111m“1hq‘ I11WIII11111 1111 I1 1 11.1. .I “1““ “1‘1 I1IIIII H1“I“".‘.1I‘‘1‘“1‘1‘ ‘1‘ I “‘1‘ “““‘1 111111 “ I“ “1111111111111 1 11 II 11-1111111'1‘1‘1‘1‘ ; 1 II I. IIIZIII1IIIII1I fit a 1 I N111.H H II I1 .1, IIIIIIIIIIIIII I . I IIIIIII III 1W .1 —._;:. fl” h 1 ‘11 .— .—-.-x:"£r . #- ”Mr— -a... m__.,_ gfifi -‘00—~vv-— kw‘)‘*‘_——. _- . _.—(o A Maw-I4": W ‘ W J. —.— . .w—a . -m a. A- w“...— . “H ”m. _— .— ~_-.-.—- ‘ ’4 ¢.'.-W'“'%-‘_— .._—— .4. *— _—.~. .- , w.- ‘ .—-.—— no .fi— .... .. .& ..._.. -"_ .4 ._ H... -- -. a” 4 . . w-..-.— *‘J _..— .4 -— fl- a”- o-.. M ’b—M _. h.‘ "2: =. ~E 1-..... :..—- M... ._._.~ up" ~. ...o .. .— w... ’7 a , _"‘L- “- - - ‘ .4." w""'<'.' W . _ .._.——d'.._..- ”—— “.— mm - “—— ..—. A’I.‘ five "x. v.” 1 .40 "M‘- ..—-—-§ “4.. n.- -5“ . _.—.. # W” W— . I I ' 2“; 3% L‘ I “ ‘ “ IIII : ., . I “:1" WI 1‘131 3;..- . “i? ' ' g . 111-1111111111; 11 - i.- 1‘ 1-. I I ‘12" IEI'In . , ' . ' I: . , 111111; .III I III 1 ' . I 1 11‘1“““1‘11‘1‘1‘“111‘111‘1 ‘ '1. {“111 ‘1‘11‘11 J.» 'I “ :11: 11'.“ . I1 . - '.‘1I‘1é‘III‘11‘I‘LI1111111,3153... “'I .‘"‘I11I‘1"“ ‘ 1 ' I 1‘ I I I It n.( 1 . . , . i1111“‘ I‘I p113k £11 I; .1 .19 1141111‘1‘ ‘ 1511111111111! 1 1.11. '13.. .1‘1311 1:1'Z: ‘ 11111111111111 ‘IIIIIIIIIIWEE‘ 111 “11‘11‘IIIIIII I...‘ 11111 1.5.1; IIIIIIII . II... ““1‘11J‘Il11‘1‘ 1,111.11. 1‘1“‘1I11"‘“““ “11* I112 $1511 ‘6. 11111111111“‘31'1‘1111‘1“‘1‘";13, 11‘1‘111:““““‘.‘:§‘ .1 11112151113111 x1111111 IIIII M II 1 I1 1111. 1‘Hl1111b. 111:1” 1“‘111‘,‘11:‘4.‘11%1j11‘ 15311111111? < . ' “.11 1‘11: 111 ““‘ I“‘;1‘1“ 11 I1I1I‘11I 111111 "1 ‘ ‘,1‘I1I1111“£-‘1‘:‘I‘1“1v'1111-1! ‘ ‘ ““1 2411““ . . 1III1 II HI 1 H%w1~IW Ifikfifiéfiflhhem H*m . 3%“ g? 1 1 113:1“ 11111111112111 ‘ ‘ ‘ 11‘ 1"!001 ‘ I11‘1 1131373111. HM1I1111‘L$‘2‘1:‘11I{11I‘A311111113355“ "1“ 11111111 '151'1 1f; ' 11‘1‘111I1“}§:I.:I;I1 I11 1 “N“ “"‘ ‘1;1111‘1“ 11“ 1M ““111‘ “““ “‘1‘“1“ “1‘ 1‘ 111W 1111‘11 “““““““,‘.1“ “ :1. : .131::I,;11'1111111111111111111111‘1‘1 11:1111111111111111911 [11111111191111 11.1111111111111111111“11}I1‘11111P1IUI11 11111111111 111111 1111:111‘31'1J: 11m x '7 " ”‘1 , ,1 ‘ -?I~,I.‘f““II’II-I‘“ “1 I, I1- 1.111 E " IIIII1‘3111‘1‘1I:."Ii11“..II~‘I1I1IIII3111111111111“ “ 1‘11“ 1““"““‘““ 1111 “1'1““ :1 I 1 111 IIII 1‘1 “1111115115111 [L1I1-r I‘ . 1 I’ . . I‘III I“"‘I‘.III:‘I‘ 1.2111 ‘ i I' ""1“‘1~1 III? I“ 1" ““ “ I11 ‘1‘“ 1" I111 I1 1; WWI ' ‘ “‘II“ Ig‘I'. 1II'1 ‘I‘I“ 1 , 1U” "I1111‘111 ‘1 “ 1 ‘ “III‘ W11I1 W1 In 'H‘1H ‘1“ 111111 ‘wn1II“11“ “‘W““““““““‘ ii» 111 I! IIIIII‘ .‘I,1IIII I“ I" IIIIII1“II II I 1“ II‘ 1111’ “2' ‘1 111‘ 1.11 11111111 111111 1 I l1111:1111 1111111111111 1111111111111 1111 1' .111 1111 1 |1 111 1.11 111111111} ‘ "' . I . III‘III. I ~.,.II 11 1 II" I“ ‘II‘ II I: “I ‘ 11 “I 111 I111'111“‘1"‘11‘1111 I1 11 1111‘“ 11.1111 1' ‘111111 1 1 111' 1 'a -III.I1..I:.. I. I '11 II III III 11.1.1 1III1IIIIIII‘II 1 11m IIIIIII11111“II1I'.| 1111 111111.11 |. LIBRARY l Michigan State University l ‘\ I, This is to certify that the dissertation entitled INORGANIC PYROPHOSPHATE:D-FRUCTOSE-6-PHOSPHATE l-PHOSPHOTRANSFERASE IN PLANTS AND ITS REGULATION BY A NATURALLY OCCURRING ACTIVATOR, D-FRUCTOSE 2,6-BISPHOSPHATE presented by Dario C. Sabularse has been accepted towards fulfillment of the requirements for Ph.D. Biochemistry degree in R.L.QMQLWN R.L. Anderson Major professor Date November 9, 1982 MS U L! an Affirmative Action/Equal Opportunity Institution 0-12771 RETURNING MATERIALS: Place in book drop to remove this checkout from your record. FINES will be charged if book is returned after the date stamped below. .___.________ MSU LIBRARIES INORGANIC PYROPHOSPHATE: D-FRUCTOSE-G-PHOSPHATE l-PHOSPHOTRANSFERASE IN PL ANTSA ITS REGULATION BY A NATURALLY OCCURRING ACTIVATOR, D-FRUCTOSE 2, 6-BISPHOSPHATE 3:! Dario C. Sabularse A THESIS Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY ' 1,:5rs“ t‘" ‘ ‘ J Deparhnent of Biothemistry ;'~ I?0 ,000, i7 l; f" v m I ' 4d 25 Pl"! 3M: "3‘ .,. .3" ,' Kn Yew-35 .’-:"‘ 7:1,. 1.37“»: nd It pH ..8 w 3 L '5 ~' 4"; G/QOJWQ ABSTRACT INORGANIC PYROPHOSPHATE:D-FRUCTOSE-G-PHOSPHATE I-PHOSPHO- TRANSFERASE IN PLANTS AND ITS REGULATION BY A NATURALLY OCCURRING ACTIVATOR, D-FRUCTOSE 2,6-BISPHOSPHATE By Dario C. Sabularse Inorganic pyrophosphate:D-fructose-G-phosphate l-phosphotransfer- ase was detected in extracts of mung bean sprouts, the first such de- tection in a C3 plant and in non-photosynthetic plant tissue. The enzyme was purified over lOOO-fold. It had an absolute requirement for fructose 6-phosphate and inorganic pyrophosphate as well as for a diva- lent metal (Mg++), and was found to be activated by hexose bisphos- phates (especially by D-fructose 2,6-bisphosphate, but also by D-fruc- tose l,6-bisphosphate and D-glucose l,6-bisphosphate), the first demon- stration of a regulatory mechanism for this enzyme from any source. It exhibited hyperbolic kinetics both in the absence and presence of a hexose bisphosphate activator. The enzyme was also demonstrated, for the first time, to exist in two forms (large, Mr = 340,000 and small, Mr = 170,000). KA values of the large and small forms, respective- ly, for fructose-2,6-P2, were 25 nM and 50 nM at pH 7.8, the pH opti- mum, and 25 nM and 140 nM at pH 7.0. In the presence of l “M fructose- 2,6-P2, Km values for fructose-6-P at pH 7.8 were 0.12 mM for both forms, and at pH 7.0 were 0.15 mM and 0.50 nM, respectively, for the large and small forms. The Km for PPi was about 0.l mM for both Dario C. Sabularse forms at both pH values. Fructose 2,6-bisphosphate (1 uM) increased the affinity of the enzyme for fructose-G-bisphosphate 167-fold and increased the Vmax 15-fold; these two effects combined to give over lOOO-fold activation at 0.12 mM fructose-G-phosphate. The enzyme was unaffected by millimolar amounts of phosphoenolpyruvate. In con- trast, ATP:D-fructose-6-phosphate l—phosphotransferase from the same source was not affected by fructose 2,6-bisphosphate but was inhibited >99% by 10 uM phosphoenolpyruvate. Other properties of the two enzymes are also described. Fructose 2,6-bisphosphate was demonstrated for the first time to be naturally occurring in mung bean and other higher plants. Inorganic pyrophosphate:D-fructose-6-phosphate l-phosphotransferase was also found to be widespread among plant species. The data support a pro- posal of this thesis, that the use of inorganic pyrophosphate by this enzyme is instrumental in carbohydrate metabolism in plants, and there- fore is of major metabolic significance. ACKNOWLEDGEMENTS I would like to express my thanks and appreciation to Dr. Richard L. Anderson, my graduate advisor, for his invaluable guidance through- out the conduct of the thesis research and the preparation of this manuscript. My thanks and appreciation to the members of my guidance committee (Dr. N.E. Tolbert, Dr. R.S. Bandurski, Dr. A. Revzin, Dr. K.R. Schubert and Dr. J.L. Fairley) for providing a general direction towards the completion of my studies and for the constructive criti- cisns on the thesis manuscript. I acknowledge the Philippine Council for Agriculture and Resources Research/U.S. Agency for International Development for the scholarship that financed the initial years of my studies; the Biochemistry Depart- ment of Michigan State University for the graduate assistantship pro- vided me; the University of the Philippines at Los Banos for allowing me to be on study leave, and my mother-in-law, Gliceria Castro, who provided financial support during the critical portion of my studies. My thanks to Dr. H.A. Hood, for allowing the use of his micro- processor-controlled spectrophotometer; to Nicholas Ringo for his technical assistance on the use of the instrument; and to Patty Kehoe for her collaborative work. My thanks to my mother, Consorcia Consuelo-Sabularse, and to my brothers and sister for their moral support; and to everyone who in one way or another contributed towards the realization of this thesis. Finally, my appreciation to my wife, Veronica, who stayed on during the critical moments of the conduct of my thesis and to my sons, Julius and Anthony, for their inspiring role as I went on with my studies. iii LIST OF TABLES . . . . . LIST OF FIGURES. . . . . LIST OF ABBREVIATIONS. . INTRODUCTION . . . . . . REVIEW OF LITERATURE . .' . . . . . . . . TABLE OF CONTENTS Enzymes That Use Inorganic Pyrophosphate as Donor I I I I I I I I I I I I I I I I I I I Distribution and Some Properties of PPizF6P ferase. Q I I I I I I I I O I I I I I O I I I I I I I I I I I th e Phosphoryl l-phosphotrans- Discovery, Chemical Synthesis and Identification of 2,6-Bisphosphate. . . . . . . . . . . Assay of Fructose 2,6-Bisphosphate. . Enzymes Regulated by fructose 2,6-Bisphosphate. ATP-dependent phosphofructokinase. . . . . . Fructose l,6-Bisphosphatase. . . . . . . . . Fructose 0 o o I 0 Enzyme for the Synthesis and Breakdown of Fructose 2,6-Bis- phosphate 0 O I I I I I I I I I I I I I O O I O O O I I I 0 Review Articles on Fructose 2,6-Bisphosphate. . . . . . . . 0 Starch Metabolism During Seed Germination . . . . . . . . . . Sucrose Metabolism During Germination of Cereals and Legumes. MATERIALS AND METHODS - I I I I I I I O I I I I I I I I I I I I O I Source of Materials . . . . . . . . . . . . . . . . Assay Methods for PP1:F6P l-Phosphotransferase. . . Method A: Method B: Method C: iv Forward reaction (standary assay) . . Forward reaction (alternative assay). Reverse reaction. . . . . . . . . . . Page viii \DmmNm Other Enzyme Assays . . . . . . . . ATP— —dependent phosphofructokinase. Inorganic pyrophosphatase. . . . . Fructose-l,6-bisphosphatase. . . . Phosphoglucose isomerase . . . . . Adenylate kinase . . . . . . . . . Assays for Molecular Weight Standards Pyruvate kinase. . . . . . . . . . Aldolase . . . . . . . . . . . . . Hexokinase . . . . . . . . . . . . Ferritin . . . . . . . . . . . . . Protein Determination . . . . . . . . . . . Pressure Dialysis Concentration of Protein. Conductivity Measurements . . . . . . . . . o O o I o 0 o o o o 0 Preparation of Fructose 2,6-Bisphosphate. . . . Preparation of Fructose-6-P Free of Pi by Chromatography ThroughsephadEXG'IOooooeuncoo-ooooooo Partial Purification and Elucidation of Some Properties of PP1:F6P l-phosphotransferase. . . . . . . . . . . . . . . . Further Purification and Elucidation of Additional Proper- ties of PP1:F6P l—phosphotransferase. . . . . . . . . . . . Final Purification and Partial Separation of the Two Forms of the PPA: .F6P l- -Phosphotransferase and r of their operties . . . . . . . . . Preparation of genninated seeds. . General. . . . . . . . . . . . . . Preparation of crude extract . . . Ammonium sulfate precipitate . . . DEAE-cellulose chromatography. . . Phosphocellulose chromatography I. Phosphocellulose chromatography II BIO-GElA-Iosm-o... Characterization Separation of two forms of the enzyme. Partial Purification and Determination of ATP-dependent Phosphofructokinase . . . . . . . . . . . . . . Some Properties Extraction of the Natural Activator (Fructose-2,6-P2) from Mung mans I I I O I I I I I I I I O I I O I I I I I I I of Survey of PP1:F6P l-Phosphotransferase and Fructose 2,6-Bis- phosphate in Other Plant Species. . . . . . . . . . . . . . . Page RESULTS I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I 40 Detection of the Enzyme in an Extract from Germinated Mung Beans I I I I I I I I I I I I I I I I I I I I I I I I I I I I 40 Partial Purification and Some Properties of PP1:F6P l-Phos- photransferase. . . . . . . . . . . . . . . 43 Requirement for specific substrates and. a .divalent metal ionI I I I I I I I I I I I 43 Activation of PP; :F6P l -phosphotransferase by a product of the reaction. . . . . . . . . 43 Direct demonstration of activation by fructose l, 6- bis- phosphate. . . . . . . . . . . . . . . . . 53 Activation by glucose 1, 6- bisphosphate . . . . . . . . . . 56 Optimization of the Assays for PP1:F6P l-Phosphotrans- ferase. . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 Forward reaction (standard assay). . . . . . . . . . . . . 59 Other assays for PP1:F6P l-Phosphotransferase. . . . . . . 64 Determination of Inorganic Pyrophosphatase. . . . . . . . . . 64 Some Properties of the ATP-dependent Phosphofructokinase in Mung Beans. I I I I I I I I I I I I I I I I I I I I I I I I I 65 Stability of PP1:F6P l-Phosphotransferase at Various Condi— tions of Storage. . . . . . . . . . . . . . . . . . . . . . . 69 Further Purification of the Enzyme and Elucidation of Addi- tional Properties . . . . . . . . . . . . . . . . . . . . . . 75 KA for fructose l ,6-bisphosphate . . . . . . . . . . . . . 78 KA for glucose l ,6-bisphosphate. . . . . . . . . 78 Activation by fructose 2, 6- bisphosphate and determination of its KA. . . . . . . . . . . . . . . . . . . . . . . 78 Km for fructose-6- P and relative Vmax of PP; :F6P l-phosphotransferase in the absence and in the presence of an activator . . . . . . . . . . . . . . . . . . . . . . . 83 Effect of Fructose 2,6-Bisphosphate on ATP-dependent PhoSpho- fructokinase. . . . . . . . . . . . . . . . . . . . . . . . . 90 Isolation and Identification of Fructose 2,6-Bisphosphate fran Mung Beans I I I I I I I I I I I I I I I I I I I I I I I 90 Comparison of the Three Methods of Protein Determination. . . 97 Final Purification and Partial Separation of Two forms of the PPizFGP l-Phosphotransferase and Characterization of Their Properties. . . . . . . . . . . . . . . . . . . . . . . . . . Final purification . . . . . . ._. . . . . . . . . . . . . 97 Separation of two forms of the enzyme. . . . . . . . . . . vi Some properties of the two forms . . . . . . . . . . . . . 111 a) Heat stability at 55°C . . . . . . . .7. . . . . . . . 111 b) Effect of pH and buffer composition. . . . . . . . . . lll c) KA for fructose 2, 6- -bisphosphate at pH 7. 8 and pH 7Io I I I I I I I I I I I 1"] d) Km for fructose 6- -phosphate at pH 7. 8 and pH 7.0 . . . l20 e) Km for PP1 at pH 7. 8 and pH 7. 0. . . . . . . . . . . . l20 f) Km for fructose l, 6- -bisphosphate . . . . . . . . . . . 120 g) fOY‘P'. o o o a o s o o o o o o 120 h; Effect of Pi on the activity of the forward reaction . l20 1 Molecular weight by gel filtration on Bio—Gel A-l.5m . l3l Apparent conversion of the large form to the small form upon incubation With DTT I I I I I I I I I I I I I I I I I I I I I 131 Survey of PP1:F6P l-Phosphotransferase and Fructose 2,62 Bisphosphate in Some Plant Species. . . . . . . . . . . . . . l38 DISCUSSION I I I I I I I I I I I I I I I I I I I I I I l I I I I I 142 LITERATURE CITED I I I I I I I I I I I I I I I I I I I I I I I I I 148 vii TABLE 10. LIST OF TABLES Some Properties of PP1:F6P l-phosphotransferase partially purified from mung bean sprouts, as determined by three different assay methods . . . . . . . . . . . . . . . . . . . Effect of various divalent metals on PPizFGP l—phospho- transferase I I I I I I I I I I I I I I I I I I I I I I I I I Effects of some compounds on PP1:F6P l-phosphotransferase . . Assay for inorganic pyrophosphatase . . . . . . . . . . . . . Effect of some metabolites on ATP-dependent phosphofructo- kinase. I I I I I I I I I I I I I I I I I I I I I I I I I I I Comparison of the mung bean natural activator and authentic fructose 2,6-bisphosphate with respect to acid- and base- Stability I I I I I I I I I I I I I I I I I I I I I I I I I I Comparison of three methods of protein determination. . . . . Purification of PPizF6P l-phosphotransferase from 500 grams of mung beans . . . . . . . . . . . . . . . . . . . . . . . . Occurence of PPizF6P l-phosphotransferase in some plant species I I I I I I I I I I I I I I I I I I I I I I I I I I I Occurence of fructose 2,6-bisphosphate in some plant species. viii Page 45 46 47 66 72 96 98 108 139 141 FIGURE 12. 13. LIST OF FIGURES Pathways for the degradation of the starch components . . . . Purification of chemically synthesized fructose-2,6-P2 by chromatography on a colunn of Bio-Rad AG l-X8 . . . . . . . . Apparent specific activities of PP1:F6P l-phosphotransferase and ATP-dependent phosphofructokinase at different times of germination prior to the discovery of activation by hexose bisphosphates . . . . . . . . . . . . . . . . . . . . . . . . Apparent time-dependent activation of PPi:F6P l-phospho— transferase in the absence of triose-P isomerase and apparent stimulation of activity by glyceraldehyde-3-P . . . . . . . . Proportionality of PP1:F6P l-phosphotransferase activity and enzMne Concent ration. I I I I I I I I I I I I I I I I I I Elution profile of fructose-6-P and Pi by chromatography throughsephadEXG-lo.................... Activation of PP1:F6P l-phosphotransferase by fructose-l, “'2. I I I I I o I I I o o o 0 o c I o a I I I I II I I I o Mg++ concentration-velocity curve for PP1:F6P l-phos- photransferase. . . . . . . . . . . . . . . . . . . . . . . . Absolute requirement of PP1:F6P l-phosphotransferase for diva] ent meta] , Mg I I I I I I I I I I I I I I I I I I I I I Effect of pH on the activity of ATP-dependent phosphofructo- kinase. I I I I I I I I I I I I I I I I I I I I I I I I I I I ATP concentration-velocity curve for mung bean ATP-dependent phosphofructokinase . . . . . . . . . . . . . . . . . . . . . Effect of micromolar concentrations of phosphoenolpyruvate on the activity of ATP-dependent phosphofructokinase . . . . . . Stability of PP1:F6P l-phosphotransferase at various condi- tions 0f storage. I I I I I I I I I' I I I I I I I I I I l I I ix Page l3 30 41 49 5l 54 57 60 62 67 70 73 76 FIGURE 14. 15. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. Determination of KA of PPizF6P l-phosphotransferase for fructose 1,6'bISphOSphate a o o u u o o o o o o c o o o o o 0 Determination of KA of PPizF6P l-phosphotransferase for gIUCOSE 1,6-bISphOSphateo o o o o o n a o o n o o a o o o o 0 Determination of KA of PPizFGP l-phosphotransferase for fructose”2,6'P2 o o o c o o I a o o o c o o o o o o o I I n 0 Effect of fructose-2,6-P2, fructose-l,6-P2 and glucose-l, 6-P2 on the kinetic constants (Km and Vmax) of PP1: F6P I-phOSphOtranSferaSE. o o o o a o I o o o o o o o o o o 0 Double reciprocal plots of the saturation curves for fructose- 6-P in the absence and presence of various activators (Data frm‘ Figure 17) I I I I I I I I I I I I I I I I I I I I I I I Substrate saturation curve for mung bean ATP—dependent phos— phosfructokinase. I I I I I I I I I I I I I I I I I I I I I I Chromatographic evidence for the occurence of fructose-2, 6-bISphOSphate. . o I o a n o o o o u a n o o o o o o o o o o DEAE-cellulose chromatography of PP1:F6P l-phOSphotrans- ferase. I I I I I I I I I I I I I I I I I I I I I I I I I I I Phosphocellulose I chromatography of PPi:F6P l-phosphotrans- ferase. I I I I I I I I I I I I I I I I I I I I I I I I I I I Phosphocellulose II chromatography of PPizF6P l-phosphotrans- ferasel I I I I I I I I I I I I 'I I I I I I I I I I I I I I I Bio-Gel A-l.5m chromatography of PP1:F6P l-phosphotrans feraseI I I I I I I I I I I I I I I I I I I I I I I I I I I I Separation of the two forms of PP1:F6P l-phosphotransferase by chromatography through Bio-Gel A-l.5m. . . . . . . . . . . Relative stability of the small and large forms of PP1:F6P l-phosphotransferase at 55°C. . . . . . . . . . . . . . . . . Effect of pH and buffer composition on the activity of the large form of PP1:F6P l-phosphotransferase. . . . . . . . . . Effect of pH and buffer composition on the activity of the snall form 0f PP1:F6P l-phosphotransferase. . . . . . . . . . Comparison of the large and small forms of PP1:F6P l-phos- photransferase with respect to the effect of pH on the KA for fructose‘2,6'P2 I o I o I o I o o o a c I a o I o o o o a Page 79 81 86 88 93 . 99 101 103 105 109 112 114 116 118 FIGURE 30. 31. 32. 33. 34. 35. 36. 