ABSTRACT GLYCOLATE METABOLISM IN GREEN ALGAE by Edward Blake Nelson Assimilation of exogenous glycolate by Scenedesmus obliguus was measured at stages in the life cycle of syn- chronized cultures. Glycolate uptake occurred only during the stage of cell division. This anion stimulated reSpir- ation and photosynthesis when the cells could assimilate it. Equal rates of glycolate metabolism occurred in the light and dark, and reSpiration of Scenedesmus was stimu- lated 25% to 100%. In the light and in the absence of 002, photosynthetic oXygen evolution increased 100% to 300% in the presence of alycolate. Glycolate and glyoxylate were equally effective in stimulating oxygen evolution. This stimulation was inhibited by CMU, an inhibitor of photo- synthetic electron tranSport. It is proposed that the oxidation of glycolate to glyoxylate and the reduction of glyoxylate to glycolate is a terminal metabolic cycle, which accepts reducing equivalents from electron tranSport in photosynthesis and thereby stimulates oxygen evolution. The ability of Chlamydomonas reinhardtii to excrete glycolate was found to be regulated by C02 availability during growth. While Chlamydomonas grown on 1% 002 in air Edward Blake Nelson were capable of glycolate excretion, the ability to excrete glycolate decreased with increasing culture den- sity. Cells grown on air (0.03% C02) did not excrete glycolate. Air grown cells would excrete glycolate in the presence of 10"2 M isonicotinyl hydrazide, an inhib- itor of the glycolate pathway. When cultures grown on 1% 002 in air were transferred to air, they lost the ability to excrete glycolate in approximately 16 hours. Levels of enzyme activities which might be involved in regulating glycolate excretion were examined in Chlamydomonas cultures grown on 1% 002 in air or on air. P-glycolate phOSphatase (phOSphoglycolate phOSphohydrolase E.C. 3.1.3.18) activity was high and did not vary signifi- cantly on transfer from 1% C02 in air to air. Glycolate oxidase (glycolate: 02 oxidoreductase E.C. 1.1.3.1) could not be detected in crude extracts of Chlamydomonas. A new enzyme, glycolate dehydrogenase (glycolate: acceptor oxidoreductase, no E.C. number), which catalysed the oxi- dation of glycolate to glyoxylate, was found in these Chlamydomonas. The activity of glycolate dehydrogenase increased two to four fold 16 hours after transfer of cells from 1% C02 in air to air. The increase in activity was inhibited by cycloheximide (5 ug/ml), an inhibitor of protein synthesis. These results suggest that C02 avail- ability during growth of the algae, regulates glycolate excretion by controlling the levels of a glycolate dehydro- genase. Edward Blake Nelson Glycolate dehydrogenase was purified 10 fold from Chlamydomonas by Triton X-100 extraction of whole cells and (NH4)ZS°u fractionation of the protein. The enzyme had no oxidase activity. No glyoxylate formation occurred in the absence of artificial electron acceptors. Only dichloroindophenol and phenazine methosulfate were found to serve as electron acceptors. The natural electron acceptors are unknown. The enzyme oxidized glycolate at pH 8.0 to 8.7 preferentially to all other a-hydroxy acids testedgand the Km was 2.2 x 10‘” N. D—lactate was the second best substrate with a pH Optimum at 8.7 and a Km of 1.5 x 10-3 M. The enzyme was sensitive to sulfhydryl inhibitors. No requirement for co-factors was established, although by precedent a flavin is suggested. Glycolate dehydrogenase activity was found in the following green algae; Chlamydomonas, Chlorella, Scenedesmus, Euglena, and Acetabularia. No glycolate oxidase was detected in these algae. The higher plants examined con- tained glycolate oxidase but no glycolate dehydrogenase. It is suggested that the enzyme oxidizing glycolate to glyoxylate in algae is a dehydrogenase and in plants an oxidase, which is located in peroxisomes. Procedures for differentiating between these two enzymes are outlined. L-glutamate: glyoxylate aminotransferase (E.C. 2.6.1.4) was measured in crude extracts of Chlamydomonas. L—glutamate and L-alanine were the preferred amino donors. Edward Blake Nelson Sufficient levels of activity were present to metabolize all glyoxylate formed by glycolate dehydrogenase. The presence of glycolate dehydrogenase and glyoxy— late aminotransferase in Chlamydomonas establishes enzymes for the glycolate pathway in green algae. This accounts for the labeling data in which glycolate-i-luc was converted to glycine-l-luc in green algae. GLYCOLATE METABOLISM IN GREEN ALGAE By Edward Blake Nelson A THESIS ' Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Biochemistry 1970 ACKNOWLEDGMENTS I thank Professor N. E. Tolbert for his guidance and cooperation. His enthusiasm for research is an inspir- ation for every graduate student. The interesting and often lively discussion with the graduate students, post- doctoral fellows, and visiting professors in the Tolbert laboratory were helpful in pursuing my research. I thank Miss Arlene Cenadella for her technical assistance. Finally I thank my wife, Kirsten, for her encouragement and support throughout the course of this work. The financial assistance of the National Institutes of Health, National Science Foundation, and the Department of Biochemistry, Michigan State University is appreciated. ii TABLE OF CONTENTS Page IN'I‘RODUCTION . O O O O O O O O O 0 O O O O O O O O O 1 LITERATURE REVIEW . . . . . . . . . . . . . . . . . 3 The Glycolate Pathway in Higher Plants . . . . 3 The Glycolate Pathway in Algae . . . . . . . . 5 The Regulation of Glycolate Metabolism by Carbon Dioxide . . . . . . . . . . . . . . . . 8 Enzymes from Green Plants and Algae Oxidiz— ing Glycolate . . . . . . . . . . . . . . . . . 10 Glycolate Excretion by Algae . . . . . . . . . 12 Effects of Oxygen on Glycolate Metabolism . . . 1h Effects of Light on Glycolate Metabolism . . . 14 Glutamate: Glyoxylate Aminotransferase . . . . 15 MATERIALS AND METHODS . . . . . . . . . . . . . . . 17 Algae . . . . . . . . . . . . . . . . . . . . . 17 Plants . . . . . . . . . . . . . . . . . . . . 18 Measurement of Glycolate Excretion . . . . . . 19 Measurement of Glycolate Assimilation . . . . . 19 Measurement of Oxygen Exchange . . . . . . . . 20 Enzyme Assay Methods . . . . . . . . . . . . . 20 P-Glycolate PhoSphatase . . . . . . . . . . . . 21 Glycolate Oxidase . . . . . . . . . . . . . . . 21 Glycolate Dehydrogenase . . . . . . . . . . . . 22 Preparation of Cell Free Extracts . . . . . . . 27 iii Page Glycolate Oxidase . . . . . . . . . . . . . . . 28 Purification of Glycolate Dehydrogenase . . . . 28 Glutamate: Glyoxylate Aminotransferase . . . . 29 Protein and ChlorOphyll Determination . . . . . 30 RESULTS AND DISCUSSION . . . . . . . . . . . . . . . 31 Assimilation of Glycolate by Scenedesmus . . . 31 Effects of Glycolate on Photosynthesis and Respiration O O O O O O O O O O O O O O O O O O 37 Glycolate Excretion by Chlamydomonas . . . . . 43 Glycolate Dehydrogenase: Detection and Assay . 48 Levels of Glycolate Dehydrogenase and P-Glycolate PhOSphatase in Chlamydomonas . . . 53 Purification of Glycolate Dehydrogenase . . . . 60 Substrate Characterization of Glycolate Dehydrogenase O O O O O O O O O O O O O O O O O 65 Electron Acceptor Specificity of Glycolate Dehydrogenase o o o o o o o o o o o o o o o o o 6R- Flavin or Pyridine Nucleotide Cofactors . . . . 7“ pH Optimum and the Effect of Ionic Strength on Glycolate Dehydrogenase . . . . . . . . . . 79 Effect of Inhibitors on Glycolate Dehydrogen- ase o o o o o o o o o o o o o o o o o o o o o 0 70. Distribution of Glycolate Dehydrogenase . . . . 85 Glutamate: Glyoxylate Aminotransferase . . . . 87 GEIIERAL DISCUSSION 0 0 O O O O O O O O O O O O 0 O 0 91 The Role of Glycolate Excretion and Assimila- tion in Nature I O O O O O O O O O O O O O O O 91 The Regulation of Enzyme Levels by 002 Availability 0 O O O O O O O O O O O I O O O 0 92 Enzymes of the Glycolate Pathway in Algae . . . 93 iv Comments on Glycolate Dehydrogenase From Green Algae . . . . . . . . . . . . . . BIBIJImRAPHY O O O C O O O O O O O O O O O O Table No. 1 10 11 12 LIST OF TABLES Comparison of Different Substrates for Stimulation of nyqen Evolution and Respiration O O O O O O O O O O 0 O O O 0 Absence of Endogenous Inhibitors of Glycolate Oxidase and Dehydrogenase in Crude Extracts of Chlamydomonas . . . . . Glycolate Dehydrogenase in Chlamydomonas Purification of Glycolate Dehydrogenase . Substrate Specificity of Glycolate Dehydrogenase . . . . . . . . . . . . . . Mixed Substrate Assays with Glycolate Dehydrogenase . . . . . . . . . . . . . . Electron Acceptor Specificity of Glyco- late Dehydrogenase . . . . . . . . . . . Effect of Enzyme Inhibitors on Glyco- late Dehydrogenase . . . . . . . . . . . Survey of Algae and Plants for Glyco- late Dehydrogenase . . . . . . . . . . . Glutamate: Glyoxylate Aminotransferase Activity in Crude Extracts of Chlamydomonas O O O O O O I O O O O O O O Specificity of Amino Group Donor for Glycine Formation by Crude Extracts of Chlamydomonas O O O O O O O O O O O 0 O 0 Activity of Enzymes of the Glycolate Pathway in Green Algae . . . . . . . . . vi 'Page no 51 62 66 67 71 Q3 86 89 90 Figure No. 1 2 10 LIST OF FIGURES Page The Glycolate Pathway . . . . . . . . . 4 Absorption Spectra of Glyoxylate Phenylhydrazone in Presence and Absence of DCIP . . . . . . . . . . . . 26 Assimilation of Exogenous Glycolate by Synchronized Scenedesmus at Differ- ent Stages of Their Life Cycle . . . . 33 Glycolate Assimilation by Synchronized Scenedesmus in the Light and Dark . . . 35 Effect of CMU on Glycolate Assimila- tion by Synchronized Scenedesmus . . . 35 Effect of Glycolate on Oxygen Evolu- tion in the Light and HeSpiration in the Dark by Scenedesmus . . . . . . . . 39 A Proposed Scheme to Account for Gly- colate Stimulation of Oxygen Evolu- tion 0 O O O O O O O O O O O O 0 O O 0 [+2 Glycolate Excretion by Chlamydomonas Grown on 1% C02 in Air Relative to the Age and Density of the Culture . . 