THE INFLUENCE or BREED.) same I ‘ AGE on ueoeemceuzma Mme: , mmwmm V . , v . Thesis for the Degree of Ph; D. MTCHE-GAN STATE UNIVERSITY ALLEN FLOYD PARR 1 973 LIBR/ZR Y Michigan State University This is to certify that the thesis entitled THE INFLUENCE OF BREED, SEX AND AGE 0N LIPOGENIC ENZYME ACTIVITIES;IN SOME OVINE TISSUES presented by Allen Floyd Parr has been accepted towards fulfillment of the requirements for Ph.D. Animal Husbandry degree in Date February 28, 1973 0-1639 I I Hi 80% BMW "l x; “K VIBRV";*\' .;F".’ y} 1m 3! I: i. "We suns T "w'mz {Mil-3936" 9; W -' T 6‘ -. . . "-- VJIZYT‘) hut . . _ " ‘1: , ' L: V “autumn. . =1 T _ c .T.e:~-w:.~ K L f! I- J g u T Y 2““ Cf . SOUL.~u-;. ..' . ‘ ! '.. 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CE! ardent“ my {filter 38! «swine; than em 1m had-tutti; hm” (I .33» 03W I «1‘4: 3“ a“ 32 “a. ~. r-— ABSTRACT THE INFLUENCE OF BREED, SEX AND AGE 0N LIPOGENIC ENZYME ACTIVITIES IN SOME OVINE TISSUES by Allen Floyd Parr An experiment involving 49 lambs was conducted to study the influence of breed, sex and age on lipogenic enzyme activities. A commercial flock of western ewes was divided into two groups based on body size and mated to either a Southdown or Suffolk ram in order to obtain two strains of lambs differing in their propensity to fatten. Lambs were slaughtered at birth, 24 hours, 8, 16 and 32 weeks of age and blood, liver, longissimus muscle (In), perirenal (PRF) and subcutaneous fat (SCF) samples were collected from the carcasses. Enzymes assayed were acetyl COA synthetase (SYN), ATP-citrate lyase (CCE), acetyl COA carb0xylase (CBX), glucose-6- phosphate dehydrogenase (G-6PDH), 6-phosphogluconate dehydrogenase (6-PGDH), NADP-malate dehydrogenase (ME), NADP-isocitrate dehydrogenase (ICDH) and lipoprotein lipase (LPL). Plasma free fatty acids (FFA) were also deter- mined. SYN activities were significantly (P <.05) higher in the adipose tissues than in liver and 1M. Tissues of Seuthdowns, which have a greater propensity to fatten, had greater SYN activities than those of Suffolks. The significant (P <.05) decrease in SYN activity observed in SCF from 16 to 32 weeks of age may reflect a trend of decreased rate of fat deposition (lipogenic activity) in lambs between 16 and 32 weeks. CCE activities were Vsignificantly (P <.05) higher in PRF than in liver and 1M. All tissues of Suffolk lambs showed greater CCE activities than those of Southdowns. PRF of neonatal and 24 hour lambs had significantly (P <.05) greater CCE acti- vities than those at 8, 16 and 32 weeks. _.- ~..—— ,H Allen Floyd Parr CBX activities were highest in SCF followed in order by liver, PRF and LM. Adipose tissues of Southdowns (greater propensity to fatten) ex- hibited higher CBX activities than those of Suffolks. PRF and SCF of rams and ewes at 32 weeks shOwed significantly (P <.05) greater CBX activities than those at 8 and 16 weeks; whereas, wethers at 32 weeks had lower activities of this enzyme than those at 16 weeks. CBX activities appeared to be closely related to the level of lipogenic activity in ovine adipose tissues. G-6PDH activities were significantly (P <.05) higher in adipose tissues than in liver and LM. There did not appear to be a relationship between G-6PDH activity and the propensity to fatten in any of the tissues studied. The activity of G-6PDH reached its maximum at 16 weeks in PRF and SCF and this observation appeared to correspond to the high level of lipogenic activity generally associated with ovine adipose tissue depots at this age. SCF possessed the greatest 6-PGDH activities followed in order by liver, PRF and LM. The tissues of Southdowns exhibited higher 6-PGDH - activities than those of Suffolks and this suggested that 6-PGDH was re- lated to the propensity to fatten. The activities of 6-PGDH in PRF and SCF increased considerably with age and this obServation.indicated that the activities of this enzyme in lamb adipose tissues were closely asso- ciated with the level of lipogenic activity. Adipose tissue ME activities were significantly (P <.05) higher than those in liver and TM. The trend toward increasing ME activities with age in lamb adipose tissues followed the general pattern for lipogenic activity in growing lambs. However, ME contributed Only a minor proportion '_‘__ Allen Floyd Parr of the total NADPH generating potential in ovine adipose tissues. Adipose tissues shaved greater ICDH activities than LM and liver. ICDH activities did not appear to be closely related to the propensity to fatten. The response of ICDH to different levels of lipogenic activity was generally associated with age between 8 and 32 weeks. However, the presence of significant ICDH activities in Ovine adipose tissues indicated that ICDH has the potential to generate NADPH for lipogenesis. SCF and PRF had significantly (P <.05) greater LPL activities than LM. LPL activities in ovine adipose tissue depots did not appear to be related to the propensity to fatten. PRF of lambs at 8 weeks exhibited significantly (P <.05) higher LPL activities than those at birth, 24 hours and 32 weeks and this observation seemed to coincide to the high rate of fat depOsition usually found in PRF of lambs at 8 weeks of age. Changes in plasma FFA levels usually reflected changes in the energy state and/or the development of rumen function. THE INFLUENCE or BREED, SEX AND AGE 0N LIPOGENIC ENZYME ACTIVITIES IN SOME OVINE TISSUES BY Allen Floyd Parr A THESIS Submitted to Michigan State University rimpartial. fulfillment of the reguirements for the degree of .»u:— E . ..t A , - 90cm (F PHILOSOPHY . at. , 5.. 3..., , Department of Animal Husbandry -1973 : ' Ir‘"- . . --m‘h-_-_. ACKNOWLEDGMENTS The author wishes to express his appreciation and thanks to his major professor, Dr. R. A. Merkel, for his support and guidance throughOut this research and for his assistance in the preparation of this manuscript. Appreciation is expressed to Dr. H. A. Henneman for his help in designing the experiment and in obtaining the experimental animals for this study. The author also expresses his thanks to Dr. R.M. Cook, Dr. R. S. Emery and W. C. Deal for contributing enzyme procedures and technical assis- tance. Appreciation is expressed to Dr. L. R. Dugan, Dr. w. G. Bergen and Dr. R. W. Dukelow for serving as members of the guidance committee. Appreciation is also expressed to Dr. G. D. Riegle for serving as substi- tute on guidance committee in the absence of Dr. R. W. Dukelow and to Dr. W. T. Magee for assistance with statistical analysis. Special thanks are extended to George Good for the care and manage- ment of the lambs and to Paul Schurman for his help in slaughtering and collecting samples. Thanks are due Mrs. Dora Spooner for assistance in the laboratory analyses. Special appreciation is expressed to Mrs. Bea Eichelberger for typing the manuscript. The author is also grateful to other faculty, staff and especially fellow graduate students in the Meat Science Laboratory for assistance extended this research study. The author wishes to express sincere appreciation to his parents, Mrs. Hattie Parr and Mr. Dewey Parr, for their understanding and dedicated assistance. ii To the Patrick Brown and Cory Phillips familieS, the author expresses appreciation for their encouragement and friendship throughout the course of this degree. TABLE OF CONTENTS Page wcTIm I U I U . I O I I I I I I I l I I O I I O 0 O I I I I 1 Sites of Lipid Biosynthesis . . . . . . . . . . . . . . . . . 4 Intracellular Localization of Fatty Acid Synthesis . . . . . 6 o o ‘1 Anatomical Location and Adipose Tissue Cellularity Lipogenesis . . . . . . . . . . . . . . . . . . . . . . . . . 12 __Substrate for Lipogenesis . . . . . . . . . . . . . . . . . . 13‘ Non-ruminant . . . . . . . . . . . . . . . . . . . . . l3 Rminan t I I I I U I O I O I I I I I I I I I I I I I I O 15 Métabolism in the Young Ruminant . . . . . . . . . . . . . . 22 . Liposeflic PrECursors - o a I o c a o I a o c o u o o i '0 s a 24 «.sourcesofACetleOA...-........'.........26 ‘ l m~minant . I I I I -. I I Q I . I I I . I I I I 0 I l 28 minant . I I . I I I . I I U . . C l O I I I I I C C I 33 .-Malonyl GOA Formation . . . . . . . . . . . . . . . . . . . . 35 U."Sourcesl of Reducing Equivalents 38 m‘m‘neport- of Lipids I I I I I I I I I I I I I I I I I I I I I 43 4,8 Potential Sources of Reducing Equivalents for Lipogenesis Glucose- 6- -Phosphate Dehydrogenase and 6- -Phosphog1uconate Dehydrogenase . . . . . . . . . . . . . . . NADP-Malate Dehydrogenase . . . . . . . . . . . . . . . NADP- Isocitrate Dehydrogenase . . . . . . . . . . . . . Lipoprotein Lipase . . . . . . . . . . . . . . . . . . Free Fatty Acids . . . . . . . . . . . . . Statistical Analysis . . . . . . . . . . . . . . . . . . . RESULTS AND DISCUSSION . . . . . . . . . . . . . . . . . . . . . Acetyl COA Generation . . . . . . . . . . . . . . . . . . . Acetyl COA Synthetase . . . . . . . . . . . . . . . ATP-Citrate Lyase . . . . . . . . . . . . . . . . . Malonyl COA Formation . . . . . . . . . . . . . . . Acetyl COA Carb0xylase . . . . . . . . . . . . . . . . Glucose- 6- -Phosphate Dehydrogenase and 6- -Phosphogluconate Dehydrogenase . . . . . . . . . . . . . . . . NADP- -Ma1ate Dehydrogenase . . . . . . . . . . . . . . . NADP-Isocitrate Dehydrogenase . . . . . . . . . . . . Circulating Triglyceride Uptake . . . . . . . . . . . . . . Lipoprotein Lipase . . . . . . . . . . . . . . . . . . LITERATURE CITED . . . . . . . . . . . . . . . . . . . . . . . . . c o a o o o o o c o c c u e a g o o c o n u .5 o a s s o 75 76 77 77 92 98 99 108 109 120 126 146 146 153 179 Table 10 11 11A 312 12A 13 13A 14 14A LIST OF TABLES Experimental design . . . . . . . . . . . . . . . . . Composition of concentrate mixtures . . . . . . . . . Reagents and assay design for acetyl COA synthetase . Reagents and assay design for acetyl COA carb0xylase Reagents and assay design for citrate cleavage enzyme Reagents and assay design for glucose— —6- -phosphate dehydro- genase and 6-phosphogluconate dehydrogenase . . Reagents and assay design for malic enzyme . . . . Reagents and assay design for isocitrate dehydrogenase Reagents and assay design for lipoprotein lipase . . Reagents and assay design for free fatty acids . . Cytoplasmic acetyl COA synthetase activity in various tissues by breed, sex and age . . . . . . . . . . . . Breed X sex, breed X age and sex X age interactions of cytoplasmic acetyl COA synthetase activity in various tissues . . o . . . . . . . . . . . . . . . . . . . . Mitochondrial acetyl COA synthetase activity in various tissues by breed, sex and age . . . . . . . . . . . Breed X sex, breed X age and sex X age interactions of mitochondrial acetyl COA synthetase activity in various tissues . . . . . . . . . . . . . . . . . . . . . . Total acetyl COA synthetase activity in various tissues breed, sex and age . . . . . . . . . . . . . . . . . Breed X sex, breed X age and sex X age interactions of total acetyl COA synthetase activity in various tissues . . u ATP-citrate lyase activity in various tissues by breed, sex and age . . . . . . . . . . . . . . . . . . . . . Breed X sex, breed X age and sex X age interactions of ATP-citrate lyase in various tissues . . . . . . . . vi by Page 55 56 60 65 67 69 71 72 74 82 84 86 88 89 91 95 97 Table 15 16 16A 17 18A 19 19A 20 20A 21 21A 22 Page Acetyl COA carb0xylase activity in various tissues by breed, sex and age . . . . . . . . . . . . . . . . . . . . 