37. Comparison of the large and small forms of PPizFGP l-phospho- transferase with respect to the effects of pH on the Km for fructose-6-PlIIIIIIIIIIIIIIIIIIIIIIII Comparison of the large and small forms of PPizFGP l-phospho- transferase with respect to the effect of pH on the Km for PP]. I I I I I I I I I I I I I I I I I I I I I I I I I I I I I Comparison of the large and small forms of PP1:F6P l-phospho- transferase with respect to the Km for fructose-l,6-P2 at pH 7.8. I I I I I I I I I I I I I I I I I I I I I I I I I I I Comparison of the large and small forms of PP12F6P l-phospho— transferase with respect to the Km for P1 at pH 7.8 . . . . . Effect of Pi on the activity of the two forms of PP1:F6P 1-phosphotransferase in the absence and presence of two con- centrations of fructose-2,6-P2. . . . . . . . . . . . . . . . Elution profile of molecular weight standards and the two forms of PP1:F6P l-phosphotransferase chromatographed on Bio'Ge1A105mooIooIIooIIIIIIoIIIIIIII Semi-log plot of the molecular weight of the standard pro- teins versus the peak fraction number from the elution pro- file on the Bio—Gel A 1.5m colunn (Figure 35) . . . . . . . . Apparent DTT-dependent change in KA for fructose-2,6-P2 for the large form of PPizFGP l-phosphotransferase, suggesting a concomitant change to the small form of the enzyme. . . . . . xi Page 121 123 125 127 129 132 134 136 LIST OF ABBREVIATIONS Abbreviations used without definition were taken from the list in the Journal of Biological Chemistry Instructions to Authors - l982. Additional abbreviations used are listed below. C3 plant a plant utilizing ribulose-Pz for initial fixation of C02. CAM crassulacean acid metabolism DEAE diethylaminoethyl DTT dithiothreitol EDTA ethylenediaminetetracetate HEPES 4-(2-hydroxyethyl)-l-piperazineethanesulfonic acid MES 2-(N-Morpholino)ethahe sulfonic acid P1 inorganic orthophosphate PPi inorganic pyrophosphate PIPES l,4-piperazinediethane sulfonic acid mU milliunit (milliunits) All sugars are of the D-configuration unless indicated otherwise. xii INTRODUCTION Inorganic pyrophosphate (PPi) is generated in many major bio— chemical processes. Among them are amino acid activation during pro- tein synthesis, nucleotide polymerization during nucleic acid synthe- sis, acyl-CoA formation via acyl-CoA synthetase during activation of fatty acids, and formation of sugar nucleotides during carbohydrate metabolism. It is a generally accepted concept that the PPi generated in many biochemical systems is hydrolyzed by inorganic pyrophosphatase, thereby providing a favorable thermodynamic condition towards the direction of PPi formation for the prior reaction. PPi hydrolysis in this manner dissipates the high energy of the pyrophosphate bond, however, whereas energy would be conserved if the PPi could be utilized by PPi-dependent phosphoryl-transfer enzymes, thereby sparing an equivalent ATP while at the same time serving to pull the PPi-generating reaction. There are a nunber of phosphoryl-transfer enzymes, primarily in microorganisms, that are known to use PPi as the phosphoryl donor. One of these enzymes is inorganic pyrophosphate:D-fructose-S-P l-phos- photransferase (PPizFGP l-phosphotransferase) which, at the start of this investigation, was known to occur only in microorganisms. Since the reactions involving PPi formation are ubiquitous in living systems, it was decided to investigate the possible occurrence of the enzyme in non-photosynthetic tissue of a C3 plant, in particular sprouted mung beans (Phaseolus aureus). The choice of a germinating starchy seed was prompted by the knowledge that PPi is formed in large amounts during the conversion of starch to sucrose, that the intracellular localization of inorganic pyrophosphatase is primarily in the chloroplast, and that the tissue would be devoid of photosynthetic processes, which could complicate analysis. In addition, it may be speculated that during the early stages of germination, when the plant is yet at its photosynthetic-independent stage, ATP-sparing reactions would be advantageous. This thesis will demonstrate (i) the existence and widespread occurrence of PP1:F6P l-phosphotransferase in plants, (ii) the first discovery of the activation of this enzyme by hexose bisphosphates, in particular a very potent activation by fructose-2,6-P2, (iii) the first demonstration of the natural occurrence of fructose-2,6-P2 in higher plants, (iv) the purification and properties of the enzyme from mung bean sprouts, (v) the first detection of two catalytically active forms of enzyme, and (vi) other findings that are relevant to a pro- posal that PPi utilization via PPizFGP l-phosphotransferase is important in energy metabolism in plants. REVIEW OF LITERATURE Enzymes That Use Inorganic Pyrophosphate as the Phosghoryl Donor. There are several phosphotransferases that use PPi as the phos- phoryl donor in at least some organisms (l, 2). They are: (a) acyl-- CoA synthetase, EC 6.2.1.1 to EC 6.2.1.3 (ATP + acylate + CoA 2 AMP + PPi + Acyl-CoA), for which in some organisms the net flux could be from right to left (1); (b) phosphoenolpyruvate carboxytransphos- phorylase, EC 4.1.1.38 (PPi + oxaloacetate I phosphoenolpyruvate + Pi + C02), discovered by Siu and Hood (3); (c) glucose-G-phos- phatase, EC 3.1.3.9 (glucose + PPi I glucose-6-P + P1), which catalyzes the indicated reaction under certain conditions (4, 5); (d) pyruvate, orthophosphate dikinase, EC 2.7.9.1 (ATP + P1 + pyruvate Z phosphoenolpyruvate + PPi), which occurs in plants and certain microorganisms (6-9); (e) pyrophosphatezserine phosphotransferase, EC 2.7.1.80 (PPi + L-serine + L-serine-P + Pi), found in propionic acid bacteria (10); (f) PP1:F6P 1-phosphotransferase EC 2.7.1.90, (PP1 + fructose-G-P 2 fructose-1,6-P2 + Pi), which will be elaborated on in the following section; and (g) PPi-dependent acetate kinase, EC 2.7.1.2, (acetyl-P + P1 : acetate + PPi), present in an anaerobic amoeba (ll). PPi has been reported to be a product of photophosphorylation in Rhodospirillum rubrum chromatophores (12). In this photosynthetic bacteriun, the PPi is an energy donor for several energy-linked reactions, including cytochrome reduction (13), transhydrogenation (14), NAD+ reduction (15), and ATP synthesis (16). Distribution and Some Properties of PPj:F6P 1-Phosphotransferase. Subsequent to the discovery of the enzyme in Entamoeba histolytica (a parasitic, anaerobic amoeba) in 1974 by Reeves and co—workers (17), the enzyme was reported to be present in four bacterial species: Propionibacterium shermanii (18), Bacteroides fragilis (l9), Pseudomonas marina (20) and a marine Alcaligenes species (20); and, during the course of this thesis research, in leaves of pineapple, a CAM plant (21). As will be demonstrated in this investigation, the enzyme is now known to be widely distributed in plant species, and in a recent abstract, it was reported to be present in rat liver (22). The enzyme from all reported sources is specific for PPi and fructose-G-P. No other mono- or diphosphorylated sugar can substitute for fructose-6-P and no nucleoside triphosphate or polyphosphate can replace PPi (17, 18, 21). The substrate affinities of the enzyme from E. histolytica and g. shermanii, respectively, are: Km for fructose-G-P, 38 uM and 100 uM; Km for PPi, 14 uM and 69 uM; Km for fructose-1,6-P2, 18 pH and 51 uM; and Km for Pi, 800 and 600 pH. The enzyme isolated from E. shermanii is a dimer of 95,000 molecular weight (18), whereas the E. histolytica enzyme has a molecular weight of 83,000 (23). No regulatory properties have been previously reported for the enzyme from any source. DiscoveryI Chemical Synthesis and Identification of Fructppe 2,6-Bis- phosphate. The discovery, chemical synthesis, and identification of fructose-- 2,6-P2 are credited to the independent but concurrent efforts of three separate laboratory groups: Hers' group - University de Louvain, Brussels (24-28), Pilkis' group - Vanderbilt University, Nashville, Tennessee (29-31), and Uyeda's group - University of Texas, Dallas, Texas (32-36). Van Schaftingen, Hue, and Hers (24, 25) reported the activation of ATP-dependent phosphofructokinase by a low-molecular-weight effector which could be isolated from rat liver extract by ultrafiltration, gel filtration or heat treatment but was rapidly destroyed in trichloro- acetic acid even in the cold. The low-molecular-weight stimulator was a nonreducing derivative of fructose-6-P and was completely destroyed upon incubation with 0.01 M HCl for 10 min at 20°C, with the consequent formation of equimolar amounts of fructose-G-P and Pi° These findings led to the tentative identification of the stimulator as fruc- tose-2,6-P2. Van Schaftingen and Hers (26) first described a chemi- cal synthesis of the low-molecular-weight stimulator by mixing fruc- tose-6-P with phosphoric acid. However, the method has very low yield and other sugar phosphates are generated. Nevertheless the chemical synthesis experiment proved that the only components of the stimulator were phosphate and fructose-6-P. Employing the method by Pontis and Fischer (37) for the conversion of fructose-l-P to fructose-Z-P, a procedure for the chemical synthesis of fructose-2,6-P2 was worked out by Van Schaftingen and Hers (27). The synthesis involved intramolecular cyclization of fructose-1,6-P2 to fructose 1,2 cyclic, 6-bisphosphate, followed by alkaline hydrolysis, digestion of the fruc- tose-1,6-P2 by fructose-l,6-bisphosphatase, and purification of the fructose-2,6-P2 by column chromatography on Dowex A01. The chemi- cally synthesized fructose-2,6-P2 had properties identical to that of the naturally occurring activator from rat hepatocyte extract. Upon exanination by 31F and 13C nuclear magnetic resonance spectro- scopy, the configuration and structure were deduced to be B-D-fructose 2,6-bisph05phate (28). From Pilkis' group, evidence for a low-molecular—weight activator from rat hepatocyte extracts for the ATP-dependent phosphofructokinase that was influenced by glucagon was reported (29, 30). The properties of the low-molecular-weight activator were similar to those that had been just reported by Van Schaftingen and Hers for fructose-2,6-P2 (30). Their subsequent paper (31) reported on the chemical synthesis of fructose-2,6-P2 from fructose-1,6-P2 by (i) cyclization utilizing the dicyclohexylcarbodiimide method of Pontis and Fischer (37), to form fructose 1,2 cyclic,6-bisphosphate, (ii) ring opening by alkaline treatment, (iii) heat treatment at alkaline pH to destroy the fructose-1,6-P2 but not the fructose-2,6-P2, and (iv) purification by chromatography on a DEAE-Sephadex colunn. The structure of the chemically synthesized sugar bisphosphate, which had properties identical to that of the activator from hepatocyte extract, was definitively identified as B-D-fructose-2,6-P2 by mass spectrometry and 13C NMR spectroscopy. Uyeda's group reported the isolation of an "activation factor“ for phosphofructokinase in liver extract in articles by Furuya and Uyeda (32, 33). They noted that the "activation factor" overcomes the ATP inhibition but had no effect on the catalytic activity under optimum assay conditions for ATP-dependent phosphofructokinase. Furthermore, they observed that AMP acted synergistically with the "activation fac- tor“ in reversing ATP inhibition. Richards and Uyeda (34) observed that the "activation factor“ for ATP-dependent phosphofructokinase in isolated hepatocytes changed its concentration in response to glucose and glucagon. Glucose increased the concentration of the “activation factor" but glucagon decreased it. In subsequent publications, Uyeda et a1. (35, 36) have shown by chemical analysis, synthesis, and 13C NMR spectroscopy that the "activation factor" for the ATP-dependent phosphofructokinase is B-D-fructose-2,6-P2. The synthetic compound was prepared from fructose-1,2 cyclic,6-P2 by alkaline hydrolysis and separation by paper chromatography. The synthetic and natural fruc- tose-2,6-P2 showed identical effects on the allosteric kinetic prop- erties of both rat liver and rabbit muscle phosphofructokinase. It should be recognized that the three laboratory groups reported practically the same findings and succeeded in an almost simultaneous publication of their work. Assay of Fructose 2,6-Bisphosphat . Pure preparations of chemically synthesized fructose-2,6-P2 can be determined by measuring the amount of fructose-6-P and/or P1 re- vealed after mild acid hydrolysis (26, 31, 35). In extracts and par- tially purified samples, fructose-2,6-P2 is detected by its ability to activate the ATP-dependent phOSphofructokinase at sub-optimal sub- strate concentration (26, 31, 34). For a quantitative determination in tissue extracts, a method has been described by Hue et al. (38), based {-4 on the measurement of the acid-revealed fructose-6-P in a coupled en- zymatic assay with bacterial NADH-linked luciferase. Another method is by extraction in hot neutral or alkaline buffers followed by determina- tion of the fructose-2,6-P2 by comparing the amount of activation of ATP-dependent phosphofructokinase with sample of standard fructose-2,- 6-P2. The activation of homogeneous rat hepatic ATP-dependent phos- phofructokinase is found to be roughly linear over the range of 3-10 nM when the enzyme is assayed with 0.2 mM fructose-6-P and 1 mM ATP (39). Enzymes Regulated by Fructose 2,6-Bisphosphate. ATP-dependent phosphofructokinase. Fructose-2,6-P2 is a potent activator of rat liver ATP-dependent phosphofructokinase with KA = 50 nM. The activation is in the form of increasing the affinity for fructose-6-P but has no effect on the Vmax of the enzyme (31, 35, 40). The hexose bisphosphate has also been shown to potentiate the activation of ATP-dependent phosphofructokinase by AMP (35, 40, 41), to act synergistically with AMP to release ATP inhibition (35), to release citrate inhibition (42), and to increase the binding affinity of the enzyme to AMP in yeast (43). ATP-dependent phosphofructokinases from other sources are also ac- tivated by fructose-2,6-P2: rabbit muscle (31, 35), yeast (44, 45), Ehrlich ascites tumor (46) and pancreatic islets of albino rats (47, 48). In higher plants, only the plastid isozyme of ATP-dependent phosphofructokinase from developing endosperm of castor bean has been reported to be activated by fructose-2,6-P2 when assayed at pH 7.0 with no activation at pH 8.0; the cytosolic isozyme is unaffected at either pH 7.0 or 8.0 (49). Fructose 1,6-bisphosphatase. This enzyme, which functions at a control point in the gluconeogenic pathway, is also regulated by fruc- tose-2,6-P2 in rat liver (31, 50-52) and yeast (45). Fructose-2,6-P2 at low concentration inhibited fructose-l,6-bisphosphatase competitive- ly with the substrate, and also potentiated AMP inhibition of the en- zyme (50); at higher concentration it transformed the shape of the sub- strate concentration-activity curve from hyperbolic to sigmoidal (51). Enzyme for the Synthesis and Breakdown of Fructose 2,6-Bisphosphate. The enzyme that catalyzes the synthesis of fructose-2,6-P2 in rat liver was first reported by Furuya and Uyeda (53), and subsequently by El-Maghrabi et al. (54), Hue et a1. (55), and Van Schaftingen and Hers (56). The enzyme catalyzed the transfer of y-phosphoryl group of ATP to the hydroxyl present in carbon 2 of fructose-6-P and has been given the trivial names of fructose-6-P,2 kinase (53), 6-phosphofructo 2-kinase (54) and phosphofructokinase 2 (56). In this literature re— view it will be referred to as fructose-6-P,2-kinase. Fructose-6-P,2-- kinase is found in liver, brain, heart muscle, kidney, testes and skel- etal muscle of rats (57), and in yeast cells (58). The fructose-G-P, 2-kinase has also been shown to be regulated by cyclic AMP-dependent, phosphorylation with the resulting activation of the enzyme (59-62). The enzyme (fructose-2,6-bisphosphatase) responsible for the hy- drolysis of the phosphate from the C-2 position of fructose-2,6-P2 was first reported by Furuya et al. (63). Richards and co-workers (64) observed that glucagon (10'11 M), epinephrine (10'5 M), or calciun (2.4 mM) and ionophore A23187 (10-5 M) administration to hepatocytes produced simultaneous activation of fructose-2, lO 6-bisphosphatase and inactivation of fructose-6-P,2-kinase within 2 minutes. They suggested that the level of fructose-2,6-P2 is con- trolled by reciprocal changes in fructose-2,6-bisphosphatase and fruc- tose-6-P,2-kinase activities. El-Maghrabi et a1. (65) had purified fructose-2,6-bisphosphatase to homogeneity and demonstrated that fructose-6-P,2-kinase activity co- purified with the bisphosphatase activity. They reported that the cat- alytic subunit of the CAMP-dependent protein kinase phosphorylated the enzyme (1 mole of phosphate/mole subunit of the dimeric enzyme) result- ing to a concomitant activation of fructose 2,6-bisphosphatase and an inhibition of the fructose-6-P,2-kinase activity. This phosphoryla- tion-dephosphorylation involved a seryl residue per subunit of the dimeric bifunctional fructose-G-P,2-kinase/fructose—2,6-bisphosphatase (66). Review Articlg§_ppgfru§tp§e 2,6-Bisphosphat . There are two recent reviews on fructose-2,6-P2. The first is by Pilkis et al., entitled "Fructose 2,6-bisphosphate: a mediator of honnone action at the fructose 6-phosphate/fructose 1,6-bisphosphate substrate" (66a) and the second is by Hers and Van Schaftingen, entitled “Fructose 2,6-bisphosphate 2 years after its discovery“ (66b). Starch Metabolism During Seed Germination. Starch metabolism has been reviewed extensively by many authors (67-72). However, this literature survey will focus only on starch metabolism during seed germination, particularly for cereals and legumes. 11 During seed maturation of cereals, starch accumulates as a major carbon-source reserve for eventual carbon-energy needs during germination. In cereal seeds, vigorous synthesis of reserve starch occurs from about six days after flowering to ripening (73). This reserve starch is, however, enzymatically degraded to low-molecular-weight carbohydrates which are further metabolized during the course of germination. In legunes such as soybean, starch is a transient reserve material in both developing and germinating cotyledon (74). During germination of several legune species, amylase activity increases and starch is depleted (75-77) but the seed apparently produces starch during imbibition and germination (78,79). In nonphotosynthetic organs, the synthesis of reserve starch oc- curs in amyloplasts, which are a specialized form of plastids that function for starch synthesis (80). Starch occurs in nature as a water-insoluble granule. Its size and shape is often characteristic of the plant species and the plant maturity (81, 82). Starch occurs in two forms. Amylase, the linear type of starch, consists of D-glucose residues in a(l-4) linkage. Amylopectin, the branched form, consists of chains containing, on the average, about 20-25 a(l-4)-linked D-glucose residues which are interlinked to a branched structure. The molecules thus contain 4-5% of a(l-6)-D-glucosidic linkages (72). According to Preiss and Levi (68), since starch of plants primari- ly occurs as water-insoluble granules in distinct plastids, starch degradation must occur in three phases. These are: (i) reduction of the granule to soluble maltodextrins, (ii) debranching and degrading the larger maltodextrins to glucose and glucose-l-P, and (iii) further metabolism of glucose or glucose-l-P and movement (translocation) of 12 the products from the site of starch storage. The cleavage of a(I-4) bonds of starch may be accomplished by amylases and maltase or by starch phosphorylase, and the a(I-6) bonds by hydrolytic debranching enzymes (71). The pathway and the enzymes involved in the degradation of starch to glucose or glucose-l-P are shown in Figure l. The reaction mediated by the various hydrolytic enzymes during starch degradation are well known. A brief description, taken fran Preiss and Levi (68) and Manners (72) of the action of these various hydrolytic enzymes is given below. 1. a-Amylases. These enzymes catalyze an essentially random hy- drolysis of non-terminal a-(l-4)-g1ucosidic linkages in both linear and branched substrates. The nonnal end products are maltose and glucose from amylose and these sugars, together with branched oligosaccharide a—dextrins, from amylopectin. At low enzyme concentration, maltotriose may also be present (72). 2. p-Amylases. This group of enzymes which occur only in cereals and in certain other higher plants such as sweet potatoes and soybeans catalyses a stepwise hydrolysis of alternate linkages starting at the nonreducing end in starch-type polysaccharides with the liberation of maltose. Enzyme action on linear substrates is usually complete; with branched substrates, the enzyme is unable either to hydrolyse or bypass the a—(l-6)-D-glucosidic inter-chain linkages so that the enzyme action is incomplete, the products being maltose and a e limit-dextrin (68, 72). 