45 Decrease in Ability of Chlamydomonas to Excrete Glycolate After Being Transferred from 1% C02 in Air to Air . U7 Effect of Isonicotinyl Hydrazide on Glycolate Excretion by Air Grown Chlamydomonas . . . . . . . . . . . . . 50 Changes in Specific Activity of P- Glycolate PhOSphatase and Glycolate Dehydrogenase (Glycolate: DCIP Oxi- reductase) in Chlamydomonas after Transfer from.I%C02 in Air to Air . . 55 vii Figure No. Page 11 Effect of Cycloheximide on Specific Activity of Glycolate Dehydrogenase After Transfer of Chlamydomonas from 1% 002 in Air to Air . . . . . . . . 59 12 Purification Scheme for Glycolate Dehydrogena se 0 o o o o o o o o o o o o o 61 13 Specific Activity of Glycolate Dehydro- genase Released from Chlamydomonas by 1% Triton X-100 o o o o o o o o o o o o o o 6”“ 1h Determinations of Km's for Glycolate and D-Lactate . . . . . . . . . . . . . . 69 15 Glyoxylate Formation in Relation to DCIP Reduction, the Requirement of DCIP for Glyoxylate Formation . . . . . . . . 73 16 Determination of the Km for DCIP . . . . 76 17 Substrate Saturation Curve for PMS . . . 78 18 pH Curve for the Oxidation of Glycolate and D-Lactate by Glycolate Dehydro- genase . . . . . . . . . . . . . . . . . 81 viii ll ll.il| III. III I ll I I All. ‘1 I I ll ' i bicine p-CMB CMU DCIP EDTA FAD FMN MES NAD+ NADP+ P MS tricine Tris LIST OF ABBREVIATIONS N,N-bis(2-hydroxyethyl)glycine p-chloromercuribenzoic acid 3-(p-chlor0phenyl)-1,i-dimethylurea 2,6-dichloroind0phenol (Ethylenedinitrilo)tetraacetic acid Flavin Adenine Dinucleotide Flavin Mononucleotide 2-(N-morpholino)ethanesulfonic acid Nicotinamide Adenine Dinucleotide Nicotinamide Adenine Dinucleotide PhOSphate phenazine methosulfate N-tris(hydroxymethyl)glycine ethanesulfonic acid tris(hydroxymethyl)aminomethane ix INTRODUCTION The use of luCOZ for research in photosynthesis led to the discovery of the path of carbon in photosynthesis. It was originally observed that glycolate (a-hydroxyace- tate) was rapidly formed during photosynthesis (10), and later work showed that it could be a major product of photosynthesis (97). The determination of the direction of glycolate metabolism led to the formulation of the glycolate pathway (Figure 1). Applying the concept of "the unity of biochem- istry," this pathway determined for higher plants was also attributed to green algae. Later work in a number of labor- atories then suggested that the glycolate pathway did not exist or was not complete in green algae. It has been the goal of the research reported in this thesis to ascertain the role of the glycolate pathway in green algae. It is felt that such information is important to our basic understanding of plant biochemistry and furthermore to our understanding of primary productiv- ity in the biOSphere. If glycolate accounts for approxi- mately fifty percent of photosynthetic carbon metabolism as determined by Zelitch (104), then the problem of glyco- 2 late metabolism in algae is eSpecially important. LITERATURE REVIEW The Glycolate Pathway in Higher Plants Glycolate metabolism in higher plants has been demon- strated in three ways; i) feeding eXperiments with glycolate- 14c. 2) 14002 fixation eXperiments and, 3) inhibitor studies. The metabolic pathway derived from these studies, commonly called the glycolate pathway (92), is shown in Figure 1. Specifically labeled glycolate-Inc fed to a number of green plants has given results consistent with the glyco- late pathway (7, 60, 87). Glycolate-i-luC gave glycine-i- luc, serine-1-140 and glycerate-i-luc, while glycolate-2- 1“C gave glycine-Z-luc, serine-2,3-1uc and glycerate-2,3- inc (76). A precursor product relationship was found for these metabolites when glycolate-Z-luc was fed to wheat (76). The labeling pattern in glucose isolated after glycolate—luC feeding was consistent with the operation of the glycolate pathway (46). Photosynthetic 1“(:02 fixation with higher plants gave uniformly labeled glycolate at short times, during which BAP-glycerate was still carboxyl labeled (10, 42). Labeling patterns of intermediates of the glycolate path- way, reflected synthesis from glycolate rather than 3-P- glycerate. Both glycine and serine have been found to be mmznpmm mpoaoomau 028 .H mhzwam mommwe mommwe mommoe _ mooo + mwelmmzlllv o..un_oe nlllv moloelm _ moooo moooo moooo mmammo . m . + omoe ro moe . m m . N All]: . m moooo mOOUo mOOUo 5 uniformly labeled. If serine were derived from 3-P-g1ycer- ate, serine-l-luC would be eXpected at short times. Metabolism of glycolate via the glycolate pathway results in the loss of a one carbon unit. This one carbon piece is released as 002 in a process commonly called photo- reSpiration. By use of an inhibitor of glycolate oxidation, an a—hydroxypyridinemethane sulfonate, Zelitch (ion) has shown the release of C02 in the light by tobacco leaf disks was dependent on glycolate oxidation. Goldsworthy (36) obtained similar results with tobacco leaf segments using hydroxypyridinemethane sulfonate and another inhibitor of glycolate metabolism, isonicotinyl hydrazide. The Glycolate Pathway in Algae While it is well established in higher plants that glycolate is rapidly synthesized and metabolized, the metabo- lism of glycolate by algae has been an area of controversy. It was originally observed with Scenedesmus by Schou et. al. (83) that glycolate-luc was assimilated both aerobically and anaerobically in both the light and dark. The products detected by chromatography were consistent with the glycolate pathway (Figure 1). Specific labeling data obtained from glycerate isolated after feeding glycolate-Z-lnc was consis- tent with the pathway. Unfortunately no kinetic studies were done by Schou et. a1. Other workers (85) have fed glycolate- 140 to Chlorella, however the data wererum clear enough for 6 evaluation. The above mentioned work supported the presence of the glycolate pathway in algae. Work in other laboratories suggested that algae lacked a glycolate pathway. Tolbert and 2111 (89) observed that most of the glycolate synthesized was excreted by algae rather than metabolized. Numerous other laboratories have confirmed this (32, 61, 74). Labeling data from photosyn- thetic lucoz fixation with algae showed serine to be carboxyl labeled rather than uniformly labeled as found in higher plants, indicating it came from 3-P-glycerate rather than glycolate (13, 43). Originally glycine was found to be labeled differently than glycolate during 1“002 fixation (43), however recent work shows them to be labeled identi- cally (13). Also, it was observed that green algae lacked glycolate oxidase (42). To obtain data to reconcile these differences, several laboratories attempted to do feeding eXperiments with glyco- late-140. Unfortunately it has not been possible to obtain reproducible glycolate assimilation by green algae grown under normal laboratory conditions (16, 43, 57). Also cell free extracts of green algae would not metabolize glycolate- 1“c or P-glycolate-luc (42). The reasons for these observa- tions are not clear. In order to obtain conditions which allow reproducible assimilation of exogenous glycolate, gly- colate assimilation by synchronous cultures of Scenedesmus obliguus has been studied. Using this algae it has been 7 possible to obtain conditions which allow reproducible gly- colate assimilation. Bruin (i3) fed glycolate-i-luC to these cells and degraded the glycine and serine formed. The labeling data was consistent with the Operation of the glycolate pathway. This confirmed the original work of Schou et. al. (83). Thus it appears as though the glycolate pathway operates in green algae at least under certain con- ditions. Consistent with this is the recent detection of an enzyme from green algae which oxidizes glycolate to glyoxylate (55, 66, 106). It remains to be determined why the serine is carboxyl labeled. Recent work by Merrett and his associates (37, 38, 58, 59) has suggested that an alternative pathway of glyco- late metabolism might exist in Chlorella. Using acetate- 1“C they observed the following pattern in short term feed- ing eXperiments: acetate ———> glycolate ———-) glycerate —.—_> serine No glycine was detected. This pathway is identical to the way glycolate is metabolized by Pseudomonas (52). The overall significance of such a pathway in algae remains to be determined. Several other items related to the glycolate pathway in algae deserve comment. If the pathway is Operational then it is expected that green algae should be able to grow heterotrOpically on glycolate if they are able to assimilate it. It has been observed that glycolate will support slow growth of a phytoplankton strain of Chlorella (84) and 8 Chlorella elliposidea but not 9. pyrenoidosa (82). Glyco- late would not support dark growth of Euglena at pH 3.9 or 6.9 (23). At a physiological pH of 8,Dropp and McGill (28) found in testing 39 strains of algae that glycolate would not support growth of any of the strains. The lack of growth could be due to the high pH, which would keep glyco- late in the anion form rather than as the free acid, which is the more readily absorbed form. Several of these strains would, however, grow on acetate at pH 8.0. The effect of glycolate on cellular reSpiration has been studied by Myers (63) who found that glycolate stimu- lated reSpiration of Chlorella at pH 3.9 but not at pH 6.9. Sen (84) observed glycolate stimulated reSpiration at pH 6.1 with a phytoplankton strain of Chlorella which grew on gly- colate. Anderson (2) observed stimulated reSpiration by glycolate using Prototheca zopfii, a colorless algae. No other studies to the author's knowledge have shown stimulated reSpiration by glycolate with algae. In this thesis and in publication (65) the stimulation of reSpiration by exogenous glycolate is reported. This occurs in cells of Scenedesmus which are capable of assimilating glycolate. The Regulation of Glycolate Metabolism by Carbon Dioxide The synthesis and excretion of glycolate by algae has been found to be very sensitive to C02 concentration and availability. Carbon dioxide effects glycolate metabolism 9 in two ways; one, by controlling the levels of metabolites and two, by regulating the activity of an enzyme which metabolizes glycolate. Numerous laboratories have observed that increasing the C02 concentration above 0.2% to 0.4% during photosynthesis lead to a decrease in glycolate syn- thesis, even though photosynthetic rates were not saturated (61, 74, 97). The effect has also been observed with higher plants (105) and isolated chloroplasts (90). It has been suggested that this phenomenon was observed because a pro- posed precursor of glycolate such as ribulose-1,5-diphos- phate could be oxidized to glycolate when the carbon reduc- tion cycle was not saturated (90. 