no longer be titrated with iodine. The longer periods of incubation show a slower but continuous decrease in titratable groups indicating a disappearance of the thiol groups through interaction with maleic acid. The data of Table VIII also show that such other thiols as cysteine and glutathione do not add readily to the double bond of maleic hydrazide. Additionally, the studies using tissue homogenates indicated that under the experimental conditions prevailing, no bio­ logical system in the tissue was capable of catalyzing the addition of thiol with maleic hydrazide or converting the maleic hydrazide into a compound that would add thiol. Morgan and Friedmann (102) in a study of the interaction of thiols with unsaturated compounds found that while maleic acid readily reacted with thiol compounds, the following unsaturated acids did not: mesconic, cis and trans cinnamic. citraconic, 96 TABLE VIII INTERACTION OF MALEIC HYDRAZIDE AND THIOL COMPOUNDS (Results expressed as milliequivalents x 10® of iodine oxidized by the thiol compound in a 1.0 ml. aliquot of solution) Time (hours) h 7 System components 0 M aleic hydrazide (blank) 2k 0 3 3 5 192 192 193 191 2 3 M aleic hydrazide and Mercaptoacetic acid 192 191 193 189 M aleic hydrazide and Glutathione (G-SH) 189 190 190 186 M aleic hydrazide and Cysteine 188 187 . 189 186 M aleic acid and Mercaptoacetic acid 191 67 52 31 Mercaptoacetic acid (blank) M aleic acid (blank) 0 - 3 Time (days) 1 2 k 182 - Mercaptoacetic acid (blank) 185 18U M aleic hydrazide and Mercaptoacetic acid 190 189 Shoot tissue homogenate - - k Boiled homogenate - - 3 Homogenate and Mercaptoacetic acid 183 183 185 182 B oiled homogenate and Mercaptoacetic acid 185 183 187 185 Homogenate , Mercaptoacetic acid and Maleic hydrazide 186 186 187 185 B oiled homogenate, Mercaptoacetic acid and Maleic hydrazide 18U 18U 185 187 187 1 - 97 These results indicate that the suggestion of Muir and Hansch (101), that maleic hydrazide inhibits growth by reacting with sulf­ hydryl groups, is not a very probable one. This assumes, of course, that maleic hydrazide remains as such in the biological system. These authors 1 suggestion was based primarily on an analogy between the interaction of thiol compounds and maleic acid and the probable interaction of thiols and maleic hydrazide. correct one. This analogy is not a 98 SUMMARY 99 SUMMARY A preharvest foliar application of a 0,02 M solution of maleic hydrazide does inhibit new shoot and root growth of topped radishes (Raphanus sativus) harvested UB hours after treatment and subsequently sbored, A study of the carbohydrase enzyme components in the radish root indicated that maleic hydrazide has had no observable effect on the beta amylase, phosphorylase or phosphatase activity in these tissues. Alpha amylase and pectin-methyl-esterase activity was not detected. Chemical analysis of the gross nutritional components of both the shoot and root tissues of control and maleic hydrazide treated plants revealed that dry weight, ether extract, Kjeldahl nitrogen, reducing and non-reducing sugars and polysaccharides other than starch had not been