3. a-Glucosidases. a-Glucosidase hydrolyzes the a(1-4) linkages of dextrins, attacking from the nonreducing end and liberating glucose. AMY LOSE ———__1. s-Amylase a-Amylase MALTOSE MALTOSE + MALTOTRIOSE Phosphorylase a-Glucosidase ' V GLUCOSE GLUCOSE-l-PHOSPHATE AMYLOPECTIN—T» B-Amylase + DBE a—Amylase MALTOSE MALTOSE, MALTOTRIOSE Phosphorylase + a-DEXTRINS + DBE : a—Glucosidase a-Glucosidase + DBE i) LUCOSE GLUCOSE-l-PHOSPiATE Figure l. Pathways for the degredation of starch components (DBE = Debranching enzyme). From: Manners, 0.0. (1974) In Plant Carbohydrate Biochemistry (Pridham, J.B., ed.) pp. 109-125. Academic Press, London and N.Y. 14 The a-glucosidases are most active in the hydrolysis of maltose, but they also hydrolyze maltotriose and maltotetrose (68, 72). 4. Starch phosphorylases. These enzymes catalyze a reversible reaction between a-D-glucose l-phosphate and starch. In leaf tissues, which have transient starch content, phosphorylase could efficiently convert starch to the hexose phosphate level and hence to sucrose, for subsequent translocation. In storage tissues, the role of phosphory- lase is not so clear-cut, since varying amounts of amylolytic enzymes are present. Several workers in this area have been reluctant to ascribe a purely degradative role to phosphorylase and there have been suggestions that it is involved in the synthesis of primers for starch synthetase, since the enzyme cannot use glucose or maltose as acceptor substrate. Several isozymes of phosphorylase are found in several plant species (72). 5. Debranching enzymes. These are active on B-limit dextrins, amylopectin a-limit dextrin, and pullulan, a fungal polysaccharide con- taining a—(1-6)-linked maltotriose residues (68, 72). Dunn (83) described a model for starch breakdown in higher plants, which suggests that a-amylase is the only degradative enzyme that has the ability to act on the starch granule ifl.!i!2- Subsequent degrada- tion is brought about by the action of s-amylase, phosphorylase, malt- ase and debranching enzymes. It has been pointed out that the initial rate-limiting step in the process of starch degradation is the action of apamylase on the starch-granule surface. Starch degradation during germination is paralleled by increase in a-amylase activity. The increase in a-amylase in germinating wheat and barley seeds has been attributed to the 3; novo synthesis of the enzyne 15 of the aleurone cells in response to gibberellic acid (84-86). Ho and Varner (87) concluded that there is no accumulation of an inactive a-amylase precursor in barley aleurone cells. Gibbons (88) observed that during germination of barley seeds the a-amylase protein moiety moves away from the entire face of the scutellum and intermediate crushed cell layers. As growth of the seedlings proceeds, the enzyme begins to be synthesized in the aleurone layer and transported away from the layer to the endosperm. Studies by Okamoto et al. (89) and Okamoto and Akazawa (90) on cereal seeds (barley, wheat, rye, oat, maize and rice) in the early stages of germination have shown that a-amylase is synthesized in the epithelial cells of the scutellun. The observation supports the report of Goswami et a1. (85) that germinating wheat seeds with intact embryo had greater a-amylase activity than those with embryo excised. A recent report by MacGregor and Matsuo (91) supports the observa- tions of Okamoto et a1. (89) and Goswami et a1. (85). They observed by means of scanning electron microscopy that starch degradation, in ker- nels of wheat and barley during the initial stages of germination, started at the endospenn-anbryo junction and moved along the junction to the dorsal edge of the kernel. From these results, MacGregor and Matsuo (91) suggested that the site of initial a-amylase synthesis in germinating cereal grains is in the embryo and not the aleurone layer. In contrast to gglppyp synthesis of a-amylase, s-amylase is pre- sent in some seeds in the latent form and is transformed to its active fonn on germination (72, 84, 85). Palmiano and Juliano (92) reported that in germinating rice, increases in a-amylase activity are blocked by protein synthesis inhibitors, but a-amylase activation is not. 16 However, as ppyp synthesis of e-amylase in rice scutellum during germination had been reported (93). Okamoto and Akazawa (93) reported that their results indicated strongly that at the onset of germination of rice seeds B-amylase is synthesized pg ppyp in the scutellum and that in the later stages there occurs activation of an inactive, latent form of the enzyme associated with the starch granules in the endosperm which become dominant in later stages of germination. Phosphorylase, debranching enzyme and a—glucosidase had been found in many mature and in germinating cereal seeds. Phosphorylase has been reported in sweet corn (94), rice (92, 95) and wheat (96). Debranching enzyme has been reported in malted barley (97, 98), sweet corn (99), waxy maize (100) and rice (92). a-Glucosidase has been reported in malted barley (101), rice (102), sorghum (103) and corn (103-105). Harris (106) studied, by means of electron microscopy, the cotyle- don cells of germinating mung bean seeds. He reported that starch grains had shown erosions from within leading to the formation of a hollow shell, and the erosion was accompanied by intrusion of cytoplasm into the shell. He also observed rougher inner surface of the shell suggesting that it was the site of starch hydrolysis; the hydrolyzing enzymes were presumed to be in the cytoplasm for no evidence was found of vesiculation associated with the inner face of the starch shell. The above sequence was for large starch grains which are present in the mature seed prior to germination. These starch grains of germinating mung bean were not surrounded by membranes, nor were there remains of plastid membranes, as in starch grains of germinating gigpm sativum (107). However, during germination cotyledon cells of legumes generally develOp plastids which synthesize small compound starch- 17 grains similar to those reported in Pisum arvense (108). The pattern of breakdown of these membrane-bound starch grains has not been clearly described. I In germinating peas, Juliano and Varner (75), have concluded that a-amylase is the major enzyme involved in the initial degradation of starch into more-soluble forms while phosphorylase and B-amylase assist in the further conversion to free sugars. a-Amylase is synthesized gg ‘ppyp during germination (75, 96, 109, 110). e-Amylase and phosphorylase appear within hours of imbibition (96, 107). Debranching enzyme was found in particles in the ungerminated pea seed (111-113). The enzyme is activated during germination. In germinating lentils, studies by Tarrago and Nicolas (77) re- ported increase of a-amylase and phosphorylase very similar to that seen in peas, but B-amylase activity was very low and no mention was made of the debranching enzyme. Much of the knowledge about starch degradation during seed germi- nation comes from studies with cereals and limited studies with legunes. It appears that there are species variations on the mechanism of the breakdown of starch with respect to the hydrolytic enzymes that predominate. However, it appears that the enzymatic degradation of starch during germination is initiated by a-amylase action. The products are then degraded by a-and B-amylase, phosphorylase, debranching and arglucosidase activities. The increase in activities of the enzymes is presumably due to pg ppyp synthesis at the onset of gennination. 18 Sucrose Metabolism During Germination of Cereals and Legumes. Sucrose is the main form of carbohydrate for translocation in nearly all higher plants (114). The nonreducing property of sucrose is regarded to be the important characteristic of a transport species (115). Arnold (116) proposed the hypothesis that sucrose acts as a protective derivative of glucose. If glucose were the translocate spe- cies, it would readily be attacked by enzymes which catalyze its metab- olism (116). Several studies have shown that during seed germination, the site of sucrose synthesis is the scutellum. In germinating wheat seeds, glucose was absorbed from the endosperm by the scutellun and synthe- sized to sucrose in that tissue (117). Similar results were observed in germinating rice (118) and in Ayepalfgppa seeds (119). By the action of phosphorylase on starch, glucose-l-P is produced. In the conversion of glucose-l-P to sucrose, the first enzyme involved is UDP-glucose pyrophosphorylase which catalyzes the reaction (120) UTP + D-glucose-l-P Z UDP-glucose + PPi. UDP-glucose pyrophosphorylase was first purified from mung bean seedlings by Ginsburg (121). It was observed that this enzyme was specific for UDP-glucose. Delmer-and Albersheim (122) found that UDP-glucose pyrophosphorylase activity is high in extracts of nonphotosynthetic tissue of mung beans. From UDP-glucose, there are two alternative routes for sucrose formation. One route involves the following reactions: UDP-glucose + fructose-6-P I UDP + sucrose-6'-P by the action of sucrose phosphate synthase; and sucrose-6'-P + sucrose + P1 catalyzed by sucrose phosphate phosphatase. The fructose-6-P is formed from glucose-l-P by the action of phosphoglucomutase and phOSphoglucose isomerase. The 19 second route involves the reaction, UDP-glucose + fructose I UDP + sucrose, which is catalyzed by sucrose synthase. Leloir and his collaborators (123, 124) discovered the existence of these enzmnes capable of synthesizing sucrose and sucrose phosphate. The generally accepted major route in the synthesis of sucrose in plants is the sucrose phosphate synthase pathway (67, 120, 125). Results of studies by Hawker (126) on germinating broad beans, maize seeds, and castor beans have all suggested that sucrose phosphate synthase and sucrose phosphatase catalyze the synthesis of sucrose via sucrose phosphate. Preiss and Greenberg (127) reported that sucrose phosphate synthase from wheat germ exhibited sigmoidal saturation curves for fructose-6-P and UDP-glucose. It has been observed that Mg++ enhances the activity of wheat germ sucrose phosphate synthase in the direction of sucrose synthesis whereas sucrose inhibited its activity (128, 129). The apparent affinity of the enzmne for Mg++ was also decreased by sucrose thereby reducing the activity induced by Mg++ (129). Sucrose degradation is achieved by the reversal of the sucrose synthase reaction, sucrose + UDP (ADP) z UDP-glucose (ADP-glucose) + fructose, which is the principal mechanisn of sucrose cleavage in plant cells (125, 130). Studies on sucrose synthase from mung beans showed that the major function of the enzyme is the catalysis of the synthesis of nucleoside diphosphate glucose from translocated sucrose in nonphotosynthetic tissue (131, 132). Sucrose breakdown is also achieved by the action of invertase. According to Akazawa and Okamoto (133), this breakdown of sucrose is also common in higher plants. The resulting free glucose and fructose .‘1' a a" It should be noted that the synthesis of every mole of sucrose W glucose-l-P would form 1 mole of PPi. .5.» L .‘J‘ V . .-L ‘4‘. A. _______4 J v g. MATERIALS AND METHODS Source of Materials Commercially available mung bean seeds and other plant materials used were obtained locally. Duckweed was harvested from a laboratory culture. All biochemical reagents used were obtained from Sigma Chemical Company, unless noted otherwise. Other chemicals commercially avail- able were of analytical grade. Special notice is given to the commercial sources of the following materials. Membrane filters (PM30) used in the concentration of protein samples by pressure dialysis were obtained from the Amicon Corporation. Phenol reagent (Folin and Ciocalteu) used in Lowry protein determination was obtained from the Harleco Corporation. DEAE-cellulose and phosphocellulose were obtained from Sigma Chemical Company. Bio-Gel A-l.5m (Agarose beads for gel filtration); AG 1-8X, 200-400 mesh, Cl' form (anion exchange resin); Bio-Rad protein assay dye reagent; and Bio-Rad protein assay standard II (bovine plasma albumin) were obtained from Bio-Rad Laboratories. Sephadex (G-10 and G-100 medium grade) were obtained from the Pharmacia Fine Chemicals. "Enzyme-grade“ ammonium sulfate was obtained from the Mallinckrodt Company. Ferritin was obtained from the Boehringer Mannheim Biochemicals. 21 22 Assay Methods for PszF6P 1-Phosphotransferase. Method A: Forward reaction (standard assay) Principl . The continuous spectrophotometric assay was based on coupling fructose-1,6-P2 formation to non-rate-limiting amounts of aldolase, triose-P isomerase, and a-glycerol-P dehydrogenase. With the three coupling enzymes in excess, the rate of fructose-1,6-P2 forma- tion was equivalent to one-half the rate of NADH oxidation, which was measured by the absorbance decrease at 340 nm. Reagents. HEPES-NaOH buffer (pH 7.8), 170 mM, containing 2.1 nM EDTA Fructose-6-P, 340 mM Sodiun pyrophosphate (Na PPi), pH 7.8, 17 mM MgC12, 102 mM Fructose-1,6-P2 aldolase, 40 mU/ul, previously dialyzed against 1 mM EDTA, pH 7.8 Triose-P isomerase, 200 mU/ul, previously dialyzed against 1 mM EDTA, pH 7.8 a-Glycerol-P dehydrogenase, 40 mU/ul, previously dialyzed against 1 mM EDTA, pH 7.8 NADH, 4.1 mM Activator (17 "M fructose-2,6-P2) Procedure. The following were added to a microcuvette with a 1.0-cm light path: 80 ul of HEPES buffer, 5 ul of fructose-6-P, 10 HI of Na PPi, 10 ul of MgClg, 5 ul of fructose-1,6-P2 aldolase, 5 ul of triose-P isomerase, 5 ul of a-glycerol-P dehydrogenase, 10 ul of NADH, 10 ul of fructose-2,6-P2, a rate-limiting amount of PPizFGP 1-phosphotransferase, and water to a volume of 170 pl. The reaction 23 was initiated by the addition of PPi:F6P l-phosphotransferase. A control cuvette minus PP; measured NADH oxidase and apparent fructose-G-P reductase activities, which was subtracted from the total rate. The rates were measured with a Gilford multiple-sample absorbance-recording spectrophotometer. 'The cuvette compartment was thermostatted at 30°C. Care was taken to confirm that the rates were constant with time and proportional to the amount of PPizF6P l-phosphotransferase. Definition of unit and specific activity. One unit was defined as the amount of enzyme that catalyzes the phosphorylation of l umol of fructose-G-P per minute in the standard assay. Specific activity (units per milligram of protein) was based on protein determinations by the method of Whitaker and Granun (134). Method 8: Forward reaction (alternative assay). Principle. For some determinations, such as measuring the activi- ty with fructose-1,6-P2 as the activator, a continuous spectrophoto- metric assay based on coupling Pi formation to non-rate-limiting amounts of glyceraldehyde-3-phosphate dehydrogenase and 3-phosphogly- cerate kinase was used. With the coupling enzymes in excess, the rate of Pi formation was equivalent to the rate of NAD+ reduction, which was measured by the absorbance increase at 340 nm. Reagents. HEPES-NaOH buffer (pH 7.8), 170 mM, containing 2.1 mM EDTA Fructose-G—P (freed from P; by chromatography on a column of Sephadex G-10), 170 nM Sodium pyrophosphate (Na PPi), pH 7.8, 17 mM MgClz, 102 mM 1 . !'!Esll§lhdl1p 24 DL-glyceraldehyde—3-P (prepared from DL-glyceraldehyde-3-P diethylacetal; Sigma Chemical Co.), 17 mM of the D-enantiomorph ADP (vanadium free), 17 nM Glyceraldehyde-3-phosphate dehydrogenase, 40 mU/ul, previously dialyzed against 1 nM EDTA, pH 7.8 3-Phosphoglycerate kinase, 80 mU/ul, previously dialyzed against 1 mM EDTA, pH 7.8 NADT, 8.2 mM Activator (3.4 mM fructose-1,6-P2) Procedure. The following were added to a microcuvette with a 1.0-cm light path: 80 ul of HEPES buffer, 10 ul of fructose-6-P, 10 ul of Na PPi, 10 u1 of MgClz, 5 ul of DL-glyceraldehyde—3-P, 10 ul of ADP, 5 ul of glyceraldehyde-3-P dehydrogenase, 5 ul of 3-phosphoglycerate kinase, 10 u1 of NADT, 5 ul of fructose-1,6-P2, a rate-limiting amount of PP1:F6P l-phosphotransferase, and water to a volume of 170 ul. The reaction was initiated by the addition of PPizFGP l-phosphotransferase. A control cuvette minus the enzyme preparation measures P1 release due to non-enzymatic hydrolysis of glyceraldehyde 3-phosphate, which was subtracted from the total rate. This assay is valid only when used with phosphatase-free preparations. In addition, the fructose-G-phosphate reagent must be essentially free of P1 so that the initial absorbance values can be kept low. The rates were measured with a Gilford multiple-sample absorbance-recording spectrophotometer. The cuvette compartment was thermostatted at 30°C. 25 Method C: Reverse reaction. Principl . This continuous spectrophotometric assay was based on coupling fructose-6-P formation to non-rate-limiting amounts of phos- phoglucose isomerase and glucose-6-P dehydrogenase. With the coupling enzymes in excess, the rate of fructose-6-P formation was equivalent to the rate of NADP+ reduction, which was measured by the absorbance in- crease at 340 nm. Reagents. HEPES-NaOH buffer (pH 7.8), 170 nM, containing 2.1 mM EDTA Fructose-1,6-P2,Ll7 mM Sodiun phosphate (Na Pi)» pH 7.8, 17 nM MgClg, 102 mM Phosphoglucose isomerase, 8D nU/ul, previously dialyzed against 1 mM EDTA, pH 7.8 Glucose-6-P dehydrogenase, 40 nU/ul, previously dialyzed against 1 mM EDTA, pH 7.8 NADP+, 8.2 mM Procedure. The following were added to a microcuvette with a 1.0- cm light path: 80 ul of HEPES beffer, 10 ul of fructose-1,6-P2, 10 ul of Na P;, 10 ul of MgClz, 5 ul of phosphoglucose isomerase, 5 ul of glucose-6-P dehydrogenase, 10 ul of NADP+, a rate-limiting amount of PP1:F6P 1-ph05photransferase, and water to a volume of 170 pl. Fruc- tose-1,6-P2 served both as the substrate and activator. The reaction was initiated by the addition of PPizFGP 1-phosphotransferase. A control cuvette minus Na P1 measured fructose-l,6-bi5phosphatase activity, which was subtracted from the total rate. The rates were measured with a Gilford multiple-sample absorbance-recording 5““ 26 Spectrophotometer. The cuvette compartment was thermostatted at 30°C. Other Enzyme Assays. All of the following assays in which pyridine nucleotides were monitored were done at 30°C and 340 nm with a Gilford automated record- ing spectrophotometer. ATP-dependent phosphofructokinase. The standard assay was a con- tinuous enzyme-coupled assay wherein the oxidation of NADH was fol- lowed. The reaction mixture (170 pl) consisted of 80 mM HEPES-NaOH (pH 7.4), 10 mM fructose-6-P, 1.0 mM ATP, 6 mM MgC12, 0.24 mM NADH, 0.2 unit of fructose-1,6-P2 aldolase, 2.0 units of triose—P isomerase, 0.2 unit of a-glycerol-P dehydrogenase, and rate-limiting amounts of the enzyme. Inorganic pyrophosphatase. Catalytic hydrolysis of PP; was mea- sured by the appearance of P1 using Josse's (135) modified Fiske and SubbaRow method. The incubation mixture (0.3 m1) contained 20 umol of 2-amino-2-methyl-l,3-propanediol-chloride (pH 9.1), 0.4 umol of MgClz, 0.2 umol of Na4P207, and enzyme preparation. The reaction was incubated for 20 min at 30°C, stopped by chilling to 0°C, followed by addition of 0.7 m1 of a mixture containing 0.1 ml of 5N H2504, 0.1 m1 of 2.5% ammonium molybdate (tetrahydrate), 0.1 ml of 3% NaHSO3 - 1% p-methylaminophenol sulfate (Elon), and 0.4 ml of water. After 10 min at 25°C, the absorbance at 660 nm was determined. Fructose-1,6-bisphosphatas . The enzyme was assayed by a phosphoglucose isomerase-glucose 6-phosphate dehydrogenase-linked assay, wherein the reduction of NADP+ was followed. The reaction mixture (170 pl) consisted of 80 mM HEPES-NaOH (pH 7.4), 2.0 nM 27 fructose-1,6-P2, 6 mM MgClg, 0.24 mM NADP+, 0.2 unit of phospho- glucose isomerase, 0.2 unit of glucose-6-phosphate dehydrogenase and rate-limiting amounts of the enzyme. Phosphoglucose isomerase. The enzyme was assayed by a glucose-6 ph05phate dehydrogenase-linked assay, wherein the reduction of NADP+ was followed. The reaction mixture (170 pl) consisted of 80 mM HEPES-NaOH (pH 7.4), 10 mM fructose-6-P, 6 mM MgC12, 0.24 mM NADP+, 0.2 unit of glucose-6-phosphate dehydrogenase, and a rate-limiting amount of the enzyme. Adenylate kinase. This enzyme was assayed by a pyruvate kinase- lactate dehydrogenase-linked assay wherein the oxidation of NADH was followed. The reaction mixture (170 p1) consisted of 80 mM HEPES-NaOH (pH 7.4), 1.0 mM AMP, 0.5 mM ATP, 3 mM phosphoenolpyruvate, 6 mM MgClz, 0.24 mM NADH, non-rate-limiting amounts of the coupling enzymes, and rate-limiting amounts of adenylate kinase. Assays for Molecular Weight Standards. All of the following assays in which the oxidation of NADH was monitored were done at 30°C and 340 nm with a Gilford automated record- ing spectrophotometer. Pyruvate kinase. Rabbit muscle pyruvate kinase was determined by measuring the production of pyruvate from phosphoenolpyruvate by a lactate dehydrogenase-linked assay. The reaction mixture (170 pl) contained 80 mM HEPES-NaOH (pH 7.8), 6 mM MgC12, 1 mM ADP, 0.5 mM ph05phoenolpyruvate, 0.24 mM NADH, non-rate-limiting amounts of lactate dehydrogenase, and 10 p1 of the column fractions. 