101 Breed X sex, breed X age and sex X age interactions of acetyl COA carb0xylase activity in various tissues . . . . . . . . 103 Glucose-6-phosphate dehydrogenase activity in various tissues by breed, sex and age . . . . . . . . . . . . . . . 113 Breed X sex, breed X age and sex X age interactions of g1ucose-6-phosphate dehydrogenase activity in various tissues . . . . . . . . . . . . . . . . . . . . . . . . . . 114 6-Phosphogluconate dehydrogenase activity in various tissues by breed, sex and age . . . . . . . . . . . . . . . 116 Breed X sex, breed X age and sex X age interactions of 6- phosphogluconate dehydrogenase activity in various tissues 118 NADP-malate dehydrogenase activity in various tissues by breed, sex and age . . . . . . . . . . . . . . . . . . . . 123 Breed X sex, breed X age and sex X age interactions of NADP- malate dehydrogenase activity in various tissues . . . . . 125 Cytoplasmic NADP-isocitrate dehydrogenase activity in various tissues by breed, sex and age . . . . . . . . . . . . . . . 131 Breed X sex, breed X age and sex X age interactions of cytoplasmic NADP-isocitrate dehydrogenase activity in various tissues . . . . . . . . . . . . . . . . . . . . . . 133 Mitochondrial NADP-isocitrate dehydrogenase activity in various tissues by breed, sex and age . . . . . . . . . . . 135 Breed X sex, breed X age and sex X age interactions of mitochondrial NADP-isocitrate dehydrogenase activity in various tissues . . . . . . u . . . . . . . . . . . . . . . 137 Total NADP-isocitrate dehydrogenase activity in various tissues by breed, sex and age . . . . . . . . . . . . . . . 139 Breed X sex, breed X age and sex X age interactions of total NADP-isocitrate dehydrogenase activity in various tissues . 140 Proportion of the total NADPH generating potential of dehydrogenases in adipOSe tissues at various ages . . . . . 142 Page . Wfiotein lipase activity and free fatty acids in various .4 .‘ssuesby-breed, sexandage . . . . . . . . . . . . . . . 148 11.! sex, breed X age and sex X age interactions of s :0 Q I m o I o m u u a I o o o e I o n o s o I o l 150 a"? ; ~~-, #:41'551‘ 'u Figure l 2 10 ll 12 13 14 15 LIST OF FIGURES Page 23 novo fatty acid synthesis from glucose and acetate . . 25 Intracellular location of the reaction sequences involved in the formation of acetyl COA from glucose and acetate . 27 Citrate cleavage pathway and its relationship to lipogenesis from glucose and acetate . . . . . . . . . . . . . . . . 3 Pathways mOSt likely involved in the transfer of oxaloacetic acid between the fat cell cytoplasm and mitochondria . . 32 Pathways of reducing pOwer (NADPH) generation for do novo fatty acid synthesis . . . . . . . . . . . . . . . . 39 Diagram of the transport and uptake of triglycerides and fatty acids by ruminant plasma and their metabolism in tissues . . . . . . . . . . . . . . . . . . . . . . . . . 44 Means and standard error of the means of cytoplasmic acety1 COA synthetase activity in various tissues . . . . . . . 78 Means and standard error of the means of mitochondrial acetyl COA synthetase activity in various tissues . . . . 79 Means and standard error of the means of total acetyl COA synthetase activity in various tissues . . . . . . . 80 Means and standard error of the means of ATP-citrate lyase activity in various tissues . . . . . . . . . . . . 94 Means and standard error of the means of acetyl COA carb0xylase activity in various tissues . . . . . . . . . 100 Means and standard error of the means of glucose-6- phosphate dehydrogenase activity in various tissues . . . 110 Means and standard error of the means of 6- -phosphog1uconate dehydrogenase activity in various tissues . . . . . . 1 Means and standard error of the means of NADP-malate dehydrogenase activity in Various tissues . . . . . . . . 121 Means and standard error of the means of cytoplasmic NADP-isocitrate dehydrogenase activity in various tissues 127 ix Page 3;“: and standard error of the means of mitochondrial ”‘ NADP-isocitrate dehydrogenase activity in various tissues 128 Means and standard error of the means of total NADP- isocitrate dehydrogenase activity in various tissues . . 129 Means and standard error of the means of lipoprotein lipase activity in various tissues . . . . . . . . . . 147 LIST OF APPENDIX TABLES Appendix Page I Ewe grain rations . . . . . . . . . . . . . . . . . . 179 II Raw data . . . . . . . . . . . . . . . . . . . . . . . 180 III Raw data . . . . . . . . . . . . . . . . . . . . . . . 187 IV Simple correlations - liver . . . . . . . . . . . . . 196 V Simple correlations - longissimus muscle . . . . . . . 197 VI Simple correlations - perirenal fat . . . . . . . . . 198 VII Simple correlations - subcutaneous fat . . . . . . . . 199 xi 431“: INTRODUCTION In our affluent society where food is plentiful and constantly avail- able, excessive adiposity is common and it represents a definite health hazard to man. In addition, animal fats are frequently incriminated in the problem of cardiovascular diseases and because of the negative attitude toward these fats they are a source of economic concern to the meat animal industry. Consequently, the need exists for minimizing the waste fat depots in livestock because of the low demand for animal fats and because production efficiency decreases during the fattening phase, waste fat results in high production costs. The U.S.D.A. in 1966 reported that about two billion pounds of fat was trimmed from beef carcasses alone. This represented about 500 million dollars in feed costs, but was worth only about 100 million dollars as a by product. However, considerable progress has been made towards the production of leaner animals as evidenced by the decrease in lard production by approximately 30 pounds per pig over the past 20 years. Even so, the average meat animal still contains ex- cessive fat especially in the subcutaneous and perirenal depots. This presents the meat animal industry with the monumental task of increasing efficiency of production coupled with the conversion of a higher proportion of the caloric intake to increased protein and the edible portion. As a consequence of the above problems there is a genuine concern in further- ing our understanding of the regulation of adiposity in man and meat animals. The controversy surrounding the relationship between animal fat and cardio- vascular disease as well as other human health aspects adds even greater impetus to the need for controlling adiposity in meat animals. The in- formation gained from the investigations of the biochemical, physiological, hormonal and metabolic control mechanisms of lipogenesis in domestic ani- mals will not only be of use in the regulation of fat deposition in meat animals, but could play a vital role in the control of adiposity and related diseases in man. while improvement in production efficiency and carcass composition of the meat animals has been made through dietary manipulation, the use of synthetic hormones and genetic selection, there is an urgent need for a thorough understanding of regulating adiposity which would involve a more direct interaction with biochemical events in adipose tissue. This need is even more critical since some growth stimu- lants such as diethylstilbestrol, are no longer permitted in meat animal feeds. The regulation of fat deposition requires an understanding of the biochemical and physiological sequence of events in lipid biosynthesis and transport. Adipose tissue is a metabolically active tissue with very adaptive and precisely regulated biochemical and physiological functions. A great deal of information has been amassed with regard to the metabolic pathways in adipose tissue and the enzymes catalyzing individual steps of these pathways (Renold and Cahill, 1965; Lehninger, 1970). Virtually all the available information has been collected from investigations involving non-ruminant animals, particularly laboratory animals. Physiological, neural and hormonal mechanisms synchronize the balance between lipogenesis and lipolysis (Adler and Wertheimer, 1967). Many of the metabolic steps are involved in the conversion of glucose or acetate into lipids and several are considered as possible rate-limiting factors. Research efforts on intracellular metabolic control have been directed towards four major areas: 1) the ability of adipOSe tissue to synthesize acetyl COA from glucose or acetate; 2) the level of gghngyg fatty acid synthesis by controlling the incorporation of acetyl COA into long chain fatty acids; 3) the ability to supply the reducing equivalents necessary for fatty acid synthesis and 4) the availability of a-glycerophosphate for triglyceride synthesis. Thus, this study was undertaken to observe the influence of breed, sex and age on enzymatic activity and lipid biosynthesis in lambs of two strains of sheep differing in their propensity to adiposity with the following specific objectives: 1. To study the influence of breed, sex and age on the ability of various tissues to convert acetate to acetyl COA. 2. To observe the effect of breed, sex and age on the capacity of various tissues that provide the reducing equivalents (NADPH) for lipogenesis. 