UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Studies on heart muscle lipases and studies on 3', 5'-cyclic nucleotide phosphodies-terase Yamamoto, Masanobu 1966

Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.

Item Metadata


831-UBC_1966_A1 Y3.pdf [ 11.41MB ]
JSON: 831-1.0104904.json
JSON-LD: 831-1.0104904-ld.json
RDF/XML (Pretty): 831-1.0104904-rdf.xml
RDF/JSON: 831-1.0104904-rdf.json
Turtle: 831-1.0104904-turtle.txt
N-Triples: 831-1.0104904-rdf-ntriples.txt
Original Record: 831-1.0104904-source.json
Full Text

Full Text

- X -STUDIES ON HEART MUSCLE LIPASES AND STUDIES ON 3«,5t-CYCLIC NUCLEOTIDE PHOSPHODIESTERASE by MASANOBU YAMAMOTO A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY In the Department of Pharmacology We accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH COLUMBIA September, 1966 In presenting t h i s thesis i n p a r t i a l f u l f i l m e n t of the requirements for an advanced degree at the University of B r i t i s h Columbia, I agree that the Library s h a l l make i t f r e e l y available f o r reference and study, I further agree that permission-for extensive copying of t h i s thesis for scholarly purposes may be granted by the Head of my Department or by his representatives. I t i s understood that copying or publication of t h i s thesis for f i n a n c i a l gain s h a l l not be allowed without my wri t t e n permission. The University of B r i t i s h Columbia Vancouver 8, Canada Supervisor: G. I. Drummond. MASANOBU YAMAMOTO. STUDIES ON HEART MUSCLE LIPASES AND STUDIES ON y,5'-CYCLIC NUCLEOTIDE PHOSPHODIESTERASE. ABSTRACT PART I STUDIES ON HEART MUSCLE LIPASES The study of the role of l i p i d s in supplying the energy requirements of the heart has attracted widespread attention, particularly within the past decade. It i s now known that the heart, under normal conditions, oxidizes l i p i d s as i t s main source of energy. Numerous investigators have studied the Jjn vivo and in  vitro uptake and u t i l i z a t i o n of exogenously supplied lipid s i n the form of triglycerides, free fatty acids and ketone bodies. However, very few have studied the u t i l i z a t i o n of endogenous li p i d s by the working heart. We have examined the relative importance of both endo-genous glycogen and triglycerides for supplying the caloric needs of the isolated beating rat heart, and found that under the perfusion conditions used, endogenous glycogen appears to supply the i n i t i a l source of energy. A lipase in rat cardiac tissue was also examined. The enzyme had a pH optimum near 6.8, and was strongly inhibited by 0.2 M NaF and by 2 x 10~^M diisopropylfluorophosphate. Most of the activity was found in the nuclear fraction of tissue homogenates. The enzyme hydrolyzed both monoolein and mono-stearin, and possessed much less activity against tripalmitin. The enzyme also rapidly hydrolyzed the monostearin component of Ediol (a commercial coconut o i l emulsion widely used in lipase studies), and the implications of these findings are discussed. It was concluded from these studies that a lipase other than lipoprotein lipase exists in rat myocardium. - i i i -PART II STUDIES ON CYCLIC 3», 51-NUCLEOTIDE PHOSPHODIESTERASE In recent years, the study of the role of cyclic 3»,5»-adenosine monophosphate (cyclic 3*,5»-AMP) in the regulation of several biological reactions and processes has received widespread attention. The presence of a physiological mecha-nism for terminating the action of cyclic 3t,5.*-AMP in biological systems would therefore be expected. Indeed, an enzyme, cyclic 3*,5"-nucleotide phosphodiesterase has been shown to exist in most mammalian tissues which have been studied for i t s activity. The central nervous system, parti-cularly the cerebral cortex, possesses a very high activity of this enzyme. In this study, cyclic 3 1,5'-nucleotide phosphodiesterase was par t i a l l y purified from rabbit brain and i t s properties were studied. The enzyme required Mg+^ions for activity and was inhibited by 2 x lO'^M theophylline. Cyclic 3*,5*-dAMP, cyclic 3*,5»-GMP and cyclic 3*,5«-dGMP were hydrolyzed by the brain diesterase at approximately one-half the rate at which cyclic 3*,5»-AMP was hydrolyzed. L i t t l e activity against cyclic 3>,5t-CMP, cyclic 3»,5«-dCMP and cyclic 3«,5«-TMP was detected, although cyclic 3,,5«-UMP was hydrolyzed at approximately 13% of the rate at which cyclic 3*,5*-AMP was hydrolyzed. The brain diesterase therefore possessed a high specificity for cyclic 3 f, 5'-nucleotides with purine bases. Optimum enzyme activity was observed near pH 7.0, and the activity was stimulated about 1.5-fold by 0.06 M imidazole. The K value of the enzyme with cyclic 3*,5}-AMP as substrate - iv -was approximately 0.8 x 10"^M. The properties of the part-i a l l y purified phosphodiesterase from brain were thus very similar to the diesterases which have been purified from beef and dog hearts. A study of the intracellular localization of the brain diesterase indicated that about 50% of the activity was loca-ted in the 105,000 x g supernate. The microsomal and mito-chondrial fractions also contained considerable amounts of diesterase activity, but l i t t l e activity was located in the nuclear fraction. A survey of cyclic 3*,5*-nucleotide phosphodiesterase activity in several available specimens of the plant kingdom indicated the absence of this enzyme activity in these orga-nisms. However, appreciable levels of diesterase activity were detected in E. c o l i . - V -TABLE OF CONTENTS Page PART I STUDIES ON HEART MUSCLE LIPASES 1 INTRODUCTION 2 EXPERIMENTAL PROCEDURE 16 Materials 16 Methods I Perfusion Studies 18 II Cardiac Lipase Studies 23 Lipase Assay 23 Preparation of Substrates - 27 Measurement of Tripalmitin-l-C 1^ Hydrolysis 27 Preparation of Enzyme Extract 28 RESULTS 30 I Perfusion Studies 30 II Cardiac Lipases 36 A. Existence of NaF-inhibited Lipase 38 B. Preparation of Partially Purified Extract from Heart Tissue 49 C. Properties of Monoglyceride-hydrolyzing Enzyme 51 1. Albumin Requirement 54 2. pH Optimum 54 3. Temperature Optimum 57 4. Effect of Physical State of ; Substrate 57 5. Inhibitor Studies 59 6. Intracellular Localization 63 DISCUSSION 65 - v i -TABLE OF CONTENTS (cont'd.) Page PART II STUDIES ON 3',5'-CYCLIC NUCLEOTIDE PHC^PWD lE^TEiRASE1 " : : 79 INTRODUCTION 80 EXPERIMENTAL PROCEDURE 94 Materials 94 Methods - Standard Diesterase Assay 94 - Partial Purification of Brain Diesterase 99 RESULTS 103 1. Preliminary 103 2. Partial Purification of Brain Diesterase 106 3. Properties of Brain Diesterase 108 (a) Metal Requirement 108 (b) Effect of Imidazole, pH Curve 108 (c) Effect of Theophylline 111 (d) Cyclic 3«,5'-dAMP/Cyclic 3«,5'-AMP Activity Ratios 111 (e) Hydrolysis Rates of Purine and Pyrimidine Cyclic 3»^'-Nucleotides 113 (f) Further Studies on Specificity of Brain Diesterase 116 4. Cellular Distribution of Brain Diesterase 121 5. Survey of Diesterase in Human Brain, Dog Nervous System, Marine Organisms, Plants and Microorganisms 125 DISCUSSION 132 BIBLIOGRAPHY 140 -' V l l -LIST OF TABLES No. T i t l e Page PART I STUDIES ON HEART MUSCLE LIPASES I Effect of Rat Serum on Tripalmitin Hydrolysis 44 II Partial Purification of Cardiac Monogly-ceride-hydrolyzing Enzyme 52 III Inhibition of Monoglyceride-hydrolyzing Enzyme by Various Compounds 61 IV Intracellular Distribution of Monogly-ceride-splitting Enzyme 64 V Relative Rates of Hydrolysis of Monogly-cerides and Triglycerides 72 PART II STUDIES ON 3'5'-CYCLIC NUCLEO- TIDE MQSPtilOblES^ERA^r — : — VI Relative Activities of 5'-Nucleotidase and Diesterase in Snake Venom 105 VII Partial Purification and Yield of Diesterase 109 VIII Cyclic 3',5'-dAMP/Cyclic 3'i5'-AMP Activity Ratios 114 IX Relative Hydrolysis Rates of Purine and Pyrimidine Cyclic 3',5'-Nucleotides 115 X Cellular Distribution of Diesterase in Rabbit Brain 122 XI Distribution of Diesterase Activity in Human Brain 127 XII Diesterase Activity in Various Areas of Dog Nervous System 128 XIII Diesterase Activity in Various Marine Organisms 129 XIV Diesterase Activity in Plants and Micro-organisms 130 - v i i i -LIST OF FIGURES No. T i t l e Page PART I STUDIES ON HEART MUSCLE LIPASES 1 Glycogen Standard Curve 21 2 Tripalmitin Standard Curve 24 3 Palmitic Acid Standard Curve 26 4 Tissue Glycogen Content - Substrate Free Perfusion 32 5 Tissue Triglyceride Content - Substrate Free Perfusion 35 6 Time Course of Lipolytic Activity 40 7 Inhibition of Lipolytic Activity in Crude Homogenates by NaF 41 8 Effect of NaF on Hydrolysis of Monostearin and Tripalmitin 43 9 Hydrolysis of E d i o l R Components 46 10 Enzyme Concentration Curve - Monostearin as Substrate 53 11 Effect of Albumin on Lipolytic Activity 55 12 pH Curve of Monoglycerlde-hydrolyzing Enzyme 56 13 Effect of Temperature on Cardiac Monoglyceride-hydrolyzing Lipolytic Activity 58 14 Effect of Monoolein Concentration on Enzyme Activity 60 15 Effect of Fasting on Myocardial Glycogen Level 67 PART II STUDIES ON -3' ,5'-CYCLIC NUCLEO-TWE PHOSFHQD IESTERASE1 ~ " — 16 Structural Formula of Cyclic 3' ,5'-AMP 82 - XX -LIST OF FIGURES (cont'd.) No. T i t l e Page 17 Mediation of Cyclic 3«,5'-AMP in the Glyco-genolytic Response of Liver to Epinephrine 85 18 Inorganic Phosphate Concentration Curve 96 19 Spectrophotometrie Assay of Phosphodies-terase 98 20 Hydrolysis of 5*-AMP by Snake Venom (Crotalus adamanteus) 104 21 Phosphodiesterase Activity as a Linear Function of Protein Concentration 107 22 pH Curve of Brain Diesterase and Effect of Imidazole 110 23 Inhibition of Diesterase by Theophylline in vitro 112 24 Hydrolysis of Cyclic 3«,5»-AMP by Brain Diesterase 117 25 Hydrolysis of Cyclic 3«,5»-dAMP by Brain Diesterase 118 26 Hydrolysis of Cyclic 3«,5'-GMP by Brain Diesterase 119 27 Hydrolysis of Cyclic 3«,5«-dGMP by Brain Diesterase 120 28 Absence of 5'-Nucleotidase Activity in Par t i a l l y Purified Preparation 123 29 Absence of Cyclic 2«,3«-AMP Hydrolytic Activity in Partially Purified Preparation 124 30 The Two-Messenger Concept for the Expression of Hormonal Control 133 31 Structural Formulae of Methyl Xanthines 137 - X -LIST OF ABBREVIATIONS FFA Free fatty acid DFP D iisopropylfluorophosphate EDTA Ethylenediaminetetraacetic acid 5«-AMP Adenosine 5»-phosphate ATP Adeno s ine 5«-triphosphate UDPG Uridine diphosphate glucose Cyclic 3« ,5t -AMP Adenosine 3*,5*-phosphate Cyclic 3» ,5« -UMP Urid ine 3 *,51-phosphate Cyclic 3« ,5« -dAMP Deoxyadenosine 31,5 *-phosphate Cyclic 3« ,5« -dGMP Deoxyguanosine 31,5 *-phosphate Cyclic 3« ,5« -GMP Guanosine 3*,5»-phosphate Cyclic 3« ,5t -dCMP Deoxycytidine 3*,5*-phosphate Cyclic 3« ,5t -TMP Thymidine 3»,5*-phosphate Cyclic 2« ,3« -AMP Adenosine 2«,3*-phosphate P i Inorganic phosphate - x i -ACKNOWLEDGEMENT I am deeply grateful to Dr. George I. Drummond, Department of Pharmacology, U. B. C. for frequently taking time off his busy schedule to give me much valuable advice and helpful criticism throughout the course of this work. I also wish to thank him for making available Medical Research Council grants which made my graduate work possible. I thank Dr. James G. Foulks, Head of the Department of Pharmacology, U. B. C. for the frequent use of his laboratory f a c i l i t i e s . My thanks also to Dr. Hans-Peter Baer, Mrs. Loverne Duncan, and Mrs. E. Hertzman for the occasional technical assistance, many interesting discussions, and personal encouragement. In addition, I am grateful to Dr. L. Druehl and Mr. M. McLaren of the Department of Botany, U. B. C., for making available several specimens of plants, and to Dr. D. Duncan (B. C. Research Council) and Dr. W. J . Polglase (Department of Biochemistry, U. B. C.)r for the generous supply of microorganisms. I am particularly indebted to Mr. G. Kent and Mr. R. Smith of this department for frequently giving me a helping hand when needed throughout these past few years. PART I STUDIES ON HEART MUSCLE LIPASES 2. INTRODUCTION It i s widely known that during,starvation, the mammalian organism derives v i r t u a l l y a l l of i t s energy requirements from fat catabolism. Less widely realized, perhaps, i s the fact that even under normal physiological conditions, mammals depend to a large extent on the oxidation of fats for energy. For example, Fredrickson and Gordon (20) measured the expired C^02 after injection of C^-labelled albumin-bound long-chain fatty acids into man and indicated that up to 50% of the energy could have been derived from fatty acids during the post-absorptive state. Various tissues and organs have been examined for their a b i l i t y to oxidize lipids jLn vivo and in  v i t r o . In 1930, Richardson et al.(65) found that the res-piratory quotients of incubated, excised renal and muscular tissues were consistently intermediate between 0.7 and 1.0, and suggested that fats may have participated in the oxida-tive process. Artom (66), Volk et al.(68) and Geyer and associates (69) demonstrated that when carboxyl-labelled C ^ - f a t t y acids were incubated with kidney, l i v e r , spleen, heart, lung, brain, skeletal muscle and testis slices, a l l of these tissues were capable of oxidizing the fatty acids to C^o^. The in vitro uptake of fatty acids by the isolated diaphragm was measured by Wertheimer and Ben-Tor (70), and the oxidation of octanoic acid by this tissue was demonstrated by Hansen (71). According to Neptune et al.(90,91), isolated diaphragms possess the a b i l i t y to u t i l i z e endogenous l i p i d s . When rat diaphragms were incubated for 4 hours in a substrate-3. free medium, there was a slight net decrease in tissue free fatty acids, triglycerides and especially in total phospho-l i p i d s , F r i t z and co-workers (72) measured the uptake and oxidation of acetate-l-C 1^, octanoate-l-C 1^ and palmitate-14 1-C by the isolated diaphragm and skeletal muscle at rest and at work. Their data indicated that e l e c t r i c a l l y stimu-lated skeletal muscles oxidized twice as much palmitate than resting muscles in the presence or absence of added glucose. Freidberg et ai.(73) showed that after 5 minutes of moderate exercise, the plasma FFA concentration in man dropped from approximately 0.82 to 0.61 mM, then increased sharply to about 1.2 mM after cessation of activity. Similar observa-tions were recorded by Issekutz and Miller (74) in dogs, who noted that the decrease in plasma FFA levels was accompanied by a 5-fold increase in oxygen uptake. F r i t z (63) has suggested, therefore, that during moderate, sustained work, i f the oxygen supply i s adequate, skeletal muscles can oxi-dize a considerable amount of lipid s for energy, Indeed, 14 when Havel and his associates (21) infused palmitate-l-C intravenously into human subjects walking on a treadmill at 3-4 miles per hour, they found a rapid mobilization of fatty acids (presumably from adipose tissue), and an increased rate of oxidation of palmitate-l-C 1^. The uptake and oxidation of labelled long-chain fatty acids by skeletal muscle in vivo was measured at rest and during e l e c t r i c a l stimulation by Spitzer and Gold (40). Their data agreed with the concept that FFA are oxidized by skeletal muscle at rest, and that 4. FFA oxidation increases during muscular activity. Andres et a l o(75) measured the differences in arteriovenous concen-trations of 0^, CO^, glucose and lactate in forearms of human subjects at rest, and concluded that under basal conditions the oxidation of glucose could account for only 7% of the oxygen uptake, Since the mean respiratory quotient of fore-arm muscle was 0;80, these investigators suggested that the major non-carbohydrate material which served as fuel under the conditions of their experiment was l i p i d . From the fore-going, i t i s clear that both diaphragm and skeletal muscles are capable of oxidizing fatty acids directly for energy. Recently, Masoro et ;al»(92) found that no decrease in endo-genous triglycerides or in any of the muscle phospholipids had occurred in monkey skeletal muscles which had been stimulated for 5 hours in si t u . They concluded, therefore, that skeletal muscles are capable of oxidizing only exo-genously supplied l i p i d s . The u t i l i z a t i o n of fatty acids for energy by the brain has been studied both _in vivo and jLn v i t r o . According to Gordon and Cherkes (77) and Quastel and Wheatley (78), the brain does not u t i l i z e l i p i d s for energy. On the other hand, a number of investigators have indicated that the brain i s capable of oxidizing lipids to a limited extent (68, 69, 79, 80). However, in view of a number of earlier observations on the R.Q. of the brain (81), i t would appear that although the brain possesses the enzymes for oxidizing l i p i d s , the amount of energy derived from this source i s insignificant 5. compared to that contributed by the oxidation of glucose. Other tissues which have been examined for their a b i l i t y to oxidize l i p i d s are adipose tissue (82) and li v e r (66, 68, 69). Isolated mitochondria from the latter tissue actively oxidize l i p i d s , especially in the presence of carnitine (83, 84, 85). It i s well known that the heart functions largely aerobically and one might expect therefore that this organ would be particularly adapted for the oxidation of l i p i d s . Perhaps $wo of the earliest investigators to suggest that the heart must u t i l i z e some substrate other than carbohydrates were Visscher and Mulder (18) in 1930. Using the isolated heart-lung preparation, these investigators discovered that even after 6 hours of work, the same amount of glycogen was found as in the normal, unworked hearts. Furthermore, they suggested that since a l l the carbohydrates in the heart-lung system could not account for the total energy requirements of the heart, some other non-carbohydrate substrate must have been utilized during the 6 hours of work. Eight years later, Visscher (19) presented quantitative evidence that the non-carbohydrate source may be fat. Again using the heart-lung preparation, he showed that during 3 hours of cardiac work, the total fat content of ventricular muscle decreased from 3.71 i 0.76 to 3.18 "± 0.68 g per 100 wet weight of cardiac tissue. In spite of Visscher*s investigations (18, 19) and Cruikshank«s review in the 1930«s (94) on myocardial 6. metabolism, interest in cardiac l i p i d metabolism appears to have subsided for almost two decades, u n t i l Bing and his associates (22, 86, 87) in 1953 and 1954 reported on the in vivo uptake of fatty acids by the myocardium in human sub-jects. They found that at blood fatty acid levels of 1.105-0.286 mEq/100 ml, the extraction of fatty acids was 0.016 ±. 0.013 mEq/100 ml. Their 4aturn i s not highly impressive, but their reports on the in vivo uptake of fatty acids by the human heart appear.: to have attracted the interest of numerous investigators to the area of cardiac l i p i d metabolism. While measuring the transport : ; of plasma FFA, Gordon (88) also noted the myocardial extraction of FFA jLn vivo. In accor-dance with earlier observations, the uptake of FFA by the myocardium in vivo was also demonstrated by Ballard et a l . (89). Their data indicated that in fasting dogs, free fatty acids accounted for only 23% of the total fatty acids extracted by the heart, while the esterified fatty acids made up the other 777.. Similar conclusions were reached by Scott and co-workers (24) who measured the myocardial removal of FFA under normal and pathological conditions in dogs. It i s perhaps pertinent to mention here that when Bragdon and Gordon (93) injected C 1^-labelled chylomicrons into fasted rats and analyzed the various tissues for radioactivity, the tissues with the highest specific activity were the l i v e r and heart, which accounted for about 50% and 25% respectively of the total a c t i v i t i e s found. More recently, Rothlin and Bing (23) showed that oleic acid was extracted by the heart to a greater 7. extent than any other long-chain fatty acids in a r t e r i a l blood. Similar results have been obtained with the isolated perfused heart (98). It must be emphasized that these in vivo studies, though important, nevertheless gave only an indication that l i p i d s may have been utili z e d by the heart, since only their uptake by the myocardium was measured. The strongest evidence that lipids in the form of FFA and triglycerides are not only taken up but also oxidized by the heart for energy has come from more recent studies u t i l i z i n g the isolated perfused heart. Evans and his associates (27) used a closed perfusion system to demonstrate that isolated rat hearts converted about 90% of the palmitate-l-C^ in the perfusion medium to C l£i02 i n 6 0 minutes. That short-chain fatty acids (C 1^-labelled acetate, propionate, n-butyrate, n-octanoate) were also readily oxidized directly by isolated dog hearts to 14 C O2 was shown e a r i l i e r by Cavert and Johnson (95). Opie >et'al»(26), Shipp (97) and Shipp and his associates (96) found that whereas palmitate-l-C"^ was taken up and oxidized by hearts obtained from both fed and fasted rats, glucose-14 U-C was oxidized only by hearts obtained from fed rats. Furthermore, when both substrates were made available to the isolated hearts from either fed or fasted rats, palmitate-1-was -preferentially taken up and oxidized over glucose-U-C^. Studies with free fatty acids as substrates for the myo-cardium demonstrated unequivocally their importance as an energy source. However, the question as to whether circu-8. lating triglycerides (which actually represent the majority of the total circulating fatty acids in vivo) are taken up and oxidized by the myocardium remained unanswered u n t i l recent years. The relative importance of esterified fatty acids in supplying the energy demands of the heart was suggested earlier by Ballard et al»(89), who found that 777. of the total fatty acids extracted by the heart in vivo was in the esterified form. The uptake and oxidation of t r i -glycerides by the isolated perfused rat heart were studied by Gousios and co-workers (33), These investigators found that 307. of the total d < 1.006 lipoproteins (labelled with C1^-tripalmitin) in the perfusion medium was extracted by hearts from starved rabbits and that 107. of the extracted t r i g l y -cerides was oxidized to C^02. Hearts obtained from fed rabbits extracted 157. of the available C 1 Z h-tripalmitin and 14 oxidized 87. of the extracted triglyceride to C Olive-crona (31) and Olivecrona and Belfrage (32) injected C1^-3 -glycerol-H -palmitate-labelled chylomicrons intravenously into rats and examined the distribution of the labels in a number of tissues, including l i v e r , heart and adipose tissues. They concluded from their studies that the heart (and adipose tissue) took up chylomicron triglycerides intact, and that extensive and rapid hydrolysis of this glyceride occurred probably near the plasma membrane. The uptake and oxidation of d < 1.006 lipoprotein triglycerides by the isolated rat heart was also demonstrated by Delcher and associates (34) who found that about 407.