28 Aldolase. Rabbit muscle aldolase was assayed by a triose-P isom- erase-a-glycerol-P dehydrogenase-linked assay. The reaction mixture (170 p1) contained 80 mM HEPES-NaOH (pH 7.8), 6 mM MgClg, 1 mM fruc- tose-1,6-P2, 0.24 mM NADH, 1.0 unit of triose-P isomerase, 0.2 unit of a-glycerol-P dehydrogenase, and 10 p1 of the colunn fractions. Hexokinase. Yeast hexokinase was assayed by a pyruvate kinase-- lactate dehydrogenase-linked assay. The reaction mixture (170 pl) con- tained 80 mM HEPES-NaOH (pH 7.8), 6 mM MgC12, 5 mM glucose, 0.5 mM ATP, 0.24 mM NADH, 2 mM phosphoenolpyruvate, 1 unit of pyruvate kinase, 1 unit of lactate dehydrogenase, and 10 pl of the colunn fractions. Ferritin. Ferritin was determined by reading the absorbance at 280 nm (room temperature) of 50-p1 column fractions diluted to 250 p1 with the buffer used to elute the column. Protein Determination. Protein was determined by the procedure of Lowry et al. (136), by the Bradford method (137) using bovine serum albumin as the standard, or by the method of Whitaker and Granun (134), which does not need a standard protein. Pressure Dialysis Concentration of Protein. Pressure dialysis was performed using an Amicon Diaflo apparatus (400-ml and 50-ml capacity), equipped with a PM30 membrane filter, hav- ing a 30,000-molecular-weight range cutoff. Concentration of the pro- tein was performed under nitrogen pressure (30 psi). When not in use, the filter was stored at 4°C in 20% ethyl alcohol after first washing it with distilled water. 29 Conductivity Measurements. Column fractions from purification steps were assayed for salt concentration using a standard conductivity meter with variable-conduc- tance control. Samples of 0.05-m1 volume were diluted lOO-fold with de-ionized water and read against a diluted buffer blank. The instru- ment was calibrated using the highest and lowest salt concentrations of the gradient in the appropriate buffer. Preparation of Fructose-2,6-Bisphosphat . Fructose-2,6-P2 was chemically synthesized as described by Van Schaftingen and Hers (27). The procedure involved (i) intramolecular cyclization of fructose-1,6-P2 to fructose 1,2 cyclic,6-bisphosphate, (ii) ring opening by alkaline treatment, (iii) neutralization and treatment with fructose-l,6-bisphosphatase, and (iv) separation by chromatography through Bio-Rad AG l-8X with elution by a gradient of NaCl. Fructose-2,6-P2 was quantified by measuring the fructose-6-P and P1 revealed upon acid hydrolysis. The fructose-6-P concentration was determined by enzymatic end-point assay using phosphoglucose isomerase and glucose-6-P dehydrogenase. The P1 concentration was determined by enzymatic end-point assay using glyceraldehyde-3-phos- phate dehydrogenase and 3-phosphoglycerate kinase. The separation of fructose-2,6-P2 from P1 and fructose-6-P through Bio-Rad AG l-8X is shown in Figure 2. 30 Figure 2. Purification of chemically synthesized fructose-2, 6-P2 by chromatography on a colunn of Bio-Rad AG l-X8. The fructose-2, 6-P2 corresponds to the fractions that gave equal concentrations of acid-revealed fructose-G-P and P- (fractions 56-68). The fructose-6-P concentration was determined by enzymatic end-point assay by coupling to phosphoglucose isomerase and glucose-6-P dehydrogenase. The P1 concentration was determined by enzymatic end-point assay by coupling to glyceraldehyde-3-P dehydrogenase and 3-phosphoglycerate kinase. 31 01 b 01 r 1 Ti mM F RUCTOSE-S-P or P; h) 1 C0 .1 IO 20 3O 4O 50 60 7O FRACTION NUMBER, 35 mI/TUBE Figure 2 II PI 0 FRUCTOSE-G-P U ACID REVEALED P; O ACID REVEALED FRUCTOSE-S-P l — 500 400 04 o 0 mM NoCl 200 I00 O 32 Preparation of Fructose-G-P Free of P; by Chromatography Through Sephadex G-10. I One-half gram of fructose-6-P was dissolved in water to 1.7 m1 and loaded onto a Sephadex G-10 column (2.6 x 100 cm) previously equilibrated with double-distilled H20. The sugar phosphate was eluted with double-distilled H20 at room temperature at a linear flow rate of 2.5 cm/hr. Two-hundred-twenty l-ml col unn fractions were col- lected. Fructose-G-P was determined by a phosphoglucose isomerase-glu- cose—6-P dehydrogenase-linked end-point assay. The reaction mixture (170 pl) contained 80 mM HEPES-NaOH (pH 7.8), 6 mM MgC12, 0.5 mM NADPT, 2 units of phosphoglucose isomerase, 1 unit of glucose-6-P dehydrogenase and 2 pl of 1/20 dilution of the column fractions. The absorbance change after the completion of the reaction was read at 340 nm. The concentration of fructose-G-P was calculated from the equiva- lent NADH formed. The P1 was determined by a glyceraldehyde-3-P dehydrogenase-3-phosphoglycerate kinase-linked end-point assay. The reaction mixture (170 pl) contained 80 mM HEPES-NaOH (pH 7.8), 6 mM MgC12, 1 mM ADP (vanadium free), 0.5 mM of the D-enantiomorph of DL-glyceraldehyde-S-P (prepared from DL-glyceraldehyde 3-phosphate diethylacetal), 0.5 mM NAO+, 1 unit of glyceraldehyde 3-ph05phate dehydrogenase, 2 units of 3-ph05phoglycerate kinase, and 15 pl of the column fraction. The absorbance change after the completion of the reaction was read at 340 nm. The concentration of P1 was calculated from the equivalent NADH formed. 33 Partial Purification and Elucidation of Some Properties of PPizFGP l-phosphotransferase. I A preliminary purification scheme devised earlier in this investi- gation has been described in the article of Sabularse and Anderson (138). This enzyme preparation (lOO-fold pure), along with three assay procedures, was utilized to elucidate the absolute requirement for fructose-G-P and PP; as well as for a divalent metal ion (Mg++), activation by a product of the reaction (fructose-1,6-P2), activation by glucose-1,6-Pz, and the effect of some compounds on the activity. Further Purification and Elucidation of Additional Properties of PP;:F6P l-phosphotransferase. Another preliminary purification scheme is described by Sabularse and Anderson (139) and elaborated on in an article by Anderson and Sabularse (140). This enzyme preparation (670-fold pure) was used in the determination of the KA for fructose-1,6-P2, glucose-1,6-Pz and fructose-2,6-P2; the Km for fructose-6-P and the relative Vmax: in the absence and in the presence of a hexose bisphosphate activator; and as a reagent in the demonstration of the natural occurrence of fructose-2,6-P2 in mung beans. Final Purification and Partial Separation of Two Forms of PPi:F6P l-Phosphotransferase and Characterization of Their Properties. Preparation of germinated seeds. Commercially available mung bean seeds (500 g) were soaked in distilled water for 12 hrs in the dark at 30°C. The soaked seeds were spread between 8 layers of moist cheese 34 cloth and allowed to germinate for an additional 12 hrs in the dark at 30°C. General. All purification steps were conducted at O-4°C. The as- say employed was Method A (described earlier) with 1 pM fructose-2,6-P2 as the activator. Preparation of crude extract. The germinated seeds were separated from the seed coat by repeated washing and hand-picking. The germi- nated beans, free of the seed coats, were suspended 1:1 (v/v) in buffer A (50 mM Tris-acetate, pH 7.0, containing 1 mM EDTA and 180 mM Na acetate) and ground with a mortar and pestle. The homogenate was filter-squeezed through 8 layers of cheese cloth and clarified by centrifugation at 13,000 x.g for 20 min. The supernatant solution was designated as the crude extract. Ammonium sulfate precipitate. To 2030 m1 of the crude extract, 369 g of ammonium sulfate was added slowly to bring the concentration to 30% saturation. The solution was stirred for 30 min and then centrifuged at 13,000 x.g for 20 min. The 2170 m1 of supernatant was decanted and 253 9 more of ammonium sulfate was added to bring the concentration to 50% saturation. After 30 min., the solution was centrifuged once again. The supernatant was discarded and the pelleted material was suspended in buffer B (10 mM Tris-acetate, pH 7.3 containing 0.1 mM EDTA and 10% glycerol) to 146 ml. DEAE-cellulose chromatography. The pooled 30-50% ammonium sulfate fraction was dialyzed against three 3-liter changes of buffer B and loaded onto a DEAE-cellulose column (4.7 x 26 cm) pre-equilibrated with buffer B. The column was washed with 5 bed-volumes of buffer 8 before the protein was eluted with 4.5-1iter linear gradient of 0 to 0.4 M KCl 35 in the same buffer. Fractions 62 to 95, containing most of the activity, were pooled for further purification. Phosphocellulose chromatography 1. The pooled fractions from the DEAE-cellulose column were concentrated to 77 ml by pressure fltration through a PM30 membrane. The concentrated pooled fractions were dia- lyzed against 3 changes of 2 liters each of buffer C (5 mM Na-PIPES, pH 6.6 containing 10 mM KCl, 0.1 mM Na EDTA and 10% glycerol), and loaded onto a phosphocellulose column (4.0 x 14.5 cm) previously equilibrated with buffer C. The loading was followed by passing through the column about 5 bed-volumes of buffer C before the enzyme was eluted with buffer C containing 34 mM Na PP; with the pH maintained at 6.6. Fractions 50-115, containing most of the activity, were pooled for further purification. Phosphocellulose chromatography II. The pooled fractions from phosphocellulose chromatography 1 were concentrated to 77 ml by pres- sure filtration through a PM30 membrane. The concentrated pooled fractions were dialyzed against three 2-liter changes of buffer C, and loaded onto a second phosphocellulose column (2.6 x 11.0 cm) previously equilibrated with buffer C. The loading was followed by passing through the column about 4 bed-volumes of buffer C before the enzyme was eluted by a 1500-m1 linear gradient of O to 0.5 M KCl in buffer C. Fractions 69-92, containing most of the activity, were pooled for further purification. Bio-Gel A-1.5m. The pooled fractions from phosphocellulose chromatography II were concentrated to 10 ml by pressure dialysis through a PM30 membrane. The concentrated pooled fractions were applied to a Bio-Gel A-l.5m colunn (3.4 x 95 cm) equilibrated with 36 buffer B with the addition of 20 mM KCl, and then eluted with the same buffer. Fractions (S-ml each) were collected and those with high specific activity for the enzyme (fractions 82 through 123) were combined. Separation of two forms of the enzyme. PP1:F6P l-Phosphotrans- ferase in the above fraction was separated into two forms by further chromatography on Bio-Gel A-l.5m as detailed in the Results section. Partial Purification and Determination of Some Properties of ATP-dependent Phosphofructokinase. ATP-dependent phosphofructokinase was extracted and partially pur- ified from 50 g of mung beans germinated in the dark at 30°C for 24 hrs. The sprouts were separated from their seed coats, suspended 1:1 (v/v) in buffer (50 mM Tris-acetate, pH 7.0, containing 5 mM dithio- threitol, 1 mM EDTA, and 180 mM K acetate, and were ground with a mor- tar and pestle. The homogenate was filter-squeezed through 8 layers of cheese cloth and clarified by centrifugation at 22,000 x‘g for 20 min. The ATP-dependent phosphofructokinase in the supernatant fluid (the crude extract) was partially purified (20 fold) by the following proce- dure: precipitation with ammonium sulfate (30-45% saturation); chroma- tography on Sephadex G-100 (the ammonium sulfate precipitate was dis- solved to 10 ml with the elution buffer and loaded onto a 500-ml bed-volume column, and was then eluted with 10 mM Tris-acetate buffer at pH 7.0 containing 1 mM DTT, 0.1 mM EDTA and 180 mM K acetate); chro- matography on DEAE-cellulose I (the active fractions from Sephadex G-100 column chromatography step were pooled, dialyzed, and loaded onto a 20-ml bed-volume column previously equilibrated with 10 mM Na-MES, pH 37 6.2, containing 1 mM DTT and 0.1 mM EDTA, and then the protein was eluted by 20 ml each of a step gradient of 0.1, 0.2, 0.3 and 0.4 M KCl in the pH 5.2 Na-MES buffer); and chromatography on DEAE-cellulose 11 (0.2 and 0.3 M KCl eluates from the DEAE-cellulose I step were pooled, dialyzed, and loaded onto a second ZO-ml bed-volume column previously equilibrated with the pH 6.2 Na-MES buffer, and the protein was eluted by a ZOO-m1 linear gradient of 0-400 mM KCl in the pH 6.2 Na-MES buffer). The tubes containing most of the phosphofructokinase activity devoid of PPizF6P l-phosphotransferase were combined, concentrated by pressure filtration through a PM30 membrane, and stored frozen. This enzyme preparation was designated as DEAE-cellulose II ATP-dependent phosphofructokinase. The DEAE-cellulose II ATP-dependent phosphofructokinase prepara- tion was used in the determination of the saturation curves for fruc- tose-6-P and ATP, and the effect of pH and some metabolites on the activity. Extraction of the Natural Activator (Fructose-2,6-P2) from Mung figppg. The natural activator was extracted from germinated mung bean (Phaseolus aureus Roxb.) seeds. Twenty-five grams of mung bean seeds, purchased locally, were germinated in the dark at 30°C for 24 hr. The sprouts were separated from their seed coats, suspended 1:1 (v/v) in buffer (100 mM glycine-NaOH, pH 10.4, containing 300 mM NaCl) and ground with a mortar and pestle. The homogenate was squeeze-filtered through 8 layers of cheese cloth and the filtrate was adjusted to pH 9.4 with NaOH, then clarified by centrifugation at 22,000 x.g for 20 38 min. The crude extract was pressure filtered through a PM30 ultrafil- tration membrane and the filtrate was diluted with water six-fold be- fore being introduced to a column ( l x 15 cm) of Bio-Rad AG 1-X8 anion exchange resin (Cl' form, 200 to 400 mesh). The column was washed with 10 bed-volunes of water and the sugar phosphate was eluted with a linear gradient (100 ml, 30 ml/h) of 100 to 400 mM NaCl. Eighty-four 1.2-ml fractions were collected. Fructose-1,6-P2 was determined by enzymatic end-point assay using fructose-1,6-P2 aldolase, triose-P isomerase and a-glycerol-P dehydrogenase. The unknown activator was located and measured by its ability to activate PPizF6P l-phosphotransferase; if the column fraction also contained fructose-1, 6-P2, PP1:F6P l-phosphotransferase was added to the assay mix after the initial rapid decrease in absorbance. A scaled—up isolation procedure starting with 1000 g of germinated beans was conducted in which fructose-l,6-bisphosphatase treatment and a second anion-exchange chromatography step were used to separate the activator from fructose-1,6-P2. The fractions containing the activa- tor were pooled and concentrated by lyophilization. A portion of the concentrate was subjected to mild acid hydrolysis (pH 2.5, room temperature, for 30 min) and the resulting hydrolysate was neutralized with NaOH, and examined for fructose-G-P by specific enzyme-coupled assays employing (i) glucose 6-phosphate isomerase (EC 5.3.1.9) and glucose-6-phosphate dehydrogenase (EC 1.1.1.49), and (ii) phosphofructokinase (EC 2.7.1.11), fructose-1,6-P2 aldolase, triose-P isomerase, and a-glycerol-P dehydrogenase. 39 Survey of PP1:F6P l-Phosphotransferase and Fructose 2,6-Bisphosphate in Other Plant Species. Assay Method A with fructose-2,6-Pz as activator was employed to determine the presence of the enzyme in several plant species. In each plant species that was surveyed for the enzyme, crude extract from about 25 g of sample was prepared in similar manner as in the prepara- tion of crude extract from sprouted mung bean. The crude extracts were assayed for the enzyme both in the presence and absence of fructose-2, 6-P2. The blank was either no PP1 or fructose-G-P. Protein was determined by the method of Whitaker and Granum (134). Using the purified PP1:F6P 1-phosphotransferase as a reagent and employing assay Method A, a survey of the occurrence of fructose-2,6-P2 in some plant species was conducted. A five to twenty-five gram sample of each plant species was used and the extraction and isolation procedures were as described earlier for the extraction and isolation of the natural activator from sprouted mung beans. The amount of fructose-2,6-P2 per unit weight of sample was estimated by noting the relative area of the elution profile from the AG l-X8 anion exchange column and relating it to the weight of plant material that was used during the extraction. RESULTS Detection of PPigFGP 1-Phosphotransferase in an Extract From Germinated Mung Beans. With minor modifications of published assay methods (17, 18), this enzyme activity was detected in the crude extract [prepared by following the procedure of Nomura and Akazawa (141) for the preparation of crude extract for enzymatic assay for sucrose-P synthase from germinated rice seeds] of sprouted mung beans. However, the apparent specific activity was very much lower than that of the ATP dependent phosphofructokinase, being 1/40 even at the 24-hour germination time, when both the enzymes were at their highest apparent specific activities (Figure 3). In contrast, in other species where the enzyme has been reported (17-21), the PP;:F6P l-phosphotransferase activity is much greater than that of the ATP-dependent phosphofructokinase. Because the existence of this enzyme in C3 plants had not been previously recognized, the research was pursued on the basis of the idea that the apparent low specific activity in mung beans was due to suboptimal assay conditions or the existence of the enzyme in an inhibited or unactivated state. 40 41 Figure 3. Apparent specific activities of PPizFGP I-Phospho- transferase and ATP-dependent phosphofructokinase at different times of germination prior to the discovery of activation by hexose bisphos- phates. The assay mixture (170 pl) for PPizF6P 1-Phosphotransferase contained 80 mM HEPES-NaOH buffer (pH 7.4), 25 mM Na fructose-G-P, 1.0 mM PPi, 3.0 mM MgC12, 0.24 mM Na NADH, 0.2 unit of fructose-1,6- P2 aldolase, 2.0 units of triose-P isomerase, 0.2 unit of glyceralde- hyde-3-P dehydrogenase, and 10 pl of crude extract. A similar assay mixture was employed for ATP-dependent ph05phofructokinase except that 1 mM ATP replaced PP;, and only 10 mM Na fructose-6-P and 5 p1 of crude extract were used. The observed velocities were divided by 2 to correct for the 2-fold amplfication. Protein was determined by the method of Lowry et a1. l-PT = l-phosphotransferase. PFK = ATP-depen- dent phosphofructokinase. 42 3 290% 335 En. m W W — ..._O >._._>_._.0< ”:..—5me 5 7. 