3. To determine the influence of breed, sex and age on the uptake of lipids by various tissues. r—vW.——.—~— LITERATURE REVIEW Sites of Lipid Biosynthesis The study of lipid metabolism has led to concentrated research efforts on the lipogenic capabilities of liver, adipose and mammary tissue. The liver has long been regarded as the primary site of lipogenesis as Pihl and Bloch (1950) in early work, using isotope tracers, found that liver had a high turnover rate of long chain fatty acids. It was concluded from this observation that the liver had a high lipogenic capacity. These investigators also stated that the appearance of labeled triglycerides in extrahepatic tissue must have resulted to a large extent from the uptake of lipids from the bloodstream. However, just prior to this finding Masoro _£._l- (1949) presented evidence indicating that the conversion of glucose to fatty acids was not limited to a single tissue in the adult rat. This evidence was based on the finding that labeled palmitic acid was isolated in extrahepatic tissues of hepatectomized adult rats. Later, Feller (1954) reported that adipose tissue was capable of active catabolic and anabolic lipid metabolism and that adipose could readily synthesize fatty acids from acetate. Furthermore, when the values obtained for fatty acid synthesis were expressed on a fat free or total nitrogen basis, adipose tissue formed fatty acids at rates equal to or greater than those found for liver slices. The biosynthetic pathways are apparently similar in both liver and adip0se tissues, but because of their differences in response to hormones the extent and control of fatty acid biosynthesis may be markedly affected. Much of the present day research efforts have been directed toward determining the relative contribution made by liver and adipose tissue to total body lipogenesis. The findings of Jansen g3 El. (1966) indicate that adipose tissue in rats and mice accounts for at least 50 percent of the newly synthesized fatty acids and O'Hea and Leveille (1968; 1969a, b) reported that pig adipose tissue possessed all the attributes of an active lipogenic tissue. Also, the almost complete absence of citrate cleavage enzyme in porcine liver is undoubtedly an important factor contributing to its low capacity for synthesis of fatty acids from glucose and is consistent with the results of O'Hea and Leveille (1969b) indicating that adipose tissue accounted for 99 percent of the newly synthesized fatty acids in the pig. Thus, these results indicate that adipose tissue plays a major role, if not nearly the exclusive role in fatty acid synthesis in pig. Similarly, Ballard _£ 21. (1969) in a review of lipogenesis stated that in ruminants, where acetate is the major precursor for fatty acid synthesis, adipose tissue appears to be the most important site of fatty acid synthesis and recently Hood (1972) reported that in Holstein steers adipose tissue was the only important lipogenic tissue. In contrast, O'Hea and Leveille (1969c) reported that in the chicken the liver is the major site of lipogenesis and Goodridge and Ball (1967) reported similar findings when studying lipogenesis in the pigeon. In ruminants, the short chain fatty acids, up to a chain length of 014 and part of the 016 acids, in milk fat Originated by g; novo fatty acid synthesis from acetate and B-hydrOxybutyrate absorbed by the mammary .IiI. ’vw—v—m-W- . - , - 5-.- _| - tissue from the blood (Bickerstaffe, 1971). Earlier Annison et 1. (1967) working with goats and Bishop _£-§l. (1969) working with cows reported that the long chain fatty acids, i.e., 18 or more carbon atoms and part of the fatty acids with 16 carbon atoms, in milk fat are derived from plasma lipids. Furthermore, Linzell (1968) showed that even though sub— stantial glucose is absorbed by the mammary gland, it does not contribute to the synthesis of milk fatty acids. In contrast, Linzell t l. (1969) and Spincer _£._l' (1969) reported that in non-ruminants, milk fatty acids (including a number of short chain fatty acids) are exclusively derived from glucose. Moreover, the latter authors found that fatty acids with chain length up to C18 were synthesized from both glucose and acetate and the balance of the fatty acids were contributed by the plasma lipids. Intracellular Localization of Fatty Acid Synthesis At least three intracellular sites of fatty acid synthesis are recognized: (I) cytoplasmic, (2) mitochondrial and (3) microsomal. The enzyme systems of the cytoplasmic and mitochondrial sites are similar, yet differences do exist since the end product of the cytoplasmic synthetic system is primarily palmitic acid and that of the mitochondria primarily stearic acid (Masoro, 1968). The extramitochondrial enzyme system is by far the most active and is probably the system chiefly responsible for the dg novo synthesis of fatty acids. The system located in the cytosol is responsible for the fig novo synthesis of the entire fatty acid molecule. This so-called fatty acid synthetase system requires acetyl COA as a "sparker" molecule, malonyl COA as substrate and NADPH as hydrogen donor (Lehninger, 1970). Harlan and Wakil (1963) reported that fatty acid synthesis innitochondria is based on at least two mechanisms. The first mitochondrial system is similar to the cytoplasmic system, while the second system consists of the elongation process and requires acetyl COA as substrate and NADPH as hydrogen donor to elongate acyl-COA esters by two carbon increments (Donaldson _£__l., 1970). In addition to the two classical mechanisms (cytoplasmic and mitochon- drial) of fatty acid synthesis and elongation, a mechanism of synthesis in microsomes was postulated by Nugteren (1963). According to Masoro (1968), this enzymatic sequence can elongate either saturated or unsaturated fatty acids by reacting malonyl COA with the long chain fatty acyl COA esters to generate a series of COA intermediates with the ultimate formatiOn of fatty acyl COA esters two carbons longer. In recent experiments de- signed to study the mechanism of microsomal fatty acid synthesis, Rous (1971) faund that the reducing equivalents were supplied by NADPH. More— over, Donaldson _£__l. (1970) suggested that the microsomes may play a key role in the control of fatty acid synthesis since the elongation and/or desaturation of palmitic acid synthesized gg_ggyg would provide a spectrum of fatty acids for subsequent estrification that would confer the proper physical characteristics upon lipids stored in various fat depots. Anatomical Location and Adipose Tissue Cellularity Recent advances in the measurement of the cellularity of adipose tissue have made it possible to study the effect of fat cell size and number upon fat metabolism. Several investigators have confirmed the hypothesis that adipOSe tissue mass can increase by either cell prolifera- tion (hyperplasia) or by cell enlargement (hypertrophy) or a combination of the two. Hirsch and Han (1969) faund that early growth of rat epidymal fat pads and retroperitoneal depots was accompanied by progressive enlarge- ment of adipose cells as well as by increases in number. Moreover, beyond the 15th week, the depot grew exclusively by the process of cellular enlargement, with no further change in cell number. In an experiment designed to study the changes in the total carcass adipose tissue of young pigs, Anderson and Kauffman (1972) faund that the changes between 1 and 2 months were primarily due to hyperplasia and between 2 and 5 months the changes were due to a combination of hyperplasia and hypertrophy. Furthermore, the work of Hood (1972) and Anderson and Kauffman (1972) indicated that after 5 to 6 months of age the increase in the total adi- pose mass of pigs was primarily due to hypertrophy. Hood (1972) also found that during growth of the bovine animal, an increase in adipose mass was accompanied by cellular hypertrophy and hyperplasia. However, by 14 months of age hyperplasia was complete in all but the late developing intramuscular adipose tissue. While working with excessively obese human subjects, Hirsch and Knittle (1970) found that both adipose cell size and number was greater in obese subjects as compared to normal subjects and the degree of obesity was directly and highly correlated to the number of adipose cells. The results of DE Girolamo and Mendlinger (1971) and D1 Girolamo t l. (1971) indicate that definite species differences exist as evidenced by the finding that the guinea pig expands its epididymal adi- pose tissue by hyperplasia with little hypertrophy; whereas, in the rat, pig and hamster it occurs mainly by hypertrophy. Moreover, the results of Hirsch and Knittle (1970) indicate that childhood obesity is primarily associated with hyperplasia; whereas, the studies of both these authors and Salans _£__l. (1971) provide evidence that adult onset obesity in humans is mainly due to hypertrophy. The work of Bjorntorp and Sjostrom (1971) showEd that moderate enlargement of fat depots in humans is mostly due to cell hypertrophy, whereas, in severe obesity hyperplasia is the dominating factor. However, recent work by Hood (1972) and Lee (1972) indicate that excessive fat accumulation in pigs is caused primarily by cell enlargement and the percent carcass fat was found to be more highly correlated to adipose cell size than with cell number. In an experiment designed to study the effects of starvation on adipocyte size and number, Hirsch and Han (1969) reported that cell size decreased in the semi—starved rat adipose tissue and observed little change in cell numbers. Hood (1972) reported similar findings when studying the effects of starvation on pig adipOSe tissue. The results of a study conducted by Knittle and Hirsch (1968) indicated that nutri- tional restrictions during the suckling period of rats resulted in a permanent decrease in adipose cell number. In contrast, Lee (1972) re- cently showed that caloric restriction during the suckling period had no effect on adipoeyte numbers of adult pigs. Knittle and Hirsch (1968) demonstrated that in rats the lipogenic capacity of the adipocyte was not related to cell size. However, more recent work has shown that in human (Smith, 1971a) and porcine (Anderson _5 _l., 1972; Anderson and Kauffman, 1972) adipose tissues, adipocytes of different size, fat depot location and animal age all affect lipogenic rates or enzyme activities. Anderson _£_al. (1972)‘ also found that pig adipose tissue from areas where fat is readily deposited (perirenal) had higher enzyme activities, larger cells, less connective tissue, greater ether extractable lipid and a lower concentration of adipocytes per gram of tissue than samples from anatomical areas where fat is deposited more slowly (hind limb subcutaneous). Moreover, Hood (1972) reported that the activity of NADP-isocitrate dehydrogenase, NADP-malate dehydrogenase, glucose-6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase in bovine adipose tissue was directly related to adipocyte volume. Recent studies involving various adipose tissue depots reveal that intramuscular fat or marbling varies in many respects from the other fat depots. Studies by Moody and Cassens (1968) and Hood (1972) with bovine intramuscular fat and by Lee and Kauffman (1971) with porcine intramuscular fat show that adipocytes of this depot are smaller than corresponding subcutaneous adipocytes. Also, the findings of Hood (1972) suggest that the increase in intramuscular fat in bovine muscle is due primarily to hyperplasia and the presence of numerous small cells indicates that intra- muscular fat is later developing than other adipose tissue depots. Intra- muscular fat possesses the same basic lipogenic capabilities as subcu- taneous fat, which allows for lfl.