-907. of the total C0 2 was derived 9 . from the exogenously supplied triglycerides, and thus con- cluded that lipoprotein-triglyceride fatty acids were the  primary"source of fatty acids for the heart. It should be mentioned b r i e f l y that other substrates have also been shown to be rapidly metabolized by the heart. Williamson and Krebs (29), Williamson (30) and Hall (99) demonstrated that acetoacetate, acetate and ^-hydroxybutyrate were, as might be expected, oxidized rapidly by the perfused rat heart in preference to glucose. In summary, the evidence presented by numerous investi-gators demonstrates clearly that a variety of mammalian tissues are capable of oxidizing lipi d s for energy. The heart, in particular, appears to depend primarily upon li p i d s as i t s source of energy. The importance of FFA in myocardial metabolism has been reviewed recently by Evans (28) and by Bing (100). The comparative aspect of muscle metabolism, with special emphasis on the importance of l i p i d metabolism in insects, birds and fishes has been reviewed by Drummond and Black (76). , The intracellular fate of FFA»s which are transported to the myocardium (either complexed with serum albumin or as chylomicron triglycerides) has been examined, and the evi-dence indicates that not a l l of the FFA fs are directly oxidized by the myocardium; a portion i s re-esterified to triglycerides and phospholipids. For example, Shipp (97) found that when rat hearts were perfused with 0.5 mM palrai-tate-1-C 1 4 f o r 30 minutes, 4.69 ± 0.16 umoles of palmitate were taken up per gram weight of tissue, 2.34 - 0.24 umoles 14 . recovered as C 0^, and 1.68 - 0.06 umoles recovered as tissue l i p i d s . In hearts perfused for 1 hour with 0.5 mM palmitate-l-C 1 4, 65.4 ± 2.3% of the C 1 4 in tissue lipids was recovered as triglyceride, 14.1 ± 1.1% as phospholipids, 6,6 i 0.5% as FFA, and 13.9- 1.7% as cholesterol. The : -synthesis of triglycerides in rat hearts 'jn vivo from injec-ted palmitate-l-C''"4 was shown earlier by Borgstrom and Olivecrona (101). According to Stein and Stein (103), the isolated perfused rat heart incorporated \ palmitate-l-C"1"4 into tissue l i p i d s , the triglycerides accounting for 70-75%, and the phospholipids, 25-30%.,of the label incorporated. Qualitatively, similar observations have been recorded by Shipp et al*(35) and by Olson (102). Hence i t is reasonable to assume that FFA*s which are taken up in excess of the immediate energy requirements are re-esterified and stored in the form of phospholipids or triglycerides. Indeed, elec-tron micrographs of cardiac muscle c e l l s often show abundant amounts of l i p i d droplets often adhering to the mitochondria. The u t i l i z a t i o n of these endogenous li p i d s for energy by the myocardium was investigated by Shipp and his asso-ciates (35, 36, 104) and also by Denton and Randle (37). Essentially, the observations made by Shipp et al»were that when glycogen-depleted rat hearts (whose intracellular lip i d s had been pre-labelled with C^" _in vivo) were perfused in a closed system with substrate-free buffer, the production of C l 40o gave direct evidence that endogenous li p i d s were 11. oxidized. Furthermore, they stated that the net decrease in endogenous C^-labelled phospholipid content alone could account for oyer 757. of the total metabolic C0 2 formed under these conditions. However, in direct contrast to these observations, Denton and Randle (37) showed that after 60 minutes of substrate-free perfusion, the triglyceride content f e l l from 18.7 - 0.8 to 8.7 ± 0.7 umoles per gram dry weight of tissue, and there was no change in endogenous phospho-l i p i d levels. These contradictory observations present much d i f f i c u l t y in assessing the exact role of endogenous li p i d s as potential fuel for the working heart muscle, and therefore this particular aspect of cardiac metabolism must s t i l l be considered open for further investigation. The role of endogenous lipids in supplying the energy requirements of the heart may be uncertain, but as mentioned earlier, the importance of exogenously supplied chylomicron triglycerides in this respect cannot be over-emphasized. Complete agreement exists among researchers (31, 32, 33, 34) that chylomicron triglycerides are taken up intact by the myocardium and subsequently hydrolyzed rapidly to FFA.*s. These observations immediately suggest that l i p o l y t i c enzymes must exist in heart c e l l s to hydrolyze triglycerides to FFA and glycerol. Indeed, an active enzyme, lipoprotein lipase, was characterized in cardiac tissue in 1955 by Korn (38, 39). This enzyme was also detected in l i v e r , kidney, spleen, aorta, lung, skeletal muscle and adipose tissue, by Korn (38, 47), as well as in post-heparin plasma (42) and diaphragm (50). 12. Lipoprotein lipase has been extensively studied by numerous investigators, and since i t has been the subject of a f a i r l y recent review (41), only the salient features of this enzyme w i l l be presented. Lipoprotein lipase has been reported puri-fied from post-heparin plasma 1480-fold by Hollett and Meng (42) using isoelectric precipitation and ammonium sulfate fractionation procedures. The purified preparation was optimally active at pH 8.5 and i t s activity destroyed by heating for 5 minutes at 50°. The natural substrate for lipo-protein lipase appears to be chylomicrons, which are composed of about 90% triglycerides, some phospholipids, cholesterol esters and about 2% protein. Coconut o i l emulsions and other a r t i f i c i a l triglyceride preparations are attacked at only a slow rate by the enzyme, unless small quantities of serum are present (39). A unique property of lipoprotein lipase i s i t s sudden appearance in the circulation after heparin administration, as f i r s t observed by Hahn (105). Indeed, the enzyme is eluted within minutes from adipose tissue and heart when these organs are perfused with buffer containing heparin and serum (43, 44, 45, 46). The pH optimum for this enzyme is near 8.5 and i t s activity is inhibited 100% by 0.2 to 1.0 M NaCl (38, 43, 45, 48) and 30-60% by protamine sulfate, 20 mg/ml (43, 48). The enzyme is very slightly (0-7%) inhibited by 0.2 M NaF (43, 48, 49). Hollenberg (50) and Alousi and Mallov (17) have noted a 2-3 fold increase in lipoprotein lipase activity in hearts obtained from 3-4 day fasted rats. A simi-lar increase in enzyme activity was observed by Nikkila est a l . 13. (51, 52) in the myocardium of rats which had been subjected to moderate exercise for 90 minutes. The mode of action of lipoprotein lipase has been investigated by Borgstrom and Carlson (53), Carlson and Wadstrom (54) and most recently, by Payza et al.(106). Carlson and Wadstrom»s data (54) indicates clearly that chylomicron triglycerides are hydrolyzed rapidly to monoglycerides, but the hydrolysis of the latter glyceride occurred very slowly. This was illustrated by the rapid (150-fold) increase in monoglyceride content within the f i r s t 5 minutes of incubation, accompanied by decreases in t r i -glyceride and diglyceride levels. Payza and co-workers (106) have similarly shown that monoglycerides accumulated when an a r t i f i c i a l coconut emulsion (Ediol^) was used as substratei It i s reasonable to conclude, therefore, that monoglycerides are not hydrolyzed to any extent by lipoprotein lipase, and that i t s action i s specific for triglycerides, and may even extend to diglycerides. The physiological importance of lipoprotein lipase in cardiac energy metabolism cannot be underestimated. In adipose tissue, a lipase possessing properties con-siderably different from that of lipoprotein lipase was shown to: exist by Rizack (107). Using E d i o l R as substrate, he showed that the enzyme was inhibited 16% by 0.6 M NaCl, 7% by 8 x 10"% EDTA, 66% by 0.2 M NaF, but not inhibited by protamine sulfate, 300 jug/ml. The optimum activity was observed near pH 6.5 as compared with pH 8.5 for lipoprotein lipase. The most interesting feature of this lipase was 14. that i t could be re-activated when incubated with epinephrine and tissue sediment, suggesting an important means of control-ling free fatty acid release from adipose tissue. It was later reported by Rizack (108) that the enzyme could be a c t i -vated i n vitro by 2 x 10"% cyclic 3* ,5*-PiSGP. Much interest is currently being directed toward the po s s i b i l i t y that this enzyme i s under hormonal control and therefore regulates the output of free fatty acids from adipose tissue. Recently, Bjorntorp and Furman (13) reported that a li p o l y t i c activity, similar to that observed by Rizack (107) in adipose tissue, existed in rat hearts. Using E d i o l R as substrate and crude extracts as the enzyme source, these investigators indicated that the l i p o l y t i c activity was opti-mal near pH 6.8, slightly inhibited (8%) by 0.5 M NaCl but strongly inhibited (68-100%) by 0.2 M NaF. It was not inhi-bited by protamine sulfate, 400 ug/ral. Furthermore, they reported that when heart tissue from fasted rats was incu-bated in the presence of epinephrine, 1 ug/ml, the activity increased from 12.70 ± 1.30 to 13.60 i 1.60 (uraoles FFA released/g tissue/hour). They concluded that in addition to lipoprotein lipase, another l i p o l y t i c component existed in rat cardiac tissue, whose function was perhaps analogous to that of the lipase found in rat adipose tissue by Rizack (107). Evidence supporting the concept that lipids play a major role in supplying the energy demands of the mammalian heart has been presented. It may be concluded that the heart in vivo derives part of i t s energy from the direct oxidation of 15. albumin-bound free fatty acids which are taken up from the ar t e r i a l circulation. However, the fuel for muscle metabolism i s to a much greater extent derived from exogenously supplied chylomicron triglycerides and very low density lipoproteins. It follows, therefore, that cardiac lipases must play a v i t a l role in providing a source of oxidizable fatty acids for the myocardium. The question as to whether endogenously stored cardiac lipi d s are readily mobilized and utilized for energy has not been unequivocally answered. The work to be des-cribed in this thesis was undertaken to further study the nature of l i p o l y t i c a c t i v i t i e s in heart muscle. Special attention has been directed toward l i p o l y t i c activity other than lipoprotein lipase activity. We have been particularly interested by the suggestion that a cardiac lipase may exist which is activated by epinephrine. An investigation to pro-vide additional insight into the possible u t i l i z a t i o n of endogenous triglycerides by isolated perfused rat hearts i s also reported. 16. EXPERIMENTAL PROCEDURE Materials Glycogen was obtained from Nutritional Biochemical Company. Diazyme , an amyloglucosidase preparation, was •a purchased from Miles Chemical Company. Glucostat^ reagent, which contains glucose oxidase and horseradish peroxidase, was purchased from Worthington Biochemical Corporation. R Ediol , which was generously provided by Dr. Martin Rizack of the Rockefeller Institute, New York, contains coconut o i l 507., sucrose 12.5%, glyceryl monostearate 1.5%, and poly-oxyethylene sorbitan monostearate 2.0%. Bovine serum albumin (Fraction V) was purchased from Sigma Chemical Company and purified before use by the method of Goodman (1) as follows: 50 g of the crude albumin was dissolved in 200 ml of glass-distilled water by simply placing the albumin powder over the water and allowing i t to dissolve overnight. The resultant dark amber solution was then lyo-philized; the residue was powdered with mortar and pestle, covered with anhydrous 2,2,4-trimethylpentane containing 5% acetic acid, and f i n a l l y placed in the cold room overnight. As much as possible of the acetic acid-trimethylpentane extraction solvent was then aspirated, and the albumin washed twice with anhydrous trimethylpentane. Agitation of the albumin suspension during the extraction process with organic solvents was kept to a minimum to reduce the extent of protein denaturation. After aspiration of the trimethylpentane, the albumin was again covered with the anhydrous 57. acetic acid-trimethylpentane mixture, and stored in the cold room over-night. The removal of the acetic acid-trimethylpentane mixture and washing with anhydrous trimethylpentane was repeated. The organic solvent was removed under vacuum, and the powder obtained was taken up in a suitable volume of glass-distilled water. To remove the last trace of acetic acid, the albumin solution was dialyzed by continuous flow for 3 days against a total volume of 60 l i t e r s of demineral-ized water, followed by 20 l i t e r s of glass-distilled water. The solution was then lyophilized and the extracted albumin stored in the deepfreeze u n t i l required. Commercial albumin (Fraction V) contains about 0.60 eq FFA/mole. After extrac-tion by the method of Goodman (1) just described, the content of FFA is reduced to about 0.14-0.18 eq/mole. Monoolein (Calbiochem, "907. pure") was made free of trace triglyceride contaminant by adsorption on 80-200 mesh s i l i c i c acid, followed by elution with chloroform:methanol, 2:1. Tripalmitin was obtained from Eastman Organic chemicals and purified ( > 997.) by s i l i c i c acid chromatography. T r i -palmitin-1-G 1^ (967. pure) was purchased from Nuclear-Chicago Corporation. Commercial monostearin was obtained from the Faculty of Pharmacy, U. B. C , and re-crystallized twice from hot ethanol before use. S i l i c a gel GF (Merck) was secured from Canadian Laboratories. 18. Methods I. Perfusion Studies Normal fed and 3-day fasted female Wistar rats weighing between 275 and 325 grams were used. The animals were stunned by a blow on the head, their hearts removed immediately and attached to a cannula of a Langendorf perfusion apparatus. The apex of the hearts was secured to a Stratham Force Dis-placement Transducer, a 5-gram tension applied, and the rate and strength of contractions recorded on a Grass Model 5D polygraph.. The flow rate was adjusted as required for maxi-mal efficiency of the heart, usually between 5 and 8 mis per minute. The perfusion medium was carbogenated Krebs-Ringer bicarbonate solution at pH 7.4, 37°. When epinephrine was added to the perfusion f l u i d , i t was injected with a Lambda Pump Driver at the rate of 0.2 to 1.0 jug per minute. When heparin was used, i t was injected at the rate of 60 ug per minute. At the end of the perfusion period, the hearts were removed from the apparatus and the auricles cut away and dis-carded. The ventricles were carefully blotted to remove excess water and divided in two in such a way as to provide approximately equal parts of the l e f t and right ventricular tissues for subsequent glycogen and triglyceride analyses. Samples thus obtained were immersed in liquid nitrogen within one minute following termination of perfusion, then assayed on the following day. Tissue Glycogen — Tissue glycogen was assayed enzymatically 19. according to Johnson et jal (2). A sample, of ventricular tissue; weighing between 150-250 mgwas placed in a graduated 12-ml centrifuge tube: containing 1.0 ml of 30% KOH. The tuber was t placed in boiling water for 20 minutes, the con-tents cooled, and 1.25 ml 95% ethanol was added to precipi-tate the glycogen. The contents of the tube were mixed thoroughly with a glass rod. The tube was chilled in ice for 15 minutes, then the contents heated to a b o i l in a water bath. The precipitate was collected by centrifugation for 15 minutes, using a bench top Model H International centrifuge. The supernatant f l u i d was decanted and the pre-cipitate dissolved in 1.0 ml glass-distilled water. Then 1.25 ml 957. ethanol was added to re-precipitate the glycogen, and the tube chilled and centrifuged as before. The sediment was taken up in 2.0 ml of glass-distilled water, and usually an 0.2 ml aliquot was taken for glycogen determination. The amyloglucosidase solution used for the glycogen assay was prepared by mixing 200 mg Diazyme with 100 ml 0.1 M potassium phosphate buffer, pH 6, and f i l t e r i n g . The enzyme solution was stored at 4°, and discarded after one week. The glucose oxidase-horseradish peroxidase reaction mixture (Glucostat^ x 4) was prepared by f i r s t dissolving the contents of the smaller (chromagen) v i a l in 4.0 ml methanol. The contents of the larger (enzyme) v i a l ware then dissolved in about 380 ml of buffered glycerol (4 volumes glycerol plus 6 volumes 0.04 M potassium phosphate buffer, pH 7), the chromagen solution added, and made up to 400 ml with buffered glycerol. This preparation was stored in the deepfreeze in small individual quantities, and was stable to repeated freezing and thawing. The incubation mixture for glycogen determination con-sisted of the following: 0.2 ml aliquot of the glycogen solution, 1.0 ml of Glucostat R reagent, 1.0 ml of DiazymeR solution, and 0.8 ml glass-distilled water. The mixture was incubated for 1 hour at 37° and the reaction stopped by the addition of 0.5 ml 2 N HC1. The optical density was read at 400 mu in a Beckman DU spectrophotometer, using a light path of 1.0 cm. The standard curve for glycogen i n the range, 5-100 ug i s illustrated in Fig. 1. Recovery experiments indicated 90-95% recovery of added glycogen. Tissue Triglycerides Tissue triglycerides were extracted by the method of Folch e_t a l (7). Ventricular tissues weigh-ing between 150 and 250 mg were minced and homogenized for at least 7 minutes in a glass mortar (with a loose f i t t i n g Teflon motor-driven pestle) with 18 volumes of chloroform-methanol mixture (2:1). The flaky suspension was fi l t e r e d through paper into a 12-ml centrifuge tube, using 2 volumes of the chloroform-methanol mixture as a f i n a l rinse. Four volumes of glass-distilled water was then added, the tube shaken by hand, and centrifuged. The upper phase was care-f u l l y removed with a pipette, and a 2.0 ml aliquot of the lower phase transferred to a screw-capped tube and evaporated to dryness under a gentle stream of nitrogen. The residual l i p i d was assayed for triglycerides by the original 21. FIG. 1 Glycogen concentration versus optical density* The assay of glycogen was performed with the coupled amyloglucosidase-glucose oxidase technique of Johnson e_t a l . (2) as described in the text, except that the quantities of glycogen were varied. 22. method of Van Handel and Zilversmit (3) as modified by Jagan-nathan (4), except that s i l i c i c acid and diisopropyl ether were substituted for zeolite and chloroform, respectively. To the l i p i d residue was added 1.2 g activated s i l i c i c acid followed by : 7.5 ml diisopropyl ether, and the contents shaken on a mechanical shaker for 30 minutes. After centri-fugation at low speed, a 4.0 ml aliquot was transferred to another screw-capped tube and the organic solvent evaporated to dryness under a gentle stream of nitrogen. The rate of evaporation was iiincjiieasel by immersing the tube in a water bath at 60°. To this residue was added 1 drop of 2.5% KOH and 1.0 ml of aldehyde-free ethanol, and the triglycerides hydrolyzed to glycerol and fatty acid salts by heating for 30 minutes at 60°. Two drops of 6% acetic acid were then added, and the contents evaporated to dryness in an oven at 55° using a gentle stream of air to hasten the process. To the dry contents were added 10.0 ml petroleum ether (b.p, 35-60°) and 2.0 ml 0.7 N H2S0^. The tube was then capped tightly and inverted 25 times. The petroleum ether layer was removed by aspiration and discarded. To remove the last traces of the organic solvent, the tube was heated in an oven for 15 minutes at 60° under a gentle stream of a i r . Three drops of 25 mM sodium metaperiodate solution were then added, and the contents of the tube mixed thoroughly. After 10 minutes, 0.2 ml of freshly prepared sodium b i s u l f i t e (10% w/v) solu-tion was added and the contents mixed thoroughly. Ten ml of chromotropic acid solution were f i n a l l y added, the tubes shaken and heated in a boiling water bath for 30 minutes. After cooling to room temperature, 1.0 ml of 5% thiourea solution was added, the contents mixed and the intensity of the colour was read at 570 raji in Beckman DU spectrophoto-meter using a light path of 1.0 cm. Aldehyde-free ethanol was prepared by heating 1000 mis ethanol under reflux for 60 minutes with 20 g zinc dust and 20 g KOH. The ethanol was then d i s t i l l e d , discarding the head and t a i l fractions. Chromotropic acid reagent was prepared in subdued light by f i r s t dissolving 2.24 g chromo-tropic acid in 200 mis H^ O, then adding this solution to 900 mis of sulfuric acid solution (300 mis H^ O plus 600 mis concentrated H^SO^). The reagent was stored in the dark and prepared fresh every two weeks. The standard curve for tripalmitin is shown in Fig. 2. Recovery experiments carried through from the s i l i c i c acid extraction step indicated recoveries in the range 95-98%. Both glycogen and triglyceride tissue levels are expressed as mg/g dry weight of ventricular tissue. This i s based on the observation that the dry/wet weight ratios were 25.4% and 20.5% for non-perfused hearts, and hearts perfused over 5 minutes, respectively. II. Cardiac Lipase Studies Lipase Assay -- Standard lipase assays were performed in screw-capped tubes at 37° on a Dubnoff metabolic shaker. Incubation time was 30 minutes. The incubation mixture con-tained 60 mM potassium phosphate buffer, pH 6.8, 20 mg 24 . E 0 100 200 300 400 500 600 TRIPALMITIN Ipv) FIG. 2 Tripalmitin concentration versus optical density. The assay was performed as described in the text except that the i n i t i a l s i l i c i c acid extraction step was omitted. Recovery experiments indicated 95-98% recovery of added t r i -palmitin . when carried through the s i l i c i c acid extraction step. 25. purified bovine serum albumin, enzyme preparation, the appro-priate substrate, and glass-distilled water to make a f i n a l volume of 1.0 ml. When Ediol served as substrate in the standard assay, 0.1 ml of a 1:19 dilution was used. The amount of monoolein as substrate was either 5 or 15 ueqs per reaction mixture. Tripalmitin was used at a concentration of 2.5 mg per ml of reaction mixture. Lipolytic activity was measured by the method of Duncombe (5), as modified by Vaughan et al<>(6) as follows: The reaction was stopped by the addi-tion of 1.0 ml of a mixture containing 0.9 M triethanolamine, 0.1 N acetic acid, and 5% cupric nitratetSH^O. The purpose of this treatment i s to convert the FFA formed to the chloro-form-soluble copper soaps. Chloroform, 6.0 ml, was added and the tubes shaken on a mechanical shaker for 15 minutes. After brief centrifugation, the aqueous copper solution and the denatured protein were removed by suction. An aliquot (0.2-2.0 ml) of the chloroform layer was removed, made up to a f i n a l volume of 2.0 ml with chloroform, and 0.25 ml of freshly prepared 0.1% diethyldithiocarbamate (prepared in n-butanol) was added. The intensity of the colour was read in a Beckman DU spectrophotometer at 440 mu using a light path of 1.0 cm. The standard curve for palmitic acid i s shown in Fig. 3. In the author*s opinion, this method i s very much superior for long chain fatty acid determination Lto. : the microtitrimetric method of Dole (117) which has been used for many years. The conditions for the assay u t i l i z i n g tripalmitin-1-C^ 4 26. e u O •40 * a. E O •30 o >-CO Z •20 Ul o -J < o •10 o. o 0 100 2 0 0 3 0 0 PALMITIC ACID ( m j i e ' q ) FIG. 3 density. Palmitic acid concentration versus optical To 1.0 ml of mixture containing 20 mg albumin and 60 mM phosphate buffer, pH 6.8, was added 6.0 ml chloroform con-taining the indicated amounts of palmitic acid. Extraction and subsequent assay were performed as described i n the text. \ \ 27. were essentially identical to the standard assay which con-tained E d i o l R as substrate, except that the volume of E d i o l R (1:19) used was 0.05 ml instead of 0.1 ml. One mp unit of enzyme activity is defined as that amount which produced 1.0 mjj equivalent FFA/60 min at 37°. Specific a c t i v i t y i s defined as the number of mueqs FFA produced/mg protein in 60 minutes at 37°. Protein was determined by the biuret method (14) and optically by the method of Warburg and Christian (61). Preparation of Substrates — Monoolein suspension was prepared by heating 570 mg monoolein in 32 ml 0.25 M sucrose containing 5% acacia (pH 7) at 70°. The mixture was homogenized in a Servall Omnimixer at maximum velocity for 30 seconds at 70°, then slowly cooled with the omnimixer operating at a lower velocity. This