97). In the eXperiments discussed above, the algae were grown under laboratory con- ditions with 1% C02 or higher in the atmoSphere and then tested for glycolate synthesis or excretion at various CO2 levels. The second way in which 002 effects glycolate metabolism has not been extensively studied although it appears to be of equal physiological importance. Watt and Fogg (99) observed that Chlorella grown in 3% C02 excreted glycolate, while cells grown in 0.03% C02 (air) were capable of excreting glycolate only when the culture was very dilute. Two hypotheses could be proposed to eXplain Watt and Fogg's data: I) growth on air caused a drastic reduction in the cells' ability to synthesize glycolate or II) growth on 3% 002 caused a repression of enzymes metabolizing glycolate 10 and thus forced excretion rather than further metabolism. To differentiate between these two hypotheses, eXperiments have been carried out on the regulation of glycolate metabo- lism in Chlamydomonas by 002. The work supports the second hypothesis, that high C02 represses a glycolate metaboliz— ing enzyme in Chlamydomonas. Enzymes from Green Plants and Algae Oxidizing Glycolate Glycolate oxidase was first described in green tissue by Clagett, Tolbert and Burris (19). This enzyme which appeared to be present in all green plant tissues oxidized glycolate to glyoxylate: glycolate glyoxylate It also oxidized L-lactate to pyruvate; however the Km for L-lactate was too high for the system to be of physiological importance. D-lactate was not oxidized by the enzyme (103). Low levels of glycolate oxidase were found in etiolated tissue, however in the light (31, 86, 87) or upon vacuum infiltration of glycolate into the tissue (53) the level of activity increased. Zelitch and Ochoa (103) showed that glycolate oxidase is a FMN requiring flavoprotein. The enzyme has been crystallized from Spinach leaves and the active Species has been shown to have a molecular weight of 270,000. No metal co-factors have been detected (34). Recent work has established the sub-cellular local- ization of glycolate oxidase to be the peroxisome (92). This 11 microbody apparently contains catalase and most of the enzymes of the glycolate pathway. The close proximity of catalase and glycolate oxidase assures that H202 formed dur- ing the oxidation of glycolate will be rapidly decomposed. The role of glycolate oxidase in algae metabolism is not clear. It was reported by Hess and Tolbert (43) that glycolate oxidase measured as an oxidase was absent in green algae that excreted glycolate. 0n the other hand Zelitch and Day (106) reported the presence of an enzyme in Chlorella and Chlamydomonas which oxidized glycolate to glyoxylate using DCIP as an electron acceptor. Lord and Herrett (55) also reported the presence of glycolate oxidase in Chlorella, but did not show that oxygen was consumed dur- ing the oxidation of glycolate to glyoxylate. Using Nitella, a Charophyte, Downton and Tregunna (27) reported a small amount of glycolate oxidase activity in crude extracts assaying with an oxygen electrode. The above mentioned work did not clarify whether gly- colate oxidase was present in green algae or whether there was a different enzyme oxidizing glycolate which did not link to oxygen. Previous work by Schou et. al. (83) demon- strated that glycolate could be metabolized anaerobically in the dark by Scenedesmus. Nelson et. al. (65) also observed anaerobic glycolate metabolism by Scenedesmus and suggested that green algae at least had an enzyme oxidizing glycolate which did not link to oxygen. To clarify this work the 12 characteristics of an enzyme from Chlamydomonas which oxidizes glycolate have been studied. The results show that it is not an oxidase as found in higher plants and," furthermore,that it is different than the higher plant enzyme by other characteristics. To avoid confusion and to follow the rules of the International Union of Biochemistry (78), the following enzyme terminology will be used. Glycolate oxidase (glycolate:02 oxidoreductase E.C. 1.1.3.1) will refer to enzymes which have been shown to link directly to oxygen, such as glycolate oxidase from Spinach. Glycolate dehydro- genase (glycolate:DCIP oxidoreductase, no E.C. number) will refer.to the enzyme isolated from green algae which does not link directly to oxygen. Glycolate Excretion by Algae One of the most interesting phenomena associated with glycolate synthesis and metabolism by algae is the observation that during photosynthesis under proper condi- tions, large quantities of glycolate are rapidly excreted into the medium. This observation has been made by a large number of research groups working under laboratory condi- tions (61, 74, 89, 97). Even of more significance, eSpeci- ally to the naturalist and ecologist is that this phenomena apparently occurs to some extent in nature. Work by Fogg and associates (32, 33) has shown that glycolate is excreted 13 by natural pOpulations of phytOplankton and that detectable quantities of glycolate can be found in some natural water bodies. Watt (98) has shown that glycolate excretion in nature was highest under conditions cptimum for photosyn- thesis. The ability of algae to excrete glycolate is appar- ently not limited to green algae (Chlorophyceae). Helle- bust (41) found excretion in all major classes of marine algae besides greens, including browns (Chrysophyceae), blue greens (CyanOphyceae), and diatoms (Bacillariophyceae). From Hellebust's work no relationship of pigment content to glycolate excretion can be made. Watt (100) has demonstrated glycolate excretion by the diatom, Stephanodiscus, under natural and laboratory conditions. Studies on the regulation of glycolate metabolism by C02 are presented in this thesis and have been published (66). There are however other factors controlling excretion which are not understood. Chang (1?) using synchronous cultures of Ankistrodesmus and Scenedesmus observed that glycolate excretion occurred during active growth, but not during cell division. This observation has since been con- firmed (35. 95). Hess et. al. (45) observed glycolate excretion only during certain phases of the life cycle of Scenedesmus. It has been suggested that the changes in the level of a glycolate oxidizing enzyme through the cell's cycle could account for this observation (95). 14 Effects of Oxygen on Glycolate Metabolism Oxygen affects glycolate synthesis and excretion and photosynthesis in an opposite manner. It was first observed by Benson and Calvin (10) that glycolate synthesis by green algae requires oxygen. Tolbert and Zill (89) demonstrated that glycolate excretion requires oxygen. Numerous other laboratories have confirmed these results. When oxygen was 90 to 100% of the atmOSphere, glycolate accounted for 10% (8), 30% to 40% (20) or 92% (97) of the newly fixed C02 by Chlorella. While absolute amounts vary, all laboratories have observed this phenomena. The relationship, if any, of enhanced glycolate synthesis to the observed inhibition of photosynthesis by high oxygen (the Warburg effect) (94) is not understood. Studies by Ellyard and Gibbs (30) with isolated Spinach chloroplasts have shown that increasing oxygen concentration leads to increased glycolate synthesis. They were unable to determine the biochemical mechanism which controls this effect. Effect of Light on Glycolate Metabolism Increased light intensity increased glycolate syn- thesis in both algae (74, 98) and higher plants (104). The increase in glycolate synthesis occurred even when photosyn- thesis had been saturated with light (74). The quality of light also affected the glycolate system. Becker et. al. (9) 15 have observed that in blue light glycolate was not excreted by Chlorella, while in red light glycolate was excreted. Hess and Tolbert (44) have observed a complex effect of red and blue light on glycolate synthesis depending on growth conditions. At present no unifying theory can be prOposed to account for the effect of different wavelengths of light on glycolate synthesis. Glutamate: Glyoxylate Aminotransferase The presence of an enzyme which converts glyoxylate to glycine is necessary for the completion of the glycolate pathway as shown in Figure 1. L-glutamate: glyoxylate aminotransferase activity has been shown in pea (21), tobacco (102), wheat (48), and Spinach leaves (49) as well as endOSperm tissue (22). Kisaki and Tolbert (49) have shown that the enzyme was localized in the peroxisome, an observation consistent with its role in the glycolate path- way. A requirement for the transaminase cofactor pyridoxal phosphate has been shown by Wilson et. al. (102) with tobacco leaf extracts. To account for the conversion of glycolate-i-luc to glycine-l-luc in algae and an equal labeling in glycolate and glycine after photosynthetic 1I"'C02 fixation (13), the participation of glutamate: glyoxylate aminotransferase is suggested. The presence of this aminotransferase to the author's knowledge has not been reported in green algae. 