changed significantly by chemical treatment. The starch content of the shoot tissue was nearly doubled in the treated samples as com­ pared to the non-treated samples. A comparative quantitative study of the ability of an acid treated extract of various control and maleic hydrazide treated radish tissues to oxidize DPNH is presented. The oxidative system was analyzed quantitatively for the activity of its known constitutive enzymes; that is (a) diaphorase, (b) DPNH-cytochrome c reductase and (c) cytochrome oxidase. 100 The DPNH oxidase a c tiv ity was the same in both the control and tre a te d root tissu e* ■ The a c tiv ity of the con stitutive enzymes was also the same in both types of tissu e. The maleic hydrazide treated shoot tissue exhibited a marked in h ib itio n of the DPNH oxidase system* This in h ib itio n resulted from p a r tia l fa ilu re s in both DPNH-cytochrome c reductase and cytochrome oxidase systems. The DPNH oxidase a c tiv ity of the treated shoot stub tissue was somewhat lower than th at of the corresponding control tissue. This in h ib itio n resu lted from a reduced a c tiv ity of the diaphorase system. The a c tiv ity of the cytochrome systems was the same fo r both the con­ t r o l and maleic hydrazide treated tissu e. An in v e s tig a tio n of the in v itro in te ra c tio n of th io ls and maleic hydrazide indicated th a t ho addition compound resulted from the in ­ cubation of m aleic hydrazide and th io ls a t a pH of 7 even though the ad d itio n o f th io l and maleic acid was noted. Tissue homogenates had no e ffe c t on the system investigated fo r *the possible in teractio n of th io l and maleic hydrazide. 101 BIBLIOGRAPHY 102 BIBLIOGRAPHY 1. P. Boysen-Jensen, Growth Hormones in Plants, McGraw-Hill Book Co., Inc., New York, N. Y., 1936. 2. A. Paal, Jahrb. wiss. Botan., 58, 1*06 (1 9 1 9 ). tr 3* . k* F. Kogl and A. J. Haagen-Smit, Proc. Acad. Sci., Amsterdam, 3k. 11*11 (1931). C.A., 26, 2755 (1932). » F. Kogl, A. J. Haagen-Smit and H. Erxleben, Hoppe-Seyler's Z. physiol. Chem., 288, 90 (193U). 5. E. SalkowskL, Z. physiol. Chem., £, 8, 23 (1885). 6. A. Ellinger, Ber., 38, 2881* (1905). 7. H. A. Foersterling, J. prakt. Chem., 51, 371 (1895)* 1*29 (1895). 8. F. Arndt, Rev. fac* sci. univ. Istanbul, 9A, 19 (191*1*). UO, 1787 (191*6). 9. F. Arndt, L. Loewe and L. Ergener, Rev. fac. sci. univ. Istanbul, 13A, 103 (19U8). C.A., 1*3, 579 (19U9). Ber. 28, C.A., 10. R. M. Mizzoni and P. E. Spoerri, J. Am. Chem. Soc., 73, 1873 (1951). 11. D. L. Schoene and 0. L. Hoffman, Science, 109, 588 (191*9). 12. C. D. Darlington and J. McLeish, Nature, 167, 1*07 (1951). 13. G. Deysson and M, Deysson, Bull. soc. chem. biol., 35* 1209 (1953). l)|, V. A. Greulach and E. Atchison, Bull. Torrey Botan. Club, 77* 262 (1950). 1 5 . V. A. Greulach and E. Atchison, Botan. Gaz., Ill*, 1*78 16. V. A. Greulach and J. G. Haesloop, Am. J. Botan., y., 1*1* (1951*). 17. R. H. Moore, Science, 112, 52 (1950). 18. A. W. Naylor and E. A. Davis, Botan. Gaz., 112, 112 (1950). 19. S. N. Hao, Ph.D. Thesis, Mich. State Coll., (1951*)* 20. F. W. Went and K. V. Thiamann, Phytohormones, The Macmillan Co., New York, N. Y., 1937. (1 953). 