procedure gave a satisfactory suspension which remained stable for a considerable length of time. Similarly, very stable suspensions of tripalmitin (25 mg/ml)... and monostearin (50 Lieq/ml) were prepared at pH 7.0. The substrate preparations were stored at room temperature. E d i o l R labelled with tripalmitin-l-C^" 4 was prepared by carefully evaporating a suitable aliquot of toluene contain-ing about 5 u curies activity, and mixing the residue with 2.5 ml of a 1:19 dilution of E d i o l R at 60° for at least 60 minutes on a mechanical shaker. Shaking was continued while the mixture was cooled to room temperature. Measurement of Hydrolysis of T r i p a l m i t i n - l - C 1 ^ — Hydrolysis of t r i p a l m i t i n - l - C 1 4 was followed by removing an aliquot of 28. the chloroform layer obtained during the standard FFA assay, and evaporating i t to dryness in a centrifuge tube. The residue was quantitatively taken up in small volumes of chloroform and spotted on a thin layer chromatograph plate, using s i l i c a gel GF (Merck) as the adsorbent. The plates were developed with freshly d i s t i l l e d chloroform, then exposed to iodine vapour to allow detection of the FFA and glycerides. The triglyceride spots were consistently and clearly defined, moving just behind the solvent front. The lower glycerides and the fatty acids did not separate consistently nor com-pletely to allow an individual quantitative analysis of these components. Therefore these products of triglyceride hydro-lysis were scraped off together and counted as one component. Fifteen mis of Liquifluor (Nuclear-Chicago) containing 0.4% PPO (2,5-diphenyloxazole) and 0.05% POPOP (p-bis [2-(5-phenyloxazolyl)J -benzene) in toluene was added to the counting v i a l s , the contents thoroughly swirled and the radioactivity counted in a Nuclear-Chicago s c i n t i l l a t i o n counter. Correction for quenching was made for each v i a l by the channels ratio method. Preparation of Enzyme Extract — A l l ventricular tissues used for the preparation of extracts were obtained from fed, female Wistar rats weighing between 200 and 300 grams. Unless other-wise indicated, a l l hearts were perfused for 5 minutes with Ringer-Tyrode solution, pH 7.4, containing heparin, 20 ug per ml, before homogenization. This procedure effectively removes blood from the tissues and also elutes : a considerable amount of lipoprotein lipase from the heart. According to Robinson and Jennings (46), about 50% of the lipoprotein lipase activity of the heart i s eluted in 60 minutes, and the rate of enzyme release is highest during the f i r s t few minutes. Ventricular tissues pooled from 10 to 15 rat hearts were homogenized for three 1-minute periods at 0° in 5 vol-umes of 0.25 M sucrose containing 0.05 M Tris, pH 7, using a Servall Omnimixer. The homogenate was centrifuged at 105,000 x g for 60 minutes, and the sediment thus obtained was re-homogenized in 507. of the original volume of 0.25 M sucrose solution, pH 7, containing 0.17. Triton X-100. The homogenate was again centrifuged at 105,000 x g for 60 minutes at 0° and the supernatant fluids combined. Four mg sodium acetate per ml were added to the combined supernatant fractions and the pH was adjusted to 5.9 with the dropwise addition of 1.0 N acetic acid. Following equilibration in an ice bath for 20 minutes, the precipitate was collected by centrifuging for 60 minutes at 37,000 x g at 0° and discarded. The supernatant f l u i d was taken to pH 5.2 with 1.0 N acetic acid and equilibrated in an ice bath for 30 minutes before centrifuging at 37,000 x g for 60 minutes at 0°. The preci-pitate was taken up in 0.25 M sucrose, pH 7. The activity of the pH 5.2-5.9 fraction was quite unstable to freezing and thawing, about 50% of the activity being lost after overnight storage at -20°. Therefore much of the work described later in the text was performed with freshly prepared pH 5.2-5.9 enzyme extracts. 30. RESULTS I Perfusion Studies It is common knowledge that the isolated mammalian heart continues to function for hours when perfused with "physio-logical" solutions (e.g. Tyrode's) which contain glucose as energy source. This extremely useful technique has been employed by numerous investigators to study the various bio-chemical aspects of cardiac metabolism, and has recently been used widely in studies on l i p i d metabolism by the heart. Our f i r s t studies were designed to determine whether hearts per-fused without substrate were capable of u t i l i z i n g endogenous triglycerides as energy source. During the course of this work, Shipp et al»(35) reported that phospholipids were utilized to a much greater extent than were triglycerides by the isolated perfused rat heart perfused without substrate. This seemed rather unlikely, considering the generally accepted view that phospholipids play primarily a structural role in mammalian tissues. Furthermore, Denton and Randle (37) have reported recently that no decrease in phospholipid levels occurred during substrate-free perfusions, whereas tissue triglyceride contents decreased. In the present studies, phospholipid levels were not measured. Instead an attempt was made to compare the relative rates of u t i l i z a t i o n of endogenous triglycerides with that of glycogen under varying experimental conditions. Fasting has been shown to increase the glycogen content 31. of rat hearts (8) and the triglyceride content of guinea pig hearts (118)• Fasting also increases lipoprotein lipase activity in rat hearts (50). We have included studies on hearts from animals fasted for 3 days before sacrifice. The purpose of fasting the animals was two-fold: F i r s t , to pro-vide a larger store of endogenous substrates so that the hearts could be perfused for longer periods of time, arid second, to examine the possibility that increased cardiac lipase activity due to fasting might increase the rate of tissue triglyceride breakdown and u t i l i z a t i o n during perfusion. The effects of epinephrine and heparin upon the u t i l i -zation of endogenous substrates were also investigated. The glycogenolytic action of epinephrine is well known. How-ever, the possibility that epinephrine might also cause an increase in l i p o l y t i c activity in cardiac tissue, as i t does in adipose tissue seemed most attractive, and was used in the perfusion studies to test this possibility. The release of lipoprotein lipase activity from cardiac tissue slices and from isolated heart by heparin has been demonstrated (46). This suggested, then, that the perfusion of rat hearts with heparin might cause a reduction in tissue lipase content, resulting in a decreased rate of disappearance of endogenous triglycerides. I n i t i a l ventricular levels of glycogen in the hearts of fed rats were 10.1 - 1.3 mg/g dry weight, and 20.9 - 0.8 mg/g dry weight in hearts from 3-day fasted rats ("CONTROL" panel, Fig. 4). The glycogen levels in hearts from fed rats decreased 0 5 10 15 20 25 3 0 PERFUSION TIME (min.) FIG. 4 Tissue glycogen contents in hearts-from fed (——) and fasted (——) rats during sub strate-free perfusion. Control hearts were perfused for the indicated period of time with substrate-free Krebs-Ringer bicarbonate media, pH 7.4 at 37°. Epinephrine, 0.2-1.0 ug/min, and heparin, 60 ug/min, were injected into the perfusion system just above the cannula. Tissue glycogen contents are shown as the means ± standard error of. the mean of the number of observations in parenthesis (vertical bars). The horizontal bars represent the mean time and range over which hearts were collected. 33. at a faster rate than those from fasted rats when perfused with substrate-free medium. In contrast to hearts from fasted rats, hearts from fed animals generally developed arrhythmia and decrease in contractile force within 10 minutes of perfusion. On the other hand, hearts from fasted rats maintained good rhythm and contractions for much longer periods of time, often longer than 25 minutes. As might be expected, the addition of epinephrine to the perfusion medium caused a rapid depletion of tissue glycogen, which was especially noticeable in the hearts from fasted animals ( see "EPINEPHRINE" panel, Fig. 4). Furthermore, epinephrine appeared to have caused a more complete depletion of tissue glycogen, as compared with the control series. In fact, the glycogen content after 10 minutes of perfusion in some of the epinephrine treated (fed) rat hearts was barely detectable. The effect of epinephrine on cardiac function was to increase both rate and force for the f i r s t 3-5 minutes, followed by a rapid decline in cardiac function. Within 8-12 minutes, most of the epinephrine treated hearts were v i r t u a l l y non-functional. Arrhythmias were very frequently encountered in the presence of epinephrine. It should perhaps be noted that when the perfusate from the epinephrine-treated fasted series was assayed for glucose, none could be detected. Addition of heparin to the perfusion medium appeared to have l i t t l e effect on the rates of glycogen depletion in both fed and fasted rat hearts ("HEPARIN" panel, Fig. 4). Whereas fasting caused a significant increase in cardiac 34. glycogen content, there was no significant increase in tissue triglyceride level in hearts obtained from fasted rats. This observation i s in contrast to the observations of Wittels (118) who found increased triglyceride levels in hearts of fasted guinea pigs. Perfusion of about 30 minutes duration (Fig. 5) appeared to cause some decrease in triglyceride levels in both fed and fasted rats. However, considering the relatively large variation in the tissue triglyceride values, and the small population of rats used, i t i s suggested that i f any changes in triglyceride levels did occur, these changes were probably not very significant during the 27-30 minutes of perfusion. No decrease in triglyceride levels were noted in hearts perfused with epinephrine or with heparin. In fact, an apparent increase in triglyceride levels appeared. This apparent increase in tissue triglyceride levels in the epine-phrine and heparin perfused series ("EPINEPHRINE" and "HEPARIN" panels, Fig. 5) i s most d i f f i c u l t to interpret. It is incon-ceivable that triglycerides were synthesized under these perfusion conditions. It seems more l i k e l y that some factor may have been produced as a direct result of perfusion with epinephrine and heparin, giving rise to anomalously high values for triglycerides. The apparent decrease in tissue triglyceride levels observed during perfusion of the control series must be sub-stantiated with more data. Since more potential energy i s contained per weight of triglycerides than i s contained in glycogen, the decrease in triglycerides w i l l probably not be 22 V- 20 * • r 18 ^ 16 E . 14 10 U J 3 CO <2 4 l - . 2 0 CONTROL (4) (8) UJ o £ ,2|(i2) T T u .-> -I o DC 8 . 1 1 ® ' 6 (6) 10 15 20 25 30 0 5 10 15 20 25 30 PERFUSION TIME (min.) FIG. 5 Tissue triglyceride contents in hearts from fed ( during substrate-free perfusion. Perfusion conditions as described under Fig* 4. 22 20 HEPARIN (8) (6) 10 15 20 25 30 ) and fasted (- — -) rats 36. evident immediately under these relatively short perfusion conditions. Hence, in order to demonstrate conclusively that tissue triglycerides do decrease during substrate-free per-fusions, i t would l i k e l y be necessary to perfuse hearts for longer periods, perhaps up to 60 minutes. However, i t was found extremely d i f f i c u l t to maintain hearts in good working order for periods much longer than 30 minutes under the perfusion conditions used in these experiments. These results unfortunately shed l i t t l e light upon the relative importance of endogenous triglycerides to the isolated perfused heart. It is tentatively concluded that the immediate endogenous source of myocardial energy is derived from the breakdown of tissue glycogen when hearts are perfused with substrate-free media, at least during the i n i t i a l phases of perfusion. Per-haps decreased levels of triglycerides become apparent after longer periods of time. II Cardiac Lipases Triglycerides in blood (in the form of chylomicrons and very low density lipoproteins) are taken up intact, i.e. with-out prior hydrolysis, by cardiac c e l l s . When one considers that perhaps 90% of the circulating lip i d s which are extracted by the heart i s in the form of triglycerides, i t i s obvious that the role of cardiac lipases must be extremely important for providing a constant source of free-fatty acids for oxi-dation by the myocardium. In many tissues, the sequence of triglyceride hydrolysis to fatty acids proceeds enzymatically in the following manner: 37. triglycerides »-diglycerides monoglycerides »- FFA, where the i n i t i a l step is considered rate-limiting. In cardiac tissues, an enzyme system exists which hydrolyzes triglycerides to diglycerides. This triglyceride-hydrolyzing enzyme, lipoprotein lipase, probably hydrolyzes diglycerides to mono-glycerides as well. Another lipase, similar to the epinephrine-sensitive lipase of adipose tissue (107), has been reported to exist in rat hearts (13). The existence of a lipase which is specific for diglycerides has not yet been shown in heart or in any other tissue, although there is some evidence to strongly indicate that lipoprotein lipase also attacks digly-cerides. On the other hand, the presence of monoglyceride-sp l i t t i n g lipases has been demonstrated in a number of tissues, including adipose tissue (6, 16), liver (55), and the intes-t i n a l mucosa (56-59), but i t s presence has not been shown in cardiac tissue. The general procedure for studying any enzyme i s to determine f i r s t whether i t s activity exists in whole homo-genates or in intact systems. The next step i s to purify i t as a discrete entity so that i t s properties may be better studied without the problem of contamination by other enzymes possessing similar properties. This general approach was followed in the present work for investigating the l i p o l y t i c a c t i v i t i e s in rat hearts. Although numerous enzymes have been extensively puri-fied, and many even crystallized, most lipases have resisted purification. In fact, no lipase has yet been crystallized. 38. Therefore, studies on most lipases up to the present time have usually been done on crude extracts of tissues, leading to much confusion in the literature with respect to some of the properties of lipases. Another major obstacle in the study of lipases is the technical d i f f i c u l t y of preparing suitable substrates for the enzymes. For example, there are no standard triglyceride substrates for triglyceride-hydro-lysing lipases. The fatty acid moiety of a triglyceride substrate may be saturated, unsaturated, long, medium or short chain, and the physical nature of the substrate may be an oil-in-water emulsion or a crude glyceride suspension in a suitable aqueous buffer. Finally, a l l known methods of assaying l i p o l y t i c activity are considerably more laborious and technically cumbersome than most commonly used enzyme assays. A l l these reasons contribute to the fact that the study of lipases has lagged far behind the study of enzymes in other areas of the biological system. In spite of the d i f f i c u l t i e s anticipated, an attempt was nevertheless made to study the lipases of cardiac tissue, with particular emphasis on lipases other than lipoprotein lipase. A. EXISTENCE OF NaF-INHIBITED LIPASE Rat hearts were perfused for 5 minutes with Krebs-Ringer bicarbonate solutions containing 20 ug/ml heparin. The ven-tricular tissues were pooled and homogenized in 10 volumes of 0.25 M sucrose, pH 7.0 at 0-4° with the aid of a Potter-Elvehjem Homogenizer. The crude homogenate was f i l t e r e d 39. through cheesecloth and used directly for assay of l i p o l y t i c activity at pH 6.8, using E d i o l R (1:19) as substrate as described in the Experimental section. A rapid liberation of free fatty acids occurred, as may be seen in Fig. 6, and the enzyme activity was proportional to incubation time up to 30 minutes. When similar experiments were performed in the presence and absence of 0.2 M NaF, again using E d i o l R as substrate, i t was noted that a constant inhibition of about 40% occurred throughout the course of the reaction (Fig. 7). This experi-ment was performed at a pH suboptimal for lipoprotein lipase and without pre-activating the Ediol with serum. Under these conditions, the activity of lipoprotein lipase would be minimized. Since NaF i s known not to be an inhibitor of lipoprotein lipase, the observed 407. inhibition in the crude system suggested that there indeed was an active lipase or lipases present in cardiac tissue in addition to lipoprotein lipase. Inhibition by NaF would indicate that the enzyme might be similar to the non-lipoprotein lipase of adipose tissue described by Rizack (107). At this point, i t must be emphasized that Ediol 8- was used as substrate for measuring l i p o l y t i c a c t i v i t y in these experiments. The major advantage of this emulsion is that i t i s an extremely smooth and stable oil-in-water t r i g l y -ceride preparation. However, a serious disadvantage of E d i o l R as substrate is that i t contains monostearin as a stabilizing agent, and this monostearin could well serve as 40. INCUBATION TIME (mln.) FIG. 6 Time course of l i p o l y t i c activity. The reaction mixture contained 60 mM phosphate buffer, pH 6.8, 20 mg purified bovine serum albumin, 0.10 ml of 1:19 dilution of Ediol, 0.04 ml whole homogenate in a total volume of 1.0 ml. Incubation was at 37° in a Dubnoff Metabolic Shaker. Each reaction mixture contained 0.42 mg protein. 41. FIG. 7 Inhibition of l i p o l y t i c activity in crude heart homogenates by NaF. The incubation mixture contained a l l components.' of the standard assay as described in the text, except that 0.15 ml of 1:19 dilution of Ediol was used as substrate. Curve "A" - control. Curve "B" - NaF, 0.2 M included in reaction mixture. Each reaction mixture contained 0.21 mg protein. The homogenate was stored overnight at -20° and thawed before use. 42. substrate for a l i p o l y t i c enzyme. Experiments were therefore performed in order to explore further the nature of the NaF-inhibited l i p o l y t i c activity in cardiac tissue by using substrates prepared from monoglycerides or triglycerides only, When monostearin (suspended in 5% acacia solution) was used as substrate, i t was rapidly hydrolyzed by the heart homogenate (Fig. 8, Curve A). Activity again was propor-tional to time. Of particular interest was the observation that in the presence of 0.2 M NaF, a constant 40% inhibition of l i p o l y t i c a c t i v i t y was obtained (Curve B). This degree of inhibition was almost identical to that observed when similar experiments using E d i o l R was performed (Fig. 7), thus strongly suggesting that the NaF-inhibited l i p o l y t i c activity measured with Ediol^ as substrate may have been due to the hydrolysis of i t s monostearin component. Furthermore, when tripalmitin alone was used as substrate, the l i p o l y t i c a c t i -v i t y was so small as to make accurate and reliable l i p o l y t i c measurements exceedingly d i f f i c u l t (Curve C). NaF (0.2 M) did not appear to inhibit the hydrolysis of tripalmitin, although the low activ i t i e s observed made the interpretation of the data d i f f i c u l t (Curve D). When, however, the incubation mixture was supplemented with rat serum, a rapid hydrolysis of tripalmitin occurred. The data in Table I shows a greater than 12-fold increase in l i p o l y t i c activity when the t r i p a l -mitin substrate was presented to the enzymes in the form of a lipoprotein complex. This experiment clearly indicated not only the presence of lipoprotein lipase activity in 4 3 . INCUBATION TIME (mln.) FIG. 8 Hydrolysis of monostearin and tripalmitin, and effect of NaF. Line "A" represents the time course of l i p o l y t i c a c t i -v i t y of the whole homogenate when the substrate was monostearin (5 ueq/ml) in the standard assay. Line "B" represents inhibition of monostearin hydrolysis by 0.2 M NaF. Line "C" represents the l i p o l y t i c activity of the homogenate (same volume used as in "A") when the standard incubation mixture contained 2.5 mg tripalmitin as sub-strate. Line "D" represents the hydrolysis of tripalmitin in the presence of 0.2 M NaF. The hydrolysis of tripalmitin was consistently so small that accurate l i p o l y t i c measure-ments were extremely d i f f i c u l t . Each reaction mixture contained approximately 0.42 mg protein. TABLE I Effect of rat serum on the hydrolysis of tripalmitin by par t i a l l y purified extract of cardiac tissue. The complete system contained 20 mg purified bovine serum albumin, 50 mM Tris buffer, pH 8.5, 2.5 mg t r i -palmitin which had been pre-incubated for 30 minutes at 37° with 0.1 ml rat serum, 0.1 ml of enzyme extract, and sufficient water to make a f i n a l volume of 1.0 ml. Reaction mixture Lipolytic Activity mueq FFA/60 min at 37° Complete system Complete system without serum 99 8 45. cardiac extracts, but also demonstrated the marked dependence of this enzyme on serum-activated triglycerides. More impor-tant, the data provided additional evidence that an enzyme other than lipoprotein lipase exists in rat hearts. The enzyme appeared to hydrolyze monoglycerides with greater f a c i l i t y than triglycerides. The a b i l i t y of the lipase to liberate free fatty acids from E d i o l R i s l i k e l y due to i t s action on the monostearin component of this preparation. The enzyme already seemed similar to that reported by Bjorntorp and Furman (13) who used E d i o l R as substrate. The pos s i b i l i t y existed that the activity these authors had measured was actually the hydrolysis of monoglyceride. Further studies were therefore necessary to c l a r i f y the source of fatty acids arising from Ediol . One could accomplish this by following the hydrolysis of either C l 4-labelled monoglycerides or C^-triglycerides when these glycerides were added to E d i o l R and subjected to lipase activity in heart extracts. Since labelled long-chain monoglycerides were not readily available, t r i p a l -mitin- 1-C*4 was used. The rate of hydrolysis of tripalmitin-1-C"*"4 and the rate of total fatty acids released were measured simultaneously. It was assumed that fatty acids produced in the absence of tripalmitin-1-C^ 4 hydrolysis must have been derived from the monoglyceride component of Ediol . Further-more, i t was thought that the effects of known lipase inhi-bitors could be more clearly observed in such a system. As may be seen in Fig. 9, the hydrolysis of trlpalmitin-l-C^ 4 in E d i o l R occurred at an appreciable rate in the presence of 46. FIG. 9 Time course of E d i o l h y d r o l y s i s i n the presence of l i p a s e i n h i b i t o r s . L i n e s "A", "B", and "C" represent the time course of t o t a l f a t t y a c i d s produced d u r i n g the experimental c o n d i t i o n s as o u t l i n e d below. Li n e s "a""and "b" represent the r a t e of t r i p a l m i t i n - 1 - C l 4 h y d r o l y s i s d u r i n g the same i n t e r v a l . L i n e s "A" and "a" - The r e a c t i o n mixture contained 120 mg bovine serum albumin, 50 mM T r i s b u f f e r , pH 8.5, 0.3 ml r a t serum, 0.3 ml of 1:1.9 d i l u t i o n E d i o l c o n t a i n i n g t r i p a l m i t i n -l - C 1 ^ (0.5 p c u r i e / n i l ) , 0.6 ml 6000 x g r a t heart supernate, and s u f f i c i e n t water to make 6.0 ml. Incubation was performed at 37° i n a Dubnoff Metabolic Shaker and 1.0 ml a l i q u o t s removed at the i n d i c a t e d time i n t e r v a l s f o r the assay of t o t a l f r e e f a t t y a c i d s produced ( l i n e "A") and a l s o t r i p a l m i -t i n - l - C 1 4 hydrolyzed ( l i n e " a " ) . L i n e s "B'Vand "b" - Reaction c o n d i t i o n s were e s s e n t i a l l y 47. the same as indicated above for "A" and wa", except that 50 mM Tris buffer pH 7.0 was used and the system also contained NaCl, 0.5 M. Line "C" - Reaction conditions were as for "B" and "b", except that the system also contained"NaF, 0.2 M. - . The rate of tripalmitin-l-Cl4 hydrolysis (line "a") was proportional to incubation time for 90 minutes. 