§i£2 fatty acid synthesis in the intra- muscular depots. However, Lee and Kauffman (1971) and Chakrabarty and A ll Romans (1972) reported lower lipogenic enzyme activities in intramuscular fat than in subcutaneous fat. Moreover, Lee and Kauffman (1972) reported that the lipogenic enzymes of porcine intramuscular fat showed a different pattern of development than that of subcutaneous adipose tissue. In their study the lipogenic enzyme activities of intramuscular fat was higher at 5 months of age than at younger ages; whereas, the corresponding enzyme activities of subcutaneous fat had already reached its maximum before 5 months of age and was lower at this age than at younger ages. This observation supplies additional evidence to support the findings of Hood (1972) who concluded that intramuscular fat was later developing than other adipose tissue depots. In an experiment designed to study the lipogenic capabilities of the three layers of the subcutaneous fat depot of the pig, Anderson and Kauffman (1972) found that the magnitude of growth and total enzyme activity was greatest in the inner subcutaneous layer followed in de- creasing order by middle subcutaneous and the outer subcutaneous layers. The greatest proportion of the evidence presented indicates that adipose tissue depots of various anatomical locations possess basic differences in cellular and metabolic characteristics. These differences and their interactions have a profound effect on lipid metabolism and much of the recent research effort is being concentrated in this area in order to obtain an explanation for the metabolic interrelationships. r 12 Lipogenesis Most animals are intermittent eaters and as such their bodies are equipped for storage of much of the energy derived from the ingested food. Fat takes up less volume and weighs less per calorie of stored chemical energy than either carbohydrate or protein. Thus, in animals whose major food nutrient is carbohydrate, the lipogenic process is the primary system and maximum efficiency means by which dietary energy is stored. The major homeostatic function of lipogenesis is to store the energy ingested in excess of innmdiate energy requirements as fat. Adipose tissue has long been considered to be a mere fat storage depot in which excess calories are deposited but in the recent years it has captured the interests of many biologists. It is now recognized as an extremely metabolically active tissue with very adaptive and precisely regulated physiological and biochemical functions. According to Adler and Wertheimer (1967), the physiological, neural and hormonal mechanisms synchronize the balance between fat deposition and mobilization with the changing requirements of the animal. The homeostatic regulation of lipogenesis is determined to a large extent by the availability of carbo- hydrate or appropriate substrates to the tissues involved in lipogenesis and by the enzymatic capacity of these tissues to utilize these substrates. The amount of fat ingested and the state of energy balance also serve in a regulatory capacity (Masoro, 1961). Energy intake in excess of imme- diate energy requirements results in an increase in adipose tissue mass. This increased adiposity is the result of triglyceride deposition within the individual adipocytes of the various fat depots. The triglycerides 13 are composed of fatty acids derived from either dietary sources or endo- genous sources (de novo fatty acid synthesis). Substrate for Lipogenesis Non-ruminant The one important metabolic difference between ruminant and non- ruminant animals is the relative inability of ruminants to use glucose as a substrate for lipogenesis. This was demonstrated in liver slices and adipose tissue by Hanson and Ballard (1967) and in mammary tissue by Hardwick (1966) when they showed that the rates of lipogenic activity in rat tissues far exceeded the rates found in comparable ovine and bovine tissues when glucose was the in yiggg substrate source. Furthermore, O'Hea and leveille (1969b) reported that when gluCOSe-u-14C was used as the substrate in EE.ZEEE2 and in yiyg studies it accounted for 99 percent of the newly synthesized fatty acids in pig adipose tissue and Linzell _£‘_l. (1969) obtained similar results when studying porcine mammary tissue. Considerable interest has been directed toward the volatile fatty acids which are produced in the rumen, but little attention has been given to the VFA production in non-ruminants until Elsden _£._l. (1946) compared the VFA content of the alimentary tract of several species including the pig. The latter authors found that VFA were produced in the pig but they were appreciably less than in ruminants and these authors as well as Barcroft t al. (1944) reported that the chief sites of VFA production in the pig were the caecum and colon. More recent work by Friend _£._l- (1962, 1963a, b) provided additional evidence that considerable quantities of VFA are produced in the digestive tract of the monogastric animal. Barcroft _£‘_l. (1944) reported that VFA were absorbed from the caecum and colon and McClymont (1951) found the following percentages of VFA in pig blood: acetic acid 75.5%; propionic acid, 9.6%; butyric acid, 3.7%; and acids higher than butyric, 11.2%. Later work by Friend 95 _l. (1964) resulted in similar findings and they reported that the VFA were removed from the blood by the liver. In addition, O'Hea and Leveille (1969b) found when using acetate-l-IAC as substrate that the liver of the pig accounted for 25 to 30 percent of the newly synthesized fatty acids and Kinsella (1970) demonstrated that dispersed mammary cells from rats could utilize acetate and hydroxybutyrate to synthesize fat. Therefore, it is conceivable that VFA may serve as substrate for lipogenesis in non-ruminants to a limited extent. Research by Barrick _£ _1. (1953) and Garton and Duncan (1954) showed that the composition of the depot fat of non-ruminants, such as the pig, could be altered by variations in the source of dietary fat. Furthermore, Tove and Smith (1960) found that when elevated depot fat levels of lino- leate and oleate were produced by dietary means in mice, specific patterns of fatty acid replacement were observed. Mason and Sewell (1967) and Babatunde _£._l. (1968) reported that many of the body tissues exhibited a rather sensitive response to changes in the quantity and source of dietary lipids supplied to pigs. Thus, it is evident that the fatty acid composi- tion of dietary fat has a marked effect upon the fatty acid composition of the depot fat of monogastric animals. -. 15 Ruminant Fermentation in the rumen, which precedes gastric digestion in ruminants makes a large proportion of the structural components (cellulose and hemi- cellulose) of plantsayailable in forms which are directly usable by the tissues of the ruminant. Carbohydrates, proteins and all other fermentable substances are simultaneously converted by microbes to volatile fatty acids, methane, carbon dioxide, ammonia and incorporated into the microbial cells (Leng, 1970). Volatile fatty acids are known to be the major utilizable products derived from ruminant digestion of complex carbohydrates and estimates by Warner (1964) and Bergman _£ _1. (1965) indicate that they account for approximately 70 to 80 percent of the energy requirements. Bergman _£ a1. (1966) reported that over 95 percent of the VFA are accounted for by acetic, propionic and butyric acids. Earlier studies by Shaw and Ensor (1959) had shown that the molar proportions of the rumen VFA, especially acetic and propionic, can be controlled by diet to a remarkable degree. Furthermore, Satter _£__l. (1964) reported that dietary changes brOught about changes in ruminal microorganisms and that both affected the VFA formation from 1[lo-labeled polysaccharides. The latter authors demon- strated that the lower ruminal acetate to propionate ratios generally observed on high grain diets were due not only to the fermentation of more soluble carbohydrate, but also to an altered fermentation pattern of cellu- lose and hemicellulose. Thus, it is apparent that ruminal VFA production from any given feed constituent is altered by a number of environmental factors, including changes in the microflora. In addition, the work of 16 Shaw and Ensor (1959) revealed an altered pattern of VFA production in cows fed unsaturated fatty acids or oils and suggested the possibility of a direct substrate effect on VFA formation. Satter _£ _1. (1967) found that the addition of sodium lactate markedly increased the molar percentage of butyrate and butyrate-14C. The dramatic change in titratable VFA and VFA-14C production caused by the addition of sodium lactate was considered evidence for the direct influence that ration constituents or feed addi- tives exert