103 21* A. E. Hitchcock and P* W. Zimmerman, Contrib, Boyce Thompson Inst., 16, 225 (1951). 22* W„ J, Mcllrath, Am. J. Botan., 3£, 8l6 (1950). 23* A. C. Leopold and V/.U. Klein, Science, llH, 9 (1951). 2) 4. A. C. Leopold and W.H. Klein, Physiol. Plantarum, 5, 91(1952). 25. W. A. Andreae, Abstract Paper presented Am. Soc. of Plant Physiol., Ithaca, N. Y., September, 1952, p. 25. 26. W, A. Andreae and S.R. Andreae, Can. J. Botan., 31, U26(1953). 27. R. J. Gautheret, Compt. rend.,23U, 2218 (1952). 28. R. S. Choudri and V. B. Bhatnagar, J. Sci. Research Banaras Hindu Univ., 3, 86 (1952-3). C.a ., U8, 9005 (195U). 29. F. Bertossi and H. Capozzi, Bull. soc. ital. biol. sper,, 28, 1117 (1952). C.A., U8, 3186 (19SU). 30. A. S. Crafts, H. B. Currier and B. E. Day, Hilgardia, 20, 57 (1950). 31. E. B. Eskew and C. J. Willard, North Central Weed Control Conference, p. 187 (1950). 32. A. W. Naylor and E. A. Davis, Proc. Am. Soc. Plant Physiol. (2l±th Meeting). Abstracts, p. 13 (19U9). 33. L. A. 3h. H. B. (195D. Currier, B. P. Day and A. S. Crafts, Botan. Gaz., 112,272 35. A. Naylor, Arch. Biochem. and Biophys.,, 33, 3h0 (1951). 36. D. S. (1952). Mikkelsen, R. B. Griffith and D. Ririe, Agron. J., l*U,533 37. S, Wittwer and C. M. Hansen, Agron. J., U3, 3U0 (1951). 38 . S. H. Wittwer, R. C. Sharma, L. E. Weller and H. M. Sell, Plant Physiol., 25, 539 (1950). W. H, Tatum and J. H. Curme, Plant Physiol., 26, 836 (1951). 39. D. R. Patterson, S. H. Wittwer, L. E. Weller and H. M. Sell, Plant Physiol., 27, 135 (1952). U0. D. R. Patterson, Ph.D. Thesis, Mich. State Coll., (1952). lOh ifl. M. E* Highlands, J. J. Licciardello and C. E. Cunningham, Am* Potato J*, 29, 225 (1952). 1<2. E. J. Kennedy and 0. Smith, Proc. Am. Soc. Hort. Sci., 61, 395 (1953). — U3. E.L. Petersen and A. W. Naylor, Physiol. Plantarum, 6, 816 liil. C.Phouphas and A. Goris, Compt. rend., 23*4» 2002 (1952). U5. W. U6. V.A. Greulach, Bot. Gaz., Ill;, 1;80 (1953). h? • F.M. R. Isenberg, M. L. Odland, H. W. Popp and C. 0. Jensen, Science, 113, 53 (1951). 1;8. , R9. 0. Griesel, Science, 119, 8U3 (1953). (19514-). F. M. R. Isenberg, C. 0. Jensen and M. L. Odland, Science, 120, U6U (195*4). t E. Marre, Atti accad. nagl. Lincei, Rend. Classe sci. fis., e nat,, 15, *433 (1953). C.A., U8, 122i;Oe (195*4). 50. E. Marre, Congr. intern, botan., Paris, Raops, et communs., 8, Sect. 11/12, 167 (195U). C.A., U8, 10852i (195U). 51. W. A. Andreae, Congr. intern, botan., Paris, Rapps, et communs*, 8, Sect. 11/12, 151 (19514). C.A., U8, ll565i (195U). 52. W, A. Gortner and M. Kent, J. Biol. Chem., 20*;, 593 (1953). 53. G* Morel and S. D. Demetriades, Annee Biol., 31, 227 (1955). Ann. Rev. Plant Physiol., 6, h75 (1955). 5)-4. A. W. Naylor and E. A. Davis, Bull. Torrev Botan. Club, 78 , 73 (1951).' 55. R. M. Smock, L. J. Edgerton and M. B. Hoffman, Proc. Am. Soc. Hort. Sci,, 58, 69 (195l). 56. D. R. 57* A. Frey-Wyssling, Growth, 12, 58. H. D. Tate, Literature Summary on Maleic Hydrazide, 6, 23 (1952), 6C, 26 (1955). Naugatuck Chemical Division, United States Rubber TIo., Bethany, Conn. 59. V. A, Greulach, J. McKenzie and E. M. Stacey, Biol. Bull., 101, 285 (1951). Goddard, Growth, 12, 17 (19U8). l5l (I9*i8). io5 60* D. H. Dewey and S. H. Wittwer, Proc. Am. Hort. Soc.. 17, In press (1956). 