48. rat serum (line "a"). The total fatty acids produced during this period was also appreciable (line "A"). In the presence of 0.5 M NaCl (and absence of serum), the hydrolysis of t r i -palmitin- 1-C 1 4 was completely abolished (line "b"). This indicated that under these conditions (i.e., 0.5 M NaCl, no serum), lipoprotein lipase activity against triglycerides was completely inhibited. However, even under conditions where the hydrolysis of tripalmitin-1-C 1 4 was completely inhibited, FFA release was s t i l l evident in the system (line "B"). Hence, thi s ' l i p o l y t i c activity was attributed to the hydrolysis of monostearin in E d i o l R . If this were the case, the addition of 0.2 M NaF to the reaction mixture should result i n further inhibition of l i p o l y t i c activity, and as may be seen, this was indeed observed (line "C"). This experiment therefore gave additional support to the original observation that when E d i o l R was used as substrate in the "non-activated" state, a large proportion of free fatty acids liberated was derived from the hydrolysis of i t s monoglyceride component. It also supported the view that i t i s the hydrolysis of the mono-glyceride component of E d i o l R which is inhibited by NaF. It was noted that the rate of monoglyceride hydrolysis (line "B") did not change after about 25 minutes of incubation. This consistent observation in these experiments was due to the fact that under the conditions of the assay, the amount of monoglyceride was limiting the reaction. E d i o l R contains approximately 1.5% monostearin. This concentration of mono-stearin i s equal to 40 peq/ml. In those experiments u t i l i z i n g 49. tripalmitin-l-C 1^, each 1.0 ml of reaction mixture contained 0.05 ml of a 1:19 dilution of E d i o l R . Hence the total amount of monostearin available as substrate was about 100 mueqs. Considering the noticeable and inevitable shrinkage in volume of E d i o l R during storage over a two year period, i t i s not unreasonable to assume that about 150 mpeqs of monostearin were perhaps available as substrate per reaction tube. This would explain the consistent plateauing of reaction rate observed at around 25-30 minutes incubation in those experi-ments which were designed specifically to show only monogly-cerides being hydrolyzed. The results of these experiments clearly demonstrated that the l i p o l y t i c activity in rat hearts possessing proper-ties different from lipoprotein lipase was an active NaF-inhibited enzyme which rapidly hydrolyzed the monostearin component of E d i o l R . Since E d i o l R has been so widely used in the study of lipase, especially in adipose tissue, one wonders i f results have not been occasionally misinterpreted. B. PREPARATION OF PARTIALLY PURIFIED -EXTRACT FROM HEART TISSUE The presence of a NaF-inhibited monoglyceride-hydrolyzing l i p o l y t i c system in rat hearts having been established, i t was considered essential that the enzyme be isolated so that i t s properties could be better understood. As anticipated, the satisfactory purification of this enzyme proved extremely d i f f i c u l t . Some of the problems encountered during the attempt to 50. purify the enzyme w i l l be noted here, but a detailed discussion w i l l be reserved for the f i n a l Discussion section. F i r s t , about 75-85% of the enzyme activity was bound to the particulate frac-tions of the c e l l s . Attempts to solubilize the enzyme using a number of standard techniques were in most instances unsuc-cessful. For example, preparations of acetone powders, repeated freezing and thawing, deoxycholate treatment, sonica-tion at 9 and 20 kilocycles for various periods of time, sonication combined with deoxycholate treatment were tried in vain. The technique which gave reasonably satisfying results was the use of 0.1% Triton X-100, a non-ionic synthetic deter-gent. When a 105,000 x g pellet was re-homogenized with Triton X-100, a 2-to 3-fold increase in the amount of enzyme activity was noted in the subsequently obtained 105,000 x g supernatant f l u i d . This technique solubilized about 30-40% of the total enzyme activity in cardiac c e l l s , and was there-fore adopted as the basis for further attempts to purify the enzyme. Purification of the enzyme from the 105,000 x g super-natant f l u i d was also extremely d i f f i c u l t . Again a number of clas s i c a l techniques of enzyme fractionation were employed. Repeated attempts using high temperatures, ethanol precipita-tion methods, calcium phosphate gels, Sephadex G-200 columns and zinc-ethanol treatment failed to give a satisfactory purification. However, both ammonium sulfate fractionation and the isoelectric precipitation methods did provide a small measure of purification. The isoelectric precipitation tech-51. nique was adopted as the second step in the proposed further purification of the enzyme. Treatment of the extract (obtained by isoelectric precipitation) with Sephadex G-200 and ammonium sulfate were tried but the enzyme resisted further purification. A major obstacle to purification was the i n s t a b i l i t y of the l i p o l y t i c system in the isoelectric precipitate fraction, particularly to freezing and thawing. Since a considerable amount of time had already been expended in efforts to solu-b i l i z e and purify the enzyme, i t was decided to use the partially purified isoelectric precipitate fraction for the study of the enzyme. The acid precipitation method gave about a 3 to 4-fold purification over the combined supernatant frac-tion, and a yield of approximately 457.. The overall yield, however, was only 107. when based on the total activity of the whole homogenate, owing essentially to the extremely insoluble nature of the enzyme. It should be mentioned that the degree of purification obtained was about the same when either E d i o l R or monoolein was used as substrate for measuring l i p o l y t i c activity. The partial purification obtained is shown in Table II. Lipolytic activity of the isoelectric precipitate was directly proportional to enzyme concentration (Fig. 10) over a range of protein which constituted a reliable assay. C. PROPERTIES OF CARDIAC MONOGLYCERIDE-HYDROLYZING ENZYME Using the p a r t i a l l y purified isoelectric precipitate preparation (hereafter referred to as "partially purified extract") of rat heart, a number of experiments were performed 52. TABLE II Partial purification of cardiac monoglyceride-splitting enzyme. The acti v i t i e s in the f i r s t and second high speed super-nate fractions are included to indicate the increase in solubilization obtained with Triton X-100. The standard lipase assay was employed with 15 ueqs monoolein as substrate. One mp unit of enzyme activity i s that amount which produced 1.0 mpeqs FFA/60 min/37°. Specific activity is defined as mu units enzyme activity/mg protein/60 min/37°. FRACTION TOTAL ACTIVITY SPECIFIC ACTIVITY (No. 1) 105,000 g supernate 220,000 956 (No. 2) " " obtained with Triton XrlOO 646,800 3480 COMBINED 105,000 supernates 766,800 1980 pH 5.2-5.9 isoelectric precipitate 333,333 7000 53. VOLUME OF EXTRACT ADDED (mla) FIG. 10 FFA production as a function of enzyme concen-tration with monostearin as substrate. The standard assay was employed, and the enzyme used . was the par t i a l l y purified isoelectric preparation* 54. to study some of the properties of this enzyme. It was hoped that information obtained from such studies would greatly assist in the assignment of a possible physiological role of the enzyme in cardiac tissues. 1. Albumin Requirement — Under physiological conditions, long chain free fatty acids are normally transported in the blood as free fatty acid-albumin complexes. This binding of free fatty acids to albumin effectively solubilizes a con-siderable amount of otherwise insoluble FFA»s which have been released into the circulation from adipose tissues. For the same reason, albumin has frequently been added to li p o l y t i c reaction mixtures to function as a FFA-acceptor in the i n vitro system. As may be seen in Fig. 11, the require-ment for albumin in the assay mixtures was not absolute, but the presence of more than 12 mg albumin increased the a c t i -v i t y about 2-fold. Routinely, 20 mg of purified bovine serum albumin was included in the assay system. 2. pH-Qptimum — When monoolein was used as substrate, the activity of the enzyme extended over a wide pH range (pH 6.0-9.0) with the maximum activity exhibited near pH 6.5-7.0 (Fig. 12). This observation i s consistent with the report of Bjorntorp and Furman's (13) that a sodium fluoride-inhibited l i p o l y t i c activity having a pH optimum near pH 6.8 existed in rat hearts. The pH optimum of lipoprotein lipase i s near 8.5. Since, in the experiment, no triglyceride or diglyceride substrate and no serum was present, and since the reaction was performed in the presence of 0.5 M NaCl, this enzyme can be o 100 > cr a. < 50 0 4 8 12 16 20 ALBUMIN ADDED (mg) FIG. 11 Effect of concentration of albumin on lipo-lytic, activity. Incubation was carried out according to the standard assay, using 0.1 ml of 1:19 dilution of Ediol as substrate, except that the albumin concentration was varied as indicated. The enzyme used was the pa r t i a l l y purified pH 5.2-5.9 preparation. . 56. pH FIG. 12 Hydrolysis of monoolein by part i a l purified lipase.as a function of pH. The incubation mixture contained 15 peq monoolein, 20 mg purified bovine serum albumin, 0.5 M NaCl, enzyme, 0.08 M Tris and 0.08 M phosphate buffers at the pH*s indicated. The incubations were performed in a total volume of 1.0 ml for 60 minutes at 37° on a Dubnoff Metabolic Shaker. 57. readily distinguished from lipoprotein lipase. 3. Temperature Optimum — The optimum temperature for the in vitro enzymatic hydrolysis of monostearin was near 45° (Fig. 13). This optimum is beyond the normal physiological range expected of any enzyme in biological systems. There-fore, the significance of this observation can only be extended to the enzymatic hydrolysis of monostearin jLn vitro, and is of academic interest only. However, i t further empha-sizes the similarity between this enzyme and the monoglycer-ide-splitting enzyme from adipose tissue because Vaughan jet _al.(6) has shown that this enzyme has a temperature optimum of 45°. 4. Effect of Physical State of Substrate — One of the c r i t e r i a used! for distinguishing between true lipases and non-specific esterases i s that the former does not normally attack esters in aqueous solution (109). In order to study the nature of the l i p o l y t i c activity in the part i a l l y puri-fied extract, the enzyme was added to various concentrations of monoolein. A serial dilution of monoolein was made such that at low concentrations, the reaction mixture was optically clear, presumably because the monoolein was in true solution. At higher concentrations, the substrate was insoluble and in the form of a suspension. Careful visual examination of a series of reaction tubes prior to addition of enzyme indicated that the turbidity of the reaction mixture became apparent at monoolein concentrations in the region of 1.5 or 2.25 ueq/ml. When the l i p o l y t i c activity was measured in the usual manner, 58. 25 30 35 INCUBATION 40 45 50 55 TEMPE RATURE FIG. 13 Effect of temperature on cardiac monoglyc.eride« hydrolyzing lipase activity. The standard assay was employed using monostearin as substrate, except that the incubations were performed at the temperatures indicated. Control tubes were also incubated, and the enzyme added after the reactions were stopped. 59. i t was observed that a marked increase in activity occurred near 2.0-4.0 ueq/ml monoolein concentration (Fig. 14). It is readily conceded that the visual estimation of turbidity gave only a rough approximation of the actual physical state of the monoolein substrate in aqueous media. Nevertheless, the marked increase in l i p o l y t i c activity over a relatively narrow monoglyceride concentration range suggested strongly that the enzyme is a true lipase, and not a simple non-specific esterase. 5. Effect of Some Enzyme Inhibitors on Monoglyceride- hydrolyzing Lipase Activity — Many enzyme inhibitors have been employed in order to identify the active sites and to study the mechanism of action of enzymes. It appears, how-ever, that the use of inhibitors in the study of lipases has been largely restricted to distinguishing one lipase activity from another. Even when used for this purpose, many inconsis-tencies are noted in the literature with respect to the effect of inhibitors on mammalian tissue lipases, presumably due to the use of crude enzyme preparations and grossly impure sub-strates. I t was hoped that the use of some known lipase inhibitors in this study would yield additional information as to whether the cardiac monoglyceride-splitting lipase was similar to those which have been reported to exist in adipose tissue and in the intestinal mucosa. The results of these studies are presented in Table III, including the data obtained by other investiga-tors for reference. 60. 1 2 3 4 5 6 7 8 9 10 // 14 MONOOLEIN CONCENTRATION (ueq/ml) FIG. 14 Effect of monoolein concentration on the activity of the part i a l l y purified cardiac lipase. The reaction mixture contained a l l components of the standard assay, except that the monoolein concentration was varied as indicated and 0.5 M NaCl was included. TABLE III Inhibition of Monoglyceride-splitting Lipase by Various Compounds The standard assay was used with 15 ueq monoolein as substrate, except that 0.5 M NaCl was included in a l l the reaction mixtures. The various inhibitors were pre-incubated for 15 minutes at 37° with the partially purified enzyme preparation prior to addition of substrate. COMPOUND THIS STUDY cone. % inhib. Kupieki (16) cone. % inhib. Vaughan et al« (6) cone. % inhib. Strand et al» (15) cone. 7o inhib. Pope et al» (597"~ cone. 7. inhib. NaF 0.15M 75 0.20M 45 0.20M 84 1 x 10"^ 100 Isopropanol DFP (0.0 2ml/ml) 2 x 10""% 0 70 1 x 10-% mm mm 39 (0.025ml/ 0 ml), 5 x 10 % 97 (0.025ml/ ml) 0 mm mm mm mm 1 X 10"% 100 EDTA 5 x 10"% 0 5 x 10"% 0 — Protamine sulfate 400 ug/ml 0 300 ug/ml 0 *• mm mm mm mm N-Ethyl Maleimide 1 x 10"% 25 1 x 10*"% 33 — mm mm mm mm Iodoacetic acid 1 x 10"% 0 1 x 10"% 0 mm mm «m mm mm mm mm P-chloromercuri-benzoate 1 x 10"% 79 mmm» tm Wm mm - — mm mm 1 x 10"% 0 Substrate Monoolein Monoolein Monostearin Monoolein Monoolein Monoglyceride-splitting lipase source Partially purified from rat heart Partially purified from rat adipose tissue Crude extract adipose tissue Crude extract adipose tissue Highly purified from rabbit intestinal mucosa 62. The most potent inhibitors of the monoglyceride hydro-lyzing activity were NaF and DFP, and the effects observed were in good agreement with those seen for monoglyceride-hydrolyzing lipases of other tissues. The mechanism of inhibition by NaF i s not known, but the relatively high concentrations normally required to show this inhibition restricts the usefulness of this compound, except perhaps to distinguish between monoglyceride-hydrolyzing activity and lipoprotein lipase activity. On the other hand, DFP inhibited the enzyme at a relatively low concentration. DFP i s known to combine irreversibly with the -OH function of enzymes (e.g. esterases), thus blocking the active site of the enzyme. In this respect, the monoglyceride-hydrolyzing enzyme appears to resemble an esterase. The complete absence of inhibition by protamine sulfate (400 ug/ml) was expected since any lipoprotein lipase a c t i -v i t y which may have been present in the extract would have been effectively inhibited under the conditions of the assay. The presence of EDTA (5 x lO'^M) did not inhibit the activity, which l i k e l y indicates that no positively changed metal cations were required for activity. The alkylating agents, N-ethylmaleimide and iodoacetate, used at the same concentrations would have been expected to give similar results. Therefore the inhibition observed with N-ethylmaleimide, but not with iodoacetate is d i f f i c u l t to reconcile, although i t must be noted that Kupieki (16) obtained essentially similar results with these compounds. The effects 63. of these inhibitors on the monoglyceride-splitting lipase activity of heart tissue is in good agreement with those observed for the monoglyceride-hydrolyzing enzymes studied in other tissues, and further indicated the similarity of these a c t i v i t i e s . 6. Intracellular Localization — It has been previously mentioned that the enzyme was bound to tissue particles. The intracellular distribution of the monoglyceride-splitting lipase was investigated in order to provide some evidence as to i t s possible physiological function in cardiac tissue. As shown in Table IV, about 85% of the activity was bound to the particulate components of the c e l l , about 50% of the total activity being located in the nuclear fraction. In contrast, when tripalmitin was used as substrate, the only cellular fraction to exhibit any l i p o l y t i c activity was the microsomal fraction (Table IV), although the activity was so low as to make accurate determinations d i f f i c u l t . In these experiments, 0,5 M NaCl was included in the assay system in order to inhibit lipoprotein lipase activity. The particle-bound nature of the monoglyceride-hydrolyzing lipase in heart tissue is similar to that reported recently by Pope et: a l . (59) for the purified monoglyceride lipase of intestinal mucosa, in which only about 10% of the total activity was locatedlin the soluble fraction. 64. TABLE IV Distribution of monoglyceride-splitting lipase and t r i -glyceride lipase act i v i t i e s in various fractions of a rat heart homogenate. Ventricular tissues were pooled from 6 rat hearts and homogenized in 12 volumes of 0;25 M sucrose containing 0.05 M Tris buffer, pH 7.0, for two 1-minute intervals using a Servall Omnimixer to obtain the whole homogenate. The nuclear fraction was obtained by centrifuging the homogenate at 250-300 x g for 15 minutes. The l i g h t l y packed sediment was washed with 15 mis 0.25 M sucrose pH 7.0, re-centrifuged and the supernatant fluids combined. The mitochondrial and micro-somal fractions were obtained by centrifuging the combined supernatant fluids at 6000 x g x 15 minutes and 105,000 x g x 60 minutes respectively. A l l procedures were carried out at 0-4°. Standard assay conditions were employed, using either monoolein or tripalmitin as substrates, except that NaCl 0.5 M was also included in each reaction mixture. The assay for triglyceride-splitting activity was performed using 10 times more extract than used for the monoglyceride-split-ting lipase activity, and incubation was for 120 mins at 37°. Enzyme activity i s as defined in the text. MONOGLYCERIDE-SPLITTING LIPASE Fraction Total Activity (mp units) % Whole Homogenate Nuclear Fraction Mitochondrial Fraction Microsomal Fraction 105,000 x g Supernate 536,500 287,500 110,000 75,000 80,600 (100%) 53.5-20.5 14.0 15.0 TRIGLYCERIDE-SPLITTING LIPASE Whole Homogenate Nuclear Fraction Mitochondrial Fraction Microsomal Fraction 105,000 x g Supernate 2590 0 0 2000 0 (100%) 0 0 77.2 0 65. DISCUSSION The object of the perfusion studies was primarily to investigate the po s s i b i l i t y that endogenous triglycerides were utilized by the heart, and to compare this u t i l i z a t i o n with that of glycogen when no exogenous substrates were available. In general, the changes observed in the glycogen levels of rat ventricular tissue during fasting and during substrate-free perfusions were not unexpected. The increase in fasting cardiac glycogen levels deserves further comment. Although Cruikshank (110) observed that pancreatectomy caused a shift in l i v e r glycogen to cardiac glycogen, the f i r s t to observe this phenomenon in fasting was Evans (8) in 1934. Evans found that glycogen content increased in rat hearts from 341 i 15 mg/100 g to 578 ± 14 mg/100 g wet weight after a 48-hour fast. Then Lackey and co-workers (111^113) found that cardiac glycogen increased in the alloxan diabetic state (111), that there was a direct relationship between blood ketone concentrations and cardiac glycogen levels (112, 113). Next, i t was shown by Russell and Bloom (114) that growth hormone was necessary for glycogen to increase in heart during fasting. Lukens (115) suggested that the increase in cardiac glycogen was due to the increased amount of FFA reaching the heart, owing to the action of growth hormone on adipose tissue. This suggestion was later con-firmed by Bowman (116). More recently, Newsholme and Randle (9) and Garland and co-workers (10) showed that ketone bodies, 66. FFA, and pyruvate effectively inhibit the glycolytic enzyme, phosphofructokinase in perfused rat hearts. Parmeggiani and Bowman (11) demonstrated further that the inhibition of phosphofructokinase was due to the increased tissue levels of citrate. The increase in cardiac glycogen content during fasting and i t s immediate decline upon re-feeding was observed during the course of this work. As may be noted in Fig. 15, cardiac glycogen increased over 2.5-fold during seven days of fasting. The immediate depletion of glycogen after one day of re-feeding to the control level indicates a very rapid u t i l i z a t i o n and/or mobilization of glycogen to other tissues of the body, prob-ably the l i v e r . The slower rate of glycogen disappearance from perfused (control) hearts from fasted rats as compared with those from the fed animals suggested that in vivo, the biological mechanisms for u t i l i z i n g l i p i d s was accelerated in fasting rat hearts. In fact, the triglyceride levels of the fasted group in the "control" series appeared to decrease during perfusion, suggesting perhaps, that endogenous triglycerides may have been utilized under these perfusion conditions. However, in view of the unexplainable increases i n t r i g l y -ceride levels found under different perfusion conditions, i t is d i f f i c u l t to state categorically at this time that t r i -glycerides were mobilized and oxidized by the working rat heart. At best, the data is interpreted as only suggestive that this might have been the case. 3 OOf 27-01 ,24-0 2 1-0 -z LU g 18-0 o ^15-0 o 12-0 -9-0 -or 6-0 a: <t _j ZD o (I) U J 3.0 > 2 3 4 DAYS FASTED FIG. 15 E f f e c t of F a s t i n g on M y o c a r d i a l Glycogen Leve1. Rats were fas t e d f o r the d u r a t i o n i n d i c a t e d . A r a t was r e - f e d ad l i b , on the s i x t h day and the v e n t r i c u l a r glycogen content determined a f t e r 24 hours r e - f e e d i n g (dotted l i n e ) . Tissue glycogen contents are shown as the means "i standard e r r o r of the mean. The number of animal used i s shown i n p a r e n t h e s i s . ~ 68. The increased rate of depletion of glycogen from epinephrine-perfused hearts was expected. It was calculated that within about 10 minutes, between 1.8 to 3.0 mg of glycogen actually disappeared from a heart of a fasted rat under the influence of epinephrine. Since i t was d i f f i c u l t to believe that a l l the glucose derived from glycogenolysis under these conditions was being converted to lactate, or even oxidized by the Krebs cycle, the perfusate was analysed for glucose, but none could be detected by the glucose-oxidase method. The mechanism of action of epinephrine on glycogenolysis is perhaps the best understood of