16 For this reason crude extracts of Chlamydomonas were examined to determine if this enzyme was present. The results presented in this thesis show the presence, level and amino donor Specificity for glutamate: glyoxylate aminotransferase found in crude extracts of Chlamydomonas, MATERIAL AND METHODS Algae Chlamydomonas reinhardtii Dangeard (-) (No. 90) was from the culture collection of algae at the University of Indiana. Cells were grown at 20°C with 1000 ft. candle light, in low-form Fernbach culture flasks in a growth chamber. The flasks containing 1500 ml culture media were shaken approximately 60 cycles/minute on a reciprocating Eberbach shaker while being aerated with gas from a tube inserted below the level of the culture media. Media were prepared as described previously (66, 71) and autoclaved before use. Flasks were gassed by bubbling into the cul- ture medium either air or approximately 1% C02 in air. Chlorella pyrenoidosa (strain 211/8p) was obtained from Prof. M. J. Merrett. This unusual strain is the same as that used by Merrett in his investigations (55, 59). The algae were routinely grown in low-form Fernback flasks on the same medium used in growing Chlamydomonas and at room temperature (23°C) and light (approximately 200 ft. candles). The cultures were continually aerated with air from a tube inserted below the level of the culture media. Acetabularia mediterranea were obtained from Dr. P. Wolk of the A.E.C. Plant Research Laboratory. They had 17 18 been grown in sun light at room temperature on Erd-Schreiber media (75). Euglena gracilis Klebs "Z" (No. 753) was obtained from the culture collection of algae at the University of Indiana. Cultures were grown at room temperature and light with continuous aeration. The media used was described by Cramer and Myers (23), Scenedesmus obliquus (No. 393) was obtained from the culture collection of algae at the University of Indiana. This strain is the same as Gaffron's D-3. For work with synchronous cultures the algae were grown on a_16 hour day, 8 hour dark regime as previously described (17, 65). When cells were grown for enzymatic analysis, the mediawere the same as used for Chlamydomonas. The cells were then grown at room temperature and light with continuous aeration. Nitella and Chara were obtained from local lakes in fall, 1969 by Dr. W. Wetzel. Nitella was also purchased from Carolina Biological Supply, Burlington, North Carolina. Plants Elodea was purchased from Carolina Biological supply house in Burlington, North Carolina and used within a day of arrival. Spinach was purchased from local markets. Marchantia polymorpha was obtained from Dr. Ole Bjorkman of the Carnegie Institute. 19 Measurement of Glycolate Excretion Chlamydomonas were harvested by centrifugation at 1000 g for 10 minutes at 4°C, washed once in distilled water and then resuSpended in buffer to approximately 2% (v/v). In most experiments 10 mm phoSphate (pH 8.0) was used as the buffer. This pH was chosen because it has been observed that glycolate excretion is promoted by high pH (71). The cells resuSpended in 15 ml were incubated at 200 for 10 min in the dark in a lollipOp container after which the lights, 3,000 foot candles from flood lamps, were turned on with Simultaneous addition of 0.5 ml of 0.2 N NaHCO3 (final concentration 6.5 mm). Throughout the entire eXperi- ment, the cells were aerated with 100% 02. At appropriate times, samples were removed and the cells Spun out at 1000 g for 5 minutes. Aliquots of the supernatant were then assayed for glycolate (14). The assay could detect as little as 50 nmoles per ml. Pretreatment and resuSpension of algae cells has been found not to affect their ability to excrete glycolate (99). Measurement of Glycolate Assimilation Scenedesmus obliquus was harvested at 1000 g, one time washed in distilled H20 and then resuSpended in 0.02 h phOSphate pH 6.5, to give approximately a 2% v/v suSpension. The cells were continually aerated and the temperature kept at 25°C. Light, when used, was at 3,000 foot candles 20 provided by GE flood lamps. Glycolate was added at zero time to give 5.5 x 10'“ M initial concentration. Glycolate was determined in the supernatant of aliquots after the cells had been Spun out at 1000 g for three minutes (14). Measurement of Oxygen Exchange For oxygen exchange eXperimentS Scenedesmus obliquus was harvested at selected stages in the life cycle, washed one time with distilled water and resuSpended in phOSphate, 0.02 M, pH 6.5 to give a 4% v/v suSpension. Oxygen exchange was measured at 27°C on a Gilson Differential ReSpirometer, photosynthesis model, with flood lamps which delivered approximately 1000 foot candles light to the algae. Flask contents were 2 m1 of the cell suSpension and 0.5 ml H20 or other additions in the main part of the flask and 0.5 ml of 0.02 M substrate in one Side arm. In most eXperiments 002 was removed by 20% KOH with a filter paper wick in the center well. Enzyme Assay Methods All assays were carried out at 25°C. The Spectro- photometric assays were done on a Gilford recording Spectro- photometer. In all cases the enzymatic assays reported in this work were tested and found to be dependent on protein concentration over the range used. For assays done with the Spectrophotometer or oxygen electrode initial rates were 21 determined. In the case of fixed time assays, they were shown to be linear with time. P-glycolate PhOSphatase (PhOSphoglycolate phOSphohydrolase E.C. 3.1.3.18) This enzyme was assayed by the method of Anderson and Tolbert (1). The assay mixture contained 0.5 m1 cacodylate 0.20 M pH 7.0, 0.6 ml EgSOu 0.01 k, 1.0 ml 0.01 M P-glycolate (tricyclohexylammonium salt, General Biochemicals), water, and enZyme in 2.0 ml final volume. The reaction was terminated by addition of 1 ml of 10% trichloroacetic acid. Glycolate Oxidase (Glycolate: 02 oxidoreductase E.C. 1.1.3.1) Glycolate oxidase was assayed by following the reduc- tion of DCIP anaerobically at 600 mu (92). The assay mix- ture in a total volume of 2.5 ml contained 200 umoles pyro- phOSphate pH 8.7. 0.3 umole DCIP, enZyme and water or addi- tional components. The reaction was initiated by addition of 20 umoles of glycolate or other substrates from the side arm of the Thunberg cuvette. Changes in O.D. were converted to nmoles using an extinction coefficient of 21.9 cm-1 x mole-1 x 10-3 for DCIP (a). Two additional methods were used to assay glycolate oxidase and to characterize the enzyme as an oxidase. The 22 formation of glyoxylate phenylhydrazone at 324 mm was mea— sured similarly to that described by Hess and Tolbert (43). In 2.5 ml were 200 umoles pyrophoSphate, pH 8.7, 10 umoles phenylhydrazine-HCl (previously neutralized), enzyme, water and other components. The reaction was initiated by addi- tion of 20 umoles glycolate. Glycolate oxidase was also assayed by measuring the disappearance of oxygen with a Clark OPE oxygen electrode at 700 millivolts. The 3.2 ml volume reaction mixture in a continually stirred, temperature controlled chamber contained 250 umoles pyrophOSphate, enzyme, water, and other components. The reaction was initiated by addition of 20 umoles glycolate. Oxygen concentration was calculated on the basis of solubil- ity of air in water. All solutions for the assay were air saturated except the enzyme. At 25°C, the reaction mixture held 685 nmoles 02 (40). Percent change in oxygen concen- tration recorded in 5 minutes was converted to nmoles 02 x minute'l. Glycolate Dehydrogenase (Glycolate: (DCIP) oxidoreductase) Glycolate dehydrogenase was routinely assayed anaero- bically by following DCIP reduction as described for glyco- late oxidase. When measuring enzyme activity with the oxygen electrode the assay described for glycolate oxidase was used, with the addition of 8 umoles of PMS (Sigma). 23 To determine electron acceptor Specificity for glyco- late dehydrogenase the following assay systems were tested. N05 reduction was assayed by looking for the appear- ance of N0§ (68). In an assay volume of 2.0 ml were 100 umoles pyrophoSphate pH 8.7, 6 umoles KNOB, 20 umoles glyco- late, water and enzyme. The reaction was terminated by boiling and a one ml aliquot was assayed for N053 Cytochrome c reduction was assayed by looking for increased absorption at 550 nm. The assay in 2.5 ml final volume contained 200 umoles pyrophoSphate pH 8.7, 2 mg cytochrome c (Sigma type II), water, enzyme, and 20 nmoles glycolate. The assay was done anaerobically. Potassium ferricyanide reduction by the enzyme was tested by looking for Fe(CN)g‘ reduction at 410 nm in an anaerobic Thunberg cuvette. The assay mixture contained in 2.5 ml final volume 200 umoles pyrophoSphate pH 8.7, 2.5 umoles K3Fe(CN)6, freshly prepared, water, enzyme and 20 umoles glycolate. NAD+ and NADP+ reduction was tested by measuring increased absorbancy at 340 nm in an anaerobic Thunberg cuvette. The assay contained in 2.5 ml, 200 nmoles pyro- phoSphate pH 8.7. 0.8 “moles NAD+ (Sigma type III) or NADP+ (Sigma type II), 20 umoles glycolate, enzyme and water. Methylene blue reduction was looked for at 668 nm in anaerobic Thunberg cuvettes. The assay in 2.5 ml 24 contained 200 umoles perphOSphate pH 8.7, 0.4 umoles methylene blue, 20 nmoles glycolate, enzyme and water. Glutathione reduction was measured by looking for the appearance of glyoxylate phenylhydrazone. The assay was carried out as described for glycolate oxidase with the addition of 2 umoles glutathione (Sigma). In order to determine the stoichiometry between DCIP reduction and glyoxylate formation both aerobically and anaerobically, a method for the determination of glyoxylate by the phenylhydrazine assay in the presence of DCIP was develOped. Normally phenylhydrazine and DCIP react at pH 8.? leading to complete loss of DCIP color which is incompatable with the enzymatic assay. To avoid this problem the activity was measured in a standard DCIP~ reduction assay, but at fixed time intervals different assays were terminated by addition of 100 ul 12 N HCl. The mixtures were left standing for 20 minutes in which time the blue DCIP color disappeared. After this time 0.5 ml of 0.1 N phenylhydrazine-RC1 was added to the assay mixture. After another 20 minutes the absorption due to glyoxylate phenylhydrazone was measured at 340 mu. Spectra in Figure 2 Show the absorption of glyoxylate phenylhydrazone in the presence and absence of DCIP. The' presence of DCIP and strong acid caused a shift in the absorption maximum of glyoxylate phenylhydrazone from 330 nm to 344 nm. The absorption Spectrum of the product formed 25 Figure 2. Absorption Spectra of Glyoxylate Phenyl- hydrazone in Presence and Absence of DCIP Spectra measured on a Beckman EB recording Spectrophotometer. Samples were prepared as des- cribed in Methods and Materials. Each sample con- tained 2.5 ml final volume. ( ). 