9 -1-9 61. J. B. Sumner, T. C. Chou and A. T. Bever, Arch. Biochenu, 26, 1 (1950). — ■ 62. J. B. Sumner, Science, 100, Ul3 (19UU)• 63. V. Ignatieff andH. 61. R. M. Sandstedt,E. (1939). Wasteneys, Biochem. J., 30, 1171 (1936). Kneen and M. J. Blish, Cereal Chem., 16,712 “ 65. Z. I. Kertesz, J. Biol. Chem., 121, 589 (1937). 66. E. Kneen and R. M. Sandstedt, Cereal Chem., 18, 237 (l9ll). 67* M.J. Blish and R. M, Sandstedt, Cereal Chem., 10, 189 (1933). 68. H.L'l. Sell, F. A. Johnson and F. S. Lagasse, J. Agr. Res., 319 (1916). 69. J.G. Dewan and D. E. Green, Biochem. J., 31, 1071 (1937). 70. E.C. Slater, Biochem. J., 16, 181 (1950). 71. M.E. Mathews, J. Biol. Chem., 189, 695 (195l). 73a 72. H. Beevtrs, Plant Physiol., 29, 265 (1951). 73. K.C. Slater, Biochem. J., 16, 199 (1950). 7l. B.C, Slater, Biochem. J., 11, 305 (I9l9). 75. D. 76. W. C. Schneiderand 969 (1950). 77. G.H. Hogeboom, J. Biol* Chem., 177, 817 (I9l9). 78. G.H. Hogeboom and W. C. Schneider, J. Nat. Cancer Inst., 10, 983 (1950). 79. J. B. Neilands and P. K. Sturapf, Outline of Enzyme Chemistry, John Piley and Sons, Inc., New York, N. Y., 1955. Keilin and D.E. Hartree, Biochem. J., ll, 500 (1950). G. H. Hogeboom, J. Nat. Cancer Inst., 10, 80. G.A. LePage, J. Biol. Chem., 168, 623 (I9l7). 81. B.L. Horecker and a . Komberg, J. Biol. Chem., 175, 385 (1918). 106 82. H. H. Hess and A. Pope, J. Biol. Chem., 20li, 295 (1953). 83. M. P. Lenta and M. A. Riehl, Cancer Res., 12, U 98 (1952). 8iu G. H. Hogeboom and W. C. Schneider, J. Biol. Chem., 19k, 513 (1952). 85. E . G. Ball, Ann. N. Y. Acad. Sci,, U5, 363 (19UU). 86. A. L. Lehninger, J. Biol. Chem., 178, 6ll, 625 (19U9). 87. E. Kun and L. G. Abood, Science, 109, ikk (19U9). 88. T. P. Singer and E. B. Kearney, J. Biol. Chem., 183, U09 (1950). 8?. P. Ohlmeyer, Biochem. Z., 297, 66 (1938). 90. E, E. Lockhart and V. R. Potter, J. Biol. Chem., 137, 1 (19Ul)• 91. ft9Adler, H. V* Euler and H. Hellstrom, Arkiv. Kemi, Mineral. Geol., 12B, No. 38 (1937). C.A., 32, 2966 (1938). 92. 0. Meyerhof, P. Ohlmeyer and VI. Mohle, Biochem. Z., 297, H 3 93. S. P. Colowick, No 0. Kaplan and M. M. Ciotti, J. Biol* Chem. 191, hh7 (1951). 9k* J . R. Hawthorne and C. D. Harrison, Biochem. J., 33, 1573 (1939). 95. A. L. Lehninger, J. Biol. Chem., 190, 3U5 (1951)• 96. V. R. Potter, J. Biol. Chem., 137, 13 ( 1 9 U D . 97. B. L. Horecker and A. Kornberg, J. Biol. Chem., 165, 11 (19U6). 98. R. H. Goodwin and C. Taves, Am. J. Botan., 37, 22lj (1950). 99. H. Veldstra and E, Havinga, Enzymologia, 11, 373 (19^9). (1938). 100. C. J. Cavallito and T. H. Haskell, J. Am. Chem. Soc., 67 , 1991 (19^5) 101 . R. M. Muir and C, Hansch, Plant Physiol., 28, 218 (1953). 102 . E. J. Morgan and E. Friedmann, Biochem. J., 32, 733 (1938). 103. F. G. Hopkins, E. J. Morgan and C. Lutv/ak-Mann, Biochem. J., 32, 1829 (1938 ). 10 b. F. G. Hopkins and F. J. Morgan, Biochem. J., 32, 611 (1938). 105. A. F. Brodie and J. S. Gots, Science, Ilk, U0 (1951).