a l l hormones investigated. Briefly, epinephrine promotes the formation of cyclic 3",5»-AMP from ATP by stimulation of an enzyme, adenyl cyclase. Cyclic 3',5'-AMP then brings about the conversion of inactive phosphorylase b to active phosphorylase a., mediated by phos-phorylase t> kinase. Phosphorylase a stimulates glycogeno l y s i s , yielding glucose-l-phosphate. This scheme of glycogenolysis has been reviewed by Sutherland (12). The data for the tissue triglyceride series i s most d i f f i -cult to interpret. If the increases in triglyceride content of epinephrine and heparin perfused hearts were real, this would be a most interesting observation, but the idea that triglycerides were actually synthesized during substrate-free perfusions i s most unacceptable to the author. It is more li k e l y that some product of glycogenolysis was extracted into the organic layer during the extraction of tissue l i p i d s . Chromotropic acid would then later combine with the compound 69. to give falsely high values for triglycerides. The experiments with epinephrine and heparin therefore yielded no useful information regarding the breakdown of triglyceride by the heart. On the other hand, triglycerides may have been hydrolyzed and metabolized in the "CONTROL" series. It would appear that in order to demonstrate clearly whether triglycerides are indeed utilized by the heart, per-fusion conditions must be altered somehow so that hearts w i l l function normally for 60 minutes or more, even under substrate-free medium. Further studies using longer periods of perfusion (without epinephrine or heparin) and using a larger population of rats would most l i k e l y be necessary before the disappearance of triglycerides in rat hearts can be conclusively demonstrated. Evidence has been presented which clearly establishes that a l i p o l y t i c activity other than lipoprotein lipase exists in rat myocardium. The properties of the enzyme differ greatly from those of lipoprotein lipase, so that the two enzymes can be distinguished even in crude preparations. In many respects, the properties of the enzyme are similar to mono-glyceride-hydrolyzing lipases reported in adipose tissue (6, 15, 16) and intestinal mucosa (56, 58, 59), and particularly to the Ediol 8--hydrolyzing enzyme system in rat hearts (13) . The pH optimum of the monoglyceride-hydrolyzing enzyme of cardiac tissue agrees with the optimum pH of 6.8 reported by Bjorntorp and Furman (13) for an unspecified cardiac enzyme which actively hydrolyzed E d i o l R . The monoglyceride-hydro-lyzing activity in adipose tissue described by Strand et a l . (15) also has a pH optimum within this range (pH 7.0). On the other hand, Kupieki (16) and Vaughan and co-workers (6) have reported pH optimums of 7.5 and 8.0, respectively, for the same monoglyceride-splitting enzyme in adipose tissue. These inconsistencies may have arisen from the fact that different buffers and monoglyceride substrates were used in these studies. The pH optimum of the cardiac enzyme is sig-nificantly lower than that of the intestinal mucosa lipase which is reported to have an optimum pH of 7.8 (56) and 8.5-9.0 (59). The monoglyceride-splitting enzyme of rat and hog liver has a pH optimum of 8.2 (55). The optimal temperature (45°) for the jLn vitro l i p o l y t i c activity of the heart lipase enzyme is consistent with that found by Vaughan and associates (6) for the enzyme in adipose tissue. No other temperature optima have been reported for further comparison. The relative activity of monoglyceride-hydrolyzing systems against d i - and triglycerides have been described for the intestinal mucosa. Senior and Isselbacher (56) observed the comparative rates of t r i - , d i - , and monoglycer-ide hydrolysis to be 0.3, 1.6, and 91.9 respectively. Pope and associates (59) found the relative rates of hydrolysis of t r i - , d i - , and monoolein to be 0..0, 0.0, and 1.0 respec-tively. McPherson jet a l . (58) observed that monoolein was hydrolyzed 16 times as fast as t r i o l e i n . In adipose tissue, the comparative rates were 1, 43, 73, and 100 for t r i o l e i n , 1,2-diolein, 1,3-diolein, and monoolein respectively as 71. reported by Strand and co-workers (15). Again in adipose tissue, Kupieki's data (16) indicates the relative rates are 0.19, 0.17, and 1.00 for t r i - , d i - , and monostearin. The accumulated evidence indicates that whereas the intestinal mucosa enzyme i s highly specific for monoglycerides, the adipose tissue enzyme i s relatively less specific in this respect. In the present study, no experiments were designed specifically to investigate the activity of the cardiac enzyme with respect to i t s activity against d i - and t r i g l y -cerides. However, from the data collected from occasional experiments in which both tripalmitin and monostearin (or monoolein) were used as substrates, i t may be readily seen that the activity against monoolein and monostearin were significantly higher than against tripalmitin (Table V). Comparison of tripalmitin and monostearin hydrolysis in one experiment (Fig. 8) showed that the rate of monostearin hydro-lys i s was as much as 20-fold greater than that of tripalmitin hydrolysis. No definite statement can be made at present regarding the absolute specificity of the cardiac enzyme. There is no doubt that the enzyme hydrolyzes monoglycerides at an appreciably faster rate than i t does triglycerides, but much more data is required to indicate i t s actual degree of substrate specificity. For example, the hydrolysis rates of mono-, d i - , and t r i o l e i n as compared with the rates of mono-, di - , and tripalmitin would be of interest. If i t i s established that the enzyme is highly specific for monoglycerides only, then i t s action on the 1- and 2-isomers of monoglycerides, TABLE V Relative rates of Hydrolysis of Monoolein, Monostearin and Tripalmitin SUBSTRATE ACTIVITY mueq FFA/60 min RELATIVE ACTIVITY (Tripalmitin-1.0) EXPT. Monoolein 34.0 12.1 A Tripalmitin 2.8 1.0 EXPT. Monostearin 42.0 3.8 B Tripalmitin 11.0 1.0 EXPT. Monostearin 208.0 5.2 C Tripalmitin 38.0 1.0 In Experiment "A", the microsomal fraction of heart tissue was used as the enzyme source. Standard assay conditions were used except that the reaction mixture containing monoolein and t r i o l e i n were incubated in 0.05 M Tris buffer, pH 7.0 and 8.5 respectively. In Experiment "B" and "C", the enzyme preparations were the isoelectric precipitate.fraction and whole homogenate (1:10) respectively. Standard assay conditions were employed. 73. and i t s activity against medium and short chain monoglycerides. must be investigated. Further studies of this nature, of  course, must await a more extensive purification of the enzyme  than has been obtained in this study. The 757. inhibition of the cardiac l i p o l y t i c enzyme by 0.2 M NaF compares favourably with the similar type of lipo-l y t i c activity reported in adipose tissue (15, 16) and intestinal mucosa (59) where inhibitions of 84%., 45%, and 100% respectively have been reported. Since NaF does not inhibit lipoprotein lipase, the potent inhibitory action of 0.2 M NaF must indicate the presence of another l i p o l y t i c system in cardiac and other tissues. The inhibition by rela-tively low concentrations of DFP appears to be a common fea-ture of monoglyceride-splitting enzymes. Thus, when discussing the possibility that the 300-fold purified mono-glyceride-splitting enzyme in the intestinal mucosa may be an esterase (rather than a lipase), Pope and associates (59) have agreed that their enzyme was, in fact, a lipase, owing to the observation that "an increasing solubility of the substrate is associated with a decreasing rate of hydrolysis". It was likewise indicated in this study that the increased activity of the cardiac enzyme was associated with the increased a v a i l a b i l i t y of insoluble substrates. The physiological role of lipoprotein lipase in heart i s not completely understood at the present time. Its major role may be to hydrolyze exogenously supplied triglycerides to diglycerides, monoglycerides and FFA. Since the enzyme i s 74. rapidly eluted from heart and adipose tissue ±n vitro by heparin, i t s location on or near the plasma membrane has been postulated. Alousi and Mallov (17) found the following dis-tribution of lipoprotein lipase in cardiac c e l l s : nuclear fraction, 1.33 ± 0.14; mitochondrial fraction, 0.83 ± 0.25; microsomal fraction, 1.16 ± 0.14; soluble fraction, 0.99 ± 0.13. The total enzyme activity was 4.14 i 0.34. The high concentrations of the enzyme activity in the nuclear and microsomal fractions give support to the concept that lipo-protein lipase i s essentially a membrane-located enzyme. The intracellular distribution of the cardiac monogly-ceride-hydrolyzing lipase (Table IV) also indicated that the enzyme is membrane-bound. Therefore, one might reasonably speculate that the primary physiological role of the mono-glyceride-hydrolyzing lipase in cardiac tissue might be that of completing the f i n a l step in the hydrolysis of triglycerides. A similar co-ordinated l i p o l y t i c system i s also thought to exist in the hydrolysis of triglycerides in the intestinal tract jln vivo. It i s known that pancreatic lipase hydrolyzes triglycerides to monoglycerides, and that i t s action essen-t i a l l y stops at this stage of the hydrolytic process in the intestinal lumen. However, after monoglycerides are absorbed into the intestinal mucosal c e l l s , they are either further hydrolyzed to glycerol and FFA, or are re-esterified to higher glycerides. It is highly attractive to speculate that perhaps in the case of cardiac c e l l s as well, the triglycerides in chylomicrons and very low density lipoproteins are f i r s t hydro-75. lyzed to monoglycerides at the outer surface of the c e l l s (including the lumen of the endoplasmic reticulum), then absorbed, and completely hydrolyzed by the monoglyceride-spl i t t i n g lipase, or re-esterified to higher glycerides. How-ever, this idea i s not supported by isotopic studies which indicated that triglycerides are taken up intact by the heart. Therefore one i s l e f t with the alternative idea that the actions of both lipoprotein lipase and the monoglyceride-spl i t t i n g lipase occur intracellularly, but on or near the inner aspect of the plasma membrane. Some of the problems encountered during attempts to purify the monoglyceride-splitting lipase activity from cardiac tissue have already been bri e f l y mentioned. However, the d i f f i c u l t i e s involved in attempting to solubilize and purify the enzyme can-not be over-emphasized. The enzyme was tightly bound to the particulate materials of the c e l l and was extremely resistant to the usual solubilizing techniques. It is noted that Pope and co-workers (59) used 0.37o sodium deoxycholate to rupture the microsomal particles and thus solubilized the enzyme prior to purification and study of the monoglyceride-hydro-lyzing enzyme in the intestinal mucosa. In our hands, 0.17o deoxycholate treatment resulted in some solubilization, but this technique also resulted in low recoveries of activity. Although butanol extraction was considered as a possible tech-nique, the reasonably satisfactory solubilization obtained with Triton X-100 obviated the necessity for i t s use. The degree of purification of the monoglyceride-hydro-lyzing enzyme in heart tissue i s admittedly very small. 76. Nevertheless, i t should be born in mind that with the possible exception of plasma lipoprotein lipase, pancreatic lipase and the most recently reported monoglyceride-hydrolyzing lipase of intestinal mucosa, extensive purification of lipase in bio-logical systems have been overwhelmingly unsuccessful. Kupieki (16) for example, succeeded in obtaining a modest 4.8-fold purification of a monoglyceride-hydrolyzing lipase from adipose tissue. Nevertheless, when one considers the importance of l i p i d metabolism in mammalian systems, i t is essential that greater efforts be directed in the future toward purifying and studying the lipase in these organisms. The indiscriminate use of Ediol as substrate for mea-suring l i p o l y t i c activity has been c r i t i c i z e d in this thesis. With the aid of t r i p a l m i t i n - l - C 1 4 incorporated into E d i o l R , the present study has shown that even when the hydrolysis of triglycerides in Ediol was completely abolished, significant amounts of free fatty acids were s t i l l being produced during the i n i t i a l 20-25 minutes of the reaction. Since the addition of 0.2 M NaF caused a further consistent and significant (70-757.) decrease of this l i p o l y t i c activity, the source of these FFA*s must have been derived from the monostearin component of Ediol . From these observations, incidentally, TD i t was suggested that the unspecified Ediol -hydrolyzing ac t i v i t y in extracts of rat hearts which was reported by Bjorntorp and Furman (13) was probably entirely due to the hydrolysis of the monoglyceride component. Other workers have recently looked upon the use of E d i o l R in lipase studies 77. with some suspicion. Very recently, Kupieki (16) reported that the l i p o l y t i c activity of an adipose tissue extract prepared as described by Rizack (107) liberated as much free fatty acids from monostearin as from E d i o l R (i.e., 10.80 versus 9.84 ueq/hr/100 mg tissue from monostearin and E d i o l R , respectively). Furthermore, Kupieki (16) found that the pH optima were v i r t u a l l y indistinguishable when either of these substrates were used. He concluded that ". . . i t appears (that) when the hydrolysis of E d i o l R is used to follow lipo-l y t i c activity, the results can be misleading; the added TJ monostearin which serves as an emulsifier in Ediol can account for a large part of the FFA released by this substrate". The f i r s t hint, however, that the monoglyceride component of Ediol^ may be rapidly hydrolyzed was offered by Vaughan et a l . (6) who observed that glycerol production was not proportional to adipose tissue homogenate concentration when E d i o l R was used as substrate. When the amount of glycerol derivable from the stated amount of monostearin present in E d i o l R was subtracted from the amounts of glycerol produced, the "corrected" glycerol production was linear with respect to homogenate concentration and a straight line was obtained through the origin. The results of the present study there-fore not only support the suspicions of Kupieki (16) and Vaughan et: al» (6) but provided direct evidence that the mono-stearin component in Ediol i s , in fact, rapidly hydrolyzed by cardiac enzymes. It is safe to conclude from these studies that a lipase 78. other than lipoprotein lipase exists in rat myocardium. Some of i t s properties have been described. Much further study w i l l be required to c l a r i f y i t s true physiological function in overall cardiac energy metabolism. The knowledge w i l l come only with success in purifying the enzyme. Whether i t w i l l be subject to regulation through epinephrine or some other hormone can only be a matter of speculation at the present time. 79. P A R T I I S T U D I E S O N 3 1 , 5 • - C Y C L I C N U C L E O T I D E P H O S P H O D I E S T E R A S E so. INTRODUCTION In recent years, the molecular regulation of cellular activity has been one of the most intensively studied areas in biochemistry. It i s not our purpose here to review the mechanisms by which the li v i n g c e l l regulates i t s metabolic activity. We wish only to point out that many small molecules, bearing no necessary structural relation to substrate or pro-duct can profoundly influence the action of an enzyme in either a positive or negative direction. The physiological implications of such allosteric effects are boundless. One such compound which has attracted much attention as an effector or mediator of important enzymatic reactions is adenosine S'^'-cyclic phosphate (cyclic 3',5,-AMP). The discovery of cyclic 3*,5*-AMP in mammalian tissues in 1958 originated from studies on li v e r phosphorylase. In 1951, Sutherland and Cori (119) noted that when liver slices were incubated with epinephrine and glucagon, the glycogen content decreased (glycogenolysis) and phosphorylase activity increased. Studies on li v e r phosphorylase, the enzyme which degrades glycogen to glucose-1-phosphate, revealed that i t existed in an active and inactive form. Epinephrine was shown to mediate glycogenolysis by shifting the balance of li v e r phosphorylase in favour of the active enzyme. The enzyme, phosphorylase phosphatase, was isolated which catalyzed the inactivation of highly active l i v e r phosphorylase (120, 121). At this time, Rail and Sutherland (122) also reported on 81. another enzyme which catalyzed the conversion of inactive l i v e r phosphorylase to the active form. This enzyme, which was given the name phosphorylase kinase, required Mg"*""^  ions and ATP for activity. From their studies, these investigators concluded that epinephrine stimulated glycogenolysis not by acting on phosphorylase, but by acting in some way on the latter enzyme, phosphorylase kinase. Rail and his associates (123) then made the important observation that when particu-late fractions of dog liver homogenates obtained by low speed centrifugation were incubated with epinephrine (or glucagon) together with ATP and Mg + + ions, a heat-stable factor was produced which stimulated the activation of inactive liver phosphorylase. It became clear that the action of epinephrine was indirect,in that i t stimulated the synthesis of a heat-stable factor in c e l l particles which in turn acted on phosphorylase kinase, with the result that phosphorylase was activated, leading to increased glycogenolysis. This factor was soon isolated from dog liver and crystallized by Sutherland and Rail (124, 125) in 1958, and was f i n a l l y characterized (126) as adenosine 3»,5»-cyclic phosphate (Fig. 16). An enzyme system in liver which catalyzed the formation of cyclic St^-AMP was f i r s t reported by Rail and Sutherland (127) in 1958. In the presence of Mg"*"* ions, ATP, epinephrine and glucagon, 1200 x g particles of li v e r (and also of heart, skeletal muscle and brain) formed significant amounts of cyclic 3»,5I-AMP. The name "adenyl cyclase" was adopted by FIG. 16 Structural formula of adenosine-3',5«-phosphate (cyclic 3',5«-AMP). 83. Sutherland et al.(128) for the enzyme. Adenyl cyclase was detected in a l l animal tissues studied (128), except in dog red blood c e l l s . The tissues possessing the highest specific activity were cerebral cortex and cerebellum of beef, calf, sheep and pig. Equally high activities were found in li v e r flukes (Fasciola hepatica) and in earthworms (Lumbricus  terrestris) . Intermediate levels of cyclic 3», 5«-AMP-forming activity were found in testis, uterus, intestinal mucosa of dog, the l i v e r of cat and rat, skeletal muscle of rabbit and blood c e l l s of pigeons. Low in adenyl cyclase activity were rat epididymal fat pads, f l y larva, and minnow. The relative act i v i t i e s (on protein basis) in dog tissues were: brain cortex (11.0), spleen (2.0), skeletal muscle (2.0), heart ventricle (2.0), lung (1.5), kidney cortex (1.0), liver (0.5), aorta (1.0), intestinal muscle (1.0), femoral artery (1.5) and adipose tissue (1.0). Studies on the intracellular localization of this enzyme indicated that i t might be derived from plasma membranes of from nuclei. Later studies by Davoren and Sutherland (129) with pigeon erythrocytes demon-strated that no adenyl cyclase activity was associated with the nuclei. It appeared, therefore, that adenyl cyclase was located on the plasma membrane of c e l l s . The preparation of the enzyme in a purified form was seriously hampered by i t s association with particulate materials of the "nuclear" frac-tion, by i t s instability, and by i t s close association with Triton X-100 after solubilization (128). The purification of adenyl cyclase was 2- to 3-fold from brain and about 15-fold from l i v e r . 84. It was previously mentioned that epinephrine stimulated the synthesis of cyclic 3*,5i-AMP, and i t i s now known that the immediate site of action of epinephrine i s adenyl cyclase. Murad and associates (130) studied the relative potencies of several catecholamines on the adenyl cyclase system of dog myocardial and liver particles. In the myocardial system, the relative potencies were as follows: L-isopropylnorepi-nephrine (7.8), L-epinephrine (1.0), L-norepinephrine (1.0) and D-epinephrine (0.12). In the liver system, the relative potencies were: L-isopropylnorepinephrine (4.0), L-epinephrine (1.0), L-norepinephrine (1.0). .Dichloroisopropylnorepine-phrine (DCI), an adrenergic blocking agent, blocked the stimulating effect of catecholamines. The effect of epine-phrine on the particulate preparations of adenyl cyclase from brain were also invetigated by Klainer ej: a± (131). These workers observed that in the presence of epinephrine, a 2-fold increase in cyclic 3*,5*-AMP formation by particu-late preparations from the cerebellum was obtained. Cyclase preparations from the cerebral cortex, pons, medulla, and spinal cord were also stimulated by epinephrine. It has become clear that cyclic 3*,5*-AMP plays an impor-tant role in mediating the metabolic effects of catecholamines. The participation of cyclic S^M-AMP in the glycogenolytic response of the liv e r to epinephrine i s shown in Fig. 17. Essentially, a similar series of enzymatic reactions occur in skeletal muscle (132) and in cardiac muscle (133). The meta-bolic role of cyclic 3*,5,-AMP i n glycogenolysis has thus Epinephrine (or glucagon) Adenyl Cyclase ATP »- Cyclic 3«,5«-AMP depho sphopho sphorylase kinase ATP Mg + + dephosphophosphorylase (inactive) phosphorylase (active) phosphorylase phosphatase Glycogen Glucose-1-P Pi *—^| (phosphatase) I Glucose FIG. 17 Mediation of cyclic 3*,5*-AMP in the glycogenolytic response of liver epinephrine (or glucagon). 86. been firmly established. Investigations by several workers have indicated that the role of cyclic 3*,5,*-AMP in biological systems extends far beyond i t s participation in the process of glycogenolysis. The activity of a number of enzyme systems (other than the phosphorylase system in live r , heart and skeletal muscle) have been shown to be influenced by cyclic 31,5*-AMP. Haynes and Berthet (134) found that the addition of adrenocortical hormone (ACTH) to adrenal tissue slices caused a rapid and specific activation of phosphorylase in these tissues. Furthermore, Haynes (135) showed that when ACTH was added to slices of beef adrenal cortex, cyclic 31 i5t-AMJ? accumulated in these tissues. When cyclic 3* ,5}-A3XE i t s e l f was added, phosphory-lase activity increased. Thus, he concluded that the action of ACTH on the adrenal cortex was mediated by cyclic 3«,5*-AMP. Another enzyme which i s affected by cyclic 3»,5*-AMP is glycogen synthetase (UDPG-ac-glucan Transglucosylase). Belo-copitow (136) observed a decrease in glycogen synthetase activity in rat diaphragms after these tissues were incubated with epinephrine. In order to investigate further the mecha-nism of action of epinephrine on glycogen synthetase, he incubated a 4000 x g supernatant of rat skeletal muscle homo-genates with cyclic 3»,5«-AMP and ATP. When cyclic 3»,5«-AMP was omitted from the system, an increase in synthetase activity was observed. When both ATP and cyclic 3«,5'-AMP were omitted, a further increase in the enzyme activity was observed. These observations led Belocopitow (136) to suggest that cyclic 87. 