0 fl ERR... am...” mega o m o I -w 24 GERMINATION TIME, HOURS O 3 szE 2.55 .E._ am“: R—u b 251‘ O O. .9. do E254 0:695 Figure 3 43 Partial Purification and Some Preperties of PP;:F6P l-Phosphotransferase. For use in a preliminary investigation, the enzyme was partially purified (100 fold) as described by Sabularse and Anderson (138). The purification steps consisted of ammonium sulfate precipitation, DEAE cellulose chromatography, Bio-Gel A-l.5m chromatography and phosphocel- lulose chromatography. The enzyme preparation was essentially free of enzymes (e.g., ATP-dependent ph05phofructokinase, adenylate kinase, ph05phoglucose isomerase and fructose-1,6-bisphosphatase) that could interfere with the assay. Requirement for specific substrates and a divalent metal ion. The partially purified enzyme had an absolute requirement for both fruc- tose-6-P and PP;. ATP could not substitute for PP; (Table 1, Assay I). Other compounds (1 mM) that could not replace PP; as the phos- phoryl donor included UTP, ADP, UDP, P;, AMP and phosphoenolpyruvate. There was also an absolute requirement for a divalent metal ion such as Mg++. Mn++ and Co++ could replace Mg++ but were less effective. Zn++, Cu++, Fe++ and Ca++ did not substitute for Mg++ (Table 2). Several compounds that were tested as possible effectors for the enzyme had little or no apparent effect with the assay conditions used (Table 3); however, the observed inhibition by P; was significant as will be documented later in this thesis. Activation of PP13F6P l-phosphotransferase by a product of the reaction. Several observations on the kinetics of the assay suggested that the enzyme was activated by a product of the reaction, fructose-l, 6-P2: (1) whereas omitting triose-P isomerase would be expected to halve the rate of NADH oxidation in an aldolase-triose-P 44 Footnotes to Table 1. aThe assay mix (170 pl) contained 80 mM HEPES-NaOH buffer (pH 7.8), 20 mM Na fructose-6-P, 1.0 mM Na PP;, 6.0 mM MgC12, 0.3 mM Na NADH, 100 mU of fructose-1,6-P aldolase, 2.0 U of triose-P isomerase, 200 mU of a-glycero -P dehydrogenase, and partially purified PP;:F6P I-phosphotransferase. The observed velocity was divided by 2 to correct for the 2-fold amplification. bThe assay mix (170 p1) contained 80 mM HEPES-NaOH buffer (pH 7.8 , 20 mM Na fructose-6-P, 1.0 mM Na PP;, 6.0 mM MgC12, 0.3 mM Na NAD , 1.0 mM glyceraldehyde-3-P, 3.0 mM Na ADP, 100 m0 of D-glyceral- dehyde-3-P dehydrogenase, 200 mU of 3-P-glycerate kinase, and partially purified PP;:F6P l-phosphotransferase. The reported velocities have been corrected for a background rate due to glyceraldehyde-3-P hydroly- sis (see Figure 7, Curve 3). cThe assay mix (170 pl) contained 80 mM HEPES-NaOH buffer (pH 7.8), 2.0 mM Na fructose-1,6-P , 2.0 mM Na P;, 6.0 mM MgC12, 0.3 mM Na NADPT, 100 mU of phospho exose isomerase, 200 m0 of D-glucose-6-P dehydrogenase, and partially purified PP;:F6P 1-phos- photransferase. dA11 assays contained 1.7 pg of the PP;:F6P l-phOSphotrans- ferase preparation. The rates observed for the three assays (i.e., 2.8, 3.2, and 0.92) are not identical because of the influence of dif- fering concentrations of contaminating and component ions on the reaction velocity. eThe maximal slope was achieved after the optimally activating concentration of fructose-1,6-P2 was formed in the reaction. 45 Table 1. Some properties of PP;:F6P l-phosphotransferase partially purified from mung bean sprouts, as determined by three different assay methods. Conditions Reaction Velocityd Assay Omissions Additions (nmol/min) Ia None None 0.14 (measures PP;:F6P l-phospho- fructose- transferase None 0.00 1,6-P2 Fructose-6-P None 0.00 formed) MgClz None 0.00 PP; None 0.00 PP; ATP (1.0 mM) 0.00 None Glucose-1,6-P2 (0.3 mM) 1.3 None Glucose-1,6-P2 (1.5 mM) 2.8 IIb None None 3.2 (max slope)e (measures PP;:F6P l-phospho- P; formed) transferase None 0.00 Fructose-6-P None 0.00 PP; None 0.00 None Fructose-1,6-P2 (0.3 mM) 3.2 None Glucose-1,6-P2 3.2 IIIc None None 0.92 (measures PP;:F6P l-phospho- reverse transferase None 0.00 reaction) Fructose-l, 5-P2 None 0.00 P; None 0.00 None Glucose-1,6-P2 (0.3 mM) 0.92 Table 2. 46 Effect of various divalent metals on PP;:F6P l-phOSpho- transferase. The assay mix (170 p1) contained 80 mM HEPES-NaOH buffer (pH 7.8), 20 mM Na fructose-6-P, 1.0 mM g, 0.24 mM Na NADH, 0.2 unit of ase, 2.0 units of triose-P isomerase, Na4P207, 6.0 mM MgCl fructose-1,6-P2 aldo 0.2 unit of a-glycerol-P dehydrogenase, 1.5 mM glucose-1,6-P2, 10 pl of partially purified PP;:F6P il-phosphotransferase and the indicated divalent metal. Activity Divalent metal Concentration (mU) None - 0.04 None + 1 mM EDTA - 0.00 Mg“ 2 mm 1.20 Co++ 2 mM 0.76 Mn++ 2 mM 0.34 Ca++ 2 mM 0.04 Fe++ 2 mM 0.04 Cu” 2 "M 0.04 Zn++ 2 mM 0.04 Table 3. 47 Effect of some compounds on PP;:F6P l-phosphotransferase. The assay mix (170 pl) contained 80 mM HEPES-NaOH buffer (pH 7.8), 20 mM Na fructose-G-P, 1.0 mM Na4P207, 6.0 mM MgCl , 0.24 mM Na NADH, 0.2 unit of fructose-l-6-P2 aldo ase, 2.0 units of triose-P isomerase, 0.2 unit of a-glycerol-P dehydrogenase, 10 p1 of partially purified PP;:F6P l-phosphotransferase and the indicated amount of the possible effector tested. The putative effector was preincubated with the reaction mixture for 10 min before the reaction initiated by the addition of fructose-6-P. Activity (nmol fructose-1,6-Pz Concentration formed/min) of effector No effector With effector Effect 1 1 1 1 1 l 1 1 l 1 1 1 mM UDP-Glucose 0.60 0.67 mM Glucose-l-P " 0.66 mM P1 " 0.45 mM ATP " 0.50 mM UDP " 0.53 mM ADP " 0.60 mM UTP " “ mM Phosphoenolpyruvate mM AMP " " mM Citrate " " mM Glucose-6-P " " mM 3-Phospho- " " glyceric acid mM 1,3-Diphospho- “ " glyceric acid 0.08 mM NAD+ " " 12% Activation 10% “ 25% Inhibition 17% " 11% " no effect 48 isomerase-a-glycerol-P dehydrogenase-linked assay, the omission actual- ly caused a time-dependent increase (Figure 4, Curves l and 2); (ii) the time required to achieve the apparent activation of PP;:F6P 1-phosphotransferase in the absence of triose-P isomerase was decreased by the addition of glyceraldehyde-3-P (Figure 4, Curve 3); (iii) in- creasing the concentration of PP;:F6P 1-phosphotransferase resulted in more than a proportionate increase in the reaction velocity (Figure 5, Curve 1), but when triose-P isomerase was omitted from the assay, the rate was essentially proportional to the PP;:F6P l-phosphotrans- ferase concentration (Figure 5, Curve 2) and (iv) delaying the addition of aldolase to the assay mixture for 5 min resulted in an absorbance decrease that was considerably greater than that in a control in which aldolase was added at the start of the reaction (data not shown). It was deduced from these observations that varying the conditions influ- enced the steady-state concentration of fructose-1,6-P2, and that when the concentration was elevated, PP;:F6P l-phosphotransferase was activated. Thus, omitting triose-P isomerase caused glyceraldehyde-B-P to accunulate, thereby increasing the fructose-1,6-Pz concentration by mass action; the addition of exogenous glyceraldehyde-B-P in the absence of triose-P isomerase potentiated this effect; increasing the PP;:F6P 1-phosphotransferase concentration at constant levels of coupling enzymes increased the steady-state concentration of fructose-1,6-P2 even in the presence of triose-P isomerase, thus accounting for the apparent enzyme-concentration-dependent activation of PP;:F6P 1-phosphotransferase; and delaying the addition of aldolase resulted in a temporary accumulation of fructose-1,6-P2, thereby activating the PP;:F6P 1-phosphotransferase. 49 Figure 4. Apparent time-dependent activation of PP;:F6P I-phos- photransferase in the absence of triose-P isomerase and apparent stimu- lation of activity by glyceraldehyde-3-P. Curve 1, complete reaction mixture (Assay I in Table l, uncorrected for 2-fold amplification); Curve 2, complete minus triose-P isomerase; Curve 3, complete minus triose-P isomerase and plus 0.16 mM glyceraldehyde-B-P. 50 1.6 - 1 1.4— - A340 1.2 — - 3 2 I 1 1 1 1 "00 5 10 15 20 MINUTES Figure 4 51 Figure 5. Proportionality of PP F6P l- -phosphotransferase acitivity and enzyme concentration. Curve 1, complete reaction mixture (Assay I in Table 1): Curve 2, complete minus triose-P isomerase. 52 15 _ _ _ _ 5 4. 3 2 z_2\m_-m..-mmobamn_-a I 10 I 5 pg PROTEIN Figure 5 53 Direct demonstration of activation by fructose 1,6-bisph05phate. A direct demonstration of fructose-l,6-P2-dependent activation of PP;:F6P 1-phosphotransferase was achieved by following the rate of P; formation using a glyceraldehyde-B-P dehydrogenase-coupled assay (Table 1, Assay II). This assay necessitates that the reagents have negligible amounts of P; so that the initial absorbance of the reac- tion mixture could be kept low. 0f the reagents in the assay mixture, conmercial 1y available fructose-6-P was found to contribute a signifi- cantly high amount of P;. The commercially available biochemical re- agent grade fructose-6-P can have from 0.5 to over 1.0 mole % P; on prolonged storage. A 0.5 mole % P; will contribute an absorbance of 0.6 to the initial value, thereby reducing the range of absorbance that is available to follow the progress of the reaction. Therefore, fruc- tose-6-P was freed of its contaminant P; by colunn chromatography through Sephadex G-10 (Figure 6). Another precaution to be observed with the assay concerns the use of a suitable ADP reagent. The assay was inoperative unless ADP that was designated "vanadium-free“ was used. The mechanism of inhibition of the assay by vanadium was not studied thoroughly. However, when vanadium-containing ADP was added to the assay mixture for PP;:F6P 1-phosphotransferase by fructose-1,6-P2 aldolase-triose—P isomerase-a-glycerol-P-dehydrogenase, no such inhibition was observed, suggesting that the vanadium inhibits either glyceraldehyde-3-P dehydrogenase, 3-phosphoglycerate kinase, or both. The inhibition by vanadium was not studied further, although it was known (142-145) that vanadium inhibits Na/K ATPase, and therefore the site of inhibition by 54 Figure 6. Elution profile of fructose-6-P and P; by chromatog- raphy through Sephadex G-10. One-half mg of fructose-6-P was dissolved in H20 to 1.7 ml and introduced to a Sephadex G-10 colunn (2.6 x 100 cm). The sugar phosphate was eluted with H20 at room temperature at a linear flow rate of 2.5 cm/hr. The fructose-6-P was determined by enzymatic end-point assay using phosphoglucose isomerase and glucose-6- P dehydrogenase, whereas the P; concentration was determined by en- zymatic end-point assay using glyceraldehyde-3-P dehydrogenase and 3- phosphoglycerate kinase. Other details of the assay are in the text. mM FRUCTOSE-G-P (“I IZOI— I00 80;- 60 4o 55 -./ ISO ITO 180 I90 ZOO 2200 FRACTION NUMBER, I mI/ TUBE Figure 6 56 vanadium may be the 3-phosphoglycerate kinase step of the enzymatic assay. Having worked out the procedure for the assay that could monitor P; formation, the experiment to demonstrate the effect of fructose-l, 6-P2 was conducted. In the presence of added 100 pM fructose-1,6- P2, the velocity was maximal and was constant with time, whereas in its absence the velocity was initially slow and increased with time (as the fructose-1,6-P2 product accumulated) to the same rate as that achieved with initially added fructose-1,6-P2 (Figure 7). ATP, 1,3-diphosphoglycerate, or 3-phosphoglycerate, did not activate the PP;:F6P l-phosphotransferase when added to the reaction mixture during the fructose-1,6-P2 aldolase-coupled assay. The conclusion was that micromolar concentrations of fructose-1,6-P2 activated PP;:F6P l-phosphotransferase, causing a many-fold increase in the reaction velocity even at saturating levels of the substrates, fructose-6-P and PP;. Activation by_glucose 1,6-bisphosphate. The glyceraldehyde-B-P dehydrogenase-coupled assay was not convenient for routine measurements of PP;:F6P l-phosphotransferase because of the need for P;-free reagents and the inapplicability of the assay to crude extracts, which contain phosphatases. So the search for other possible activators was continued and the logical approach was to try other hexose bisphos- phates. Glucose-1,6-P2 was found able to replace fructose-1,6-P2 as an activator in the aldolase-coupled assay (Table 1, Assay I). Glu- cose-1,6-P2 (1.5 mM) activated about 20 fold, and the reaction veloc- ity was proportional to PP;:F6P l-phosphotransferase concentration. Glucose-1,6-P2 also activated PP;:F6P l-phosphotransferase in the 57 Figure 7. Activation of PP;:F6P l-phosphotransferase by fruc- tose-1,6-P2. Curve 1, complete reaction mixture (Assay II of Table 1) plus 0.10 mM fructose-1,6-P2; Curve 2, complete reaction mixture (no fructose-1,6-P2 added); Curve 3, BIank (complete minus fructose- 6-P and PP;). ‘— 58 0.4,. i T 0.3 " I A3410 0.2 '- O.I - I I 00 2 4 MINUTES Figure 7 59 glyceraldehyde-B-P dehydrogenase-coupled assay (Table I, Assay II), thus abolishing the lag. Glucose-1,6-P2 did not further activate the enzyme when the reaction was run in the reverse direction (Table l, Assay 111) because in this case fructose-1,6-P2 served both as the substrate and the activator. Glucose-1,6-P2 could not replace fruc- tose-1,6-P2 as the substrate. Optimization of the Assays for PP;:F6P l-Phosphotransferase. Forward reaction (standard assay). A preliminary determination of the KA for glucose-1,6-P2 indicated it to be about 0.5 mM. The magnesium ion required for half-maximal activity was 0.3 mM (Figure 8), and 6 mM Mg++ gave the highest Vmax of the metal ions tested. The activity as a function of pH was maximal at about pH 7.8 in HEPES-NaOH buffer. One millimolar PP; was optimal and the activity at 20 mM fructose-G-P approached the plateau for the saturation curve, which started to decrease beyond 30 mM. The addition of 0.4 mM EDTA (Figure 9) completely abolished the background activity, whereas the activity was constant in the presence of 6 mM Mg++ even when EDTA was present at 2 mM. In addition, it was determined that (NH4)ZSO4 from the coupling enzymes contributed an inhibitory effect. From the above data, the conditions for the tentative standard assay using the aldolase-coupled assay that was adopted was as follows: an assay mixture (170 pl) contained 80 mM HEPES-NaOH buffer (pH 7.8), 1 mM Na EDTA, 20 mM Na fructose-6-P, 1.0 mM Na PP;. 6.0 mM MgC12, 0.24 mM Na NADH, 0.2 unit of fructose-1,6-P2 aldolase, 2.0 units of triose-P isomerase, 0.2 unit of a-glycerol-P dehydrogenase, 1.5 mM glucose-1,6-P2, and rate-limiting amounts of PP;:F6P 60 Figure 8. Mg++ concentration-velocity curve for PP;:F6P l- phosphotransferase. The assay mixture (170 p1) contained 80 mM HEPES- -Na0H buffer (pH 7.8), 20 mM Na fructose-6-P, 1.0 mM Na PP;. 0.24 mM Na NADH, 0.2 unit of fructose-1,6-P2 aldolase, 2.0 units of triose-P isomerase, 0.2 unit of a-glycerol-P dehydrogenase, 1.5 mM glucose-1, 6-P2 and 5 pl of the lOO-fold purified enzyme. The velocity is expressed as nmol of fructose-1,6-P2 formed per min. 61 .--1 IO 1.25 - I.OO - P E. O >._._004m> 0.50 O. 25 3 4 5 [Mgr], mM 2 Figure 8 62 Figure 9. Absolute requirement of PP; :F6P 1- -phosphotransferase for divalent metal, The assay mixture (170 pl) contained 80 mM HEPES- NaOH bufferM (pH 7. 8), 20 mM Na fructose-6-P, 1.0 mM Na PP; 0. 24 mM Na NADH, O. 2 unit of fructose-1, 6- -P2 aldolase, 2. 0 units tri- ose- -P isomerase, 0. 2 unit of o-glycerol- -P dehydrogenase, 1. 5 mM glu- cose-l, 6- -P2 and 5 pl of the 100- fold purified enz e. The activities were determined at no added Mg++ , and at 6 mM Mg+ at indicated Na EDTA concentrations. 63 Z :5 T7 I 1 r rah—‘11 8’] 1.5— - 0' 16 mM Mg"+ % H—a—+—a—a.—+/x——A E g.“ 1.0— - (D 1: '9 E O.5r ‘ E if . :5 ¢¢«DAMNNEDIWQ+I 5 ohm—.265... <1 O 100 200 300 400 50 1000 M No-EDTA Figure 9 64 l-phosphotransferase. The coupling enzymes were dialyzed against 1 mM EDTA at pH 7.8 to remove (NH4)2504 or citrate which may be present. Using the tentative standard assay described above, a specific activity of 0.018 unit of PP;:F6P 1-phosphotransferase per mg protein was detected in the crude extract. This observed specific activity was an increase of about 40 times over that found when it was first detected, making the level of the enzyme about equal to that of ATP-dependent phosphofructokinase. The final standard assay (Method A as described in Materials and Methods) was essentially the same as the above tentative standard assay except for halving the concentration of Na fructose-G-P to 10 mM and the use of 1 pM fructose-2,6-P2 instead of 1.5 mM glucose-1,6-P2, which were adopted after the discovery of fructose-2,6-P2 activation (see below). Using fructose-2,6-P2 as the activator, the specific activity of the enzyme in mung bean sprouts was about twice the apparent specific activity of the ATP-dependent phosphofructokinase. Other assays far.PP;:F6P I-phosphotransferase. Assay 8 (forward reaction alternative assay) and assay C (reverse reaction), as described in Materials and Methods, were also optimized by the approach described in the above section. Determination of Inorganic Pyrophosphatase. Determination of inorganic pyrophosphatase activity in the crude extract and in the partially purified PP;:F6P 1-phosphotransferase was conducted employing Josse's modification (135) of Fiske and SubbaRow method for P; determination. In the crude extract, 65 inorganic pyrophosphatase was apparently absent, with specific activity of less than 0.002 unit/mg protein (Table 4). Simmons and Butler (146) reported that mature seeds of maize have no inorganic pyrophosphatase, also. The subcellular localization of the inorganic pyrophosphatase in higher plants is in the chloroplast (146, 147). Table 4 also shows that partially purified PP;:F6P 1-phosphotransferase was devoid of inorganic pyrophosphatase activity. Some Properties of the ATP-dependent Phosphofructokinase in Mung Beans. The enzyme that is usually responsible for the formation of fruc- tose-1,6-P2 during glycolysis is the ATP-dependent phosphofructo- kinase, normally a regulatory enzyme. Since PP;:F6P l-phosphotrans- ferase may also be involved in glycolysis, it was decided to determine some kinetic properties of the mung bean ATP-dependent ph05phofructo- kinase for comparison with those of the pyrophosphate-dependent enzyme. ATP-dependent ph05phofructokinase was partially purified (20-fold) from mung beans by ammonium sulfate precipitation, chromatography on a Sephadex G-100 column, and two successive steps of column chromatography on DEAE-cellulose. The pH dependence of the activity of the enzyme is shown in Figure 10. It is characterized by a nearly dome-shaped curve with an optimal pH range of 7.2 to 7.8. The saturation curve for fructose-6-P gave hyperbolic kinetics (the data will be presented in Figure 19). In contrast, ATP-dependent phosphofructokinase from other sources (muscle, brain, heart, liver, kidney cortex, sperm, carrots, Brussels sprouts, N, ‘pppgpg, g. pasteurianum, E, £911 and yeast) gave sigmoidal kinetics in the absence of an added allosteric effector (148). The effect of 66 Table 4. Assay for inorganic pyrophosphatase. The crude extract and partially purified PP;:F6P l-phosphotransferase (1.52 units/mg) were assayed for possible inorganic pyrophosphatase activity employing Josse's modification (135) of the Fiske and SubbaRow method. The assay mixtures were incubated for 20 min at 30°C. The sensitivity of the assay was 0.04 pmol of P;. Therefore the specific activity of inorganic pyrophosphatase in the crude extract was less than 0.002 pmol of PP; conswned per min per mg protein. Volume Protein P; Specific Preparation Protein Liberated Activity Sample (pl) (mg) (pmol) (U/mg protein) Crude extract 10 0.15 0.00 (0.007 20 0.30 0.000 (0.003 30 0.45 0.00 (0.002 PP;:F6P l-Phosphotransferase 20 0.26 0.00 (0.004 40 0.52 0.00 (0.002 67 Figure 10. Effect of pH on the activity of ATP-dependent phospho- fructokinase. The reaction mixture (170 pl) contained 1.0 mM ATP, 10 mM fructose-6-P, 6 mM MgC12, 0.24 mM NADH, 0.2 unit of fructose-l, 6-P aldolase, 2.0 units of triose-P isomerase, 0.2 unit of a—glyc- ero -P dehydrogenase, 1.3 mU of the DEAE-cellulose II ATP-dependent phosphofructokinase preparation and 100 mM of the indicated buffer. 