200 umoles pyrophOSphate pH 8.7, and 10 umole phenylhydrazine' HCl; ("°-°), 200 umoles pyrophOSphate pH 8.7, 0.30 umoles DCIP, 100 ul 12 N HCl and 10 umole phenol- hydrazine-HCl; ( ----- ), 200 umoles perphOSphate pH 8.7, 0.1 umole glyoxylate, and 10 umoles phenyl- hydrazine-HCl; (-----), 200 nmoles pyrophoSphate pH 8.7, 0.1 umole glyOXylate, 0.3 nmoles DCIP, 100 ml 12 N HCl and 10 umoles phenylhydrazine-HCl. 26 ooc 8.634 340 380 300 Wavelength in nm 27 from the oxidation of glycolate by glycolate dehydrogenase in the presence of DCIP was identical with authentic gly- oxylate phenylhydrazone. The absorption at 340 nm was linear with glyoxylate concentrations between 0 and 0.5 umoles when measured using the assay conditions described above. Preparation of Cell Free Extracts For eXperimentS to determine the level of glycolate dehydrogenase and P-glycolate phoSphatase in Chlamydomonas, the cells were harvested at 1000 g for 10 minutes at 2°C, washed once in distilled water and resuSpended in 0.001 M phoSphate pH 7.0, to give approximately 20-40% v/v cell suSpension. The suSpension was passed through a pre-cooled French pressure cell at 8000 to 12000 lb-inchz. Cell debris was removed by centrifugation at 29,000 g for 10 minutes at 2°C. The supernatant was used for the assays. For determination of glycolate dehydrogenase in other algae, cells were treated as described above except that they were suSpended in 0.1 M phoSphate pH 7.5. Acetabularia, Nitella, Chara, Elodea and Marchantia were washed with cold tap water, rinsed with cold buffer and cut into small pieces. The pieces were ground in a pre-cooled Potter-Elvehjem homogenizer with 0.1 M phOSphate pH 7.5 at 2°C. The cell debrkswas removed by centrifugation at 10,000 g for 10 minutes at 2°C. The supernatant was used for the assays. 28 Glycolate Oxidase Spinach glycolate oxidase was the gift of Dr. S. L. Vandor. This preparation was prepared by the method of Zelitch and Ochoa (103) through the acid precipitation step. The enzyme was kept frozen until use. Purification of Glycolate Dehydrogenase For the purification of glycolate dehydrogenase from air grown Chlamydomonas the enzyme was removed by a nonionic detergent treatment. Cells were harvested at 1,000 g, washed once in distilled water, washed once in 0.1 M phOSphate pH 7.5 and resuSpended approximately 10% v/v in 0.1 M phOSphate pH 7.5 at 2°C containing 1% w/v Triton X-100 (Hohm and Haas). The mixture was stirred for 45 to 60 minutes in the cold room, after which the cell debris was removed by centrifugation at 29,000 g for 10 minutes at 2°C. The supernatant was used for further purification. All Operation was carried out at 20 to 4°C. To the Triton X-100 extract, solid (NH4)ZSOu was added with constant stirring to give 35% saturation. The mixture was centrifuged at 39,000 for 10 minutes and yielded a green residue and a clear light yellow superna- tant. The green residue was removed and discarded. 'The supernatant was 50% saturated with solid (NH4)2804 and the precipitate, containing the enzyme was removed by cen- trifugation at 39,000 g for 15 minutes. The precipitate 29 was suSpended in 1/10 the original volume of 0.1 M phOSphate pH 7.5. This preparation was used for most studies concern- ing the characterization of glycolate dehydrogenase. Glutamate: Glyoxylate Aminotransferase (E.C. 2.6.1.4) This enzyme was assayed at 25°C by following forma- tion of glycine-luC in the manner described by Kisaki and Tolbert (49). In a final assay volume of 1.25 ml were 20 umoles glyoxylate-1,2-1uC, 12 nmoles amino donor, usually L-glutamate, 15 nmoles phOSphate (pH 7.5), 0.1 umole pyridoxal- 5-phOSphate (Sigma) and enZyme. The reaction was initiated by addition of glyoxylate, and terminated after 15 minutes by boiling. Glycine-Inc was separated from unreacted glyoxy— late by passage of the boiled reaction mixture over a Dowex-l acetate column (6 x 50 min) which was then washed with 2 ml of water. From the combined effluents 0.4 m1 aliquots were counted for glycine-14C in 15 ml Kinards solution (47) in a Packard Tri-Carb Scintillation counter. The counting effi- ciency was determined to be 71% with standard benzoic acid. Observed cpm were converted to umoles glycine-luc The glyoxylate-1,2—1uc (98% pure glyoxylate-1,2-14C, Calbiochem) was prepared by dilution with a 1000 fold excess cold glyoxylate to give a 0.2 M solution containing 1 uc/ml. Control experiments demonstrated that glyoxylate-1,2-14C was retained by the Dowex-i acetate. Glycine-1,2-14C was not 30 retained significantly by the column as control recovery eXperiments gave a minimum of 95% of the eXpected yield. Enzyme was prepared for this assay by harvesting air grown Chlamydomonas, washing in distilled H20 and resuSpending 20-40% v/v in 0.02 M phoSphate pH 7.5. The cells were put through French.Press twice at 10,000 to 12,000 lb-inchz. Cell debris was removed at 29,000 g for 10 minutes. The supernatant was used in the assay. Protein and ChlorOphyll Determination Protein was determined by the method of Lowry et. al. (56) using bovine serum albumin as a standard. Chloro- phyll was determined by the method of Arnon (5). RESULTS AND DISCUSSION Assimilation of Glycolate by Scenedesmus The ability of Scenedesmus obliquus to assimilate exogenous glycolate was found to be related to the life cycle of the cells. As shown in Figure 3, glycolate assim- ilation was maximum at the stage of cell division and shortly afterwards. No assimilation was detected during the middle of the light phase of growth. Furthermore, glycolate assimilation could not be detected in random growth cultures of Scenedesmus, Ankistrodesmus, Chlorella (Warburg), or Chlamydomonas. Glycolate assimilation by dividing cells was linear with time over the periods tested in both the light and dark (Figure 4). Assimilation was equally rapid in light or dark. Addition of 1 x 10-5 n cwu, an inhibitor of photo- synthetic electron tranSport (101), had no effect on glyco- late assimilation in the light (Figure 4). An inhibition by low pH on assimilation was examined. At pH 2.9 in 0.02 M phOSphate no assimilation was observed while assimilation was observed at pH 6.5. This observation is inconsistent with Schou et. al. (83), who observed assimilation at pH 2.9. PhOSphate was not required as equal assimilation was observed with 0.1 M HES pH 6.5 in both the light and dark. 31 32 .SOfimeHEmsm oaaoomoaoaa an pmsaaaopmo was apnoam -Hmo mo 0 mp0 .4 duo« x m. m was soapmapcoomoo mpmaoomam HoHpaQH .mamaaopmg dam mpocmpm- . c2 omnaaommm mm consonants psu j” 95 SH cacao. nmaaaammm mpmaoo>au pso m. b m0 mpmSQmozp a No. 0 SH >\> RH poozmpmSm .oaomo mafia map mo mowmpm Snoaomme pm oopmo>am£ macs wHHoo macaw opaq aaone mo mowmpw pcoammmdm pm msamooocoom pomasoasosmm an opmaoomao mSosmwoxm mo zodpwaaaammw .m mammam 33 vm ON m_ N_ m v 0 d \\ O '''' 0’ _ \\\ l \\ \ / p a l x ’ \ a \ a . at N a 1 ~ a; ~ ’ ~ I . / .. z o A) 20: 4 all {on Illuv. \ \ \ N. m \ \“t. Q. “Q 302» £320 :3 $02» c0336 :3 '0. <\! —. o o 0 V. 0 900m I_|tu x Fugw 017 x ewoidn ewpofllb se|0ulr1 34 Figure 4. A. Glycolate Assimilation by Synchro- nized Scenedesmus in the Light and Dark Scenedesmus was harvested at the stage of cell diviSion, suSpended to 2% v/v in 0.02 M phOSphate pH 6.5 and glycolate assimilation in the light or dark measured as described in Methods and Materials Initial glycolate concentration was 5.5 x 10—° H: ,A - assimilation in dark; (3 - assimilation in the light. B. Effect of CRU on Glycolate Assimi- lation by Synchronized Scenedesmus §genedesmus was harvested at the stage of cell division resuSpended to 2% v/v in 0.02 M phOSphate pH 6.5 and glycolate assimilation mea- sured in light t 5 x 10-5 m own as described in Methods and Materials Initial glycolate concen- tration was 5.5 x 10-4 M. C) - control; A - 1 x 10-5 M CIIEU. nmoles glycolate x ml" umoles glycolate x ml" 0.6 0.5 0.4 0.3 0.2 DJ 0.6 I; 0.5 0.4 0.3 0.2 0.! 35 J I 1 IO 20 minutes \b t \& L 1 IO 20 minutes _J 30 36 This data confirms the ability of Scenedesmus to assimilate exogenous glycolate. The process does appear however to be limited to certain stages of the life cycle of the cells, at least for Scenedesmus. It is not yet clear why some workers have obtained glycolate assimila- tion with random cultures (61, 83), while others have not (16, 43, 57). The suggestion is made on the basis of my data that the cultures used previously were actually in a stage of unrecognized synchrony which permitted glycolate uptake. The relationship between glycolate assimilation and glycolate excretion is of interest. Chang (17) using the same cultures of Scenedesmus as used in this thesis and also using synchronous Ankistrodesmus observed that glycolate excretion was maximal during the light or active growth stage, and that little or no excretion could be detected when the cells were in the dark or division stage. These observations have been confirmed (35, 95). The active stage of growth is the stage where no glycolate assimila- tion occurs (Figure 3). Thus there appears to be an inverse relationship between glycolate excretion and assim- ilation. It is not known whether glycolate assimilation is regulated by factors involved in uptake or whether ability or inability to be metabolized by the cell is the regulating factor in assimilation. 