3»,5»-AMP may have been inhibiting the synthetase system. Recently, Rosell-Perez and Larner (137) showed that the action of cyclic 3*,5*-AMP on the inhibition of glycogen synthetase activity was closely associated with enhancing the conversion of the "I" (Independent,active) form of the enzyme to the "D" (Dependent,inactive) form. It has become evident, therefore, that the inactivation of glycogen synthetase coupled with the simultaneous activation of phosphorylase by cyclic 3*,5*-AMP would constitute an important role for the cyclic nucleotide in the regulation of glycogen breakdown and synthesis. In 1960, Mansour and his co-workers (138) made an interesting observation relating to the phosphorylase system in the li v e r fluke, Fasciola tiepatica. These workers demonstrated that the activation of phosphorylase in this organism was obtained not with epinephrine, but with 5-hydroxytryptamine (serotonin). Furthermore, their studies showed that serotonin caused a rapid increase in cyclic 3*,5*-AMP levels, and also that a c t i -vation of phosphorylase was mediated by the cyclic nucleotide. Mansour (139) also demonstrated that glycolysis in homogenates of liver flukes was regulated by the activity of phosphofruc-tokinase (PFK), and that stimulation of glycolysis by seroto-nin resulted in a marked increase in PFK activity. Mansour and Mansour (140) reported that cyclic 3,»,5,»-AMP could a c t i -vate l i v e r fluke PFK which had been inhibited by ATP, and that the cyclic nucleotide could also activate an inactive preparation of PFK. Similar effects of cyclic 3*,5*-AMP in PFK activity have been demonstrated with the enzyme isolated from mammalian cardiac tissue (141, 142) and skeletal 88. muscle (143) • The observation that cyclic 3*,5*-AMP mediated the glyco-genolytic response of the liver to epinephrine led investiga-tors to study the possibility that cyclic 3«,5»-AMP might also mediate the li p o l y t i c response of adipose tissue to epinephrine. Indeed, Rizack (144) showed that an epinephrine-sensitive lipase in cell-free extracts of adipose tissue could be activated by the addition of cyclic 3*,5»-AMP in the presence of ATP and Mg"*"** ions. Butcher and his co-workers (145) found that when a derivative of cyclic 3«,5«-AMP, N 6-2t-O-dibutyryl cyclic AMP was incubated with intact isolated fat c e l l s , l i p o l y s i s was stimulated some 10-fold. Hence, another role for cyclic 3*,5*-AMP, that of increasing the mobiliza-tion of free fatty acids from fat depots, has been established. The enzyme, tryptophan pyrrolase, which opens the indole ring of tryptophan to yield formylkynurenine, appears to exist in an active and an inactive form. Although studies on the effects of cyclic 3»,5»-AMP on this enzyme system have not been extensive, there are indications that this enzyme can be converted from the inactive to the active form by cyclic 3*,5«-AMP (146, 147), although a recent report (148) indicates that 5*-AMP, guanine, guanosine and 5«-GMP were also highly effec-tive. There are several other biological processes in which cyclic 3»,5»-AMP has been implicated. For example, the posi-tive inotropic response of the heart to epinephrine and other catecholamines has been widely studied and the indications 89. are that this process may also be mediated by cyclic 3»,5*-AMP (149). Recent studies had shown that the inotropic response and the activation of cardiac phosphorylase were apparently unrelated (150, 151). Then Robison and his associates (149) observed that a single dose of epinephrine caused a 4-fold increase in cyclic 3*,5»-AMP levels in cardiac tissue within 3 seconds after injection, while the contractile force increased about 1.4-fold at 20 seconds. These observations therefore favour the hypothesis that cyclic 3«,5»-AMP may mediate the inotropic response of the heart to catecholamines. It was mentioned earlier that the activation of adreno-cortical phosphorylase by ACTH was shown to be mediated by cyclic 3*,5»-AMP. Since ACTH action on the adrenal cortex stimulates the synthesis of corticosteroids, one might expect that cyclic 3«,5»-AMP i t s e l f might mimic the action of ACTH. Indeed, Haynes et al.(152) found that when they added cyclic 3»,5«-AMP to fragments of incubating rat adrenals, the total corticoid production was stimulated almost 5-fold. Recent studies have indicated that the stimulation of steroid pro-duction by the adrenal cortex was not solely due to the activation of phosphorylase. Roberts and co-workers (153, 154, 155) found that cyclic S'j^-AMP selectively stimulated C-11/& hydroxylase activity in rat adrenal homogenates f o r t i -fied with glucose-6-phosphate and NADP, resulting in increased formation of corticosterone from exogenous 11-deoxycorticos-terone or progesterone. They also showed that the conversion of progesterone to 11/5 -hydroxyprogesterone was increased. 90. These workers concluded that the action of 3»,5*~AMP on the stimulation of steroidogenesis was independent of phosphory-lase activation, NA13PH generation and the presence of endo-genous corticosteroid precursors. Roberts et alo(156, 157) showed that cyclic 3»,5*-AMP also stimulated the hydroxylation of 11-deoxycorticosterone to 18-hydroxy-ll-deoxycorticosterone, as well as the conversion of exogenous cholesterol to pregnen-olone by isolated rat adrenal mitochondria. Karaboyas and Koritz (158) recently observed that cyclic 3*,5*-AMP stimulated the incorporation of acetate into corticosterone, and the conversion of cholesterol to corticosterone. A report by Darrington and Kilpatrick (159) indicates that cyclic 3»,5«-AMP stimulated the synthesis of two progestational steroids, 4-pregnen-20 oc-ol-3-one and progesterone by ovarian tissues of rabbits. Pryor and Berthet (160) reported that the incorporation of leucine into protein of rat l i v e r slices was inhibited when these tissues were incubated with cyclic 3«,5*-AMP or with glucagon. Exton and Park (161, 162) have indicated that the effect of these hormones on glueoncogenesis appears to be mediated by cyclic 3«,5»-AMP. Furthermore, these authors have suggested that the direct site of cyclic 3*,5*-AMP action may be the activation of phosphopyruvate carboxylase and the hexose phosphate phosphatases. Strong evidence also exists that cyclic 3*,5*-AMP may mediate the secretion of enzymes . from rat parotid glands. Bdolah and Schramm (163) have indi-cated that when rat parotid slices are incubated with dibutyryl 91. cyclic AMP, the release of amylase from the glands i s stimu-lated almost to the same extent as that observed with epine-phrine. Orloff and Handler (164) reported that the addition of cyclic 3*,5*-AMP to media in which toad bladders were immersed, caused a significant increase in the permeability of the membrane to water. Since vasopressin (antidiuretic hormone) is also known to e l i c i t the same response, these authors suggested that the antidiuretic action of vasopressin might also be mediated by cyclic 3*,f>*-AMP. Recent studies on the action of vasopressin on the toad bladder by Strauch and Langdon (165) and by Handler and associates (166) have indicated that the action of this hormone may be a direct stimulation of adenyl cyclase. The cellular mode of action of vasopressin has been recently reviewed by Orloff and Handler (167). The decreased incorporation of acetate into fatty acids and cholesterol of liver slices, and the increased production of ketone bodies by epinephrine, glucagon and by cyclic 3»,5«-AMP has been reported by Berthet (168). Cyclic 3*,5*-AMP has also been implicated in sugar transport in the thyroid gland (169), and in the stimulation of hydrochloric acid secretion by gastric mucosa (170, 171). The widespread interest in cyclic 3*,5"-AMP has led to the appearance of several review articles, the most recent of which are those on the metabolic effects of catecholamines by the following authors: Sutherland and Robison (172), Butcher (173), Krebs et al.(132), Mansour (174), and Exton and Park (162). The participation of cyclic S'j^'-AMP in glycogenolysis, 92. s t e r o i d o g e n e s i s , ketogenesis, l i p o l y s i s , a n t i d i u r e s i s and pos-s i b l y other p h y s i o l o g i c a l processes demonstrates c l e a r l y the exceedingly d i v e r s i f i e d and important r o l e played by t h i s c y c l i c n u c l e o t i d e i n r e g u l a t i n g v a r i o u s b i o l o g i c a l processes. I t would f o l l o w that an e q u a l l y important p h y s i o l o g i c a l mecha-nism i s necessary f o r the ter m i n a t i o n of the a c t i o n of c y c l i c 3',5'-AMP i n b i o l o g i c a l systems. Indeed, an enzyme which hydrolyzes c y c l i c 3',5*-AMP to 5'-AMP i s present i n most t i s s u e s . The exist e n c e of such an enzyme a c t i v i t y was f i r s t suggested by Sutherland and R a i l (124) i n 1958 w h i l e these authors were studying the p r o p e r t i e s of c y c l i c 3',5'-AMP formed by t i s s u e p a r t i c l e s . The enzyme has been subsequently p u r i f i e d from beef heart by Butcher and Sutherland (175) and r e c e n t l y from dog heart by N a i r (176). E a r l i e r , Drummond and Perrott-Yee (177) had studied the d i s t r i b u t i o n of the d i e s -terase i n v a r i o u s mammalian t i s s u e s , and had found t h a t nervous t i s s u e s , p a r t i c u l a r l y the b r a i n , possessed by f a r the highest a c t i v i t y . The kidney, heart, spleen and l i v e r of r a b b i t contained o n l y 10 - 257o of the a c t i v i t y of the b r a i n . Studies made on the p r o p e r t i e s of the d i e s t e r a s e from b r a i n revealed i t s absolute requirement f o r magnesium ion s , and that the product of h y d r o l y s i s was e x c l u s i v e l y 5'-AMP. Considering the f u n c t i o n of c y c l i c 3',5'-AMP i n so many di v e r s e b i o l o g i c a l processes, i t f o l l o w s that the d i e s t e r a s e must p l a y an important r o l e i n r e g u l a t i n g the a c t i o n of the c y c l i c n u c l e o t i d e i n va r i o u s t i s s u e s . The enzyme i s more a c t i v e i n b r a i n than i n any other t i s s u e . Adenyl c y c l a s e i s also more active in brain than elsewhere in nature. The precise physiological function of cyclic 3«,5»-AMP in nerve tissue is a problem of paramount importance. The work des-cribed in this part of the thesis constitutes a further study of the properties and part i a l purification of cyclic 3*,5'-nucleotide phosphodiesterase from mammalian brain. Some studies on the distribution of the enzyme throughout the plant and animal kingdom are also reported. 9 4 . EXPERIMENTAL PROCEDURE  Materials Cyclic 3»,5'-AMP and ca l f intestinal adenosine deaminase were purchased from Sigma Chemical Company. Cyclic 3',5'-GMP and cyclic 3',5»-UMP were prepared by Smith, Drummond and Khorana (178). Cyclic 3',5«-dAMP, cyclic 3«,5«-dGMP, cyclic 3',5»-dCMP and cyclic 3* ,5*-1MB were prepared by Drummond, Gilgan, Reiner and Smith (179). Cyclic 2«,3«-AMP was pre-pared by the method of Smith, Moffatt and Khorana (180). The cyclic nucleotides 3»,5«-dAMP, 3«,5«-GMP and 3',5«-dGMP were chromatographically pure. Crotalus adamanteus venom was purchased from Ross Allen's Reptile Institute, Silver Springs, Florida and from Sigma Chemical Company. Methods Standard Assays - Routinely, cyclic 3«,5'-nucleotide phos-phodiesterase activity was assayed according to Butcher and Sutherland (175) by measuring the release of inorganic phos-phate when Crotalus adamanteus venom was included in the assay system. The snake venom contains a potent 5'-nucleoti-dase which hydrolyzes 5'-nucleotides to give the corresponding nucleoside and inorganic phosphate. The reaction mixture contained 0.54 umoles cyclic 3',5'-AMP, 0.90 umoles MgS04, 95. 0.1 to 0.4 rag snake venom, 88 umoles Tris buffer, pH 7.5, with an appropriate dilution of the phosphodiesterase sample being assayed, in a total volume of 0.9 ml. Incubations were per-formed at 30° for 30 minutes, and the reaction terminated by the addition of 0.1 ml cold 557. trichloroacetic acid. The reaction tubes were centrifuged for 10 minutes at 10,000 x g to sediment the denatured proteins, and 0.5 ml aliquots of the supernatant f l u i d taken for inorganic phosphate analysis based on the method of Fiske and SubbaRow (181) as modified by Butcher and Sutherland (175). Thus, to 0.5 ml aliquots of the supernate were added 0.1 ml 2.57. ammonium molybdate solu-tion in 5 N I^SO^, 0.35 ml glass-distilled water and 0.05 ml reducing agent. Colour was allowed to develop for 15 minutes before reading at 720 mu in a Beckman Model DU spectrophoto-meter, using a light path of 1.0 cm. The standard curve for inorganic phosphate as measured by this method is shown in Fig. 18. One unit of enzyme activity i s defined as that amount which caused the liberation of 1 umole of inorganic phosphate in 30 minutes at 30°. The specific activity of the enzyme is defined as the umoles of inorganic phosphate released per mg of enzyme protein in 30 minutes at 30°. For kinetic experiments, the assay was based on the con-version of cyclic SS^'-AMP to inosine in the presence of brain extract, snake venom and intestinal adenosine deaminase. (One unit of deaminase activity i s defined as the number of umoles of adenosine deaminated per minute at 30° in 0.1 M citrate buffer at pH 6.5 at an adenosine concentration of INORGANIC PHOSPHATE ADDED (mjumoles) FIG. 18 Inorganic phosphate c o n c e n t r a t i o n curve. To a s e r i e s of tubes c o n t a i n i n g 0.2 ml 0.4 M T r i s , pH 7.5, 0.05 ml 18 mM.MgSO^, 0.10 ml 55% t r i c h l o r o a c e t i c a c i d , were added the i n d i c a t e d amounts of inorganic phos-phate and s u f f i c i e n t g l a s s - d i s t i l l e d water to make a f i n a l volume of 1.0 ml. An 0.5 ml a l i q u o t was taken from each tube and assayed f o r inor g a n i c phosphate as described i n the t e x t . 97. 0.45 x 10~4 M.) The concentrations of cyclic 3,*,5»-AMP were between 13 to 52 uM. The reaction mixture also contained 1.0 mM MgSO^ and 88 mM Tris buffer at pH 7.5 in a f i n a l volume of 1.5 ml. After the addition of cyclic 3*,5*-nucleotide phos-phodiesterase, the decrease in absorbancy was followed at 265 mp at 1-minute intervals using a Beckman Model DU spec-trophotometer and a light path of 0.5 cm (Fig. 19). For certain experiments where the identification of the reaction products by paper chromatography was considered advantageous, the assay system consisted of 0.25 umoles cyclic 3»,5»-AMP, 0.8 mM MgS04, 75-150 mM Tris buffer, pH 7.5, and enzyme in a total volume of 0.2 ml. In this system, no snake venom was used. The reaction was stopped after 15 minutes incubation at 30° by the addition of 0.02 ml glacial acetic acid. The tubes were centrifuged at 10,000 x g for 15 minutes and 0.02 ml aliquots of the supernate spotted on Whatman No. 1 f i l t e r paper. The chromatograms were developed by descending technique with isopropanol-ammonium hydroxide-0.1 M boric acid (7:1:2). This solvent system effectively separates adenine, adenosine, and the adenosine nucleotides. Stock solutions of snake venom (8 mg/ml) used in the diesterase assay were prepared by dissolving the lyophilized powder in 0.02 M Tris, pH 7.5. Centrifugation was occasionally required to sediment insoluble particles. The snake venom activity was completely stable to repeated freezing and thaw-ing over a period of several months. Protein was measured by the biuret method (182) and by the optical method of War-98. If—. e L/l 1 1 L 1—i 1 1 1 1 L. 0 2 3 4 5 6 T 8 9 10 M I N U T E S FIG. 19 Spectrophotometrie Assay of Phosphodiesterase. The i n c u b a t i o n mixture contained 0.03 mM c y c l i c 3',5'-AMP, 1.0' mM MgS0 4, 88 mM T r i s , pH 7.5, 0.15 u n i t s of i n t e s t i n a l adenosine deaminase, 0.32 mg snake venom, and 27 pg r a b b i t b r a i n phosphodiesterase i n a f i n a l volume of. 1.5 ml. The r e a c t i o n was s t a r t e d by the a d d i t i o n of the d i e s t e r a s e and ~Zb>265 w a s recorded at 1 minute i n t e r v a l s . 99. burg and Christian (183). Enzyme Purification - Mature rabbits were stunned by a blow behind the head and the neck vessels severed immediately. The brains were removed, placed in ice and usually frozen before use. Only the cerebral lobes were used for the prepa-ration of the extract. The tissue was homogenized in 10 volumes of 0.25 M unbuffered sucrose for 5 minutes at 0-4° using a glass mortar fitted with a motor-driven teflon pestle. A l l subsequent procedures, unless otherwise noted, were per-formed at 0-4°. The homogenate was centrifuged at 105,000 x g x 30 minutes, and the supernatant fl u i d thus obtained was set aside. The 105,000 x g sediment was re-homogenized with 5 original volumes of 0.25 M sucrose containing 0.1% deoxy-cholate. The homogenate was centrifuged for 30 minutes at 105,000 x g, and the supernatant fluids combined for the f o l -lowing ammonium sulfate fractionation step. It was found later that the centrifugation could be more conveniently performed at 37,000 x g x 60 minutes, giving; essentially the same degree of purification. Step 1 - Ammonium Sulfate Fractionation - The supernatant fl u i d was adjusted to 0.3 saturation by the addition of solid enzyme-grade ammonium sulfate with constant s t i r r i n g over a 15-minute period. The pH was maintained between 6.9 and 7.1 by the dropwise addition of 1.0 N KOH. After at least 15 minutes equilibration, the precipitate was collected by cen-trifuging for 20 minutes at 37,000 x g. The precipitate was taken up to approximately 15% of the original combined super-100. natant f l u i d volume using 1 mM imidazole, pH 7.5 containing 1 mM MgSO^ .^  The milky extract was dialyzed against 300 vol-umes of the same buffer (pH 7.5) for 3 hours in the cold room. Step-2 - Repeated"Freezing-and Thawing - After dialyzing, the extract was centrifuged at 37,000 x g x 15 minutes to remove the heavy flocculent material, and the slightly cloudy super-natant fluid was stored at -20°. Upon thawing the extract, more flocculent material always appeared, which was readily removed by centrifugation. The supernate was re-frozen, thawed and centrifuged once more before taking the prepara-tion to the next purification step, or was stored at -20°. About 5% of the enzyme activity was lost with the sediment, but no attempt was made to recover this activity. The repeated freezing and thawing of the f i r s t ammonium sulfate fraction usually gave a 6- to 10-fold purification of the enzyme, and an overall yield of about 15 to 207.. Step 3 - Heat Benaturation and Acid Precipitation - After the repeated freezing and thawing procedure, 0.11 volumes of 0.5 M imidazole, pH 7.5, and 0.02 volumes of. 0.5 M glycine, pH 10, was added to the clear supernate. The pH was taken to 10 by the addition of 1 N KOH, and the temperature of the solution brought quickly to 45°. After maintaining this temperature for about 15-16 minutes, the solution was immediately chilled by immersion in an ice-bath. The solution was then slowly taken to pH 5.8 with 0.3 N acetic acid and stirred for at least 15 minutes before centrifuging at 37,000 x g x 45 min-101* utes. The precipitate was discarded, and the pH of the super-nate brought back to 7.5 with 0.5 N KOH. The solution thus obtained was dialyzed against 300 volumes ofl.jmM imidazole, pH 7.5 containing 1 mM MgSO^ pH 7.5, with constant s t i r r i n g for at least 6 hours. Although the f i r s t ammonium sulfate step and the freezing and thawing gave reasonably consistent degrees of purification, the alkaline-heat, acid-precipitation step gave results which varied from one preparation to another. Unless otherwise indi-cated, this preparation was used for studying the properties of the brain phosphodiesterase. The overall yield at this step was about 5 to 10%, and the purification obtained ranged from 8- to 16-fold. The enzyme became exceedingly unstable with increasing purification, probably owing to the dilution of the enzyme during and after the alkaline-heat step. Con-centration of the 6-hour dialysate obtained from the alkaline-heat step was accomplished by immersion of the dialysis bag into 1 l i t r e of a 60% solution of sucrose. This technique resulted in a 90%. decrease in volume of the dialysate within a few hours. The concentrated enzyme was stored at -20° for one week with no appreciable loss of activity. Further purification of the enzyme could be obtained by taking a second (0.3 to 0.6 saturation) ammonium sulfate frac-tion after the alkaline-heat step. However, despite the 15 to 25-fold purification obtained by this method, the f i n a l yield of enzyme activity was low; hence the use of this step as a routine procedure was impractical. Nevertheless, this 102. preparation was occasionally used in some of the experiments where the use of a more highly purified preparation was i n d l cated. 103. RESULTS 1. Preliminary - Before purification of the cyclic 3',5'-nucleotide phosphodiesterase from brain was attempted, the enzymatic components of the diesterase assay system were examined. The snake venom used in the diesterase assay con-tains a potent 5*-nucleotidase which hydrolyzes 5!-AMP to adenosine and phosphate. It was therefore necessary to deter-mine the minimum quantity of snake venom required to hydrolyze a l l of the 5*-AMP produced by the diesterase under standard assay conditions. As may be seen in Fig. 20, 0.36 umoles of 5,-AMP was almost completely hydrolyzed by 30 ug snake venom at 30° in 10 minutes. When the standard assay system was subsequently developed which contained 0.54 umoles cyclic 3*,5»-AMP, an excess (100-400 ug) of snake venom was routinely used in the incubation mixture in order to eliminate any pos-s i b i l i t y of the 5*-nucleotidase limiting the overall reaction rate. When 100 pg of snake venom was used, no cyclic S 1 ^ 1 -nucleotide phosphodiesterase could be detected in the venom preparation. However, in some later experiments, larger amounts of snake venom were used in the assay system. Analy-sis for the presence of cyclic 3*,5*-AMP hydrolyzing activity at these higher concentrations of venom indicated that a trace of diesterase activity was present, as shown in Table VI. Although the presence of diesterase activity in the snake venom had insignificant effect on the results of most experi-ments where brain diesterase activity being measured was high, data from those few experiments where the activity was low 104. O a. o < CO 0 10 20 30 40 50 60 70 SNAKE VENOM ADDED 80 90 100 ( H e ) FIG. 20 H y d r o l y s i s of 5'-AMP by snake venom (C r o t a l u s  adamanteus). The i n c u b a t i o n mixture contained 0.36 umoles 5'-AMP, 20 mM MgSO^, 40 mM T r i s b u f f e r , pH 7.5, i n a f i n a l volume of 1.0 ml. The i n d i c a t e d amounts of snake venom were added to i n i t i a t e the r e a c t i o n . The r e a c t i o n was terminated by the a d d i t i o n of 0.1 ml 55% t r i c h l o r o a c e t i c a c i d a f t e r 10 minutes- i n c u b a t i o n at 30°, described i n the t e x t . Phosphate was analyzed as 105. TABLE VI Relative Ac t i v i t i e s of 5'-Nucleotidase and Cyclic 3',5'-Nucleotide Phosphodiesterase in Snake Venom. The incubation mixture contained either 0.72 umoles cyclic 3',5'-AMP or 5'-AMP, 18 mM MgS04, 0.04 M Tris, pH 7.5, the indicated amounts of Crotalus adamanteus venom (Sigma) and sufficient water to make 0.9 mT~. The reaction was stopped by the addition of 0.1 ml 55% trichloroacetic acid after 30 minutes incubation at 30°. Inorganic phosphate was assayed according to the standard procedure as described in the text. Amount of Snake Venom added (ug) jamoles Pi released from 5'-AMP & from cyclic 3',5'-AMP 0.43 0.01 0 0.86 0.03 0 2.15 0.09 0 4.3 0.24 0 8.6 0.45 0 21.5 0.75 .002 43.0 0.75 .004 129.0 0.77 215.0 0,74 .010 106. was corrected for the inherent cyclic 3 s,5'-nucleotide dies-terase in the venom. Experiments indicated that 400 ug snake venom was capable of hydrolyzing 0.02 umoles cyclic 3*,5«-AMP in 30 minutes at 30°, and therefore this correction factor was used. Preliminary experiments designed to test the v a l i d i t y of the coupled enzyme assay indicated that the rate of cyclic 3«,5«-AMP hydrolysis was directly proportional to the amount of diesterase used (Fig. 21). The enzyme preparation used for these experiments was an extract of an acetone powder which had been prepared 2 years previously and stored at -20°. The experiments therefore indicated, in addition, that the brain diesterase was quite stable to storage when prepared in acetone powder form. 2. Partial-Purification of Brain Phosphodiesterase - Drummond and Perrott-Yee (177) reported that a partial purification of rabbit brain diesterase was readily obtained by taking a 20,000 x g supernatant fraction of the whole homogenate to 0.4 saturation with ammonium sulfate. The enzyme activity was recovered from the precipitate. The present study also found most of the activity associated with the 0.4 saturated ammo-nium sulfate fraction, although the highest specific activity was recovered in the 0.40-0.45 saturated fraction. It was observed, however, that when 0.17. sodium deoxycholate was included in the 0.25 M sucrose solution used for re-homogen-izing the i n i t i a l 105,000 x g sediment, the diesterase was precipitated at lower ammonium sulfate concentrations. 