68 '50- T 1 l -‘I-I’. Z 3 1.25 — ‘1- Lo“ [:3 8 LOO- 13 D 0: L1. '5 0.75- E CI >_.. t o No MES > _ - I3 050 a No-PIPES <1? I No-HEPES o TRIS-CHLORIDE 025; 1:174 . I 6 7 8 pH Figure 10 69 ATP concentration is shown in Figure 11. An increasing ATP concentration up to 2 mM stimulated the activity whereas a further increase was inhibitory. This behavior of inhibition by a high concentration of ATP was also observed for ATP-dependent phosphofructokinases from other sources (32, 149). The effect of some metabolites on the activity of the ATP-depen- dent phosphofructokinase is shown in Table 5. Orthophosphate activat- ed, whereas PP;, AMP, ADP, citrate and phosphoenolpyruvate inhibited. The most pronounced inhibition was observed with phosphoenolpyruvate. In a separate experiment, it was observed that 10 pM phosphoenolpyru- vate inhibited >99% (Figure 12). The phosphoenolpyruvate concentration that gave 50% inhibition was 2 pM, making ph05phoenolpyruvate a very potent inhibitor of ATP-dependent ph05phofructokinase. The potent inhibition by ph05phoenolpyruvate has been previously observed for the ATP-dependent phosphofructokinase in peas (150). It should be recalled that phosphoenolpyruvate does not inhibit mung bean PP;:F6P l-phosphotransferase even at 1 mM. Stability of PP;:F6P l-Phosphotransferase at Various Conditions of Storage. It was observed initially that the partially purified enzyme lost activity on storage both at 4°C and at -20°C. To find a storage condi- tion that would preserve the activity, varying storage temperatures and the addition of possible protecting reagents (glycerol, DTT, fructose-G-P and PP;) were tried. The enzyme (in 10 mM Tris-acetate buffer - pH 7.3, 0.1 mM EDTA, 1.0 mM DTT, and 17 mM KCI) was stable (100% in 4 months) on storage at -20°C with added 20% (v/v) glycerol, 70 Figure 11. ATP concentration-velocity curve for mung bean ATP-de- pendent phosphofructokinase. The assay mixture (170 pl) contained 80 mM HEPES-NaOH buffer (pH 7.4), 10 mM fructose-6-P, 6.0 mM MgC12, 0.24 mM Na NADH, 0.2 unit of fructose-1,6-P aldolase, 2.0 units of tri- ose-P isomerase, 0.2 unit of a-glyceroT-P dehydrogenase, 0.43 m0 of DEAE-cellulose II ATP-dependent ph05phofructokinase, and the indicated amounts of ATP. 71 _ Ru O. 4. O 238.2%... Rermmoeoamn. .62: 5.254 Figure 11 72 Table 5. Effect of some metabolites on ATP-dependent ph05phofructoki- nase. The reaction mix (170 pl) contained 80 mM HEPES buffer (pH 7.4), 10 mM fructose-G-P, 1.0 mM ATP, 6 mM MgC12, 0.24 mM Na NADH, 0.2 unit of fructose-1,6- aldolase, 2.0 units of triose-P isomerase, 0.2 unit of a-egcerol-P dehydrogenase and 10 p1 of the partially purified DEAE-cellulose II ATP-de- pendent phosphofructokinase. Concentration of Activity (nmol fructose-1,6-P2 effector in assay formed/min) Mixture No effector With effector Effect 2 mM P; 0.131 0.179 37% Activation 2 mM PP; " 0.093 29% Inhibition 2 mM AMP ' " 0.083 37% " 2 I1“ ADP " 0.107 18% II 2 mM Citrate " 0.098 25% " 2 mM Phosphoenolpyruvate " 0.000 100% " 73 Figure 12. Effect of micromolar concentrations of phosphenolpyru- vate on the activity of ATP-dependent ph05phofructokinase. The assay mixture (170 pl) contained 80 mM HEPES-NaOH buffer (pH 7.4), 1 mM Na fructose-6-P, 0.5 mM ATP, 6.0 mM MgC12, 0.24 mM Na NADH, 0.2 unit of fructose-1,6-P2 aldolase, 2.0 units of triose-P isomerase, 0.2 unit of a-glycerol-P dehydrogenase, and 0.43 m0 of the DEAE-cellulose II enzyme preparation. RELATIVE ACTIVITY 74 IOO 75r- 50~ C) J I I L o 2 4 6 8 IO 12' [PHOSPHOENOLPYRUVATE], mm Figure 12 75 and at -80°C without added glycerol (Figure 13). It was unstable to storage at -20°C in the absence of glycerol (90% decrease in one week). At 4°C in the absence of glycerol, it retained its activity (100%) for about a week and then the activity decreased slowly (51% remaining after 13 weeks). At 4°C in the absence of glycerol, 17 mM PP; did not provide any protection (44% remaining after 13 weeks). The addition of 170 mM fructose-6-P, at 4°C in the absence of glycerol, accelerated the decrease in activity (only 17% left compared to 70% for the control after 6 weeks). The addition of 16 mM DTT also accelerated the decrease in activity (only 14% left compared to 88% for the control after 3 weeks). The storage-stability experiment established two storage condi- tions in which the enzyme retained its activity by 100% for at least 4 months (at -80°C, or at -20°C in the presence of 20% glycerol). In ad- dition, the results suggested a possible stabilizing role of glycerol if it were added in the buffer during the purification of the enzyme, and indicated that DTT is deleterious to the stability of the enzyme. Further Purification of the Enzyme and Elucidation of Additional Properties. The enzyme was purified (670-fold) by a procedure described in the article by Sabularse and Anderson (139) and elaborated on in an article by Anderson and Sabularse (140). This purification was conducted before the effects of glycerol and DTT were fully appreciated. The purified enzyme was stored at -80°C. This enzyme preparation was used to determine the KA for fructose-1,6-P2; the activation effect of fructose-2,6-P2 and its KA; the effect on the Km for fructose-6-P 76 Figure 13. Stability of PP;:F6P l-phOSphotransferase at various conditions of storage. The enzyme preparation purified about lO-fold was from a pooled fraction after DEAE-cellulose chromatography. The enzyme was dialyzed against 10 mM Tris-acetate buffer (pH 7.3) contain- ing 0.1 mM EDTA, 1.0 mM DTT and 17.0 mM KCL. Storage temperature and addition of glycerol, PP;, fructose-6-P, and DTT were as indicated. For the frozen samples, separate vials were used for the determination of each data point. The assay mixture (170 p1) contained 20 mM fructose-6-P, 1.0 mM PP;, 6 mM MgC12, 0.24 mM Na NADH, 0.2 unit of fructose-1,6-P aldolase, 2.0 units of triose-P isomerase, 0.2 unit of a-glycerol- dehydrogenase, 1.5 mM glucose-1,6-P2, and 6.6 pg protein (5 pl, 1.0 m0 of activity) of the enzyme preparation. LOG PERCENT ACTIVITY 77 o O “-80°C A L6 —. A -20°c, PLus 20% 7 . 4°c GLYCEROL o A 4°C, PLUS 17 mM PPI 1.4— o 4°C, PLUS I70mM - o C) ‘¥KLF11K5FH%Um6-P ., -20°c l6mM DTT 1.2 -‘ - 1.0:; I l I l I o 4 6 8 10 12 WEEKS OF STORAGE Figure 13 78 and the relative Vmax by the activators; and as a reagent to determine the natural occurrence of fructose-2,6-P2 in mung beans. A more extensive purification of the enzyme will be detailed in a later section of the thesis. 5A:for fructose 1,6-bisphosphate. The assay employed was a gly- ceraldehyde-B-P dehydrogenase-linked assay. Because fructose-1,6-P2 is a product of the reaction, the accurate determination of its KA required special instrumentation. The activity and the amount of the accunulated fructose-1,6-P2 during the progress of the reaction were determined using a Gilford 2600 microprocessor-controlled spectrophotometer, programmed for the determination of enzyme kinetic parameters in a single assay mixture (151). The resulting graph (Figure 14) showed a hyperbolic response curve from which the KA for fructose-1,6-P2 was determined to be 17 pM. KA for glucose 1,6-bisphosphate. A fructose-1,6-P2 aldolase- triose-P isomerase-a-glycerol-P dehydrogenase-linked assay was used, again employing the Gilford 2600 microprocessor-controlled spectrophotometer. The resulting graph (Figure 15) showed a hyperbolic response curve from which the KA for glucose-1,6-P2 was determined to be 0.4 mM. Activation by fructose 2,6-bisphosphate and determination of its EA: The discovery of fructose-2,6-P2 as an activator of ATP-dependent phosphofructokinase in animal tissues (27, 31, 35) prompted us to test its effect on PP;:F6P 1-phosphotransferase. Nanomolar amounts of fructose-2,6-P2 were sufficient to activate PP;:F6P l-phosphotransferase to the same degree that was effected by pM amounts of fructose-1,6-P2 or mM amounts of glucose-1,6-P2. A 79 Figure 14. Determination of KA of PP;:F6P l-phosphotrans- ferase for fructose 1,6-bisphosphate. The assay employed was glyceral- dehyde 3-P dehydrogenase-3-phosphoglyceric acid kinase-linked assay. The assay mixture (2000 pl) contained 80 mM HEPES-NaOH buffer (pH 7.8), 1 mM EDTA, 10 mM fructose-6-P (P;-free), 1 mM Na PP;, 6.0 mM MgCl , 1.0 mM ADP (vanadiun-free), 0.5 mM glyceraldehyde-B-P, 5 units of egceraldehyde-B-P dehydrogenase, 0.5 mM NAD+ and 3.5 pg protein of the enzyme preparation. The enzyme preparation was used to start the reaction. The instrument used was a Gilford 2600 microprocessor-controlled spectrophotometer programmed for determination of kinetic parameters for enzymatic reaction in a single assay mixture (153). The instrument read and stored the data for instantaneous velocity and total fruc- tose-1,6-P2 formed every 2 sec for a 4-min program. Each data point was an integral of 11 readings resulting in 110 data points for 120 readings. A print out of the 110 data points of the recorded velocity (nmol fructose-1,6-P2 formed per min) and the corresponding amounts of fructose-1,6-P2 formed was obtained. A plot of every seventh data point of the set of velocity versus fructose-1,6-P concentration was made along with the corresponding double reciprocaT plot after subtracting the initial unactivated velocity from each of the velocity values to estimate the KA. The inset is the corresponding double reciprocal plot of the fructose-1,6-P2 concentration and the velocity. 80 0 0 w mu m 0 O 4 2.59.25... $038535 .8... foods 20 30 4O 50 fl-‘RUCTOSE-Ie-e], mM IO Figure 14 81 Figure 15. Determination of K of PP;:F6P l-phosphotransfer- ase for glucose-l,6-bisphosphate. Ihe assay employ fructose-1,6-P2 aldolase-triose-P isomerase-a-glycerol-P dehydrogenase-linked assay. The assay mixture (2000 p1) contained 80 mM HEPES-NaOH buffer (pH 7.8), 1 mM EDTA, 20 mM fructose-6-P, 1.0 mM Na PP;, 6.0 mM MgC12, 0.3 mM NADH, 2.0 units of fructose-1,6-P2 aldolase, 20 units of triose—P isomerase, 2.0 units of a-glycerol-P dehydrogenase, 2.1 pg protein of the enzyme preparation, and glucose-1,6-P injected in increasing amount in the reaction mixture equipped w1th a stirrer. The instrunent used was a Gilford 2600 microprocessor-controlled spectrophotometer programmed for determination of kinetic parameters for enzymatic reaction in a single assay mixture (153). During the de- termination of the activity, the reaction mixture minus glucose-1,6- P2 was read for the initial velocity (unactivated) collected as an average of 120 readings in l min. The glucose-1,6-P2 (25.5 mM) was then injected into the reaction mixture at a flow rate predetermined to reach about 2.5 mM in 8 min. The instrunent read and stored the data for the velocity and the corresponding concentration of the glucose-l, 6-P2 added, every 4 seconds for an 8-min program. Each data point was an integral of 11 readings resulting in 110 data points for 120 readings. A print out of the 110 data points of the recorded velocity (equivalent nmol of NADH oxidized per min) and the concentration of glucose-1,6-P2 was collected. A plot of every 4th point of the set of velocity versus the glucose-1,6-P2 concentration was made along with the double reciprocal plot after subtracting the initial unactivated velocity from each value, to estimate Km. The inset is the corresponding double reciprocal plot of glucose-1,6-P2 concentration and the velocity. VELOCITY, nmol FRUCTOSE-l,6-PZ FORMED/MIN 25 20 IS IO 82 0.16 - 0.12 I/ V .08 0.04 0 2 4 14:3] 05 1.0 L5 2.10 [GLUCOSE-1,6432], mM Figure 15 25 83 plot of reaction velocity versus fructose-2,6-P2 concentration (Figure 16) gave a hyperbolic curve from which the KA for fructose-2,6-P2 was determined to be 50 nM. ;§m_for fructose-6-P and relative Vmaxof PP;;F6P 1-phospho- transferase in the absence and in theypresence of an activator. The hexose bisphosphates (glucose-1,6-P2, fructose-1,6-P2 and fructose- 2,6-P2) activate the enzyme both by decreasing the Km for fructose-6-P and by increasing the Vmax° Figure 17 shows a comparison of the saturation curve for fructose-6-P with no added activator and in the presence of a hexose bisphosphate activator. The corresponding double reciprocal plots (Figure 18) revealed that the enzyme follows hyperbolic kinetics both in the absence and in the presence of an activator. Km values for fructose-6-P, and the relative Vmax values, respectively, in the absence and presence of various activators, were as follows: no hexose bisphosphate, 20 mM, 1.0; glucose-1,6-P2, 5.0 mM, 9.2; fructose-1,6-P2, 0.56 11M, 9.2; and fructose-2,6-P2, 0.12 mM, 12 mM, 15.2. The hexose bisphosphate concentration and the increase in the affinity (decrease in Km) for fructose-6-P relative to that with no added activator, respectively, were as follows: 1.5 mM glucose-1,6-P2, 4-fold; 100 pM fructose-1,6-P2, 36-fold; and 1 pM fructose-2,6-P2, 167-fold. It is significant to note that of the three hexose bisphOSphates, fructose-2,6-P2 is the most potent activator. 84 Figure 16. Determination of KA of PP;:F6P l-phosphotransfer- ase for fructose-2,6-P2. The assay employed was as in the standard assay (Method A) except that varying concentrations of fructose-2, 6-P2 were added. Enzyme protein amounting to 60 ng was used in each assay mixture. The inset is the corresponding double reciprocal plot of the fructose-2,6-P2 concentration and the velocity after subtracting the initial unactivated velocity from each of the velocity values. 85 2.0 VELOCITY, nmol FRUCTOSE'--I,6'-P2 FORMED/MIN: O 00 0.02 0.04 0.06 0.08 01 1/[s] I 111 I ‘ O I l l 200 400 600 800 1000’ 20 [FRUCTOSE-2,6-P2], nM Figure 16 86 Figure 17. Effect of fructose-2,6-P2, fructose-1,6-P2 and glucose-1,6-P2 on the kinetic constants (Km and Vmax) of PP;:F6P l-ph05photransferase. Velocity was expressed as nmol of fructose-1,6-P2 formed per min per 64 ng protein. Other details are given in the text. Double reciprocal plots are shown in Figure 18. For no activator, glucose-1,6-Pz as the activator and fructose-2, 6-P2 as the activator, the activity was determined as in standard assay (Method A) with modifications indicated. For fructose-1,6-P as the activator, the activity was determined as in the alternative assay (Method B) with modifications indicated. 87 I.5 ‘ I2 VELOCITY Q 05 I .O (0 w 013 I 1 I + PLUS IpM r-'PIUCTOSI-:-2,6--P2 PLUS 100 pM PPUCTOSE—I,6—P,_ PLUS 1.5 mM GLUCOSE-1,64; + NO ACT IVATOR + 5 IO I5 ERUCTOSE-G-P], mIVI Figure 17 2O 88 Figure 18. Double reciprocal plots of the saturation curves for fructose-6-P in the absence and presence of various activators (Data from Figure 16). 89 I00 80 60 . 1/v40_ NO ADDED ACT ImTOR Km=20 mM 20 We 0.1 00 0.2 0.4 5 4 3 I/V2 I O PLUS 1.5 mM GLUCOSE-LG-PZ I- F 1 Km=5 mM ' Vm=0.9 O 0.2 0.4 1/fi-‘RU-6—Fj, mlvi' I/[f-‘RU-S-P], mM' PLUS 100 pM FRUCTOSE-l,6-P2 1.0 19.50.56 mM 0.5_ v...=0.9 C’0 1.0 270 1.0 08~ 0.6- 0.4-- 0.2r 0 . . . . . PLUS I 11M F RUCTOSE-2,6-Pz Km=0.|2 mM Vm=l.5 012345 I/lFRU-G-P], mM‘ 1/[FRU-6-P], mMI Figure 18 90 Effect of Fructose 2,6-Bisphosphate on ATP-dependent Phosphofructoki- ygggy Fructose-2,6-P2 had no apparent effect on the partially purified ATP-dependent ph05ph0fructokinase from mung beans, which contrasts markedly with its recently reported potent activation of this enzyme in animal tisues (25, 30, 33). Plots of velocity versus fructose-6-P concentration yielded hyperbolic curves both in the absence and presence of 1.0 pM fructose-2,6-P2 (superimposable curves; Figure 19). In both cases, the Km for fructose-6-P was 0.9 mM and the Vmax was 147 nmol of fructose-1,6-P2 formed per min per milligram of protein. Subsequent to the report of Sabularse and Anderson (139), it was reported by Miernyk and_0ennis (49) that in the case of the ATP-dependent phosphofructokinase from developing castor bean, the plastid ATP-dependent phosphofructokinase isozyme was activated by fructose-2,6-P2 but only at pH 7.0. No activation by fructose-2,6-P2 was observed at pH 8.0 and the cytoplasmic ATP-dependent phosphofructokinase isozyme was not activated either at pH 8.0 or 7.0. In the case of the mung bean ATP-dependent phosphofructokinase, no activation by fructose-2,6-P2 was observed even at pH 7.0. Isolation and Identification of Fructose 2,6-Bisphosphate from Mung B_ea_n_s_. Chromatography of a mung bean extract on an anion-exchange colunn yielded a gradient-elution profile with two peaks in the hexose bis- phosphate region (Figure 20A). The first peak was identified as 91 Figure 19. Substrate saturation curve for mung bean ATP-dependent phosphofructokinase. Two sets of data are shown; in the absence (0) and presence 03) of 1.0 pM fructose-2,6-P2 (identical curves). The 8-fold purified enzyme (10.5 pg of protein) was used in each case. Velocity is expressed as nmol of fructose-1,6-P2 formed per minute per 10.5 pg of protein. VELOCITY 92 I.2 D (D 0.4 ‘ 6 8 IO Ip-FRUCTOSE-S-P], mM Figure 19 93 Figure 20. Chromatographic evidence for the occurrence of fruc- tose 2,6-bisphosphate in mung beans. A, a mung bean extract was prepared and chromatographed on Bio-Rad AG 1-X8 as described in Materials and Methods. B, a synthetic mixture of authentic fructose-1,6-P2 and fruc- tose-2,6-P2 was chromatographed as in A. Fructose-1,6-P (O) was determined with an enzymatic end-point assay initiated by ructose-l,6-P2 aldolase, whereas fructose-2, 6-P (O) was measured by its ability to activate PP;:F6P l-pfiosphotransferase (see Materials and Methods). 94 3 2.25538 90-08530-.. .8: 5.5.54 nu nu nu nv nu II A 10.. _ n/ -...\/./ 5 0 Ma m _ _ d _ j . B .14” _ _ _ m m w .. . I 64 72 80 I 48 I 56 FRACTION NUMBER ’52 40 LO;— 0.5— O 5 O. 5. 2 2 I .o. m.-w._-m.moeo:mn.a .25 _ m . 5. O 0 Figure 20 95 fructose-1,6-P2 by its reaction in a coupled enzymatic assay initiated by fructose-1,6-P2 aldolase. The second peak was located by its ability to activate PP;:F6P 1-phosphotransferase and was identified as fructose-2,6-bisphosphate as described below. Anion exchange chromatography of an authentic mixture of fructose 1,6-Pz and fructose-2,6-P2 (Figure 208) yielded an elution profile similar to that observed with the mung bean extract (Figure 20A). The unknown activator in fraction 56 (Figure 20A) also behaved like authen- tic fructose-2,6-P2 with respect to its stability in 0.25 N NaOH at 90°C and its room temperature lability at pH 2.5 (Table 6), thus locat- ing one of the ph05phate groups at carbon 2, assuming a ketohexose bis- phosphate (25, 31). The unknown activator was further identified as fructose-2,6-P2 following a scaled-up procedure in which fructose-l,6-bisphosphatase treatment and a second anion-exchange chromatography step were used to separate the activator from fructose-1,6-P2. The activator was further concentrated by lyophilization, subjected to mild acid hydrolysis (pH 2.5 at room temperature for 30 minutes), and the hydrolysis product was identified as fructose-6-P by specific coupled enzyme assays employing (i) 0-fructose-6-phosphate isomerase (EC 5.3.1.9) and D-glucose-G-phosphate dehydrogenase (EC 1.1.1.49) and (ii) phosphofructokinase (EC 2.7.1.11), D-fructose-l,6-bisphosphate aldolase, triose-P isomerase, and a-glycerol-P dehydrogenase. 96 Table 6. Comparison of the Mung Bean Natural Activator and Authentic Fructose 2,6-Bisphosphate with Respect to Acid- and Base- stability. Enzyme activitya in the presence of: Treatment Fructose-2,6-P2 (1 pM) Natural activator None 100 100 Acid hydrolysisb 0 0 Base hydrolysisc 100 100 aActivity is expressed as percent of the velocity observed in the standard assay using 3.1 milliunits of PP;:F6P l-phosphotrans- ferase and the modifications indicated. pr 2.5 (adjusted with HCl) at 24°C for 30 min., then neutral ized with NaOH. cNaOH (0.25 N) at 90°C for 30 min., then neutralized with HCl. 97 Comparison of the Three Methods of Protein Determination. The pooled fractions from various steps of purification of PP;:F6P l-phosphotransferase were assayed for protein using the method of Whitaker and Granum (134), Lowry et al. (136), and Bradford (137). The result showed a close agreement on the protein values (Table 7). The Whitaker and Granunlnethod, a new procedure that was reported in 1980, has the advantage of being non-destructive, quick, and economical. However, the method is only 40% as sensitive as those of Lowry et a1. and of Bradford. 