37 Effects of Glycolate on Photosynthesis and Respiration The effects of exogenously added glycolate on the rates of photosynthesis and reSpiration by Scenedesmus measured at a time when cells were capable of assimilating glycolate have been published (65). These results will' only be summarized here. As shown in Figure 5, glycolate stimulated reSpiration approximately 100% over controls. When compared to other substrates (Table 1), acetate and glucose were two to three times more effective in stimulat- ing reSpiration. Similar results have been obtained by Sen (8#) with Chlorella. The level of stimulated reSpira- tion by glycolate was much higher with Scenedesmus than with Chlorella however. Glycolate stimulated oxygen evolution during photo- synthesis by Scenedesmus taken at a time when they were capable of glycolate assimilation (Figure 5). Compared to other substrates tested, Table 1, glycolate was the best substrate. The observation that glucose and fructose can stimulate oxygen evolution has been confirmed (72). Glyco- late stimulated oxygen evolution was inhibited by 6 x 10'6 h CNU, an inhibitor of photosynthetic electron tranSport (101). A scheme to account for these observations is shown in Figure 6. The necessary enzymes to Operate the postu- lated cycle have been found in green algae (12, 66). It should be noted that the prOposed cycle will only account for stimulated oxygen evolution if the enzyme oxidizing 38 - - .: muoH N m.m mpmaoomam m msm m.o mm .mpmzomosp ; No.0 SH sofimfibap Haoo mo mwwpm mSp pm mnemoposmom pmudsoa£osmm do SoamsmamSm &N.N m no HE m pmsampzoo Mmdam 20mm msammpmsmom an xamQ ms» sa scapmaammwm psm unwaq mgp 2H Scapsaobm smwmxo Co mudaoomao mo pomwmm .m ostHm 39 on Amy—52:5 NEE. 0v Om ON 0. xmdo A _ _ _ _ F d \ \ \ \ FIG... wh<400>40 r \_ / 54588 oz / _ _ _ _ _ ./ _ _ n + /. ,x O O N O 10 xSV1a 83d aewvaoxa 2o m 0 Q on 40 Table 1. Comparison of Different Substrates for Stimulation of Oxygen Evolution and ReSpiration Data are eXpressed as ratio of activity with each substrate as compared with glycolate. Final substrate con- centrations were 3.3 mm and rates were corrected for con- trols without added substrates. Oxygen Evolution Oxygen Uptake Substrate in Light in Dark Activity ratio as substrate/glycolate Organic Acids Glycolate 1.0 1.0 Glyoxylate 1.0 1.0 Acetate 0.5 2.5 P-Glycolate (0.1 <0.1 D,L-Lactate 0.2 1.0-1.2 Formate <0.1 0.5-1.0 D,L~Glycerate 0.5-0.7 0.5-1.0 3-P-Glycerate 0.5-0.7 0.5-1.0 Amino Acids L-Serine 0.5 1.0 Glycine 0.5 1.0 Sugars D—Glucose 0.4 2.0-3.0 D—Ribose 0.2 ‘ 0.3 ID-Fructose 0.2 0.8 41 scapsaobm Cmumxo mo Cofipmasadpm mudaoozac you pssooo< op mamsom ommoaoam w .0 madwfim 1+2 n.._.< ~zx /\ 22:36 /\ .1 + xaoqz / \ «om. + of $0500.. a o 30 o: o toamcot. c8805 0 c o . n 1 28553.05. x \ / 32°26 \/ .842 \ / cuzm L13 glycolate is not an oxidase. Otherwise stimulated oxygen evolution and oxygen uptake due to glycolate oxidation would be equal and no net stimulation would be observable. Since the prOposal of this cycle in algae (65), the enzyme in green algae oxidizing glycolate has been found to be a dehydrogenase (this thesis) and not an oxidase. A scheme similar to this has been prOposed by Asada et. al. (6) to account for glyoxylate stimulated photophOSphorylation in isolated chlorOplasts. Glycolate Excretion by Chlamydomonas Chlamydomonas reinhardtii, grown on air supplemented with 1% 002 was capable of excreting glycolate when tested under the cptimum conditions described in Methods and Materials. Glycolate excretion at detectable levels was not found when cells were grown on air. The ability of cultures grown on air supplemented with 1% 002 to excrete glycolate drOpped off with increasing culture density (Figure 7). This also occurred in older cultures, an observation which confirmed a previous report (89). This phenomenom may be partially related to availability of C02 per cell as the culture density increases. It was found that transferring cultures from C02 supplemented air to only air resulted in the disappearance within about 15 hours of glycolate excretion by the cells during the test assay (Figure 8). Likewise when cultures were transferred an .oMSpHSo do HE 9mm HHmSQoaoHSo an mo mammn exp :0 ommmmaexo ma mpdmsmo casuaso ”mamaampmz one woospm; ma oopahommp mm Aopmnomosm : No.0v 0.x mp pm pswaa 05p CH scapmaoxo opdaoomam new venue» one oopmobads mums maamo mwspaso one no spamemp,mmm owe ms» op mpdpmamm wd< 2H moo RH 20 cacao monosoomamaso an sodpoaoxm mpmaoomao .m maswam 45 qIO -\> &m on poochmSmoh use oopmmbaw .ombofima was manuaso one do scapaoa d .mmaap mumahmoaaaw p4 .A 00 Mmo.ov has on omhammmsdap mama was Ca moo ma :0 wsasoam mohfipazo aa< op saw 2“ Nooima Eon“ omhammmsmae msamm hopwg opmaoomam mpoaoxm op wmsoaoomadano no mpaaanw SH mmwmaomm .m wadsam 1+7 a: E O (D C"z <1 a: ‘— a: m of. ‘ ’4 m a: 3 o I _.__‘l L l O 8 8 § 0 V Mudwoluo '_6w x I_ugu.l x emIOOMO se|owu 48 from air to 002 supplemented air, they regained the ability to excrete glycolate within about 15 hours. In order to determine whether the absence of glyco- late excretion in air grown cultures was due to the absence of glycolate synthesis or the inability of the cells to excrete glycolate, eXperiments with isonicotinyl hydrazide were performed. Pritchard et. al. (7%) have shown that iso- nicotinyl hydrazide stimulated glycolate excretion by Chlorella grown on air supplemented with H% C02. They pro- posed that isonicotinyl hydrazide acted as an inhibitor of the glycolate pathway, thus forcing excretion of glycolate. Air grown Chlamydomonas, when treated with 0.02 M isonico- tinyl hydrazide in 1 mm phOSphate (pH 7.0), excreted glyco- late (Figure 9), but not in the absence of this inhibitor. Therefore the absence of glycolate excretion by air grown cultures was not due to their inability to synthesize glyco- late, but rather their failure to excrete it. Glycolate Dehydrogenase: Detection and Assay Cells of Chlamydomonas were harvested and the crude extracts tested for glycolate oxidase activity using the three assays described in Methods and Materials. ho activ- ity in crude extracts could be detected with the oxygen electrode or phenylhydrazone assays (Table 2). Glycolate dehydrogenase (glycolate: DCIP oxidoreductase) activity was found. All glycolate dehydrogenase activity was soluble after centrifuging crude extracts 10 minutes at 29,000 g. .oeppano we: moauenomn Handpooanomd penp paeoxe maaeoapnepd cepeeap mes one ennpano eaew enp Song mez Honpnoo e29 .maefinepen one moonpes n“ penanowep me unwaa one na epeaoomfiw epenoxe on mpaadne Mom empmmp use no.5 mov-mpesomoga a Hoo.o .menumness Hesspoodcomn 2 No.0 na >\> MN on peoneomnm one oepme>nen enez nae no nzonm maaeo 49 menoaoowaeanu nzoau Man an noapenoxm epeaoomao no eofiwenomm HanapooanomH no poemnm .0 ennwfim 50 mm...32:2 on Cc Om ON 0. . iv .. . . Jomkzoo mo_NI ..._>z_._.oo_zom_ + 9 O N 0 f0 .-IU1 x ammo/('6 salowu 51 Table 2. Absence of Endogenous Inhibitors of Glchlate Oxidase and Dehydrogenase in Crude Extracts of Chlamydomonas Activity determined in 29,000 g supernatant of French Press extracts buffered with 1 mm phoSphate pH 7.0. Assays described in Methods and Materials. nmoles x min"1 Phenylhydrazone Assay crude extract Chlamydomonas Spinach glycolate oxidase combined Oxygen Electrode Assay crude extract Chlamydomonas Spinach glycolate oxidase combined DCIP Reduction Assay air grown Chlamydomonas crude extract 1% C02 grown Chlamydomonas crude extract combined extracts 15.1 15.3 20.14. 20.u 57.5 ,0 , I . 64.? 52 Tests were run to determine if endogenous inhibitors of glycolate dehydrogenase or glycolate oxidase were present in the crude extracts of Chlamydomonas cells (Table 2). Glycolate oxidase from Spinach leaves was tested with an equivalent amount of algal enzyme on the basis of DCIP reduction, and no inhibition of oxygen uptake or glyoxylate formation by the Spinach enzyme was observed. When extracts of Chlamydomonas grown on 1% COZ in air, which had DCIP reductase activity were mixed with extracts of air grown cells, no inhibition of DCIP reduction with glycolate was observed (Table 2). Therefore it appears that natural inhibitors of glycolate dehydrogenase or glycolate oxidase are not present in Chlamydomonas extracts. During the course of this investigation glycolate dehydrogenase was recognized as a DCIP reductase (106), and although assayed for by DCIP reduction it had been called glycolate oxidase in Spite of the fact that no oxidase activity was observed. On the basis of the data presented in Table 2 and in the Electron Acceptor Specificity section of this thesis the glycolate dehydrogenase of Chlamydomonas is not an oxidase as is found in higher plants. Thus while Chlamydomonas lacks glycolate oxidase, it has a glycolate dehydrogenase which carries out the oxidation of glycolate to glyoxylate. 53 Levels of Glycolate Dehydrogenase and P-Glycolate PhOSphatase in Chlamydomonas In view of the above data on glycolate excretion, it seemed reasonable to examine the levels of enzymes involved with glycolate metabolism to see if alterations in their activity could account for the difference in excretion ability. P-glycolate phOSphatase has been suggested to be involved in glycolate synthesis (91). The level of this enzyme did not change significantly when cells were trans- ferred from 1% C02 supplemented air to air (Figure 10), therefore it does not appear as though the phOSphatase is involved in the regulation of excretion ability. The effect on glycolate dehydrogenase of transferring Chlamydomonas from 1% C02 supplemented air to air was deter- mined (Figure 10). The enzyme level increased three fold on the basis of either protein or chlorOphyll, simultaneous with the loss of the ability of the cells to excrete glyco- late. The levels of glycolate dehydrogenase determined on the basis of chlorOphyll and protein in five separate experi- ments are shown in more detail in Table 3. Glycolate dehyd- rogenase activity was at least two to four times higher in air grown cells than in cells that were capable of glycolate excretion during growth on air with C02. To determine if the increased activity of glycolate dehydrogenase was due to enzyme activation or new protein 5h .naepona HIwa x HInHe N meHoEn .emeponoen I .Hamnnonoano HIwa N «Inna N meaoan .emeponoenooaxo mHoQ "epeHoome "namponn HIme N aInHa x emaoa: .emepennmona "damnaonoano HIwa x HInHB x meaoaa .emepenneona epeaoohHwIm I I OCTQ.’ "mHeHHepe; one moonpes nH oenanomeo me oememme one: weamnnm .o.m ma openawona : Hoo.o nH oenenean enez mpoenpxe one .mpoenpxe eonno \. mmenm nonenm Bonn unevenneanm u ooo.dN na oenannepeo one: naeponn one Hamnaonoano no mamen enp no meeznne enp mo mefipabdpoe onHoeam nan op nan na moo RH Bonn nemmneaa nepne menoaomwaeano na nemeposoen Iooaxo mHoQ “epeaoomaov emenemonomnem epeaoomau one emepenamonm epeaoomHuIm no mpabapon ouhaoenm na newneno .3 when; 55 BSVLOHCIBH dIdOO :31V‘IOOA‘IO Mudomluo I_6uI x I_quI x salowu x 2-0' uIaIOJd I”6w x '_quI x 39mm x 9'0 [o _ r l I <1 on) Si 3 -0“- u.I < m cum 0) l— a) z 4 4 4 q I— I l— o 3) g a: 8 o o *- III I In I: a. I: \ a: LIJ I- <1 10 b -Qu. \ 4 (D / \ m D \ O I In N — uIeIOJd I"6m x '_U!Lu x 99'0er x z0| Mudwomo 'jw x I_qu.I x se|ouIrI BSVLVHdSOHd 31V'100A19-d 56 Table 3. Glycolate Dehydrogenase in Chlamydomonas Grown on Air or 1% C02 in Air Specific activity of glycolate dehydrogenase in 29,000 g supernatant of French.Press extracts determined by DCIP reduction on basis of chlorOphyll or protein as described in Methods and Materials. Data from five cultures of air grown and five grown on 1% C02 in air capable of excreting glyco- late. Air Crown 1% C02 in Air Chlorophyll Protein ChlorOphyll Protein nmoles x min"1 x mg"l nmoles x min"1 x mg"l range 260-520 2.8-4.9 80-150 0.7-1.7 average 388 3.8 122 1.1 57 synthesis, an inhibitor of protein synthesis, cycloheximide (29), was used at a concentration of 5 ug/ml. This inhibi- tor should prevent the increase in levels of glycolate dehydrogenase if new protein synthesis is involved, when cultures are transferred to air from air supplemented with 002. It has been shown that cycloheximide does not effect the C02 fixation ability of Chlorella (62). In the presence of cycloheximide levels of glycolate dehydrogenase did not increase (Figure 11). This result suggested that the increase in glycolate dehydrogenase was due to de novo syn- thesis of the enzyme. This data suggests that the level of glycolate dehyd- rogenase present in Chlamydomonas controls whether the algae will excrete glycolate or metabolize it. Cells grown on high C02 excrete glycolate and have low dehydrogenase levels, while cells that do not excrete glycolate have elevated dehydrogenase levels. Both types of cells synthesize glyco- late as shown by the isonicotinyl hydrazide data. This is further supported by the observation that levels of P-glyco- late phOSphatase did not vary significantly between high and low 002 grown cells (Figure 10). Therefore the availability ofCOz during growth regulates glycolate excretion by enhanc- ing further_glycolate metabolism rather than effecting glycolate synthesis. 58 .Hannnonoano ann N nInnn H moaonn .oonanxonoaono I unnoponn nImn x nInna N onoan .oonannonoaoho I "Hamnnonoano nImn w nInna x ooaonn .Honpnoo I unnoponn nImE N «Inna x moHan .Honpnoo I "0.5 m9 opennmonm 040+ : noo.o nn oonemonn onoz mpoenpxm .maennopea one moonpom nn oonnnomoo me mpoenpno moonm nononn Bonn mpnepennonnm w ooo.mN on» nn memme non» Ionoon mHoQ onp an oonnanopoo mez omenowonomnoo opeHoomHm no mpn>npoe onnnoonm .mnnon : nopne ooooe mes Annnooa onnpano Ha\mj mv oonaHXon Ioaomo one nne op nne nn moo NH Bonn oonnonmnenu one: monnpano nae op nne on Nov Rn aonn menonoohneano no nonmnene nopn< omenowonomnom ooonoonno no npn>npo< onnnooom so monanwosonono no poonnm .nn onswnn 59 S Mudwomo ,_bw x '_quI x 99mm 8 8 l0 mummzasmh mm...m< mmDOI mm OW n. o. m o .l w— + )H‘mAI «-01. -m IId I «I s I . mo_s:meOJO>o “.0 29.2094 In ugeIOId I“5m x I,_u_IuI x 99mm 60 Purification of Glycolate Dehydrogenase Glycolate dehydrogenase was purified from air grown Chlamydomonas according to the scheme shown in Figure 12. The enzyme was recovered in relatively good yield in a preparation free of chlorOphyll and other major interferring pigments. The results of a typical purification are shown in Table 4. While this table shows only a 3.3 fold purifi- cation, it must be remembered that the preparation begins with the Triton X-100 extract. The Triton X-100 extract preparation was a partial purification step in itself. When compared with the activity found in crude extracts (Table 3), it can be seen that Triton X-100 extraction gave approximately a three fold purification. Using this method of calculation, purifications as high as 30 fold have been obtained. The use of Triton X-100 to release glycolate dehyd- rogenase from cells was studied. As shown in Figure 13, there was an initial releaSe of approximately 50% of the enzyme. The rest of the activity was more slowly released until a maximum of 82% of the total in French Press extracts was obtained. Whether examined on the basis of protein or chlorOphyll a Similar release rate was obtained. As can be seen from.Pigure 13, shorter time periods of Triton X-100 extraction could lead to increased purification. This occurs, however, with a decrease in enzyme recovery, and generally the longer extraction time for more recovery was used. Figure 12. 61 Purification Scheme for Glycolate Dehydrogenase* cells in 1% w/v Trition X-100, 0.1 M phOSphate pH 7.5 stir #5 min to 60 min centrifuge 29,000 g (10 min) pellet discard green supernatant make 35% (NH4)2804 saturated centrifuge 39,000 g (10 min) clear yellow green residue supernatant (discard) make 50% (NH ) so saturatgd2 h centrifuge 39,000 g (15 min) I supernatant (discard) pellet resuSpend in 1/10 volume 0.1 M phOSphate pH 7.5 *All work done at 2°-4°C. 62 m.m mm m.n no In- oon nOnpeonnnnnm mno>ooom & n.mm oomm oonv m.mn ooom m.nn omos Annoponn HIwa a HInns x moaonnv Aooodomm mHUQV nonpnooe onnnooom mango nopon oaono> NH OHH OHH OOH HE ponnoo sommnsmzv one unevennonnm sommnemzv one unevennonnm sommnsmzv eon poenpxo ooalx nOpHHB nanpoenm omenomonomnom opeaoomau no nonpeoananm y: oases 63 Figure 13. Specific Activity of Glycolate Dehydro- genase Released from Chlamydomonas by 1% Triton X-100 Glycolate dehydrogenase activity was deter- mined by DCIP reduction in the 29,000 g supernatant of Triton X-100 extracts. Extracts were prepared by stirring cells (10% v/v) in 1% Triton X-100 w/v 0.10 M phOSphate pH 7.5. (3- umoles x min"1 x mg- chlorOphyll; 15- nmoles x min"1 x mg“1 protein. 61+ 228.620 79: x TEE x $.08: Am \\ A\\ la\m\m a 0.. £295 79: x E? x 8.02.96: 75 60 45 minutes 30 IS 65 Further purification of glycolate dehydrogenase was not obtained in many attempts. This was due to the insta- bility of the preparations and the lack of large amounts of starting material. Consequently studies with the enzyme were run on the (NH4)2304 fractionated preparation and referred to as the partially purified dehydrogenase. Substrate Characterization of Glycolate Dehydrogenase The affinity of partially purified glycolate dehyd- rogenase for glycolate and other substrates was tested. The results showed glycolate to be the preferred substrate, while D-lactate, L-lactate, glyoxylate and D,L-a-hydroxy- butyrate also served as substrates (Table 5). The affinity of the enzyme for D-lactate is of Special interest since other workers have shown that glycolate oxidase from higher plants does not oxidize D-lactate, while it does oxidize L-lactate at a rate of about 60% that of glycolate (79, 103). This rate of oxidation with Spinach glycolate oxidase was confirmed. The affinity of glycolate dehydrogenase for glyoxylate and D,L-a-hydroxybutyrate was similar to that found for higher plants for glycolate oxidase (79). Mixed substrate experiments were carried out to determine if activities observed with glyoxylate, D and L- lactate could be attributed to one or more enzymes. The rates of oxidation of mixtures of substrates were not addi— tive (Table 6). The results indicated that all substrates 66 Table 5. Substrate Specificity of Glycolate Dehydrogenase Relative rates of DCIP reduction with glycolate taken as 100. Substrates at 8 mM final concentration. 16 umoles of D,L mixtures were added to give 8 mM final concentration of each. Substrate Relative Activity Glycolate 100 D-Lactate 50-70 L-Lactate 10-25 Glyoxylate 30-50 D,lpd-Hydroxybutyrate 50-75 Glycine 0 meso-Tartrate D,L-Malate D,L_aAPhenyl Lactate D,L-BAPhenyl Lactate P-Glycolate OOOOOO D,L-Glycerate 67 Table 6. Mixed Substrate Assays with Glycolate Dehydrogenase Relative rates of DCIP reduction with glycolate taken as 100. All substrates at 8 mM final concentration. Substrate Relative Rates Found Predicted Glycolate 100 Glyoxylate 34 Glyoxylate + Glycolate 82 67 L-Lactate 22 L-Lactate + Glycolate 63 61 D-Lactate 7 66 D-Lactate + Glycolate 83 83 68 were oxidized by the same enzyme. The slightly high value obtained with glyoxylate can be eXplained by contamination of the glyoxylate with glycolate. The Km's for glycolate and D-lactate were determined using the method of Lineweaver and Burke (5h). The Km for glycolate oxidation by glycolate dehydrogenase from Chlamydomonas was 2.2 x 10“” M (Figure 14). A high Km of 1.5 x 10'3 M for D-lactate oxidation was obtained (Figure 14). Thus the oxidation of D—lactate may be of no physio- logical importance to the glycolate dehydrogenase system. Electron Acceptor Specificity of Glycolate Dehydrogenase The ability of the partially purified glycolate dehydrogenase to reduce various electron acceptors was investigated. The results indicated that only DCIP and PMS, of all the acceptors tested, would link to glycolate dehydrogenase (Table 7). No oxidase activity could be detected with either the oxygen electrode or the phenyl- hydrazone assay. Glyoxylate formation was measured in relationship to DCIP reduction anaerobically and also measured aerobically in the presence and absence of DCIP (Figure 15). Aerobic glyoxylate formation was dependent on the presence of DCIP, no glyoxylate formation was detected in its absence. This experiment confirmed the results of Zelitch and Day (106), who reported a 1:1 stiochiometry between DCIP reduction and glyoxylate forma— tion. (ill! II III] ll|lll1§ [1:11 ll 69 Figure 1h. Determination of the Km's for Glycolate and D-Lactate Glycolate dehydrogenase activity measured by following DCIP reduction as described in Methods and Materials. 