107. 0 0-22 0-44 0-66 0-88 I'TO PROTEIN ADDED (mg) FIG. 21 Phosphodiesterase A c t i v i t y as a L i n e a r Function of P r o t e i n Concentration. The r e a c t i o n mixture contained 0.36 pinoles c y c l i c 3',5 1 -AMP, 20 mM MgSO^, 40 mM T r i s , pH 7.5, 60 pg snake venom, the i n d i c a t e d amounts of p r o t e i n and s u f f i c i e n t water to make a f i n a l volume of 0 . 9 ml. The r e a c t i o n was stopped by the a d d i t i o n of 0.1 ml t r i c h l o r o a c e t i c a c i d a f t e r 30 minutes i n c u b a t i o n at 30°. The d i e s t e r a s e p r e p a r a t i o n used was an acetone powder e x t r a c t of beef b r a i n . P i was assayed as described i n the t e x t . 108. Furthermore, repeated freezing and thawing of the dialyzed extract obtained from the 0.3 saturated ammonium sulfate frac-tion consistently resulted in a 6-10-fold purification. The alkaline-heat treatment followed by the acid precipitation frequently gave an additional 2-fold purification. An example of the purification of the diesterase from rabbit brain i s shown in Table VII. The specific activity of the f i n a l prepa-ration from brain was about half of that reported by Butcher and Sutherland (175) who purified the enzyme from beef heart. While this work was in progress, Nair (176) also reported purifying the diesterase from dog heart. The specific a c t i -vity of the purified enzyme from dog heart was around 27.5, which is in the same range as that obtained for the beef heart enzyme. Attempts to purify the brain enzyme by the use of Sephadex G-200 columns, or by adsorption on calcium phosphate gels under various conditions were unsuccessful. Several attempts to purify the enzyme on DEAE-cellulose columns were also unsuccessful, owing to the in s t a b i l i t y of the enzyme in dilute solutions. 3. Properties of the Partially Purified Phosphodiesterase -(a) The i n i t i a l studies on brain diesterase by Drummond and Perrott-Yee (177) indicated that the rabbit brain dies-terase had an absolute requirement for Mg"*"1" ions and was com-pletely inhibited by EDTA (1.0 mM). These observations were f u l l y confirmed in the present study. (b) The effect of 0.06 M imidazole on brain diesterase activity was investigated. As may be seen in Fig. 22, imi-109. TABLE VII Partial Purification and Yield of Cyclic 3*,5'-nucleotide Phosphodiesterase from Rabbit Brain. The cerebral cortex from a rabbit brain was fractionated as described in the text. Activities are also defined in the text. Fraction Total Activity Specific Activity 7o Yie°ld P u r i f i -cation Homogenate 728 0.9 (100) 1.0 Combined Supernate (37,000 x g) 660 2.3 90 2.4 0.3 Ammonium sulfate 159 4.3 22 4.6 Fro zen- Thawed twic e 156 9.0 21 9.2 Alkaline-Heat and Acid precipitation 20 14.0 2.7 14.8 110. p H F I G . 22 pH Curve of B r a i n C y c l i c 3«,5'-Nucleotide Phosphodiesterase and E f f e c t of Imidazole. The r e a c t i o n mixture contained 0.8 mM c y c l i c S'j^'-AMP, 1.3 mM MgSOA, 4 ug enzyme p r o t e i n of s p e c i f i c a c t i v i t y 19 u n i t s , and 0.06 M T r i s p l u s imidazole or 0.12 M T r i s . A f t e r a 20-minute i n c u b a t i o n at 30°, the r e a c t i o n was terminated by heating the tubes i n b o i l i n g water f o r 1 minute. The pH of the r e a c t i o n mixtures were adjusted to n e u t r a l i t y , snake venom (400 ug) was added, and the tubes incubated f o r 10 minutes at 30°. The r e a c t i o n xvas stopped by the a d d i t i o n o f 0.1 ml c o l d t r i c h l o r o a c e t i c a c i d . Inorganic phosphate was assayed as described i n the t e x t . 111. dazole caused a significant increase in diesterase activity over the entire pH range examined. Peak enzyme activity was near pH 7.0, whether imidazole was present or not. Stimula-tion of the beef heart enzyme by imidazole has also been reported by Butcher and Sutherland (175), although their data indicated l i t t l e stimulation at pH 8.5. (c) Inhibition by Theophylline in vitro - The methyl xanthines, particularly theophylline, are known to inhibit cyclic 3*,5*-nucleotide phosphodiesterase. The effect of z x 10*"% theophylline on the par t i a l l y purified brain dies-terase activity was investigated, and these results are shown in Fig. 23. The inhibition of brain diesterase by theophylline appears to be competitive in nature. The of the enzyme was about 0.8 x 10*"4M with cyclic 3*,5»-AMP as substrate. The value obtained indicated a marked similarity to those values reported for the beef heart enzyme. Nair (176), how-ever, has reported K m values near 4.9 x 10"% for the dog heart diesterase. (d) Cyclic 3t,5«-dAMP/cyclic 3«,5«-AMP activity ratios  in successive fractions obtained during purification - It was reported (177) that the brain diesterase hydrolyzed cyclic 3*, 5«-dAMP at about 50% of the rate at which cyclic 3«,5«-AMP was hydrolyzed. The present studies showed that the cyclic 3*,5»-dAMP/cyclic 3«,5»-AMP activity ratios were about 0.45. In order to determine whether cyclic 3*,5*-dAMP and cyclic 3*,5*-AMP might be hydrolyzed by the same or different enzyme, the cyclic 3* ,5*-dAMP/cyclic 3»,5*-AMP activity ratios were 112. FIG. 23. Nature of B r a i n C y c l i c 3•,5•-Nucleotide Phosphodiesterase I n h i b i t i o n by Theophylline. The p a r t i a l l y p u r i f i e d b r a i n d i e s t e r a s e p r e p a r a t i o n ( s p e c i f i c a c t i v i t y , 9.1) used i n these experiments was stored at -20° i n a concentrated sucrose s o l u t i o n . A f t e r thawing and making the appropriate d i l u t i o n , the enzyme p r e p a r a t i o n was incubated i n the presence of 2 x 10"^M t h e o p h y l l i n e as described i n the t e x t f o r the spectrophotometrie assay. \ \ 113. determined in the various fractions obtained during p u r i f i -cation. These results are shown in Table VIII. Although there was a significant difference between the ratios obtained for the whole homogenate and the combined supernatant fractions, the succeeding fractions showed insignificant differences. It was also observed that when equimolar concentrations of cyclic S^S'-dAMP were included with cyclic 3',5*-AMP in the standard phosphodiesterase assays, a 15-197. inhibition of diesterase activity was consistently obtained. These obser-vations strongly indicate that both cyclic 3»,5»-AMP and cyclic 3,,5,-dAMP are hydrolyzed by the same enzyme. However, further investigations are necessary in order to show more conclusively whether these compounds are indeed hydrolyzed by the same diesterase. (e) Hydrolysis Rates of other Purine and Pyrimidine  cyclic 3*,5«-nucleotides - The rate of hydrolysis of other cyclic 3 1,5^-nucleotides were investigated in order to further determine the specificity of the brain phosphodiesterase. The relative hydrolysis rates of a l l the cyclic 3*,5'-nucleotides which were investigated are listed in Table IX. Included for comparison are the rates obtained for the brain and heart enzyme extracts by other investigators. The data clearly demonstrate:;, the high specificity of the enzyme for purine cyclic 3*,5*-nucleotides. Cyclic 3»,5»-UMP was the only pyrimidine cyclic nucleotide which was hydrolyzed appreciably by the brain diesterase preparation, although the activity was only about 137. of cyclic 3*,5*-AMP hydrolysis rates. 114. TABLE VIII Cyclic 3S5«-dArtP/cyclic 3«,5»-AMP activity ratios. The cyclic 3*,5*-nucleotide phosphodiesterase was par-t i a l l y purified as described in the text. The specific activity of the f i n a l preparation was 15.0. Assays were performed by the standard method as described in the text, except that cyclic 3»,5»-dAMP was used at a concentration of 0.50 umoles/ml. Fraction cyclic 3',5'-dAMP Activity cyclic 3«,5»-AMP Ratio Whole Homogenate .76 Combined 105,000 x g supernatant .53 0.3 Ammonium sulfate Frozen-Thawed x2 .52 Alkaline-heat, Acid treatment .45 TABLE IX R e l a t i v e H y d r o l y s i s Rates of Purine and Pyrimidine C y c l i c 3«,5«-Nucleotides. Di e s t e r a s e a c t i v i t i e s were assayed by the standard procedure, except that the other c y c l i c 3»,5"-nucleotides as i n d i c a t e d were s u b s t i t u t e d f o r c y c l i c 3»,5«-AMP i n the assay i n equimolar c o n c e n t r a t i o n s . COMPOUND THIS STUDY (Rabbit b r a i n ) DRUMMOND ( X 7 7 ) (Rabbit b r a i n ) BUTCHER & SUTHERLAND (175) (Beef heart) NAIR (176) (Dog, heart) DRUMMOND et al(189) T B e e f bra i n ) 3*,5»-AMP 1,00 1.00 1.00 1.00 1.00 3»,5«-dAMP 0.45 1.30 0.50 3»,5»-GMP 0.50 0.33 0.33 3»,5«-dGMP 0.48 0.44 3»,5»-CMP 0.0 0.0 0 3«,5«-dCMP <0.04 0.10 3»,5»-UMP 0.13 0.11 0.17 0.12-0.15 3«,5«-TMP <0.07 0.10 116. Hardman and Sutherland (184) have recently reported purifying a cyclic 3*,5*-nucleotide phosphodiesterase from beef heart that hydrolyzed cyclic S^^-IMP at a much faster rate than i t hydrolyzed cyclic 3»,5*-AMP. Their findings suggest the possi b i l i t y that the appreciable hydrolysis of cyclic 3l,5»-UMP by the partially purified fraction of rabbit brain may be due to the presence of a separate enzyme for hydrolyzing cyclic 3»,5«-UMP. The hydrolysis of cyclic 3»,5*-AMP, cyclic 3«,5*-GMP and their deoxy analogues by the brain diesterase preparation were followed by paper chromatography (Figs. 24-27). Only one hydrolysis product could be detected for each of the cyclic nucleotides. It should be mentioned here that the isopropanol-ammonium hydroxide-0.1 M boric acid (7:1:2) sol-vent system effectively separates 3*- from 5'-nucleotides. Although no reference standards are shown for these compounds on these chromatograms, i t was noted that the product of cyclic 3 s, 5'-AMP hydrolysis was indistinguishable from authen-t i c 5'-AMP. Furthermore, earlier studies by Drummond and Perrott-Yee (177) have shown that the hydrolysis products of cyclic 3*,5*-GMP and cyclic 3«,5«-AMP were their corresponding 5"-nucleotides. Therefore, i t i s reasonable to suggest that the products of cyclic 3*,5*-deoxy-nucleotides were also their corre spond ing 5 *-deoxynucleo tide s• (f) Further studies on Specificity of Brain Diesterase -The presence of 5"-nucleotidase activity in the pa r t i a l l y purified preparation of brain diesterase was investigated. 117. . '7 >T! i m FIG. 24 H y d r o l y s i s o f C y c l i c 3«,5»-AMP by P a r t i a l l y P u r i -f i e d B r a i n D i e s t e r a s e . The r e a c t i o n mixture contained 0.25 umoles c y c l i c 3*,5*-AMP, 0 . 9 mM MgSO-4, 150 mM T r i s , pH 7.5, 45 ug p a r t i a l l y p u r i f i e d enzyme p r o t e i n , i n a t o t a l volume, o f 0.11 ml. Incubation at 30° was stopped a t 15 minutes by the a d d i t i o n o f 0.02 ml g l a c i a l a c e t i c a c i d . An a l i q u o t (0.02 ml) was spotted on Whatman No. 1 f i l t e r paper and the chromatogram developed w i t h i s o p r o p a n o l -ammonium hydroxide-0.1 M b o r i c a c i d (7:1:2). The "0" and "15" i n d i c a t e d u r a t i o n of i n c u b a t i o n (rain). The solvent f r o n t extended 16 cm from the o r i g i n . 5»-nucleo-t i d e s always remain near the o r i g i n under the c o n d i t i o n s used. 118. FIG. 25 H y d r o l y s i s of C y c l i c 3«,5«-dAMP. The enzyme i n c u b a t i o n c o n d i t i o n s and chromatographic pro-cedures were as described under F i g . 24, except that the sub-s t r a t e was 3«,5*-dAMP. The sol v e n t f r o n t extended 16 cm from the o r i g i n . 119. -v '• V^ ..\..\v.v>' F I G . 26. H y d r o l y s i s of C y c l i c 3»,5*-GMP. The en2yme i n c u b a t i o n c o n d i t i o n s and chromatographic procedures were as described tinder F i g . 24, except that the substrate was c y c l i c 3*,5»-GMP. The so l v e n t f r o n t was 16 cm from the o r i g i n . 120. •K, •i;s JHBHBMHFHHFm FIG. 27 H y d r o l y s i s o f C y c l i c 3»,5»-dGMP. The enzyme i n c u b a t i o n c o n d i t i o n s and chromatographic procedures were as described under F i g . 24, except that the substrate was c y c l i c 3»,5*-dGMP. The solv e n t f r o n t was 14.5 cm from the o r i g i n . 121. As shown in Fig. 28, no 5'-nucleotidase activity was present in the preparation. The presence of, a cyclic 2«,3»-nucle0tide phosphodies-terase activity was reported by Drummond and Perrott-Yee (177) in their ammonium sulfate preparation from rabbit brain. The par t i a l l y purified preparation obtained as described in the present investigation also contained a considerable amount of cyclic 2*,3'-nucleotide phosphodiesterase activity. However, when the fraction obtained by the alkaline-heat, acid preci-pitation method was further fractionated with ammonium sulfate (0.3-0.6 saturation), the cyclic 2«,3"-nucleotide phosphodi-esterase activity was completely eliminated (Fig. 29). Furthermore, the 0.3-0.6 saturated ammonium sulfate fraction yielded up to a 2-fold increase in purification of the 3',5*-nucleotide phosphodiesterase activity having specific a c t i -v i t i e s in the range 15-25. However, the f i n a l ammonium sulfate step gave extremely low yield of enzyme and was not s u f f i -ciently reproducible to merit consideration as a routine tech-nique for purifying the brain diesterase. 4. Cellular Distribution of Cyclic 3,»,5'-nucleotide Phospho- diesterase - Fractionation of rabbit brain into several cellular components was performed as described by De Robertis et a l (185). The results (Table X) indicated that about 50% of the diesterase activity was located in the 105,000 x g supernatant fraction. The microsomal and mitochondrial frac-tions contained considerable amounts of diesterase activity, but l i t t l e activity was located in the nuclear fraction. TABLE X Cellular Distribution of Cyclic 3»,5*-Nucleotide Phos-phodiesterase in Rabbit Brain. The cerebral cortex from a rabbit was homogenized for 5 minutes in 8 volumes of 0.33 M sucrose (unbuffered) with the aid of a glass homogenizer fitt e d with a teflon pestle and centrifuged for 10 minutes at 900 x g. The sediment was washed twice with 0.33 M sucrose, and after re-centrifu-gation at 900 x g, the supernates were combined with the original 900 x g supernate. The mitochondrial fraction was obtained by centrifuging the combined supernates for 20 minutes at 11,500 x g. The sediment was again washed twice with 0.33 M sucrose. The microsomal fraction was obtained by centrifuging the combined 11,500 x g supernates for 30 minutes at 105,000 x g. The microsomal fraction thus obtained was washed once only. A l l procedures were carried out at 0-4°. Enzyme activity is defined in the text. The standard assay was used to determine diesterase activity. Fraction Total Activity (units) % Total Activity Whole homogenate 780 (100%) Nuclear 22 2i8~ Mitochondrial 63 8.1 Microsomal 103 13.2 105,000 x g supernate 394 50.5 Recovery 582 74.9 FIG. 28. Absence of 5'-Nucleotidase A c t i v i t y i n P a r t i a l l y P u r i f i e d B r a i n D i e s t e r a s e F r a c t i o n . The r e a c t i o n mixture contained 1.8 umoles 5»-AMP, 0.9 mM MgSO^ 80 mM T r i s , pH 7.5, p a r t i a l l y p u r i f i e d b r a i n d i e s t e r a s e ( s p e c i f i c a c t i v i t y , 14) and s u f f i c i e n t water to make 0.2 ml. Incubation was f o r 30 minutes at 30°, and the e n t i r e contents o f the r e a c t i o n mixture were spotted on Whatman No. 1 f i l t e r paper and developed as described i n the t e x t . From the l e f t , spot 1 - reference 5»-AMP; spot 2 - r e f -erence adenosine; spot 3 and 4, substrate 51-AMP incubated i n the absence and presence of 13.5 ug enzyme p r o t e i n r e s p e c t i v e l y . The s o l v e n t f r o n t was 17 cm from the o r i g i n . 124 F I G , 29 Absence o f C y c l i c 2«,3*-AMP H y d r o l y t i c A c t i v i t y i n the F i n a l (Ammonium s u l f a t e 0.3-0.6 s a t u r a t i o n ) P r e p a r a t i o n o f B r a i n D i e s t e r a s e . The r e a c t i o n mixture contained 0.24 umoles c y c l i c 2»,3»-AMP, 0.9 mM MgSO,, 80 mM T r i s , pH 7.5, 6 ug b r a i n d i e s t e r a s e p r e p a r a t i o n , i n a t o t a l volume of 0.2 ml. Incubation was f o r 15 minutes a t 30°. A l i q u o t s o f 0.02 ml were taken, spotted on paper and developed as described i n the text. S p e c i f i c a c t i -v i t y of the d i e s t e r a s e p r e p a r a t i o n was 27. The s o l v e n t f r o n t was 14.5 cm from the o r i g i n . 125. While this work was in progress, Cheung and Salganicoff (186) reported that 40% of the total activity was located in the mitochondrial fraction of rat brain. On the other hand, Drummond and Perrott-Yee (177) reported that the activity was localized entirely in the 100,000 x g supernatant fraction. It is therefore d i f f i c u l t to reconcile the differences observed between the results of the present study and those of the other authors. De Robertis e_t a l . (185) have indicated that the f i t t i n g of the teflon plunger had to be specially deter-mined in order to produce a minimal breakage of nerve termi-nals. It is therefore l i k e l y that the differences in the results reported on the cellular distribution of brain dies-terase arise essentially from the extent to which cellu l a r components are disrupted during homogenization. The present study indicated l i t t l e diesterase activity in the nuclear fraction (which also contains fragments of plasma membrane). These findings are in good agreement with those of Cheung and Salganicoff (186) who found only 77. of the activity in this fraction. Since the adenyl cyclase sys-tem which synthesizes cyclic 3*,5*-AMP is located on the plasma membrane, i t would appear that the adenyl cyclase system and the phosphodiesterase activities are spatially separated in the c e l l . It seems that such a spatial arrangement may per-haps be biologically important, since cyclic 3*,5»-AMP must be given time to act before i t s destruction by the diesterase. 5. Distribution of Cyclic 3',51-Nucleotide Phosphodiesterase  in Various Areas of the Central Nervous System and in Lower 126. Organisms - As described earlier, cyclic 3*,5*-AMP is involved in the regulation of several important biological processes. The enzyme, cyclic 3»,5*-nucleotide phosphodiesterase, which terminates the action of cyclic SS^-AMP has been shown to exist in most of the higher organisms which have been examined for i t s activity. Because the diesterase must play a role in the regulation of intracellular cyclic 3*,5*-AMP levels, the distribution of the enzyme was investigated in several tissues; namely, the human brain, the nervous system of the dog, in marine organisms and in the plant kingdom. As may be observed in Table XI, the diesterase activity was highest in the cere-bral cortex of the human brain. The survey of the distribution of diesterase activity in various areas of the dog nervous system (Table XII) also indicated that the enzyme activity was highest in the cerebral cortex. The high content of diesterase activity found in the cerebral cortex i s consistent with that observed by other investigators. The distribution of the enzyme in several available marine organisms which were exa-mined i s shown in Table XIII. The survey of diesterase a c t i -v i t y in various areas of the human brain, dog nervous system and in marine organisms were performed by Mr. Lorne K. Massey, a medical student working in this laboratory. Several available plants were also investigated for diesterase activity. As may be seen in Table XIV, no diester-ase activity was detected in any of the plant specimens which were examined. Several techniques were used to ensure complete disintegration of yeast c e l l s , but no activity was detected. 127. TABLE XI Distribution of cyclic 3»,5*-Nucleotide Phosphodiesterase Ac t i v i t y in Human Brain. Specific activity i s as described in the text. Area of Brain Specific Activity Cerebral cortex - grey 1.6 - white 1.4 Cerebellar cortex 1.0 Pons 0.4 Corpus Callosum 1.0 Thalamus 1.0 Caudate nucleus 1.0 Vermis 0.8 Hypothalamus 0.4 128. TABLE XII Cyclic 3*,5*-Nucleotide Phosphodiesterase Act i v i t i e s in Various Areas of Dog Nervous System. Specific activity is as described in the text. Nervous Tissue Specific Activity Cerebral cortex 3.2 Cerebellum 0.4 Basal ganglion and internal capsule 2.0 Medulla 1.0 Pons 0.3 Spinal Cord - cervical - upper thoracic 0.4 - lower thoracic 0.6 - lumbar 0.2 Midbrain 0.5 Hypothalumus 2.1 Caudate nucleus 2.9 Stellate ganglion 0.4 Thoracic sympathetic ganglion 0.05 Phrenic nerve 0.3 Sciatic nerve 0.2 Thoracic sympathetic axons 0.5 Cervical sympathetic axons 0.1 Cervical superior ganglion 0.1 Nodose ganglion 0.2 Vagus nerve 0.5 Thalamus 3.1 1 2 9 . TABLE XIII Distribution of Cyclic 3*,5"-nucleotide Phosphodiesterase Activity in Several Marine Organisms. Specific activity is as defined in the text. Organism Genus Tissue Specific Activity Steelhead Trout Salmo brain 0 . 2 1 it skeletal muscle 0 . 0 7 Salmon Oncorhynkus heart 0 . 3 4 Sea anemone Metridium musele 0 . 5 4 Tubeworm Nereis (whole organism) 0 . 4 8 Sea urchin Stronglyo-centrotus gonads 0 . 2 6 II n ii intestine 0 . 8 5 Oyster Crosostrea adductor muscle 0 . 0 Clam Mytilus adductor muscle 0 . 0 Snail Thais (whole organism) 0 . 4 5 Hermit Crab Pagarus (whole organism) 0 . 0 Sea cucumber Stichopus longitudinal muscle 0 . 2 5 Crab Cancer g i l l 0 . 0 it II l i v e r 0 . 0 it n pancreas 0 . 0 it it heart 0 . 0 it ti aorta 0 . 0 130. TABLE XIV Distribution of Phosphodiesterase Activity in Plants and Micro-organisms. Plant tissues were washed with 0.15 M KC1, frozen in liquid nitrogen, ground to a powder in a chilled mortar, and further homogenized in suitable volumes of 0.5 M Tris, pH 7.5. Whole homogenates were used for assaying diesterase a c t i v i t i e s , unless indicated otherwise. The assays were performed as des-cribed in the text for the chromatographic method of detecting diesterase activity. A l l negative results were re-examined by using larger volumes of tissue homogenates or extracts and increasing the incubation time to 2 hours. JB. ferrooxidans and E_. c o l i c e l l s were disrupted by sonication at y kc/sec for 30 and 10 minutes, respectively. Specific activity as described in the text. Organism Specific Activity Plants Higher plant leaf - (Genus Tra-descantia) 0 Moss (Liverwort) - (Genus Lunularia) 0 Fungus (mycelium) - Coprinus macrorhizus 0* Yeast - Saccharomyces cerevisiae 0* Algae (Red) - Gymnogongrus norve-gicus 0* Algae (Green) - Spongomorpha coalita 0* Algae (Blue) - Phaeostrophion irregulare - 0* Micro-organisms Bacteria - Bacillus ferrooxidans 0* Bacteria - Escherichia c o l i 0.27* * indicates 37,000 x g supernate and sediment were examined for activity. 