0n the basis of these considerations the Whitaker and Granuninethod was adopted as the protein assay for subsequent protein determinations during the purification steps. Towards the later stages of purification, when the protein becomes very dilute, the microassay of Bradford may be used. Final Purification and Partial Separation of Two Forms of PP;;F6P 1-Phosphotransferase and Characterization of Their Properties. Final purification. Because DTT accelerated the time-dependent decrease in the activity of the enzyme upon storage at 4°C and glycerol stabilized the activity of the enzyme on storage at -20°C, the following changes from earlier purifications were made: DDT was not added to the buffers starting with the grinding buffer, and 10% glycerol was added to the buffer after the ammonium sulfate precipitation step. The entire purification procedure is described in detail in the Materials and Methods section. The elution profiles of the enzyme and protein during the chromatography steps are shown as follows: Figure 21, DEAE-cellulose; Figure 22, phosphocellulose I; Figure 23, phosphocellulose II; and Figure 24, Bio-Gel A-1.5m. The 98 .Nmic.Piwmou=pm we: Loam>_uue we» can 25 cm we: mioimmoposem on» use» uamuxm Axumme usewceum .copuumme accrues. < uozumz ma toxemme we: mmecmemcmeuogamosai_ mounpaa mo auw>_pu_au< amam copuuuwaweam oozed: .:o_uecpaemumo :_muoee so mcoguoz moss» mo compgagEou .N m—nep 99 Figure 21. DEAE-cellulose chromatography of PP;:F6P l-phosphotransferase. The protein was eluted by a 11near gradient of KCl in the eluting buffer. All other details are given in the text. 100 a. .535 2ng a m . E _ . _ , T. m 020 CT 1 w // .0/ / 21m.. / / O T m2 R, / De... 1 1 0 / ..mme f // N x N / T1 // C A _ _ 0m 0 051- m. w m. 3 .Ebtz: 5.2.6.5 25 Figure 21 101 Figure 22. Phosphocellulose I Chromatography of PP;:F6P l-phos- photransferase. Na4P207 (34 mM) was included in the eluting buffer. All other details are given in the text. L75 1.50 L25 ‘5 0 ACTIVITY, UNITS/ml (9) a) 102 _ ; -6 - «5 i — -4 b J3 - -2 — -1 0 20 40 Ola I . - 60 80 I00 FRACTION NUMBER, 22 ml/ TUBE Figure 22 PROTEIN, mg/ml (I) 103 Figure 23. Phosphocellulose II Chromatography of PP;:F6P l-ph05photransferase. A linear gradient of KCl was incorporated in the eluting buffer. All other details are given in the test. 104 h) I ACTIVITY, UNITS/ml (C) I 20 4O 60 80 FRACTION NUMBER, Figure 23 T \ g \ PROTEIN, mg/ml (I) .I. ,9 mM KCI (”"1 ‘inta o () IOO I20 IO ml/ TUBE 105 Figure 24. Bio-Gel A-l.5m chromatography of PP;:F6P l-phospho- transferase. All other details are given in the text. 106 1.61- ACTIVITY, UNITS/ml I.) ‘ 1 T T * 1 1 2.01— -62.5 I.2~ . \ - 50.0 37.5 10 (fl '0 PROTEIN, pg/ml ('1 12.5 I i 00 60 81) 100 120 140 FRACTION NUMBER, 5 ml/ TUBE Figure 24 0 107 enzyme in the pooled fractions from Bio~Gel A-l.5m step was l030-fold purified with a 35% recovery. A summary of the purification is shown in Table 8. Separation of two forms of the enzyme. As can be seen in Figure 24 or Figure 25a, the enzyme gave two peaks during the Bio-Gel A-l.5m step, indicating that there are two forms of the enzyme, a large form and a small form. Because the two peaks overlapped, the possibility of a dynamic equilibrium between the large and the small form existed, aside from the possibility that complete separation of the two forms may have been beyond the resolving capacity of the column. To determine if a dynamic equilibrium existed (151a), the pooled fraction from the first Bio-Gel A-l.5m step was rechromatographed on the same column at higher salt concentration; this also resulted in two peaks (Figure 25b). The fractions corresponding to peak I (Figure 25b, fractions 80-102) were pooled and rechromatographed on the same Bio-Gel A-l.5m column. Likewise, fractions corresponding to peak II (Figure 25b, fractions 103-l25) were also pooled and rechromatographed separately on the same Bio-Gel A-l.5m column. The results are shown in Figure 25c. Peak I did not generate peak 11, and peak 11 did not generate peak I, indicating that, indeed, they represent distinct forms of the enzmne which do not interconvert in an equilibrium situation. The previous purifications gave only one peak during the Bio-Gel A-l.5m step, and corresponded only to the small form. The main difference was that the earlier preparations had DTT included in the buffer and no glycerol was added. Since these results indicated that the enzyme exists in two forms (large and small), further experiments were conducted to investigate their properties. 108 Table 8. Purification of PPizFGP l-Phosphotransferase from 500 g of mung beans. Total Total Specific Fraction Volume proteina activity activity Recovery (units/mg (mg) (mg) (unitsb) protein) (%) l. Crude extract 2030 26800 1020 0.038 (lOO) 2. (NH4)ZSO4 ppt. l46 8000 ll40 0.l43 lll 3. DEAE-cellulose 77.0 1330 774 0.582 76 4. Phosphocellulose I 77.0 20l 543 2.70 53 5. Phosphocellulose II l0.0 2l.0 429 20.4 42 6. Bio-Gel A-l.5m l0.0 9.20 359 39.0 35 aProtein was assayed by the procedure of Whitaker and Granum. bOne unit of activity is equivalent to l pmol per min of fructose l,6-bisphosphate produced in the standard assay. 109 Figure 25. Separation of the two forms of PP :F6P l-phospho- transferase by chromatography through Bio-Gel A-l. m. The pooled frac- tion after phosphocellulose II chromatography (Figure 24) was chromato- graphed through a Bio-Gel A-1.5m column (3.4 x 95 cm) with a linear flow rate of 9.4 cm per hr. (a) In 10 mM Tris-acetate, pH 7.3 contain- ing 10% glycerol, 0.1 mM Na EDTA and 20 mM KCl. (b) Rechromatography of the concentrated pooled fractions (80-125) from (a), with the K01 concentration in the buffer increased to 100 mM. (c) Rechromatography of the pooled fractions from (b) corresponding to peak I (fractions 80-102), and peak II (fractions 103-125), both in the same buffer as in run (b). Peaks I and II represent the large and small forms of the enzyme, respectively. All other details are given in the text. ACTIVITY, UNITS/ML 110 N O '5 O 2.0 ~ C A I I LO 3 - O I . 50 75 I00 l25 150 FRACTION NUMBER, 5 MJTUBE Figure 25 lll Some properties of the two forms. The enzyme preparations used for the determination of properties a_through.h below were from the third chromatographic run through Bio-Gel A-l.5m (Figure 25c), fraction number 90 (0.7 unit/ml, ll30-fold pure) representing the large form and fraction nunber 117 (0.7 unit/ml, 842-fold pure) representing the small form. The protein in fractions 90 and 117 was determined by the Bradford method (137). a) Heat stability at 55°C. The small form was found to have a longer half-1ife (tkf 26 min) than the large form (t8? 14 min) as shown in Figure 26, indicating that the large form was more heat labile. b) Effect of pH and buffer composition. The activity-pH response of both the large form (Figure 27) and the small form (Figure 28) were basically the same, having an optimum pH of 7.6-7.8 for both, whether the buffer used was Na-HEPES or Na-PIPES/Tris combination buffer. How- ever, it may be noted that at pH 7.0-7.2 Na-PIPES buffer gave a higher activity that Na-HEPES for both forms. At pH 7.6-7.8 the Na-PIPES/ Tris combination buffer was also slightly better than Na-HEPES buffer for both forms. Because the increase in the activity at pH 7.8 by using Na-PIPES/Tris combination buffer was only about 5% over the activity using Na-HEPES which was previously adopted as the buffer for the standard assay, no change in the buffer composition for the standard assay was made at this point. However, such information may be considered for future work on the enzyme. c) EA for fructose 2,6-bisphosphate at pH 7.8 and pH 7.0. Figure 29 shows that the large form had a higher affinity for fructose-2,6-P2 (KA = 25 nM, both at pH 7.8 and 7.0) than the small 112 Figure 26. Relative stability of the small and large forms of PP1:F6P l-phosphotransferase at 55°C. Two hundred microliters of each of the enzymes (large form, 0.70 unit/ml with a specific activity of 43 units/mg; and small form, 0.70 unit/ml with a specific activity of 32 units/mg) in 10 mM Tris-acetate buffer (pH 7.3, 100 mM KCl, 10% glycerol and 0.1 mM EDTA) was incubated in a water bath at 55°C. At each incubation period indicated, a 20 pl sample was transferred to a 1-ml tube, cooled in an ice bath for 1 min and 5 ul sample was assayed for its activity by standard assay (Method A). LOG OF PERCENT ACTIVITY 113 1 r 1 1 1 1 T r 2.0 ‘ A SMALL FORM H (DLJVKEIFORNI 0 L8- - 1V2=26 MIN C 1 44 M11. 1.6? V2 -< O 1.4- - 1.2; -* £1 1 1 1 1 1 1 1 O 5 IO 15 20 25 30 35 MIN AT 55°C Figure 26 114 Figure 27. Effect of pH and buffer composition on the activity of the large form of PP1:F6P l-phosphotransferase. The reaction mixture (170 pl) contained 1.0 mM PPi, 10 mM fructose-6-P, 1.0 mM Na EDTA, 6.0 mM MgCl , 0.24 mM Na NADH, 1.0 uM fructose-2,6-P2, 0.2 unit of fructose-l, -P2 aldolase, 2.0 units of triose-P isomerase, 0.2 unit of a—glycerol-P dehydrogenase and 5 u1 (2.5 mU at standard assay) of PP1:F6P l-phosphotransferase. The buffer concentration was 80 mM in each case; when PIPES and Tris were used in combination, the concentra- tions were 40 mM of each. 115 1 1 T 100- O O E 75—- 2 '1— . 2 . 1.1.1 2 '2: 5o- _J E A No-MES A No-PIPES I No-PIPES/TRIS 25- O TRIS-CHLORIDE - O No-HEPES J? L L at 7 a pH Figure 27. 116 Figure 28. Effect of pH and buffer composition on the activity of the small form of PP1:F6P l-phosphotransferase. The reaction mixture (170 u1) contained 1.0 mM PPi, 10 mM fructose-6-P, 1.0 mM Na EDTA, 6.0 mM MgC12, 0.24 mM Na NADH, 1.0 uM fructose-2,6-P2, 0.2 unit of fructose-1,6-P aldolase, 2.0 units of triose-P isomerase, 0.2 unit of a-glycerol- dehydrogenase and 5 u1 (2.5 mU at standard assay) of PP1:F6P l—phosphotransferase. The buffer concentration was 80 mM in each case; when PIPES and Tris were used in combination, the concentra- tions were 40 mM of each. T T l 100' .4 >_ 75 - - t: Z i.- o \_ 1 m o .28... 8%.. a SEQ“. 1.32m < l N._ 127 Figure 33. Comparison of the large and small forms of PPizF6P 1-ph05photransferase with respect to the Km for P, at pH 7.8. The activity was determined by Method C assay, described in Materials and Methods. The velocity is in nmol fructose-1,6-P2 consumed per min by 10 pl of the enzyme preparation in each case. 128 A SMALL FORM 1.5 OLARGEFORM 1.2 I/V Km=0.28mM 0.3 1 w 1 1 1 A 1 1 1 1 1 ‘5‘4'3'2'1 O 1 2 345 I/[PJ,mM" Figure 33 129 Figure 34. Effect of Pi on the activity of the two forms of PP1:F6P l-phosphotransferase in the absence and presence of two con- centrations of fructose-2,6-P . The activity was determined as in the standard assay, Method A forward reaction), with modifications indicated. RELATIVE ACTIVITY 80 60 4o 20 100 80 6O 20 130 L. 1 1 1 LARGE FORM 0 N0 PRU-2,6-P2 - zoo nM 1=Ru-2,6-R2 .1 1000 nM PRU-2,6432 1 J 1 fi— 1 1 1 SMALL FORM 0 N0 PRU-2.64:, I zoo nM FRU-2,6-F§ A1000 nM PRU-2,643 l l I 0.5 1.0 15 [P1]. mM Figure 34‘ 2.0 131 the large and small forms of the enzyme were severely inhibited by P1 (large form, 50% inhibition at 0.78 mM P1; and small form, 50% inhi- bition at 0.70 mM P1). In the presence of 200 nM fructose-2,6-P2, the degree of inhibition by Pi was less severe for both the large and the small forms (large form, 50% inhibition at 1.90 mM P1; and small form, 50% inhibition at 1.95 mM P1). At saturating amounts of fruc- tose-2,6-P2 (1000 nM), the P1 concentration that was necessary to inhibit by 50% was greater than 2 mM for both the large and the small forms of the enzyme. 1) Molecular weight by gel-filtration on Bio-Gel A-l.5m. Determination of the molecular weights of the two forms of the enzyme by column chromatography yielded values of 170,000 for the small form and 340,000 for the large form (Figures 35 and 36). Apparent conversion of the large form to the small form upon incubation with DTT. During previous purification procedures, when DTT was included as one of the components of the buffers, only the small form of the enzyme was detected, suggesting that DTT may be responsible for the conversion of the large form to the small form. In addition, the two forms of the enzyme had a large difference in their KA values for fructose-2,6-P2 at pH 7.0 (large form, KA = 25 nM; and small form, KA a 140 nM). From these observations, it should be possible to obtain a presumptive evidence for the transformation of the large form to the small form by monitoring the KA after incubation with DTT. Figure 37 shows an apparent DTT-dependent change in KA for fructose-2,6-P2 as reflected by a change in the activation ratio of the large form to that of the small form, suggesting a concomitant change from the large form to the small form. The "activation ratio" 132 Figure 35. Elution profile of molecular weight standards and the two forms of PP1:F6P l-phosphotransferase chromatographed on Bio-Gel A-l.5m. A 5.6—ml sample containing 40 units of rabbit muscle pyruvate kinase, 20 units of rabbit muscle aldolase, 40 units of yeast hexoki- nase and about 10 units each of the two forms of PP-:F6P l-phospho- transferase were chromatographed using a column (3.1 x 94.5 cm) of Bio- Gel A-l.5m equilibrated with a buffer (10 mM Tris-acetate, pH 7.3, con- taining 100 mM KCl, 10% glycerol and 0.1 mM Na EDTA). Elution of the proteins was done using the same buffer, collecting 5.0-m1 fractions at a flow rate of 80 ml per hr. A separate run, on the same colunn, of 15 mg horse spleen ferritin and 40 units of yeast hexokinase was conducted to determine the elution profile of ferritin, the concentration of which was determined by its absorbance at 280 nm. The yeast hexokinase served a a countercheck for any possible variation of the separate run. Enzymatic assays are described in the text. Symbols: 0 horse spleen ferritin, O PP1:F6P l-phosphotransferase, A rabbit muscle pyruvate kinase, .rabbit muscle aldolase, and deast hexokinase. 20 133 ACTIVITY (ARBITRARY UNITS) ('5 G I I 01 1 01% 1 90 / l l 1 100 110 120 130 FRACTION NUMBER Figure 35 2.0 1.5 A 280 (D) 0.5 140 134 Figure 36. Semi-log plot of the molecular weight of the standard proteins (horse spleen ferritin, Mr = 450,000; rabbit muscle pyruvate kinase, Mr = 237,000; rabbit muscle aldolase, Mr = 160,000, and yeast hexokinase, Mr = 102,000) versus the peak fraction number from the elution profile on the Bio-Gel A-l.5m column (Figure 35). The mo- lecular weights of the large and the small forms of PP;:F6P l-phos- photransferase were interpolated from their peak fractions. LOG MOLECULAR WEIGHT 5.6 5.4 52 5.0 5001,0001 I I I I I I '400,000 ‘ LARGE FORM (3.4 x 105) -300,000 - PYRUVATE - KINASE “200,000 SMALL FORM - ALDOLASE ‘ (1.711105); HOO’QOO HEXOKINASE _ I l I l I l I l 95 100105 110 115 120 125 I30 FRACTION NUMBER Figure 36 136 Figure 37. Apparent DTT-dependent change in KA for fructose-2, 6-P2 for the large form of PP;:F6P l-phosphotransferase, suggesting a concomitant change to the small form of the enzyme. Aliquots of pooled fractions representing the large and small forms of the enzyme were incubated in 15 mM DTT. The activity was determined as in the standard assay (at 0, 35 and 1000 nM fructose-2,6-P2) after timed periods of incubation. In each assay, 2.0 p1 (original activity of 2.5 mU) of the large and small forms were used giving a final concentra- tion of 0.18 mM DTT in the assay mixture. The "activation ratio“ is the fold activation when assayed in the absence and presence of 35 nM fructose-2,6-P2 divided by the fold activation when assayed in the absence and presence of 1000 nM fructose-2,6-P2, and therefore is a reflection of KA. ACTIVATION RATIO AT 35 nM TO 1000 nM FRU-2,6-P2 ,O 03 Q 01 Q .rs .O 01 Q N 9. O 137 A SMALL FORM 0 LARGE FORM if 42‘. l 1 1 l 1 L 2 4 e a 10 12 INCUBATION TIME, HOURS Figure 37 138 is the fold activation when assayed in the absence and presence of 35 nM fructose-2,6-P2 divided by the fold activation when assayed in the absence and presence of 1000 nM fructose-2,6-P2, and therefore is a reflection of KA. At zero time of incubation, the activation ratio for the large and small forms were 0.46 and 0.23, respectively, which were consistent with what is predicted by a low KA for the large form and higher KA for the small form. Upon incubation with DTT, from 1 hour on, the activation ratio for both the large and small forms were essentially equal, about 0.21, suggesting that the large form may have been transformed to the small form by the action of DTT. The 0.21 activation ratio was about equal to the theoretical value of the activation ratio that would be predicted by Michaelis-Menten equation for the small form as follows: (V at 35 nM) - (V at zero nM) Activation ratio (V at 1000 nM) - (V at zero nM) Vm ($1)/[KA + (51)] Vm (521/[KA + (52)] 0.228 (at Vmax = 1, KA = 140 nM, 51 = 35 nM and $2 = 1000 nM) Survey of PP;:F6P l-Phosphotransferase and Fructose 2,6-Bisphosphate in Some Planthpecies. A survey of the occurrence of the enzyme in plants indicated that the enzyme is widespread among plant species (Table 9). The enzyme could also be found in various plant parts: seeds, leaves, bulb, roots, stalks and fruits. 139 Table 9. Occurrence of PP;:F6P 1-Phosphotransferase in Some Plant Species SPECIES PART SPECIFIC ACTIVITYa (nil/"19 protein)b MUNG BEANS Phaseolus aureus GERMINATED SEED 36.0 SOYBEAN, Glycine max GERMINATED SEED 9.1 PEAS, Pisgm‘gatixa GERMINATED SEED 5.2 SPINACH, Spinacia oleracea LEAVES 7.3 CORN, g m GERMINATED SEED 8.4 SCALLION, (GREEN ONIONS) BULB 30.0 Allium schoenOprasum ROOTS 1.5 LEAVES 4.9 CARROT, Daugus‘gargta ROOTS 0.3 ONION, m c_ega. BULBS 4.7 CELERY, Apium graveolens STALKS 6.5 LEAVES 1.4 GREEN PEPPER, Capsicum aggggm FRUIT 13.0 NALNUTS, Juglans 5p. SEED 22.0 BANANA, Mug; acuminata RIPE FRUIT 0.1 I DUCKHEED, Lgmna minor, NHOLE PLANT 8.2 aPPi:F6P 1-Phosphotransferase activity was assayed by method A (forward reaction, standard assay). ( )bProtein was determned by the method of Whitaker and Granun 134 . ‘ 140 Fructose-2,6-P2 was also found in several plant species (Table 10). It is present in seeds, leaves, bulb, roots and fruits. 141 Table 10. Occurrence of Fructose 2,6-Bisphosphate in Some Plant Spe- cies. PP;:F6P 1-Phosphotransferase was used as a reagent for determining the presence of fructose-2,6-P2. The assay was basically Method A (forward reaction, standard assay) except for the omission of fructose-2,6-P2. The reSponse of the enzyme to activation by column fractions of extracts was the basis for determining the presence of fructose-2,6-P2. A SPECIES PART RELATIVE AMOUNTS MUNG BEANS Phaseolus aureus GERMINATED SEED ++ DRY SEED + PEAS, Pisum sativa GERMINATED SEED ++ ONION, Allium cepa BULB ++ SCALLION, (GREEN ONIONS) BULB + Allium schoenOprasum ROOTS ++ LEAVES ++ CARROT, Daucus carota LEAVES ++ BANANA, Musa acuminata RIPE FRUIT ++ SPINACH, Spinacia oleracea LEAVES ++ ++ Significant amount + Trace amount DISCUSSION The detection of PP;:F6P l-phosphotransferase activity in mung beans establishes the presence of the enzyme in C3 plants. Prior to the report by Sabularse and Anderson (138), the enzyme had only been found in a few microbes and in leaves of pineapple, a crassulacean plant (21). In addition, no regulatory mechanism for the enzyme from any source, namely its activation by hexose bisphosphates, has been previously reported. The activation of the enzyme by fructose-l,6-Pz (a product of the reaction it catalyzes) was established indirectly by a series of experiments, and directly by following the rate of Pi formation using a glyceraldehyde-3-P dehydrogenase-coupled assay wherein the response of the enzyme to activation by the accumulated fructose-l,6-P2 during the reaction was demonstrated. It was also found that millimolar amounts of glucose-l,6-P2 could substitute for micromolar amounts of fructose-l,6-P2 to the same degree of activation. For a time, the aldolaSe-coupled assay with glucose-1,6-P2 as an activator was conveniently employed to assay for the enzyme even in the crude extract. Fructose-1,642 and glucose-1,642 are known activators of ATP-dependent phosphofructokinase from animal sources and some microorganisms (148, 151b, 151c). During the progress of this thesis research, however, fructose-2,6-P2 was reported to be a naturally 142 143 occurring activator for ATP-dependent phosphofructokinase in animal tissues (see Review of Literature section). when fructose-2,6-P2 was tried as a possible activator for PP;:F6P l-phosphotransferase, it was found that nanomolar amounts (KA a 50 nM) could replace micro- molar amounts of fructose-l,6-P2 (KA = 17 pM) or millimolar amounts of glucose-l,6-P2 (KA = 0.4 mM) to give the same or even greater degree of activation. Fructose-2,6-P2 was also demonstrated in this investigation to be a naturally occurring compound in mung bean (139) and other plants, in addition to being the most potent activator of PP;:F6P l-phosphotransferase, suggesting that it is the physiolog- ically significant activator for the enzyme. Because the enzyme activ- ity is inhibited by P1 even in the presence of fructose-2,6-P2, the Vmax for the reverse reaction is lower than that of the forward reaction, favoring a glycolytic role for the enzyme. In g, histolytica and P, shermanii, the PP1:F6P l-phosphotransferase appeared to play a major role in glycolytic pathway (l7, 18). However, when P. shermanii was grown on lactate or glycerol, the activity of the PPi-dependent phosphofructokinase was 15-20 times that of the fructose bisphospha- tase, which led to the speculation that the enzyme may function in a gluconeogenic as well as glycolytic capacity (18). 