on the rumen fermentation via altering the metabolic routes used by the existing microbial population. The changes in the molar proportions of VFA and acetate to propionate ratios are mainly due to changes in propionate production in the rumen (Davis, 1967). Church (1969) concluded that the ingestion of immature grass, increasing the amount of carbonaceous or protein supplements, increasing feed intake and pelleted roughages tend to result in increased propionate levels; whereas, increased levels of acetate are usually seen with silage and mature pastures. The influence of the acetate to propionate ratio on lipogenesis was first demonstrated by Armstrong _£__l, (1957, 1958) and Armstrong and Hexter(l957) in a series of experiments in which steam VFA were infused into the rumen of sheep fed maintenance rations. They found that when the mixture of steam VFA contained about 70 percent acetic acid on a molar basis, the efficiency with which the absorbed products of digestion were utilized for lipogenesis was 33 percent. Similarly, when the rumen liquor cantained about 45 percent acetic acid on a molar basis, the efficiency with which the absorbed products of digestion were used to synthesize body fat was 56 percent. Later work by Blaxter (1966) using roughage rations (high molar concentration of rumen acetic acid) demonstrated a drastic decrease in the efficiency of body fat synthesis. Moreover, grain rations (lower molar concentration of rumen acetic acid) resulted in a marked increase in the efficiency of body fat synthesis. When given as the sole source of energy, acetic acid is poorly utilized and acid accumu- lates in the systemic blood causing severe acidosis, decreased blood sugar and a marked increase in protein breakdown (Blaxter, 1962). In experiments with labelled compounds, Davis _£._l' (1960a, b) demonstrated that acetic acid provided a high efficiency energy source when propionic acid was also present and Blaxter (1962) concluded that acetic acid is not readily utilized unlesscarbohydrate or a carbohydrate precursor is given at the same time. Thus, it appears that as the proportion of energy derived from acetic acid is increased, the efficiency with which the metabolizable energy of the ration is used for lipogenesis declines. Substantial amounts of acetic, propionic and butyric acid with smaller amounts of long chain fatty acids, are formed in the rumen and absorbed through the rumen wall. Pennington and Sutherland (1956) reported that the epithelium of the rumen wall is capable of metabolizing butyrate, propionate and to some extent acetate. Cook and Miller (1965) showed that the mixture of fatty acids that is transported to the liver is de- pleted of its butyrate as compared to the mixture in the rumen fluid. Halter _£_§l. (1963) demonstrated that acetate was not metabolized in perfused goat liver when propionate and butyrate were present and Lang and Annison (1963) concluded that both propionate and butyrate in the portal blood are largely removed during the passage through the liver, A "r N.» s U 18 the former giving rise to lactate, glucose and carbondioxide and the latter producing chiefly B-hydroxybutyrate. Moreover, the work of Cook and Miller (1965) indicated that the propionate in the portal blood was almost quan- titatively removed during passage through the liver. They concluded that propionate is mainly metabolized by the liver while acetate metabolism occurs mainly in the extrahepatic tissues. These conclusions agreed with those of Reid (1950) who had earlier concluded that in general, acetate is the only short chain VFA present in significant concentrations in the peripheral blood. Recent results of Smith (1971b) clearly showed that of the VFA produced most abundantly in the rumen, sheep liver mitochondria most effectively metabolized a mixture of propionate and butyrate. Acetate was metabolized only in the absence of the other two, but the presence of acetate did not modify the metabolism of the other two. These results are consistent with the appearance of acetate being the only VFA present in significant quantities in peripherial blood. Thus, it is evident that acetate is essentially the only VFA available to the tissues and is the major substrate for the metabolic processes in extrahepatic tissues in ruminants. The amount of glucose transported from the ruminant digestive tract to the blood, at least under normal feeding conditions, is usually con- sidered negligible (Roe g£_al., 1966). One consequence of this differ- ence between monogastric animals and ruminants is the need to produce glucose in the latter group. Glucose synthesized in the liver from pro- PiOnate, amino acids, lactate and glycerol is used as an energy source in brain and some other tissues, but neither ruminant liver nor adipose [fig—'— 2‘ 19 tissues can convert significant amounts of glucose into triglycerides for storage (Ballard _t _l., 1972). Some enzymes which are necessary for lipogenesis to occur, from glucose as substrate, are absent in ruminants. These enzymes are glucokinase (Ballard and Oliver, 1964), ATP-citrate lyase and NADP-malate dehydrogenase in ruminant liver (Hanson and Ballard, 1967) and ATP-citrate lyase, NADP-malate dehydrogenase and pyruvate car- b0xylase in adipose tissue (Hanson and Ballard, 1967). Bergman _£ _1. (1970) concluded that it was unlikely that the low activities of hexo- kinase and glucokinase in sheep liver regulate fatty acid synthesis from glucose; however, the mere presence of these enzymes indicate a physiolo- gical role in sheep liver for producing glucose in both the fed and starved states. Baldwin and Ronning (1966), Baldwin _£ _1. (1966) and Young 25 31. (1969) measured the activities of NADP-malate dehydrogenase and ATP- citrate lyase in liver and adipose tissues of cattle fed high carbohydrate diets. These experiments showed little if any adaption to the diets in either liver or adipose tissue and the activities of both enzymes remained very low compared to the corresponding enzyme activities in rat tissues. Furthermore, Hardwick _£ _1. (1963) and Hardwick (1966) demonstrated that only acetate supplied carbon for milk fatty acid synthesis. The results of a similar study by Bauman _£ _1. (1970) indicated that the lack of glucose utilization for fatty acid synthesis in ruminant mammary gland was due to a nonfunctioning citrate cleavage pathway or more specifically 10w citrate cleavage enzyme and NADP-malate dehydrogenase. Hood (1972) also concluded that acetate was the only substrate incorporated into fatty acids in bovine adipose tissue, but glucose was quantitatively the more 20 important substrate for the synthesis of triglyceride glycerol in adipose tissue and intestinal mucosa. However, Ballard _£ _1. (1972) using radio- isotope incorporation studies with isolated adipose tissues showed an l8-fold increase in fatty acid synthesis from glucose in sheep with abomasal infusions and a 34-fold increase when glucose was injected intravenOusly. The glucose infusion caused a different effect in the liver from that observed in adipose tissue; the primary effect in the liver was to stimu- late acetate metabolism. Moreover, the enzymes of the citrate cleavage pathway showed much greater adaption in adipose tissues than in liver and they concluded that ATP-citrate lyase and NADP-malate dehydrogenase adapt to dietary stimuli, The low rates of fatty acid synthesis in hepatic or adipose tissues from glucose in these studies emphasize' the lesser im- portance of this lipogenic pathway in the ruminant compared to monogastrics. Nonetheless, glucose plays an important role in supplying reduced coenzyme ‘nicotinamide adenine dinucleotide phosphate via the hexose monophosphate shunt and a-glycerophosphate via the Emden Meyerhof pathway for lipid biosynthesis in the ruminant. Thus, gluconeogenesis is of obvious importance in ruminant metabolism. Propionate, a major VFA produced from dietary carbohydrate is a major substrate for g1uconeogenesis in ruminants (Armstrong, 1965; Ford, 1965). Cook and Miller (1965) concluded that propionate is absorbed from the rumen of sheep and goats largely as such and presented to the liver for metabolism. In contrast, Ieng _£__l. (1967), who infused radioactive PEOPiOnate in the rumen of sheep, estimated that up to 70% of the propionate “33 metabolized to lactate before it was converted to glucose and they 21 suggested the rumen epithelium was the site of this conversion of propionate to lactate. However, Weigand _£ _l. (1972) concluded from propionate in- fusion studies that the direct conversion of propionate into lactate by bovine rumen reticulum epithelium was small and probably amounm to less than 5 percent. Also, the majority of the propionate reached the liver where it was converted to glucose without being converted to lactate. However, in earlier studies, Bergman _£__l. (1966) found that propionate accounted for only about 40 percent of the glucose and they suggested that protein probably serves as a substrate for glucose formation in the adult ruminant. The findings of Ford and Reilly (1969) who compared the mean specific activities of plasma amino acids and glucose after the intravenous infusion of a mixture of 14C-amino acids into sheep, substan- tiated the earlier work of Bergman _£._l- (1966). They concluded that 11 to 17 percent of the glucose was derived from plasma amino acids. None- theless, considerable controversy still exiSts as to the specific sub- strates for g1uconeogenesis and their physiological importance to lipo- genesis in ruminants. Ruminant depot fats are relatively insensitive to changes in the fatty acid composition of the diet when the usual.fats rich in C18 fatty acids are fed (Tove, 1965). This unique feature of ruminant metabolism is the result of dietary lipid modification in the rumen before absorption. HYdrolysis effected in the rumen includes the release of fatty acid from ester combination and the release of galactose from galactoglycetide (Carton, 1965). The rumen lipase system responsible for this hydrolysis is dUe to lipolytic bacteria (Robson and Summers, 1966). Wood t al. (1963) A r--- 22 and Polan (1964) concluded that dietary fatty acids are not degraded to VFA in the rumen, while McCarthy (1962) reported that only small