7O 0.: L- A/ / A / A l A/ V 0053/ l J 5 IO l/Glycolote (mM) 0.3 — /A A .L A/ #3 O.| — 1 l .50 .IO l/D-Loctote (mM) 71 Table 7. Electron Acceptor Specificity of Glycolate Dehydro- genase Assays carried out as described in Methods and Materials. Activity eXpressed relative to DCIP reduction. .__1 _ Electron Acceptor Relative Affinity DCIP 100 PMS 78 K3Fe(CN)6 NAD+ NADP+ Methylene Blue Oxygen Glutathione Cytochrome c N03 FMN OOOOOOOOOO FAD 72 Figure 15. Glyoxylate Formation in Relation to DCIP Reduction, the Requirement of DCIP for Glyoxylate Formation Glyoxylate formation measured by the glyoxy- late phenylhydrazone assay in presence and absence of DCIP as described in Methods and Materials: A- anaerobic glyoxylate formation + DCIP; V— anaerobic DCIP reduction; C>— aerobic glyoxylate formation + DCIP I - aerobic glyoxylate formation - DCIP. 73 m 4 OAV 0v / ADV r m m 3 8038 Eve .6 8.58 223020 3.9:: 20 I5 IO minutes 74 The Km for DCIP reduction with glycolate as the sub- strate was 1.7 x 10-5 M (Figure 16) determined by the method of Lineweaver and Burke (5“). For PMS the Km was 3 x 10'5 M (Figure 17) determined by a V versus S plot. It is note- worthy that the KIn for DCIP is approximately 20 times lower for glycolate dehydrogenase as compared to the Km of DCIP for glycolate oxidase whose DCIP Km is 3.8 x 10‘“ M. Flavin or Pyridine Nucleotide Cofactors Glycolate oxidase is a flav0protein (103) as are vari- ous other d-hydroxyacid dehydrogenases (3, 15, 24, 70, 80). Attempts were made to demonstrate a flavin co-factor for glycolate dehydrogenase from Chlamydomonas. Addition of 2 x 10'“ M FMN or FAD to the assay had no effect on the rate of DCIP reduction using partially purified enzyme. Treatment of the (NH4)2304 purified enzyme with acid (NH4)280u at pH “.5 or 5.0 following the method of Zelitch and Ochoa (103) resulted in the complete loss of activity. Neither FMN nor FAD ‘was able to restore activity. Addition of NAD+ or NADP+ did not stimulate the rate of DCIP reduction. Treat- ment of the enZyme with acid washed charcoal did not cause a loss of activity. These results do not demonstrate that either a flavin or a pyridine nucleotide is a cofactor for glycolate dehydro— genase. By precedent the involvement of a flavin co-factor is indicated, however, this remains to be established in the case of glycolate dehydrogenase from Chlamydomonas. 75 .mpa>apom mademmd oaomop zHHmodhpmaopozaonpooam pmsaaampop codpmapsoocoo mHom .mawanmpmz pad mUOSpmz ca popdaomop mm sodpoSpma mHom waazoaaom pmmdmmd mmmamwOHUmsmp mpmaoohao mHoo wee ea esp ae modemsaaaepmm .ea easmam 76 On. AEEV a_ooh 00. On. 0. ~ _ _ _.O Nd 77 .Aa muoa x m.pv He>ea mnapdaspdm d pd mm: opmaoomao .mdwwd on» on poops maomop game pmox dam Smonm dogmaman mm: mam .mamaampma paw muonpoz :« ponaaommp use opoapooam amwmxo Gd Spas acapospma zmwmxo waaSOHHom oomdmmm mmmammonpznop opmaooaau mam you mbaso soapQHSpmm mpmnpmnsm .ma madwam 78 4 4 q .— <1\ 8 O ' If) -exoidn uefiflxo 9mg aAgiDIGH Final Concentration PMS Mx IO"4 l fin-Ill. i ll 1' 79 pH Optimum and the Effect of Ionic Strength on Glycolate Dehydrogenase The pH curves for the oxidation of glycolate and D- lactate are shown in Figure 18. The pH cptimum for glyco- late extended over a broad range between 8.0 and 9.0 and was similar to that observed for partially purified glyco- late oxidase (19, 103). When determining these optima the change in extinction coefficient of DCIP with decreasing pH was taken into account (u). The Optimum for D-lactate oxidation was at pH 8.7. As glycolate oxidase purified from Spinach was very sensitive to ionic strength when DCIP reduction was assayed (3h), the effect of increasing ionic strength on glycolate dehydrogenase was tested. Enzyme was dialysed against 0.1 M phOSphate pH 7.5 to remove (NHu)ZSOu. The dialysed enzyme was then assayed with NaCl or (NHu)2804 added to the assay mixtures. No effect on the reduction of DCIP was observed with these salts even when the ionic strength of the assay mixture was doubled. No significant difference in the rate of DCIP reduc— tion by glycolate dehydrogenase was observed when the fol- lowing buffers were tested at pH 8.7, 0.1 M pyrophOSphate, bicine, tricine, glycylglycine and Tris. Effect of Inhibitors on Glycolate Dehydrogenase Glycolate dehydrogenase was sensitive to inhibitors 80 Figure 18. pH Curve for the Oxidation of Glycolate and D-Lactate by Glycolate Dehydrogenase Rates were determined by measuring DCIP reduc- tion. Buffer used was 0.1 M pyrophOSphate/phOSphate adjusted to desired pH. Final pH was determined after completion of the assay. Relative Activity 81 IOO — ’45-'“5-‘A‘ A’ ’ ‘ ~ ~ 1” I, “ ‘\ gt ycolate’, A ,’ \ ‘\‘ 80 ’- ’1’ I, \ ‘\ I ” \ A IA’ ’ Q‘ ’ ,0 s ,I' I ‘0 o I I I 60 'A I” D-Iactate I’D ” o" 40 - 20 b L i 7.0 8.0 9.0 82 which effect sulfhydryl groups such as Cusou, N-ethylmale- imide and, p-CMB (Table 8). Glycolate oxidase from higher plants has been shown to be relatively insensitive to p-CMB (34, 69), however, glycolate oxidase from rat liver (64) and renal cortex (80), have been found to be inhibited by these sulfhydryl inhibitors. The effect of metal complexing agents on glycolate dehydrogenase gave equivocal results. o-Phenan- throline was found to inhibit 70% of the activity at 4 mM (Table 8) when added directly to the DCIP assay mixture (pH 8.7). Incubation of the enzyme at 4°C with 10 mM 0- phenanthroline at pH 7.5 (0.1 M phOSphate) gave only 50% inhibition after 24 hours. o-Phenanthroline was not separated from the enzyme before assaying, making the o- phenanthroline concentration in the assay 2 x 10’“ M. Dialysis against 10 mM o-phenanthroline for two hours at pH 8.1 (0.1 M phoSphate) caused a complete loss of activ- ity, compared to controls lacking o-phenanthroline. After removal of the o-phenanthroline by continued dialysis, no reactivation occurred by incubation of the inhibited enzyme at 4°C up to two hours with 5 x 10'“ M ZnClg or 5 x 10'” M FeClB. Also addition of 5 x 10-5 M ZnClZ, FeC13 or MgClZ to the assay directly with inhibited dialysed enzyme did not lead to reactivation. In both types of reactivation eXperiments, 0.1 M phOSphate pH 8.1 was used as the assay buffer rather than 0.1 M perphosphate pH 8.7 in order to Table 8. Effect of Enzyme Inhibitors on Glycolate Dehydro- genase 83 Inhibition measured after addition of inhibitor directly to the DCIP reduction assay mixture. in the presence of inhibitor for 10 minutes before addition of glycolate. Enzyme was Inhibitor Concentration % Inhibition N-Ethylmaleimide 5 10-5 m 35% 1 10-4 M 60% 5 10"i M 100% Cuso4 1 io-Lt M 20% l 10"3 M 60% p-CMB 1 10-5 M 35% 2 10-5 M 73% 5 10-5 M 100% o-Phenanthroline 10-3 M 25% u 10-3 M 71% 8 10-3 m 90% m-Phenanthroline 8 10-3 M 0% EDTA 1.5 10-2 m 0% KCN 5 10'” M 47% 1 10-3 m 85% B-Hydroxyquinoline Sulfonate 1.5 10"2 M 30% 84 prevent binding of the cation by pyrophOSphate. Inhibition by o-phenanthroline could be avoided by addition of 10 mM mercaptoethanol or cysteine to the dialysing mixture. m-Phenanthroline had no inhibitory effect (Table 8), indi- cating o-phenanthroline inhibition was not due to non- Specific interactions. Work in Horecker's laboratory with rabbit muscle fructose-1,6-diphOSphate aldolase has shown similar inhibitory results with o-phenanthroline at pH's above 8.0 (50). This aldolase is not a metalloenzyme. It was inhibited, however, by o-phenanthroline and the o- phenanthroline inhibition was relieved by addition of sulf- hydryl reagents. On the basis of the aldolase work it is suggested that o-phenanthroline inhibition of glycolate dehydrogenase could be due to enhanced sulfhydryl oxidation rather than metal complexing. No inhibition of glycolate dehydrogenase was observed with EDTA after incubation in 10 mM EDTA for up to 24 hours at pH 7.5. Addition of EDTA directly to the assay mixture up to 16 mM had no effect on the rates. 8-Hydroxyquinoline sulfonate gave 30% inhibition at 15 mM when added directly to the assay (Table 8). The high concentrations necessary for inhibition indicated non-Specific interactions rather than metal complexing. Cyanide caused inhibition of activity (Table 8). Cyanide has been shown to inhibit the Zn—flavoprotein—a- hydroxy acid dehydrogenase of yeast (24). Cyanide did not 85 inhibit glycolate oxidase isolated from Spinach, however it inhibited glycolate oxidase from liver, an enzyme which has no metal co-factor (64). This type of inhibition by cyanide is felt to be due to sulfhydryl interactions rather than metal complexing (26). Thus the use of cyanide does not distinguish between metalloenzymes and sulfhydryl sensitive enzymes. On the basis of the data it is not possible to state definitely that glycolate dehydrogenase is or is not a metalloprotein. Distribution of Glycolate Dehydrogenase The work reported so far in this thesis has been done exclusively with glycolate dehydrogenase isolated from Chlamydomonas. To determine if glycolate dehydrogenase is present in other green algae and perhaps in lower forms of plants, tests were run on several other green algae and plants. Both glycolate oxidase and glycolate dehydrogenase were assayed from crude extracts by measuring DCIP reduction, D and L lactate Specificity and the effect of cyanide on DCIP reduction. These assays should distinguish between the two enzymes as glycolate oxidase oxidizes L-lactate but not D- lactate, and is not effected by cyanide, while glycolate dehydrogenase oxidizes D-lactate and is inhibited by cyanide. The results of the survey are presented in Table 9. .oMSpNHS mmmmm op mapooaap poops mpfizmmo .mamaaopmm was mpozuma SH ponaaomop mm soapospma mHom an muomapxo opsao ad dopammma mpabapo