'E. histolytica apparently contains no fructose bisphosphatase (2), so that the enzyme in this amoeba may have both a glycolytic and a gluconeogenic role. The enzyme in microorganisms apparently does not require an activator. PP; is generated in many biochemical processes (see Introduction). In germinating starchy seeds, such as mung bean, a major source of PP; would be the conversion of starch to sucrose: 144 one mole of PP; is formed by UDP-glucose pyrophosphorylase for each mole of sucrose synthesized. Because sucrose is the main transport form of sugars (114), considerable PP; would be generated in sucrose synthesis alone. In addition, inorganic pyrophosphatase activity in germinating mung bean seeds is undetectable (Table 4), suggesting that mung bean seeds grown in the dark would contain levels of PP; sufficiently high to serve as a phosphoryl donor. In support of this position, Sinmons and Butler (146) and Bucke (147) reported that the intracellular localization of inorganic pyrophosphatase in the leaves of higher plants is in the chloroplast. Furthermore, Simmons and Butler (146) reported that mature seeds of maize have no pyrophOSphatase. The observed effect of P;, AMP, citrate and ADP on the activity of the ATP-dependent phosphofructokinase in mung bean is in agreement with what is generally reported for the enzyme in other plants (148). ADP and AMP are both positive effectors for mammalian ATP-dependent phosphofructokinase, but are negative effectors for the plant enzyme (148). The potent inhibitory effects of phosphoenolpyruvate observed for the mung bean ATP-dependent phosphofructokinase has also been observed for the pea seed enzyme by Kelly and Turner (150). It is interesting to note that P;, which activates the ATP—de- pendent phosphofructokinase, inhibits PP;:F6P l-phosphotransferase. Likewise, phosphoenolpyruvate, which inhibits the ATP-dependent phosphofructokinase at micromolar levels, does not affect the activity of PP;:F6P l-phosphotransferase even at 100 times the concentration that inhibits the ATP-dependent phosphofructokinase. ATP at 1 mM is non-inhibitory for the ATP-dependent phosphofructokinase but is 145 slightly inhibitory to PP;:F6P l-phosphotransferase. Conversely, 2 mM PP;, which is not inhibitory to PP;:F6P l-phosphotransferase, is inhibitory for ATP-dependent phosphofructokinase. ADP, AMP, and citrate inhibit the ATP-dependent phosphofructokinase but do not affect PP;:F6P l-phosphotransferase activity. Furthermore, PP;:F6P l-phosphotransferase is activated by nanomolar amounts of fructose-2, 6-P2, whereas no effect is observed for the ATP-dependent phospho- fructokinase even at substrate concentrations that are below the Km. PP;:F6P l-phosphotransferase exhibited Michaelis-Menten kinetics both in the absence and presence of an activator. Fructose-2,6-Pz (1 pM) increased the Vmax lS-fold and also increased the affinity (decreased the Km) for fructose-6-P 167-fold. From the Michaelis-Menten equation these two effects can be seen to combine to give more than a thousand-fold increase in reaction velocity at 0.12 mM fructose-6-P, which is the Km of the activated form of the enzyme at optimum pH. In contrast, fructose-2,6-P2 increases the affinity of mammalian ATP-dependent phosphofructokinase for fructose-G-P, but does not affect the Vmax (31, 35, 40). From the above considerations, it is conceivable that even if the two enzymes (ATP-dependent and PP;-dependent) were present in the same intracellular location, the attendant conditions would determine which of the two enzymes should have the active role in the conversion of fructose-6-P to fructose-l,6-P2. The role of the PP;:F6P l-phosphotransferase could be particularly important during the early stages of germination; during these stages, a process which conserves ATP may be advantageous, since the system is still at the photo- synthetic-independent stage. 146 Two forms of PP;:F6P l-phosphotransferase are present in mung beans - the large and small forms. They differ with respect to their molecular weight (large form, Mr 8 340,000 and small form Mr = 170,000) and are much larger than the enzymes from either P. shermanii (Mr = 95,000; ref. 18) or E. histolytica (Mr = 83,000; ref. 23). They also differ in their stability at 55°C (large form has t3 = 14 min whereas small form has t8 = 26 min); KA for fructose-2,6-P2 (the large form has KA = 25 nM at pH 7.8 and pH 7.0 whereas the small form has KA = 50 nM at pH 7.8 and KA = 140 at pH 7.0); and Km for fructose-6-P (the large form has Km = 0.15 mM whereas the small form has Km = 0.50 mM at pH 7.0). Some kinetic properties are the same: Km for PP;, Km for fructose-6-P at pH 7.8, Km for fructOse-l,6-P2, Km for P;, and the inhibition by P; both in the presence and absence of fructose-2,6-P2. A comparison of the kinetic properties of the two forms of the enzyme suggests that the large form may be the more physiologically significant form. I vitro, a presumptive evidence for the possible conversion of the large form to the small form by the action of DTT was demonstrated. DTT was also found to be deleterious to the stability of the enzyme. Light and DTT treatment had been shown to inactivate glucose-6-P dehydrogenase (152) and ATP-dependent phosphofructokinase (153) in peas, and to activate a nunber of enzymes of photosynthetic metabolism (154-159). It is therefore probable that the action of DTT on PP;:F6P l-phosphotransferase simulates a possible in £119 reductive process mediated by light. A possible inactivation of PP;:F6P l-phosphotransferase by light would be consistent with 147 preventing the breakdown of hexose-phosphates and the storage of carbohydrates during photosynthesis. The detection of the two forms of PP;:F6P l-phOsphotransferase, which are both catalytically active, has never been shown for the enzyme from other sources. It merits future studies by purifying the two forms to homogeneity and further characterizing their properties. In addition, the formation of fructose-2,6-P2 and the mechanism of control of its levels should also be studied. In animal tissue the levels of fructose-2,6-Pz is under hormonal control (66a, 66b). It- is tempting to speculate that a hormonal mechanism of control may also occur in plants. A survey of the occurrence of PP;:F6P l-phosphotransferase in plants indicates that the enzyme is widespread among plant species. The enzyme could be found in various plant parts: seed, leaves, bulb roots, stalks and fruits. Likewise, the activator fructose-2,6-P2 is also widespread among plant species. It is present in seeds, leaves, bulb, roots and fruits. The widespread occurrence of both the PP;:F6P 1-phosphotransferase and the activator fructose-2,6-Pz supports the idea (138, 139) that PP;:F6P l-phosphotransferase is instrunental in energy metabolism in plants. 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. LITERATURE CITED Reeves, R.E. (1976) Trends Biochem. Sci. 1) 53-55. Wood, H.G., O'Brien, W.E., and Michaels, G. (1977) Adv. Enzymol. ,1§, 85-155. 5111, POM-Lo, and "00d, H06. (1962) J. 81010 C116“. £31, 3044‘30510 Nordlie, R.C. (1974) Curr. 19p. Cell. Regul. 8, 33-117. Nordlie, R.C., Arion, W.J., and Glende, Jr., E.A. (1965) J. Biol. Chem. gig, 3479-3484. Hatch, M.D., and Slack, C.R. (1968) Biochem. J. 196, 141-146. Evans, H.J., and Wood, H.G. (1968) Proc. Nat. Acad. Sci. (USA) 61, 1448-1453. ' Reeves, R.E. (1968) J. Biol. Chem. 243, 3202-3204. Buchanan, 8.8. (1974) J. Bact. 112, 1066-1068. Cagen, L.M., and Friedmann, H.C. (1968) Biochem. BiOphys. Res. Conmun. 111, 528-533. Reeves, R.E., and Guthrie, J.D. (1975) Biochem. Biophys. Res. Commun. gg, 1389-1395. Baltscheffsky, H., van Stedingk, L.-V., Heldt, H.-W., and Klingenbery, M. (1966) Science 151, 1120-1121. Baltscheffsky, M (1967) Biochem. Biophys. Res. Commun. 18; 270-276. Keister, D.L., and Yike, N.J. (1967) Biochem. g, 3847-3857. Baltscheffsky, M. (1969) Arch. Biochem. Biophys. 119, 646-652. Keister, D.L., and Minton, N.J. (1971) Biochem. Biophys. Res. Conmun. 4_2_, 932-939. Reeves, R.E., South, D.J., Blytt, H.J., and Warren, L.G. (1974) J.Biol. Chem. 119, 7737-7741. - 148 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 149 O'Brien, N.E., Bowien, S., and Wood, H.G. (1975) J. Biol Chem. 150, 8690-8695. Macy, J.M., Ljungdahl, L.G., and Gottschalk, G. (1978) 1, Bacteriol. 111, 84-91. Sawyer, M.H., Baunann, P., and Baumann, L. (1977) Arch. Microbiol. 11;, 169-172. Carnal, N.W., and Black, C.C. (1979) Biochem. Biophys. Res. Commun. 86, 20-26. Gitomer, W.L., and Veech, R.L. (1982) Abstract 3569 Fed. Proc. 11, 880. Reeves, R.E., Serrano, R., and South, D.J. (1976) J. Biol. Chem. g§1, 2958-2962. Van Schaftingen, E., Hue, L., and Hers, H.-G. (1980) Biochem. J. 1_92_, 887-89 50 Van Schaftingen, E., Hue, L., and Hers, H.-G. (1980) Biochem. J. 32’ 897-90] 0 Van Schaftingen, E., and Hers, H.-G. (1980) Biochem. Biophys. Res. Commun. 96, 1524-1531. Van Schaftingen, E., and Hers, H.-G. (1981) Eur. J. Biochem. 117, 319-323. Hesbain-Frisque, A.-M., Van Schaftingen, E., and Hers, H.-G. (1981) Eur. J. Biochem. 111, 325-327. Claus, T.H., Schlumpf, J.R., El-Maghrabi, M.R., Pilkis, J., and Pilkis, S.J. (1980) Proc. Natl. Acad. Sci. (USA) 11, 6501-6505. Claus, T.H., Schlumf, J. Pilkis, J., Johnson, R.A., and Pilkis, Pilkis, S.J., El-Maghrabi, M.R., Pilkis, J., Claus, T.H., and Cunning, D.A. (1981) J. Biol. Chem. _2_5_§_, 3171-3174. Furuya, E., and Uyeda, K. (1980) Proc. Natl. Acad. Sci. (USA) 11, 5861-5864. Furuya, E., and Uyeda, K. (1980) J. Biol. Chem. ggg, 11656-11659. Richards, 0.5., and Uyeda, K. (1980) Biochem. Biophys. Res. Commun. 21, 1535-1540. Uyeda, K., Furuya, E., and Luby, L.J. (1981) J. Biol. Chem. _2_5_6_, 8394-8399. 36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50. 51. 52. 53. 54. 150 Uyeda, K., Furuya, E., and Sherry, A.D. (1981) J. Biol. Chem. 999, 8679-8684. Pontis, H.G., Fischer, C.L. (1963) Biochem. J. 99, 452-459. Hue, L., Blackmore, P.F., Shikama, H., Robinson-Steiner, A., and Neely, P., El-Maghrabi, M.R., Pilkis, S.J., and Claus, T.H. (1981) Diabetes 99, 1062-1064. Van Schaftingen, E., Jett, M.-F., Hue, L., and Hers, H.-G. (1981) Pilkis, S.J., El-Maghrabi, M.R., McGrane, M., Pilkis, J., and Claus, T.H. (1982) Fed. Proc. 11, 2623-2628. $011ng, H.D., Kuduz, J., and Brand, I.A. (1981) FEBS Lett. 130, 309-313. Kessler, R., Nissler, K., Schellenberger, W., and Hofmann, E. (1982) Biochem. Biophys. Res. Commun. 107, 506-510. Avigad, G. (1981) Biochem. Biophys. Res. Commun. 102, 985-991. Lederer, B., Vissers, S., Van Schaftingen, E., and Hers, H.-G. (1981) Biochem. Biophys. Res. Commun. 103, 1281-1287. Bosca, L., Aragon, J.J., and $015, A. (1982) Biochem. Biophys. Res. Commun. 106, 486-491. Sener, A., Malaisse-Lagae, F., and Malaisse, W.J. (1982) Biochem. Biophys. Res. Commun. 104, 1033-1044. Malaisse, W.J., Malaisse-Lagae, F., Sener, A., Van Schaftingen, E., and Hers, H.-G. (1981) FEBS Lett. 125, 217-219. Miernyk, J.A., and Dennis, D.T. (1982) Biochem. Biophys. Res. Commun. 199, 793-798. Pilkis, S.J., El-Maghrabi, M.R., Pilkis, J., and Claus, T.H. (1981) J. Biol. Chem. 999, 3619-3622. Van Schaftingen, E., and Hers, H.-G. (1981) Proc. Natl. Sci. (USA) 19, 2861-2863. Morikofer-Zwez, S., Stoecklin, F.B., and Walter, P. (1981) Biochem. Biophys. Res. Conmun. 191, 104-111. Furuya, E., and Uyeda, K. (1981)'J. Biol. Chem. _2§_5_, 7109-7112. EI-Maghf‘abi, M.R., CIaUS, T.H., P11k15, Jo, and P11k15, S.J. (1981) Biochem. Biophys. Res. Comnun. 101, 1071-1077. 55. 56. 57. 58. 59. 60. 61. 62. 63. 64. 65. 66. 66a. 151 Hue, L., Blackmore, P.F., and Exton, J.H. (1981) J. Biol. Chem. 999, 8900-8903. Van Schaftingen, E., and Hers, H.-G. (1981) Biochem. Biophys. Res. Commun. 191, 1078-1084. Kuwajima, M., and Uyeda, K. (1982) Biochem. Biophys. Res. Commun. 191, 84-88. Furuya, E., Kotaniguchi, H., and Hagihara, B. (1982) Biochem. Biophys. Res. Commun. 105, 1519-1523. Richards, C.S., Furuya, E., and Uyeda,K. (1981) Biochem. Biophys. Res Commun. 100, 1673-1679. Van Schaftingen, E., Davies, D.R., and Hers, H.-G. (1981) Biochem. Bioplus. Res. Conmun. 103, 362-368. Furuya, E., Yokoyama, M., and Uyeda, K. (1982) Proc. Natl. Acad. é-Slo (USA) E, 325-3290 El-Maghrabi, M.R., Claus, T.H., Pilkis, J., and Pilkis, S.J. (1982) Proc. Natl. Acad. Sci. (USA) 19, 315-319. Furuya, E., Yokoyama, M., Uyeda, K. (1982) Biochem. Biophys. Res. Commun. 105, 264-270. Richards, C.S., Yokoyama, M., Furuya, E., and Uyeda, K. (1982) Biochem. Biophys. Res. Commun. 104, 1073-1079. El-Maghrabi, M.R., C1aus,T.H., Pilkis, J., Fox, E., and Pilkis, El-Maghrabi, M.R., Fox, E., Pilkis, J., and Pilkis, S.J. (1982) Biochem. Biophys. Res. Conmun. 199, 794-802. Pilkis, S.J., El-Maghrabi, M.R., McGrane, M., Pilkis, J., Fox, E., and Claus, T.H. (1982) Molecular and Cellular Endocrinology 29, 245-266. 66b. Hers, H.-G, and Van Schaftingen, E. (1982) Biochem. J. 999, 1-12. 67. 68. 69. 70. Preiss, J. (1982) Ann. Rev. Plant Physiol. 99, 431-454. Preiss, J., and Levi, C. (1980) In The Biochemistry of Plants (Preiss, J., ed.) v. 9, pp. 371-423, Academic Press, N.Y. Preiss, J., and Levi, C. (1979) In Enc clopedia of Plant Ph siolo v. 6, Photosynthesis II (Gibbs, M. and Latzko. E., eds.) pp. 2824312, Springer-Verlag, Berlin, Heidelberg, New York. Dennis, D.T., and Miernyk, J.A. (1982) Ann. Rev. Plant Physiol. E, 27'500 71. 72. 73. 74. 75. 76. 77. 78. 79. 80. 81. 82. 83. 84. 85. 86. 87. 88. 89. 152 ap Rees, T. (1974) In Plant Biochemistry (Northcote, D.H., ed.) v. 11, pp. 89-127 MTP Int. Rev. Sci. Biochem. Ser. One, fifitterworth, London and Univ. Park Press, Baltimore, Maryland. Manners, D.J. (1974) In 91ant Carbohygrate Chemistry (Pridham, J.B., ed.) pp. 109-125, Academic Press, N.Y. Akazawa, T., Minamikawa, T., and Murata, T. (1964) Plant Physiol. 19., 371-3780 Adams, C.A., Rinne, R.W., and Fjerstad, M.C. (1980) 999. 991;. _4_5, 577-582. ‘ Juliano, 8.0., and Varner, J.E. (1969) Plant Physiol. 141, 886-892. Yomo, H., and Varner, J.E. (1973) Plant Physiol. 91, 708-713. Tarrago, J.F., and Nicolas, G. (1976) Plant Physiol. 99, 618-621. Hizukuri, S., Fujii, M., and Nikuni, Z. (1961) Nature 199, 239-240. “""' Webster, 8.0., and Leopold, A.C. (1977) Amer. J. Bot. 91, 1286-1293. Akazawa, T. (1976) In Plant Biochemistry (Bonner, J., and Varner, J.E., eds.) pp. 381-403, Academic Press, New York. Manners, D.J. (1974) In Essays in Biochemistry (Campbell, P.N. and Dickens, F. eds.) v. 19 pp. 37-71, Academic Press, London and New York. Banks, W., and Muir, 0.0. (1980) In The Biochemistry of Plants (Preiss, J., ed.) v. 9, pp. 321-369} Dunn, G. (1974) Phytochemistry‘19, 1341-1346. Daussant, J., and Corvazier, P. (1970) FEBS Lett. 1, 191-194. Goswami, A.K., Jain, M.K., and Paul, B. (1977) Biol. Plant. (Praha) 19, 469-471. Varner, J.E., and Ram Chandra, G. (1964) Proc. Natl. Acad. Sci. (USA) 99, 100-106. Ho, T.-H.D., and Varner, J.E. (1978) Arch. Biochem. Biophys. 187, 441-446. Gibbons, G.C. (1979) Carlsberg Res. Commun. 11, 353-366. Okamoto, K., Kitano, H., and Akazawa, T. (1980) Plant and Cell Physiol. 31, 201-204. 90. 91. 92. 93. 94. 95. 96. 97. 98. 99. 100. 101. 102. 103. 104. 105. 106. 107. 108. 109. 153 Okamoto, K., and Akazawa, T. (1979) Plant Physiol. 91, 337-340. MacGregor, A.W., and Matsuo, R.R. (1982) Cereal Chem. 99, 210-216. Palmiano, E.P., and Juliano, 8.0. (1972) Plant Physiol. 19, 751-756. Okamoto, K., and Akazawa, T. (1980) Plant Physiol. 99, 81-84. Tsai, C.Y., and Nelson, O.E. (1969) Plant Physiol. 11, 159-167. Murata, T., Akazawa, T., and Fukuchi, S. (1968) Plant Physiol. 19, 1899-1905. Abbott, I.R., and Matheson, N.K. (1972) Phytochemistry 11, 1261-1272. Manners, D.J., Marshall, J.J., and Yellowlees, D. (1970) Biochem. g, 195, 539-541. _ Manners, D.J., and Rowe, K.L. (1971) J. Inst. Brewing 11, 358-365. Manners, D.J., and Rowe, K.L. (1969) Carbohydr. Res. 9, 107-121. Drummond, G.S., Smith, E.E., and Whelan, W.J. (1970) FEBS Lett. 9, 136-140. ""' Jorgensen, 0.8. (1964) Acta Chem. Scand. 19, 1975-1978. Takahashi, N., Shimomura, T., and Chiba, S. (1971) Agric. Biol. Chem. 99, 2015-2024. Adams, C.A., Watson, T.G., and Novellie, L. (1975) Phytochemistry, 1_4_, 953-956. Marshall, J.J., and Taylor, P.M. (1971) Biochem. Biophys. Res. Conmun, g, 173-179. Harris, N. (1976) Planta (Berl.) 199, 271-272. Bain, J.M., and Mercer, F.V. (1966) Aust. J. Biol. Sci. 19, 69-84. Smith, D., and Flinn, A.M. (1967) Planta (Berl.) 11, 72-85. Swain, R.R., and Dekker, E.E. (1966) Biochim. Biophys. Acta 122, 75-86. 110. 111. 112. 113. 114. 115. 116. 117. 118. 119. 120. 121. 122. 123. 124. 125. 126. 127. 128. 129. 130. 154 Swain, R.R., and Dekker, E.E. (1966) Biochim. Biophys. Acta 122, 87-100. Shain, Y., and Mayer, A.M. (1968) Physiol. Plant. 91, 765-776. Cohen, E., Shain, Y., Ben-Shaul, Y., and Mayer, A.M. (1971) 999. J. Bot. 19, 2053-2057. Mayer, A.M., and Shain, Y. (1968) Science 199, 1283-1284. Pontis, H.G. (1978) Trends Biochem. Sci..9, 38-39. Giaquinta, R.T. (1980) In The Biochemistry of Plants (Preiss, J., ed.) v99, pp. 271-320, Academic Press, N.Y. Arnold, W.N. (1968) J. Theoret. Biol. 91, 3-20. Edelman, J., Shibko, 5.1., and Keys, A.J. (1959) J. Expt. Bot. 19, 178-189. Nomura, T., Kono, Y., and Akazawa, T. (1969) Plant Physiol.;91, 765-769. Chen, 5.5.0., and Varner, J.E. (1969) Plant PhysioI.‘91, 770-774. Davis, D.R. (1974) In Plant Carbohydrate Biochemistry (Pridham, J.B., ed.) pp. 61-81, Academic Press, London and N.Y. Ginsburg, v. (1972) .1. Biol. Chem. _2_1_11, 55-51. Delmer, D.P., and Albersheim, P. (1970) Plant Physiol. 19, 782-786. Cardini, C.E., Leloir, L.F., and Chiriboga, J. (1955) J. Biol. Chem. 915, 149-155. Leloir, L.F., and Cardini, C.E. (1955) J. Biol. Chem. 214, 157-165. Akazawa, T. (1976) In Plant Biochemistr (Bonner, J., and Varner, J.E., eds.) pp. 62-81, Academic Fress, N.Y. Hawker, J.S. (1971) Phytochemistry 19, 2313-2322. Preiss, J. and Greensberg, E. (1969) Biochem. Biophys. Res. Commun. 99, 289-295. Salerno, G.L., and Pontis, H.G. (1976) FEBS Lett. 91, 415-418. Salerno, G.L., and Pontis, H.G. (1978) FEBS Lett. 99, 263-267. Gander, J.E. (1974) In Plant Biochemistr (Bonner, J., and Varner, J.E., eds.) pp. 337-380 AEaaem1c Press, New York. 131. 132. 133. 134. 135. 136. 137. 138. 139. 140. 141. 142. 143. 144. 145. 146. 147. 148. 149. 155 Grimes, W.J., Jones, B.L., and Albersheim, P. (1970) J. Biol. Chem 919, 188-187. Akazawa, T., and Okamoto, K. (1980) In The Biochemistry of Plants (Preiss, J., ed.) v. 3, pp. 199-220, Academicj Press, NIYZ Whitaker, J.R., and Granum, P.E. (1980) Anal. Biochem. 109, 156-159. Josse, J. (1966) 0. Biol. Chem. _24_1, 1938-1947. Lowry, D.H., Rosenbrough, N.J., Fair, A.L., and Randall, R.J. (1951) J. Biol. Chem. 199, 265-275. Bradford, M.M. (1976) Anal. Biochem. 19, 248-254. Sabularse, D.C., and Anderson, R.L. (1981) Biochem. Biophys. Res. Sabularse, D.C., and Anderson, R.L. (1981) Biochem. Biophys. Res. COMIIUTI. 1_051, 848-8550 Anderson, R.L., and Sabularse, D.C. (1982) Methods in Enzymology, in press. Nomura, T., and Akazawa, T. (1974) Plant and Cell Physiol. 19, 477-483. Cantley, L.C. Jr., Josephson, L., Warner, R., Yanagisawa, M., Lechene, C., and Guidotti, G. (1977) J. Biol. Chem. 252, 7421-7423. Josephson, L., and Cantley, L.C. Jr. (1977) Biochemistry_99, 4572-4578. Hudgins, P. M., and Bond, G. H. (1977) Biochem. Biophys. Res. Commun. 77, 1024- 1029. Beauge, L. A., and Glynn, I.M. (1977) Nature 999, 355-356. Simmons, 5., and Butler, L. G (1969) Biochem. Biophys. Acta 119, 150-157. Bucke, C. (1970) Phytochemistry 9, 1303-1309. Bloxham, D.P., and Lardy, H.A. (1973) In Enzymgs (Boyer, P.D., ed.) V. 9, pp. 239-278, Academic Press, N.Y. Dennis, D.T. and Coultate, T.P. (1967) Biochem. Biophys. Acta 146, 129-137. ‘50. ‘5]. 151a. 151b. 151c. 152. 153. 154. 155. 156. 157. 158. 159. 156 Kelly, G.J., and Turner, J.F. (1969) Biochem. J. 119, 481-487. LeBlond, D.J., Ashendel, C.L., and Wood, W.A. (1980) Anal. Biochem. 191, 355-369. Gilbert, G. (1955) Disc. Faraday Soc. 99, 68-71. Beitner, R. (1979) Trends Biochem. Sci. 1, 228-230. Kirtley, M.E., and McKay, M. (1977) Mol. Cell. Biochem. 19, 141-149. Anderson, L. E., Ng, T.-C.L., and Park, K.-E.Y. (1974) Plant Physiol. 99, 835-849. Kachru, R.B., and 1. E. Anderson (1975) Plant Physiol. 99, 199-202. Anderson, L.E. (1974) Biochem. Biophys. Res. Commun. 99, 907-913. Anderson, L.E., and Lim, T.-C. (1972) FEBS LETT. 91, 189-191. Buchanan, B.B., Schurmann, P., and Kalberer, P. (1971) J. Biol. Chem. 919, 5952-5959. Hatch, M.D., and Slack, C.R.(l969) Biochem. J. 112, 549-558. Johnson, H.S., and Hatch, M.D. (1970) Biochem. J. 119, 273-280. Latzko, E., Garnier, R.V., and Gibbs, M. (1970) Biochem. Biophys. Res. Conmun. _3_9_, 1140-1144. TATE “11111111111111111111111111111115 4 6732