amounts of the long chain fatty acids are absorbed through the rumen wall. One of the most important fates of dietary long chain unsaturated fatty acids is their hydrogenation in the rumen by both bacteria and protazoa (Wright, 1960). The net effect of hydrolysis and hydrogenation results in long chain saturated free fatty acids which constitute the major lipid class present in digesta and they are passed from the rumen to the lower diges- tive tract. These studies also indicate that stearic acid is the major free fatty acid present (Keeny, 1970). Metabolism in the YOung Ruminant Many metabolic parameters in lambs and calves during the first few weeks of age, when the rumen is non-functional and they are dependent primarily on milk, resemble those of adult non-ruminants more than those of adult ruminants. Carbohydrate metabolism in young calves and lambs shows a pattern characteristic of monogastrics. Jarrett _E El' (1964) and House and Phillips (1968) observed a gradual decline in plasma glucose concentration in growing lambs and this decline was associated with a progressive decrease in the glucose entry rate into the vascular system. Ballard _£__l. (1969) suggested that this gradual decline may be partly due to the development of ruminal fermentation which diminishes the abs sorption of glucose from the gut. The data of Filsell t al. (1963); Goetsch (1966) and Bartley g£_§l. (1966) imply that the activities of the 23 gluconeogenic enzymes increase with age, while the activities of the gly- colytic enzymes decrease with age. More recent work by Leat (1970) indi- cates that the development of a functional rumen results in a reduced capacity for hepatic glucose oxidation via the pentose and glycolytic pathways and an increase in g1uconeogenesis in the liver. Thus, as the ruminant matures it becomes more dependent on g1uconeogenesis for its supply of glucose rather than dietary carbohydrates. The blood concentration of VFA, principally acetic acid, increases gradually in young ruminants, although it is variable and less striking than the change in the blood glucose level (Reid, 1953). It was suggested by Keech and Utter (1963) that the COA esters of these acids may stimulate g1uconeogenesis by activating pyruvate carboxylase and Ballard _£ _l. (1969) emphasized that this enzyme plays an important physiological role in g1uconeogenesis. Also, Randle g3 31. (1966) suggested that the increased acetyl COA to COA ratio inhibits some of the glycolytic enzymes. Ballard et al. (1969) reported substantial gluconeogenic activity in the liver of fetal calves and lambs and this was in contrast to low glu- COneogenic activity found in fetal rat livers. The capacity to synthesize BIUCose allows the fetal ruminant to be less dependent on maternal glucose than fetuses from non-ruminants. In all ruminant species studied by Ballard _£ _l. (1969) the overall activity of the gluconeogenic pathway increased rapidly at birth and was accompanied by increases in the acti- vities of the key gluconeogenic enzymes (phosphoenolpyruvate carb0xylase) Pyruvate carb0xylase, fructose 1,6-diphosphatase, glucose-b-phosphatase). Furthermore, when lambs were maintained on a diet containing milk, but no ,_v—.._ 4 24 solid food, the progressive increase in the conversion of propionate to glucose characteristic of normal maturing rumen function was delayed. This demonstrates that the biochemical development of a mature ruminant can be delayed experimentally and provides further evidence that many metabolic systems are adaptive to dietary changes. The enzymes involved in lipogenesis exhibit marked changes during the development of the young ruminant. The fetal calf liver has ATP-citrate lyase and NADP-malate dehydrogenase present as well as a functioning citrate cleavage pathway as measured by the substantial amount of incor- poration of the carbon 3 of aspartate and the carbon 5 of glutamate into fatty acids in this tissue (Ballard 2£.él-: 1969). However, citrate cleavage enzyme activity in the fetal ruminant diminishes during the first few weeks after birth and continues to decrease to the low level of activity observed in the adult ruminant liver (Hardwick, 1966; Hanson and Ballard, 1967). The fetal ruminant and to a certain degree the young ruminant possess the capacity to utilize glucose carbon for lipogenesis, but this ceases shortly after birth with the vauisition of the rumen micro- flora which results in the production of VFA to serve as the major source 0f energy. Lipogenic Precursors The $3 novo synthesis of fatty acids diagrammatically depicted in figure 1 involves the fatty acid synthetase complex which catalyzes the following overall reaction, in which one molecule of acetyl COA and seven molecules of malonyl COA are condensed to form one molecule of palmitic cwcosa l l GUJCOSE -6- PHOS PHATE NADP NAD PH 6 - PHOS Pl-KXELUCONATE _k RIBULOSE-S-PHOS PHA'I'E l r NADP NADPH 25 TR 10 LYCER IDE v \ FATTY AC ID 4 -GLYCEROPHOSPHATE NADP \ mops ACETATE ACETYL COA I 4 TR rose PHOSPHATE mun: -o—— OXALOACETATE e E1 mm mm mm mm: 1 mm NADP oxochmn-z % ISOC 1mm: f elmn: l crronasu I A‘X‘ \ K \\ /Y\ X\X\\\i\\\\\\\\X\\\‘\)\\\\“\X\\\‘XXXXSK sumac-moan /-o- - — — oxoc__1.!1_'r__mrs T—rsocimrs 2* CITRATE NADPH mp muvmr —__>—. cmum: ACETYL COA ACETATE Figure l. 23 novo fatty acid synthesis from glucose and acetate. (G-6PDH)-Glucose-6-phosphste dehydrogenase; (6-PGDH)~6-phosphogluconate dehydrogenase; (ME)-NADP-ma1ate dehydrogenase; (ICDH)-NADP-isocitrate dehydrogenase; (Syn)-acety1 COA synthetase; (CBX)-acetyl COA carb0xylase; (FAS)-fatty acid synthetase; (CCE)-NADP- citrate lyase. 26 acid. The coenzyme NADPH supplies the reducing power to drive the reaction to form palmitic acid and one molecule of acetyl COA serves as a primer. + Acetyl COA + 7 Malonyl COA + 14 NADPH + 14 H -——C> Palmitic acid + 7 C02 + 8 COA + 14 NADP + 6 H20 When discussing the research efforts on the precursor supply for lipogenesis in animal tissues it is convenient to divide these studies into four major areas: 1) those concerned with the pathway of carbon flow from glucose or acetate to acetyl COA; 2) studies involving the g2, 3219 synthesis of fatty acids from acetyl COA; 3) studies of reducing equivalent (NADPH) supply and 4) studies of the availability of a-glycero- phosphate for triglyceride synthesis. Sources of Acetyl COA All the carbon atoms for the gg_ngxg synthesis of fatty acids are supplied by acetyl COA molecules which are considered to be the building blocks. One important metabolic difference between ruminant and non- ruminant tissues is the source of carbon for lipogenesis (figure 2). In non-ruminant tissues the source of carbon for acetyl COA is glucose, whereas, in ruminant tissues the source of acetyl COA carbon is acetate. Therefore, the metabolic pathways for acetyl formation are somewhat differ- ent and will be discussed separately. 27 Non-Ruminant Ruminant I Glucose I IAcetatg I l | ems I | H.M.P.S. v I Oxaloacetate Mgctyl COA :I I 4 I l l '0' *? Pyruvate -(: Malate Citrate J Cytoplasm \\\\\$\\\\\x\\\\\3\\\\\\\\3\c\\\\‘ Mitochondria xaloacetat v I UByruvate Citrate Acet l COA Acetate Figurke 2. Intracellular location of the reaction sequences involved in the formation of Acetyl COA from glucose and acetate. 28 Non-ruminant The conversion of glucose to fatty acids involves a complex series of enzymatic reactions. In the first phase,glucose in metabolized to pyruvate in the cytosol via the Embden-Meyerhof pathway and the hexose monophosphate shunt pathway (figure 2). Pyruvate then enters the mito- chondria, where it is oxidatively decarboxylated to acetyl COA. Fatty acid synthesis from acetyl COA occurs within the cytoplasm of the cell, but uses acetyl COA formed in the mitochondria (Srere, 1965). However, Spencer and Lowenstein (1962) reported earlier that the rate of acetyl COA diffusion out of the mitochondria was too slow to meet the demands for fatty acid synthesis. Fatty acid biosynthesis thus requires the production of extramitochondrial acetyl COA from mitochondrial acetyl COA. The ob- servation of Srere (1959) that citrate cleavage enzyme was located in the cytosol of the cell, together with the demonstration that citrate was an effective substrate for fatty acid synthesis (Srere and Bhaduri, 1962; Spencer and Lowenstein, 1962; Formica, 1962) led to the conclusion that citrate played an important role in the generation of acetyl COA in the cytosol (Srere, 1965). The most tenable indirect pathway for the genera- tion of cytoplasmic acetyl COA in mammalian tissues involves the following sequence: formation of citrate within mitochondria from acetyl COA and oxaloacetic acid by citrate synthetase, transfer of citrate to the cytosol zuui the production of extramitochondrial acetyl COA and oxaloacetic acid fronn citrate by the action of the citrate cleavage enzyme (Hanson and Ballaard, 1968). The above sequence is generally referred to as the citrate 29 cleavage pathway. Recent work by Watson and Lowenstein (1970) and Martin and Denton (1970a, b) supplied substantial evidence that the citrate cleavage pathway is the major pathway whereby acetyl units for fatty acid synthesis are transferred to the cytoplasm in mammalian tissues. The citrate cleavage pathway and its relationship to lipogenesis from glucose and acetate is diagrammatically depicted in figure 3. Citrate cleavage enzyme has been implicated by numerous researchers as an important control point in the regulation of extramitochondrial acetyl COA for fatty acid synthesis in non-ruminants (Kornacker and Lowenstein, 1964, 1965; Kornacker and Ball, 1965; Ballard g£_al,, 1969). This implication was based on three observations. First, Srere (1959) noted that citrate was localized in the soluble fraction of the cell (cytosol). Secondly, Srere and Bhaduri (1962); Spencer and Lowenstein (1962) and Formica (1962) observed that citrate was an effective precursor for fatty acid synthesis. Thirdly, Brown and McLean (1965); Howanitz and Levy (1965) and Brown _£._l, (1966) found that under certain physiological conditions, citrate cleavage enzyme and fatty acid synthesis appeared to be altered in a parallel fashion. In contrast to these earlier conclusions, Srere and Foster (1967) provided some prelimin- ary evidence that no necessary relationship existed between citrate enzyme and fatty acid synthesis by showing that during a 24-hour fast the latter decreased to negligible levels without change in citrate cleavage enzyme activity. Moreover, Foster and Srere (1968) demonstrated that in recovery from fasting, fatty acid synthesis increased markedly without change in citrate cleavage enzyme activity and after alloxan administration, fatty acid synthesis decreased prior to any change in citrate cleavage enzyme. 