131. On the other hand, Cheung (187) has reported recently that extremely low diesterase activity was present in the yeast, Saccharomyces carlsbergensis. Two available microorganisms were also examined for diesterase activity. It was noted that _E. c o l i possessed an appreciable level of phosphodiesterase activity. In this organism, the entire activity was located in the 37,000 x g supernate. After the presence of diesterase activity in E_. c o l i was demonstrated in the present study, Brana and Chytil (188) reported similar observations. These authors also noted that the diesterase activity was located in the supernatant fraction obtained after centrifugation of sonicated E. c o l i c e l l s at 20,000 x g. 132. DISCUSSION The elucidation of the roles of cyclic 3*,5*-AMP is cur-rently under intense study in many laboratories. Experiments with the adenyl cyclase system has led Sutherland (190) to propose a general picture of a two-messenger system for the expression of hormonal control in biological systems. He suggests that a hormone ( f i r s t messenger) interacts with specific effector c e l l s at the plasma membrane. This inter-action results in the formation of a second messenger within the c e l l to modify intracellular enzyme activity. As a specific example, Sutherland cites the stimulation of adenyl cyclase by epinephrine ( f i r s t messenger) which results in the increased biosynthesis of cyclic 3»,5*-AMP (second mes-senger) and consequently, the several physiological effects of epinephrine which are observed. The scheme proposed by Sutherland (190) i s illustrated in Fig. 30. In such a system, the activity of the phosphodiesterase must be equally impor-tant as that of adenyl cyclase in maintaining the required intracellular levels of cyclic 3*,5*-AMP. As yet, no hormonal mechanism for controlling the activity of the phosphodiesterase has been detected. The role of cyclic 3»,5*-nucleotide phosphodiesterase in the brain i s being actively investigated. Cheung and Sal-ganicoff (186) have reported that the diesterase activity was located mainly in the cholinergic nerve endings and in the soluble synaptic neuroplasm. They suggested that the diesterase in brain was probably more closely associated with 133. (EXTRACELLULAR) HORMONE • (FIRST MESSENGER) ATP (INTRACELLULAR) I PHYSIOLOGICAL I RESPONSES. J Glycogenolysis \ Glycolysis Lipolysis V Cardiac inotropic response cyclic 3*,5»-AMP— (SECOND MESSENGER) DIESTERASE — >-5«-AMP FIG. 30 The Two-Messenger Concept for the Expression of Hormonal Control as Modified from Sutherland(190). 134. the regulation of glucose metabolism than in synaptic trans-mission. Studies of several properties of the pa r t i a l l y purified cyclic 3*,5'-nucleotide phosphodiesterase from rabbit brain revealed that the properties of the brain enzyme are very similar to the diesterase which has been purified from cardiac tissue. As reported earlier by Drummond and Perrott-Yee (177), brain diesterase catalyzed the conversion of cyclic 3»,5»-AMP specifically to 5*-AMP. No other product was formed. It was also confirmed that the enzyme requires Mg"*"* ions for activity and was completely inactive in the presence of 1 mM EDTA. Earlier studies on brain and heart diesterases suggested that the activity of the enzyme was more selective for the cyclic 3»,5'-nucleotide which contain purine bases than those with pyrimidine bases. Indeed, the present studies have shown that the enzyme has vi r t u a l l y no activity against pyrimidine cyclic 3» ,5*-nucleotides, with the exception of cyclic 3»,5»-UMP. However, Hardman and Sutherland (184) recently reported that a phosphodiesterase was present in heart that hydrolyzed cyclic 3',5*-UMP at a much faster rate than cyclic 3«,5'-AMP and other available cyclic 3',5«-nucleotides. Their obser-vation therefore suggests that the slight but significant activity against cyclic 3',5«-UMP found in the pa r t i a l l y puri-fied brain preparation may have been due to another diesterase which was specific for cyclic 3*,5'-UMP. It was noted that the hydrolysis of cyclic 3',5'-AMP was consistently inhibited by the presence of cyclic 3»,5«-.dlMP. Preliminary experiments 135. also indicated the inhibition of cyclic 3»,5'-AMP hydrolysis by cyclic 3«,5'-GMP and cyclic 3»,5«-dGMP. The data strongly indicates that cyclic 3',5'-nucleotides possessing purine bases are hydrolyzed by the same enzyme. ATP (0.125 mM and 1.25 mM) did not appear to inhibit the brain diesterase. This i s in contrast to a recent report by Cheung (191) who indicated that brain diesterase was inhibited by ATP and pyrophosphate, and have attributed to ATP a regu-latory role on the enzyme. The specificity of brain diesterase for the cyclic 3«,5»-diester linkage was unequivocally demonstrated when the f i n a l ammonium sulfate fractionation (0.3-0.6 saturation) was per-formed. This f i n a l step effectively removed the cyclic 2»,3«-nucleotide phosphodiesterase activity by precipitating the enzyme into the 0-0.3 saturated ammonium sulfate fraction, leaving a highly purified preparation of cyclic 3*,5»-nucleo-tide phosphodiesterase in the higher ammonium sulfate fraction. The present study also demonstrated that the brain diesterase is stimulated by imidazole. The quantitative response to imidazole i s in agreement with that observed for.heart dies-terase. The KJJJ. value of brain diesterase for cyclic 3*,5»-AMP was about 0.8 x 10"4M, which i s similar to that observed for beef heart diesterase,.but lower than that observed by Nair (176) for dog heart diesterase. The in vitro inhibition of brain diesterase by theophylline observed in the present investigation is also a common feature possessed by the brain and heart enzymes. The nature of theophylline inhibition 136. appears to be competitive in both instances. As one might expect, caffeine and theobromine also inhibits diesterase activity, although i t has been reported (175) that these compounds were only about 1670 as potent as theophylline in this respect. The a b i l i t y of methyl xanthines to inhibit diesterase activity is most l i k e l y due to the similarity in the structures of the methyl xanthines to cyclic 3,,5,-AMP, as indicated in Fig. 31. One might even speculate that the lower potency of caffeine.and theobromine may be due to the presence or -methyl groups on these compounds, which may interfere with their binding to the diesterase. Theophylline has been known for many years to produce a number of pharmacological effects. The compound stimulates the central nervous system, particularly the cerebral cortex, although to a lesser extent than caffeine. However, theo-phylline is the most potent of a l l the methyl xanthines in i t s diuretic action on the kidney, i t s stimulation of cardiac and skeletal muscle, and i t s effect on the relaxation of smooth muscle. These pharmacological responses to theophyl-line administration is l i k e l y due to the inhibition of phosphodiesterase in vivo, which in turn would result in increased levels of cyclic 3',5*-AMP. For example, i t i s known that when skeletal muscle is exposed to methyl xanthines (e.g., caffeine), large quantities of lactic acids are pro-duced. These observations indicate that glycogenolysis and glycolysis were stimulated owing to increased cyclic 3*,5»-AMP levels in the tissues. Similarly, theophylline has been 137 CH 3 CH 3 Theophylline Theobromine (1,3-Dimethyl xanthine) (3,7-Dimethyl xanthine) 0 0. • \ Cyclic 3«,5»-AMP FIG. 31? Structural Formulae of Methyl Xanthines 138. reported to potentiate the cardiac inotropic response to norepinephrine (192), increase l i p o l y t i c activity in adipose tissue (193), stimulate steroidogenesis (159), increase amy-lase secretion from rat parotid gland (163) and increase the permeability of toad bladders to water (166). It would be of particular interest to investigate the possibility that the observed pharmacological effects of caffeine and theo-phylline upon the central nervous system might arise from the in vivo inhibition of brain phosphodiesterase, resulting in higher levels of cyclic 3',5'-AMP in the brain. The method described in this study for the partial puri-fication of cyclic 3 1,5'-nucleotide phosphodiesterase from brain is considerably more rapid and technically easier than that described for the purification of the enzyme from beef and dog hearts. However, this advantage i s offset by the fact that the enzyme preparation generally has a specific activity equal to about one-half that reported for the heart preparations. The use of purified diesterase preparations has aided in the measurement of cyclic 3',5'-AMP in biological materials. For example, Butcher and Sutherland (175) have used the diesterase as a biological tool in order to destroy cyclic 3',5'-AMP in extracts of urine, thus obtaining "tissue blanks" in the assay for the cyclic nucleotide in urine. The universal distribution of cyclic 3',5'-nucleotide phosphodiesterase in higher organisms of the animal kingdom contrasts sharply with the absence of diesterase activity in 139. any of the plant specimens examined in this study. Suther-land (194) reported that no adenyl cyclase activity could be detected in plants. Therefore, these observations suggest that the biological importance of cyclic 3',5'-AMP may be confined largely to those organisms within the animal kingdom. The detection of diesterase activity in marine organisms indicates that cyclic 3',5'-AMP is also widespread in lower animal organisms. The biological importance of cyclic 3',5'-AMP in these organisms is of no less interest than i t s role in higher organisms. The biological role played by cyclic 3',5'-AMP in the physiology of the nervous system is presently attracting widespread attention. Elucidation of i t s precise role in this tissue must await further investigation. 140. BIBLIOGRAPHY 1. Goodman, D.S. Jr., Science 125, 1296 (1957). 2. Johnson, J.A., Nash, J.D. and Fusaro, R.M., Anal. Biochem. _5, 379 (1963). 3. Van Handel, E., and Zilversmit, D.B., J . Lab. C l i n . Med. 50, 152 (1957). 4. Jagannathan, S.N., Can. J . Biochem. 42, 566 (1964). 5. Duncombe, W.G., Biochem. J . 83, 6p (1962). 6. Vaughan, M., Berger, J.E., and Steinberg, D., J . Biol. Chem. 239, 401 (1964). 7. Folch, J., Lees. M., and Stanley, G.H.S., J . Biol. Chem. 226, 497 (1957). 8. Evans, G., J . Physiol. _82, 468 (1934). 9. Newsholme, E.A., and Randle, P.J., Nature 193, 270 (1962). 10. Garland, P.B., Randle, P.J., and Newsholme, E.A., Nature 200, 169 (1963). 11. Parmeggiani, A., and Bowman, R.H., Biochem. Biophys. Res. Comm. 12, 268 (1963). 12. Sutherland, E.W., and Rail, T.W., Pharm. Rev. 12, 265 (1960). 13. Bjorntorp, P., and Furman, R.H., Am. J . Physiol. 203, 323 (1962). 14. Gornall, A.G., Bardawill, C.J., and David, M.M., J . Biol . Chem. 177, 751 (1949). 15. Strand, 0., Vaughan, M., and Steinberg, D., J . Lipid Res. 5, 554 (1964). 16. Kupieki, F.P., J . Lipid Res. 230 (1966). 17. Alousi, A.A., and Mallov, S., Am. J . Physiol. 206, 603 (1964). " 18. Visscher, M.B., and Mulder, A.G., Am. J . Physiol. 94, 630 (1930). 19. Visscher, M.B., Proc. Soc. Exptl. Biol. Med. 38, 323 (1938). 141. 20. Fredrickson, D.S., and Gordon, R.S. Jr., J. Cl i n . Invest. 37, 1504 (1958). 21. Havel, R.J., Naimark, A., and Borchgrevink, C.F., Cl i n . Invest. 42, 1054 (1963). 22. Bing, R.J., et al.Am. J . Med. 16, 504 (1954). 23. Rothlin, M.F., and Bing, R.J., J . C l i n . Invest. 40, 1380 (1961). — 24. Scott, J.C., Finkelstein, L.J., and Spitzer, J.J,, Am. J. Physiol. 203, 482 (1962). 25. Shipp, J.C., and Opie, L.H., Circulation 22, 809 (1960). 26. Opie, L.H., Evans, J.R., and Shipp, J.C., Am. J . Physiol. 205, 1203 (1963). 27. Evans, J.R., Opie, L.H., and Shipp, J.C., Am. J . Physiol. 205, 766 (1963). 28. Evans, J.R., Circulation Res. Suppl. _15_, 96 (1964). 29. Williamson. J.R., and Krebs, H.A., Biochem. J . 80, 540 (1961). . ~ 30. Williamson, J.R., Biochem. J . 93, 97 (1964). 31. Olivecrona, T., J . Lipid Res. 3, 439 (1962). 32. Olivecrona, T., and Beltrage, P., Biochim. et Biophys. Acta. 98, 81 (1965). 33. Gousios, A., Felts, J.M., and Havel, R.J., Metabolism 12, 75 (1963). 34. Delcher, H.K., Fried, M., and Shipp, J.C., Biochim. et Biophys. Acta. 106, 10 (1965). 35. Shipp, J.C., Thomas, J.M., and Crevasse, L., Science 143, 371 (1964). 36. Shipp, J.C., et al.Am. J . Physiol. 207, 1231 (1964). 37. Denton, R.M., and Randle, P.J., Nature 208, 488 (1965). 38. Korn, E.D., J . Biol. Chem. ^ 215, 1 (1955). 39. Korn, E.D., J . Biol. Chem. 215, 15 (1955). 40. Spitzer, J.J., and Gold, M., Am. J . Physiol. 206, 159 (1964). 142. 41. Robinson, D.R., and French, J.E., Pharm. Rev. 12, 241 (1960). 42. Hollett, C , and Meng, H.C., Biochim. et Biophys. Acta. 20, 421 (1956). 43. Cherkes, A., and Gordon, J., J . Lipid Res. 1, 97 (1959). 44. Nakatani, M., Nakamura, M., and Torii,.S., Proc. Soc. Exptl. Biol. Med. 107, 853 (1961). 45. Crass, N.F., and Meng, H.C., Am. J . Physiol. 206, 610 (1964). 46. Robinson, D.S., and Jennings, M.A., J . Lipid Res. 6, 222 (1965). ~ 47. Korn, E.D., and Quigley, T.W., J . Biol. Chem. 226, 833 (1957). 48. Schnatz, J.D., Ormsby, J.W., Williams, R., Am. J . Physiol. 205, 401 (1963). 49. Mallov, S., and Alousi, A., Proc. Soc. Exptl. Biol. Med. 119, 301 (1965). 50. Hollenberg, C.H., J . C l i n . Invest. _39, 1282 (1960). 51. Nikkila, E.A., Torsti, P., and Pentila, 0., Metab. 12, 863 (1963). 52. Nikkila, E.A., Torsti, P., and Pentila, 0., Life Sci. 4, 27 (1965). 53. Borgstrom, B., and Carlson, L.A., Biochim. et Biophys. Acta. 24, 631 (1957). -54. Carlson, L.A., and Wadstrom, L.B., C l i n . Chem. Acta. 2, 9 (1957). 55. Belfrage, P., Biochim. et Biophys. Acta. 98, 660 (1965). 56. Senior, J.R., and Isselbacher, K.J., J . C l i n . Invest. 42, 187 (1963). 57. Tidwell, H.C., and Johnson,. J.M., Arch. Biochim. et Biophys. 89, 79 (1960). 58. McPherson, J.C., Askins, R.E., and Pope, J.C., Proc. Soc. Exptl. Biol. Med. 110, 744 (1962). 59. Pope, J.C. et a l . J . Biol. Chem. 241, 2306 (1966). 143. 60. Adrouny, G.A.,.and Russell, J.A., Endocrinology 59, 241 (1956). •— 61. Warburg, 0., and Christian, W., Biochem. Z. 310,.384 (1941). 62. Desnuelle, P., and Savary, P., J . Lipid Res. 4, 369 (1963). 63. F r i t z , I.B., Physiol. Rev. 41, 52 (1961). 64. Lundsgaard, E., Bull. Johns Hopkins Hosp. 63, 15 (1938). 65. Richardson, H.B., Shorr, E., Loebel, R.O., J . Biol. Chem. 86, 551 ( 1 9 3 0 ) . . . . . . 66. Artom, C , J . Biol. Chem. 213, 681 (1955). 67. Havel, R.J., and Fredrickson, D.S., J . C l i n . Invest. 35, 1025 (1956). 68. Volk, M.E., et a l . J . Biol. Chem. 195, 493 (1952). 69. Geyer, R.P., Matthews, L.W., Stare, F., J. Biol. Chem. 180, 1037 (1949). 70. Wertheimer. E., and Ben-Tor, V., Biochem. J., 50, 573 (1952). — 71. Hansen, R.G., and Rutter, W.J., J . Biol. Chem. 195, 121 (1952). 72*. F r i t z , I.B., et al.Am. jl Physiol. 194, 379 (1958). 73. Friedburg, S.J., et a l . J . C l i n . .Invest. _39, 215 (I960). 74. Issekutz, B. Jr., and Miller, H.I., Proc. Soc. Exptl. Biol.. Med. 110, 237 (1962).. 75. Andres, R., Cader, G., Zierler, K.L., J . C l i n . Invest. 35, 671 (1956). 76. Drummond, G.I., and Black, E.C., Ann. Rev. of Physiology 22, 169 (1960). 77. Gordon, R.S. Jr., and Cherkes, A., J . Clin . Invest. 35, 206 (1956)., — 78. Quastel, J.H.,,and Wheatley, A.H.M., Biochem. J . 2?, 1753 (1933). 79. Vignais, P.M., Gallagher, C.H., and Zabin, I., I. Neurochem..2, 283 (1958). 144. 80. Geiger, A., Physiol. Rev. 38, 1. (1958). 81. Himwich, H.E., and Naham, L.H., Am. J . Physiol. 101, 446 (1932). 82. Shapiro, B., Chowers, I., Rose, G., Biochim, et Biophys. Acta. 23, 115 (1957). 83. Ontko, J.A., and Jackson, D., J . Bio l . Chem. 239, 3674 (1964). 84. Bode, C , and Klingenberg, M., Biochim. et Biophys. Acta. 84,- 93 (1964). 85. Bjorntorp, P., J., Biol. Chem. 241, 1537 (1966). 86. Bing, R.J., J . Mt. Sinai Hosp. (N.Y.) 20,. 100 (1953). 87. Bing, R.J., Am. J . Med. ,15, 284.(1953). 88. Gordon, R.S. Jr., J . C l i n . Invest. _36, 810 (1957). 89. Ballard, F.B., et al»J. Cl i n . Invest. J39, 717 (1960). 90. Neptune, E.M., et a l . J . Biol. Chem. 234, 1659 (1959). 91. Neptune, E.M., et aloJ. Lipid Res. JL, 229 (1960). 92. Masoro, E.J., et a l . J . Biol. Chem. 241, 2626 (1966). 93. Bragdon, J.H., and Gordon, R.S. Jr., J . C l i n . Invest. 37, 574 (1958). 94. Cruikshank, E.W.H., Physiol. Rev. Ij3, 597 (1936). 95. Cavert, H.M., and Johnson, J.A., Am. J . Physiol. 184, 582 (1956). 96. Shipp, J.C., Opie, L.H., and Challoner, D., Nature 189, 1018 (1961). 97. Shipp, J.C., Metabolism 13, 852 (1964). 98. Willebrand, A.F., Biochim. et Biophys. Acta. 84, 607 (1964). • ~~ 99. Hall, L.M., Biochim. Biophys. Res. Comm. j), 177 (1961), 100. Bing, R.J., Physiol. Rev. 45, 171 (1965). 101. Borgstrom, B., and Olivecrona, T., J . Lipid Res. 2, 263 (1961). ~ 145. 102. Olson, R.E., Nature 195, 597 (1962). 103. Stein, 0., and Stein, Y., Biochim. et Biophys. Acta. 70, 517 (1963). 104. Shipp, J.C., Thomas, J.M., Crevasse, L., Circulation 28, 805 (1963). 105. Hahn, P.F., Science .98, 19 (1943). 106. Payza, A.N., Eiber, H., Tchernoff, A., Proc. Soc. Exptl. B i o l . Med. 122, 509 (1966). 107. Rizack, M.A., J . Biol. Chem. 236, 657 (1961). 108. Rizack, M.A., J . Biol. Chem. 239, 392 (1964). 109. Sarda, L., and Desnuelle, P., Biochim. et Biophys. Acta. 30, 513 (1958). 110. Cruikshank, E.W.H., J . Physiol. 47, 1 (1913). 111. Lackey, R.W., Bunde, C.A., G i l l , A.J., Harris, L.C., Proc. Soc. Exptl. Biol. Med. 57, 191 (1944). 112. Lackey, R.W., Bunde, C.A., Harris, L.C., Am. J . Physiol. 145, 470 (1946). 113. Lackey, R.W., Bunde, C.A., Harris, L.C., Proc. Soc. Exptl. Biol. Med. 66, 433 (1947). 114. Russell, J.A., and Bloom, W.C., Endocrinology 58, 83 (1956). ~ 115. Lukens, F.D.W., Am. J. Physiol. 192, 485 (1958). 116. Bowman, R.H., Am. J. Physiol. 197, 1017 (1959). 117. Dole, V., J . Clin . Invest. J35, 150 (1956). 118. Wittels, B., Breseler, R., J . Lab. Invest. 13, 794 (1964). -" 119. Sutherland, E.W., and Cori, C.F., J . Biol. Chem. 188, 531 (1951). 120. Sutherland, E.W., and Wosilait, W.D., Nature 175, 169 (1955). 121. Wosilait, W.D., and Sutherland, E.W., J . Biol. Chem. 218, 469 (1956). 122. Rail, T.W., Sutherland, E.W., and Wosilait, W.D., J . Biol. Chem. 218, 483 (1956). 146. 123. R a i l , T.W., Sutherland, E.W., and Berthet, J . , J . B i o l . Chem. 224, 463 (1957). 124. Sutherland, E.W., and R a i l , T.W., J . B i o l . Chem. 232, 1077 (1958). 125. Sutherland, E.W., and R a i l , T.W., J . Am. Chem. 79, 3608 (1957). 126. L i p k i n , D., Cook, W.H., and Markham, R., J . Am. Chem. Soc. 81, 6198 (1959). 127. R a i l , T.W., and Sutherland, E.W., J . B i o l . Chem. 232, 1065 (1958).. 128. Sutherland,, E.W.., R a i l , T.W., and Menon, T., J . B i o l . Chem. 237, 1220 (1962). 129. Davoren, P.R., and Sutherland, E.W., J . B i o l . Chem. 238, 3016 (1963). 130. Murad, F., C h i , Y.M., R a i l , T.W., and Sutherland, E.W., J . B i o l . Chem. 237, 1233 (1962). 131. K l a i n e r , L.M., C h i , Y.M., F r i e d b e r g , S.L., R a i l , T.W., and Sutherland, E.W., J . B i o l . Chem. 237, 1239 (1962). 132. Krebs, E.G., DeLange, R.J., Kemp, R.G., and R i l e y , W.D., Pharm. Rev. 18, 163 (1966). 133. Drummond, G.I., Duncan, L., and F r i e s e n , A.J., J . B i o l . Chem. 240, 2778 (1965). 134. Haynes, R.C. J r . , and Be r t h e t , L., J . B i o l . Chem. 225, 115 (1957). 135. Haynes, R.C. J r . , J . B i o l . Chem. 233, 1220 (1958). 136. Belocopitow, E., Arch. Biochem. Biophys. 93, 457 (1961). 137. R o s e l l - P e r e z , M., and Lamer, J . , Biochem. J . 3, 81 (1964). . " 138. Mansour, T.E., Sutherland, E.W,, R a i l , T.W., and Beuding, E., J . B i o l . Chem. 235, 466 (I960). 139. Mansour, T.E., J . Pharm. 135, 94 (1962). 140. Mansour, T.E., and Mansour, J.M., J . B i o l . Chem. 237, 629 (1962). . . . 141. Mansour, T.E., J . B i o l . Chem. 238, 2285 (1963). 147. 142. Mansour, T.E., J . Biol. Chem. 240, 2165 (1965). 143. Passoneau, J.V., and Lowry, O.H., Biochem. Biophys. Res. Commun. 10 (1962). 144. Rizack, M., J . Biol. Chem. 239, 392 (1964). 145. Butcher, R.W., Ho, R.J., Meng, H.C., and Sutherland, E.W., J. Biol. Chem. 240, 4515 (1965). 146. Knox, W.E., Piras, M., and Tokuyama, K., Fed. Proc. 24, 474 (1965). 147. Chytil, F., and Skrivanova, J., Biochim. Biophys. Acta. 67, 164 (1963). 148. Gray, G.D., Arch. Biochem. Biophys. 113, 502 (1966). 149. Robison, G.A., Butcher, R.W., Oye, I., Morgan, H.E., and Sutherland, E.W., J . Mol. Pharmacol. 1,. 168 (1965). 150. Mayer, S.E.,. Cotten, M. deV., and Moran, N.C., J . Pharm. 139, 275 (1963). 151. Drummond, G.I., Valadares, J.R.E., and Duncan, L., r Proc. Soc. Exptl. Biol. Med. 117, 307 (1964). 152. Haynes, R.C., Koritz, S.B., and Peron, F.G., J. Biol. Chem. 234, 1421 (1959). 153. Roberts, S., Creange, J.E., and Fowler, D.D., Nature 203, 759 (1964). 154. Roberts, S., Creange, J.E., and Young, P.L., Nature 207, 188 (1965). 155. Creange, J., and Roberts, S., Biochem. Biophys. Res. Commun. 19, 73 (1965). 156. Roberts, S., Creange, J.E., and Young, P.L., Biochem. Biophys. Res. Commun. 20, 446 (1965). 157. Creange, J.E., and Roberts, S., Steroids J5, 13 (1965) „ 158. Karaboyas, G.C., and Koritz, S.B., Biochem. 4, 462 (1965). 159. Darrington, J.H., and Kilpatrick, R., J . Physiol. 182, 16p (1966). 160. Pryor, J., and Berthet, J., Biochim. Biophys. Acta. 43, 556 (1960). 148. 161. Exton, J.H., and Park, C.R., Fed. Proc. 24, 537 (1965). 162. Exton, J.H., and Park, C.R., Pharmacol. Rev. 18, 181 (1966). — 163. Bdolah, A., and Schramm, M., Biochem. Biophys. Res. Commun. 18, 452 (1965). . 164. Orloff, J., and Handler, J.S., J . C l i n . Invest. 41, 702 (1962). 165. Strauch, B.S., and Langdon, R.G., Biochem. Biophys. Res.-Commun. JL6, 27 (1964). . 166. Handler, J.S., Butcher, R.W., Sutherland, E.W., and Orloff, J., J . Biol. Chem. 240, 4524 (1965). 167. Orloff, J., and Handler,,J.S., Am. J . Med. 36, 686 (1964) . "~" 168. Berthet, J., Proc. 4th Int. Cong. Biochem. 15, 107 (1958). 169. Tarui, S., Nonaka, K., Ikura, Y., and Shima, K., Biochem. Biophys. Res. Commun. 13, 329 (1963). 170. Harris, J.B., and Alonso, D., Gastroenterology 44, 830 (1963). ~"~ 171. Harris, J.B., and Alonso, D., Fed. Proc. 24, 1368 (1965) . "~ 172. Sutherland, E.W., and Robison, G.A., Pharmacol. Rev. 18, 145 (1966). 173. Butcher, R.W., Pharmacol. Rev. 18, 237 (1966). 174. Mansour, T.E., Pharmacol. Rev. 18, 173 (1966). 175. Butcher, R.W., and Sutherland, E.W., J . Biol. Chem. 237, 1244 (1962). 176. Nair, K.G., Biochem. 5, 150 (1966). 177. Drummond,, G.I., and Perrott-Yee, S., J . Biol. Chem. 236, 1126 (1961). 178. Smith, M., Drummond, G.I., and Khorana, H.G., J . Am. Chem. Soc. J33, 698 (1961). 179. Drummond, G.I., Gilgan, M.W., Reiner, E.J., and Smith, M., J. Am. Chem. Soc. 86, 1626 (1964). . 149. 180. Smith, M., Moffatt, J.G., and Khorana, H.G., J . Am. Chem. Soc. 80, 6204 (1958). 181. Fiske, C.H., and Subbarow, Y., J . Biol. Chem. 66, 375 (1925). — 182. Gornall, A.G., Bardawill, C.S., and David, M.M., J . Biol . Chem. 177, 751 (1949). 183. Warburg, 0., and Christian, W., Biochem. Z. 310, 384 (1941). 184. Hardraan, J.G., and Sutherland, E.W., J . Biol. Chem. 240, 3704 (1965). 185. De Robertis, E., De Ir a l d i , A.P., De Lores Arnais, G.R., .and Salganicoff, L., J . Neurochem. _9, 23 (1962). 186. Cheung, W.Y., and Salganicoff, L., Fed. Proc. 25, 714 (1966). 187. Cheung, W.Y., Biochim. Biophys. Acta. 115, 235 (1966). 188. Brana, H., and Chytil, F., Folia Microbiol. 11, 43 (1966). 189. Drummond, G.I., Iyer, N.T., and Keith, J., J . Biol. Chem. 237, 3535 (1962). . . . 190. Sutherland, E.W., Oye, I., and Butcher, R.W., Rec. Prog. Hormone Res. 21, 623. (1965). 191. Cheung, W.Y., Biochem. Biophys. Res. Commun. 23, 214 (1966). . 192. Rail, T.W., and West, T.C., J . Pharm. Exptl. Therap. 139, 269 (1963). 193. Hynie, S., Krishna, G., and Brodie, B.B., J . Pharm. Exptl. Therap, 153, 90 (1966). 194. Sutherland, E.W., Harvey Lectures 57, 17 (1961). 


Citation Scheme:


Citations by CSL (citeproc-js)

Usage Statistics



Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            async >
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:


Related Items