3O CITRATE CLEAVAGE-PATHWAY F-—--—_- ---x E>TPYRUVATE \‘ PYRUVATE CARBOXY SE NAD- LATE DEHYDROCENASE I l I I I I a“: I OXALOACETATE g OXALOACETATE ACETYL COA | E I I z I I 2% I ATP-CITRATE LYASE E I I | P CITRA « SYNTHET a | I I E I | O | I I > I | I | l ICITRATE SYNTHETASE IT E ft CITRATE I I I(CYTOPLASMI I I I L~ TP CITRATE LYASE I I \x [I z/ \ I \ \ // \ |0A FATTY ACID 4.x? ACETYL COA I / A’CETYL ‘\_ _________ ______, SYNTIIETS ACETYL COA SYNTHETASE ACETATE ACETATE Figure 3. Citrate cleavage pathway and its relationship to lipogenesis from glucose and acetate. 31 These authors also found that substitution of a high carbohydrate, low fat diet resulted in large increases in citrate'cleavage enzyme activity without concomitant change in lipogenesis. Moreover, they showed that the addition of purified citrate cleavage enzyme from rat liver to the fatty acid synthesizing system did not restore fatty acid synthesis. Del Boca and Flatt (1969) found fatty acid synthesis to be higher in the presence of glucose plus acetate than with glucose alone, demonstrating that the acetyl COA transport mechanism is not rate-limiting in adipose tissue. Fatty acid synthesis via the citrate cleavage pathway requires the continual replenishment of oxaloacetic acid within the mitochondria. Figure 4 diagrammatically depicts the pathways most likely to be involved in the transfer of oxaloacetic acid between the adipocyte cytoplasm and the mitochondria. Regeneration of mitochondrialoxaloacetic acid from Oxaloacetic acid formed by citrate cleavage in the cytoplasm may occur in Part by‘the pathway suggested by Martin and Denton (19703) and Ballard g£ 31° (1972): 0AA cyto ---) malate cyto ---> malate mito ---> 0AA ndto However, Kornacker and Ball (1965) suggested that the most important path- way appeared to be: 0AA cyto ---> malate cyto ---> pyruvate cyto ---> pyrUVate mito ---> 0AA mito. This latter pathway involves the transfer of (3+) from cytoplasmic NADH to cytoplasmic NADPH which appears to be necessary as Martin and Denton (1970a) demonstrated that the rate of pro- duCtion of NADPH by the hexose monophosphate shunt pathway is not suffi- cient t0 account for high rates of lipogenesis. Additional evidence to Support the suggestion that the latter pathway is the major means for the 32 GLUCOSE I I I PHOSPHOENOL ,. L o PYRUVATE - OXALCACETATE ACETY C A GHWNMfli NADH OXOGLUTARATE ASPARIATE NAD CITRATE I I l PYRUVATE MALATE NADP I \ NAQP I CYTOPLASM “‘ L s s‘““““‘ “ MITOCHONDRIA f MALATE ASPARTATE OXOGLUTARATE GLUTAMATE v OXALOACETAT PYRUVATE CITRATE ACETYL COA Figure 4. Pathways most likely involved in the transfer of oxaloacetic acid between the fat cell cytoplasm and mitochondria. 33 replenishment of mitochondrial oxaloacetic acid was supplied by Ballard and Hanson (1967). They reported that pyruvate carb0xylase was present in adipose tissue and was active enough to supply the intramitochondrial oxaloacetate to support the citrate cleavage sequence. These authors pro- posed that pyruvate carboxylase should be considered to be a component of the citrate cleavage pathway in adipose tissue. The physiological importance of acetyl COA synthetase which converts acetate to acetyl COA has not been totally elucidated in non-ruminant tissues. However, Baldwin _£._l. (1966) reported the presence of acetyl COA synthetase in the liver of guinea piglets, rats and piglets and Barth __t._l. (1971) found considerable acetyl COA synthetase in rat liver, kidney, heart and adipose tissue. Although the acetyl COA synthetase mechanism for acetyl COA synthesis has been identified in non-ruminant tissues, it only indicates that the potential of a functional system exists but does not provide unequivocal evidence of its functionality. Ruminant In ruminant tissues where acetate is the single most important meta- bolite from a quantitative standpoint (Davis 3; gl., 1960a; Lee and Williams, 1962; Kronfeld, 1968) the conversion of acetate to acetyl COA for gg_ggxg fatty acid synthesis is of profound physiological importance. The initial enzymatic step in the conversion of acetate into fatty acids is the formation of acetyl COA via acetyl COA synthetase reaction (figure 2). Acetyl COA synthesis from acetate can be summarized by the equation: 34 l Acetate + COASH + ATP A§§t e 222 Acetyl COA +'AMP X PPi Acetyl COA synthetase allows the ruminant to form acetyl COA directly in the cytoplasm, thus, bypassing the need for citrate to move acetyl units across the mitochondrial membrane. The almost negligible activities of both ATP-citrate lyase and NADP-malate dehydrogenase in both bovine and sheep liver as well as low levels of key enzymes of the citrate cleavage pathway in ruminant adipose tissue reflects the almost total inactivity of this pathway in adult ruminants (Ballard g£_§l., 1969). The occurrence of acetyl COA synthetase activity was first demonstrated by Beinert _£‘_1. (1953) in pig heart and Hele (1954) in bovine heart mitochondria. Hanson and Ballard (1967) showed this enzyme to be much more active in both bovine and sheep adipose tissue than in rat adipose tissue. The latter authors and Ballard t l. (1969) found much higher acetyl COA synthetase activities in ruminant liver than in adipose tissue and Bauman _£H_1. (1972) reported that high acetyl COA synthetase activity in rump adipose tissue of wether lambs decreased 7 to 14 fold with fasting. Even though acetyl COA synthetase is present in several ruminant tissues, adipose tissue apparently possesses the most active acetyl COA synthetase system. There has been considerable discussion as to the intracellular loca- tion of acetyl COA synthetase. The work of Aas and Bremer (1968) and Aas (1971) indicated that acetyl COA synthetase was preferentially a mito- chondrial enzyme. In contrast, Kornacker and Lowenstein (1965) and Hanson and Ballard (1967) found acetyl COA synthetase activity not only in the udtochondria but also in the cytoplasmic fraction of animal cells. Martin and Denton (1970a) and Barth g£_al, (1971) used acetate as a substrate and 35 found 80 percent of the acetyl COA synthetase activity of liver and epidi- dymal fat tissues in the cytoplasmic fraction and only 20 percent in the mitochondria. They concluded that these findings demonstrated a prefer- ential cytoplasmic location in lipogenic tissues. The latter authors reasoned that this disparity could be attributed in part to the use of propionate as substrate which is also activated by butyrl COA synthetase. Malonyl COA Formation The malonyl COA required as the immediate precursor of fourteen of the sixteen carbon atoms of palmitic acid is formed from cytoplasmic acetyl COA and carbon dioxide by the action of acetyl COA carb0xylase, which catalyzes the reaction shown below: H Acetyl COA + C0 + ATP !2-> malonyl COA + ADP + Pi 2 Acetyl COA carb0xylase (figure 1), a biotin dependent enzyme, is the first enzyme involved in the sequence of the cytoplasmic pathway of fatty acid synthesis and it has been implicated by several researchers as the key enzyme in the regulation of fatty acid synthesis in animal tissues (Vagelos, 1964; Numa 25 $1., 1965; Lane and Moss, 1971; Moss and Lane, 1971). The results presented by numerous researchers show a definite relationship between the rate of fatty acid synthesis and acetyl COA carb0xylase activity. Korchak and Masoro (1962) studied the effect of fasting on fatty acid synthesis in rat livers and found that a 50 percent reduction in acetyl COA carb0xylase activity accompanied a 99 percent depression of fatty acid 36 synthesis. In a similar study with pigs, O'Hea and Leveille (l969d) reported that following a 4 day fast, the activity of the enzyme was decreased by almost 66 percent, whereas, lipogenesis fell by more than 99 percent. Bauman_gt.al. (1972) studied the lipogenic capacity of adipose tissue in wether lambs and reported that of all the enzymes measured the activity of acetyl COA carb0xylase was most closely related to l!.!i££2 lipogenic rate. Chang g£_al. (1967) and Chakrabarty and Leveille (1969) found approximately equal activity of fatty acid synthetase and acetyl COA car- b0xylase in rat liver and the latter authors reported the response of fatty acid synthetase to meal feeding was similar to that of acetyl COA carb0xy- lase in rat adipose tissue. These latter authors and Dakshinamurti and Desjardins (1969) demonstrated that changes in acetyl COA carb0xylase activity in response to dietary alterations were evident to a much greater extent in adipose tissues than in liver of rats. Chang g£_a1. (1967) also reported that fatty acid synthetase and acetyl COA carb0xylase activities were both elevated in obese mice compared to nonobese controls. In addi- tion, Chakrabarty and Leveille (1969) reported that acetyl COA carb0xylase activity in rat adipose tissues decreased with age and corresponded to the decrease in lipogenic capacity of rat adipose tissue with age as shown by Gellhorn and Benjamin (1965). The role of acetyl COA carb0xylase as a regulatory enzyme has been attributed to its complex regulatory prOperties. Tubbs and Garland (1963) and Bortz and Lynen (1963a, b) reported that the tissue concentration of long Chain fatty acyl COA esters increased with fasting and fat feeding and they suggested that these long chain fatty acyl COA derivatives may 37 reduce fatty acid synthesis through an inhibitory effect on acetyl COA carb0xylase. Cahill _£__1. (1960) found that the addition of free fatty acids to in_yi££g systems containing either liver homogenates or epididymal fat pads depressed the fatty acid synthetic activity and Korchak and Masoro (1964) demonstrated that free fatty acids primarily inhibited the acetyl COA carb0xylase reaction. Certain tricarboxylic acids, notably citrate and isocitrate stimulate the acetyl COA carb0xylase catalyzed reaction (Martin and Vagelos, 1962; Matsuhashi t al., 1962; Kallen and Lowenstein, 1962; Waite, 1962; Gregolin _£ _1., 1966). The activation of acetyl COA carb0xylase involves an equilibrium between an active polymeric form and an inactive polymeric form of the enzyme (Numa g; _l., 1966; Gregolin gg‘al., 1968a, b). Citrate and isocitrate are capable of shifting this equilibrium in favor of the catalytically active form (Lane_g£“§l,, 1971) and this has led to the suggestion that citrate activation of acetyl COA carb0xylase has a role in the physiological control of fatty acid synthesis. However, Lynen (1967) questioned the physiological significance of citrate activation of the enzyme on the grounds that the intracellular Citrate Concentration is be- low that required for the activation of the enzyme ig’yiggg. Moreover, Denton and Halperin (1968) have shown that citrate concentration in adi- Pose tissue is not related to fatty acid synthesis in the manner which is required if fatty acid synthesis were regulated by citrate activation of acetyl COA carb0xylase. The results of a study by Iliffe and Myrant (1970) on the sensitivity of rat liver acetyl COA to citrate stimulation indicated that the enzyme is relatively insensitive to citrate in the intact liver 38 cell. This raises the possibility that citrate may not participate in the physiological control of fatty acid synthesis via regulating the activity of acetyl COA carb0xylase despite the undoubted influence of Citrate on this enzyme in the purified state. Therefore, it becomes apparent that the cause and effect relationship between the activity of acetyl .COA carb0xylase and the rate of lipogenesis is currently one of the many sub- jects of controversy surrounding lipogenesis. Sources of Reducing Equivalents The synthesis of long chain fatty acids requires large amounts of NADPH (Langdon, 1957; Brady, 1958; Wakil g£_§1., 1957) and Tepperman and Tepperman (1958) suggested that the availability of this reducing equiva- lent may be a possible controlling factor in fatty acid synthesis. The fatty acid synthetase complex preferentially oxidizes fourteen moles of NADPH during the formation of one mole of palmitic acid from acetyl COA and malonyl COA. In non-ruminants, NADPH is generated from glucose via the hexose monophosphate shunt and by the conversion of malate to pyruvate via the Citrate cleavage pathway (Ballard and Hanson, 1967; Young gt_a1., 1964; Ball, 1966). The sources of NADPH are diagrammatically depicted in figure 5. A primary source of NADPH for lipogenesis has been considered to be the hexose monophosphate shunt with glucose-6-phosphate dehydrogenase and 6-phosphog1uconate dehydrogenase supplying the cytoplasmic NADPH. The concept that NADPH formed in the oxidative pathway was essentially 39 June—3.5a 30- »uuqu go: «Iv new scan-noevu Gran—<5 n25.— nuuuavuu .«o Snug—us; .m 0.3»: mag; If IIIIIII A. IIIIIIIIIII J. IIIIIII fiI IIIIIIIII mk<>=s . _ V P :MQEISHH: tfl// ///////////////—//////¢/////// //////// %¢/////////V/////// /////V/H £35883 I IE: I I E55503 j 3.2538 mn E %=mwh .5 SE85 « <00 Cram”: . ILl magmoEInémSB: 40 and the exclusive source of NADPH for lipogenesis was accepted for a long time (Tepperman and Tepperman, 1958; Wakil, 1961; Abraham SE al., 1961). This concept is substantiated by the fact that a parallelism does exist between the observed rates of fatty acid synthesis and NADPH production by the hexose monophosphate shunt pathway for a wide range of incubation conditiomifor normal tissues and also tissues taken from animals varying in dietary status (Pande _£__l., 1964; Katz _£_gl., 1966; Saggerson and Greenbaum, 1970). Baldwin and Milligan (1966) and later Gul and Dils (1969) showed that the activities of glucose-6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase in rat mammary tissue increased markedly with the onset of lactation. Furthermore, Young gt _1. (1964); Leveille and Hanson (1966) and Leveille (1970) reported that the NADPH generating enzyme involved in the pentOSe pathway adapted to conditions of decreased and increased lipogenesis in non-ruminant tissues. However, Tepperman and Tepperman (1963) demonstrated that increased pentose pathway dehydrogenase activity was secondary to increased fatty acid synthesis. Moreover, Flatt and Ball (1964), Wise and Ball (1964) and Landau and Katz (1965) showed that intact rat adipose tissue NADPH generated in the conversion of the hexose monophosphate to pentose phosphate supplies only 50 to 60 percent 0f the reducing equivalents used in fatty acid synthesis and Saggerson and Greenbaum (1970) also found that hexose monophosphate pathway NADPH Production does not always parallel fatty acid synthesis. Several authors have suggested that the rest of the required NADPH is obtained from cyto- Plasmic NADH through the malate transhydrogenation cycle in which the NAD and NADP-malate dehydrogenases are coupled with the net result that NADH 41 is used to generate NADPH (Pande t 1., 1964; Wise and Ball, 1964; Leveille and Hanson, 1966; Rognstad and Katz, 1966). In addition, Young t 1. (1964), Saggerson and Greenbaum (1970), Allee g£_§1, (1971) and Hood (1972) reported that alterations in the activity of NADP-malate dehydrogenase in response to different lipogenic levels were similar to those of the NADPII generating enzymes of the hexose monophosphate pathway. Lowenstein (1961) suggested that cytoplasmic NADP-isocitrate dehydro- genase could provide an alternative pathway for NADPH generation. In con- trast, Young t l. (1964) reported that the correlation between isocitrate dehydrogenase and lipogenesis was almost negligible and Pande- _£__1. (1964) and Wise and Ball (1964) found that isocitrate dehydrogenase did not show a close association with lipogenesis. The results of Bauman _£'_1, (1970) and Leveille (1970) indicate that in the non-ruminant, NADP-isocitrate dehydrogenase does not appear to be of major importance in the generation of reducing equivalents because the activity is low and not adaptable to dietary changes that affect lipogenesis. The source of reducing equivalents in ruminants is less clear. Under physiological conditions enough glucose may be metabolized via the hexose monophosphate pathway to supply adequate amounts of NADPH. This suggestion is supported by the presence of significant activities of g1ucose-6-phos- Phate dehydrogenase and 6-phosphogluconate dehydrogenase in bovine and ovine adipose tissue as reported tnrRaggi t 1. (1961); Filsell et 1. (1963); Hanson and Ballard (1967); and Opstvedt t al. (1967). Beitz (1972) reported significant levels of g1ucose-6-phosphate dehydrogenase aCtivity in inner and outer subcutaneous perirenal, omental and intermuscular 42 adipose tissue depots of Holstein steers. In a similar study, Hood (1972) concluded that hexose monophosphate dehydrogenases particularly glucose-6- phosphate dehydrogenase appear to be important regulators of NADPH pro- duction for fatty acid synthesis in bovine adipose tissues. In contrast, Bauman _£._1, (1970) found that glucose-6-phosphate dehydrogenase activity in bovine mammary gland was only 1/6 of that in rat mammary tissue and they suggested that the ruminant has yet another means of generating re- ducing equivalents. The almost complete absence of the citrate cleavage pathway in ruminant tissues rules out an active malate transhydrogenation cycle as a major NADPH generator. Bauman 25 a1, (1970) have suggested that an alternate source of NADPH generation in the ruminant is via NADP- isocitrate dehydrogenase. They reported that the ratio of this enzyme to glucose-6-phosphate dehydrogenase was 13 to l for the bovine animal and l to 9 in the rat. Hood (1972) indicated that NADP-isocitrate dehy- drogenase is capable of supplying NADPH for lipogenesis in bovine inter- muscular fat. Beitz (1972) found considerable NADP-isocitrate dehydro- genase activity in adipose tissue of fasted and normal steers. Beitz (1972) also found that glucose-6-phosphate dehydrogenase activity was reduced during fasting to 50 percent of normal, but was less than the reduction in lipogenesis; whereas, NADP-isocitrate dehydrogenase activity was not reduced to the same degree as g1ucose-6-phosphate dehydrogenase activity during fasting. Bauman _£._l. (1972) studied subcutaneous adipose tissue from wether lambs and reported similar results. They found that fasting reduced the activity of the two pentose pathway enzymes by 50 percent while isocitrate dehydrogenase activity remained unchanged. It 43 appears that isocitrate dehydrogenase becomes more significant as a source of NADPH and glucose less important as a source of carbon and NADPH for lipogenesis as the availability of acetate is increased. NADP-isocitrate dehydrogenase may be the enzyme system for the generation of reducing power from acetate for lipogenesis in ruminant tissues. Beitz (1972) suggested that the supply of reducing equivalents does not seem to be the rate limiting factor in fatty acid synthesis in rumin- ants, rather fatty acid synthesis tends to regulate the amount of NADPH that is generated from glucose oxidation. Nonetheless, there are numerous unanswered questions concerning the NADPH generating system in ruminant tissues. Transport of Lipids The transport of lipids by the plasma to the various tissues and the general metabolic routes of fatty acids and triglycerides are diagrammatically depicted in figure 6. Lipid transport in ruminants appears to occur similarly to lipid transport in the nonruminant. In ruminants, as in other mammals, the long chain fatty acids enter the circulatory system via the thoracic duct in the form of chylomicrons. These chylomicrons are composed primarily of triglycerides (Felinski g£_al,, 1964; Wadsworth, 1968). About one-third of the chylomicron triglyceride is absorbed by the liver (Di Luzio, 1960), one-third by adipose tissues (Felinski g£_al., 1964) and the remaining one-third by other tissues including mammary tissue (Robinson, 1963). The liver hydrolyzes triglycerides taken up as chylomicron triglyceride and these fatty acids along with free fatty acids THORAC IC DUCT 1 J 1 CHYLOMICRONS _l LIVER MAMMARY 1 ILFA‘ PLASMA ADIPOSE MUSCLE i * ‘1 1 V.F.A. J Figure 6. l RUMEN Diagram of the transport and uptake of triglycerides and fatty acids by ruminant plasma and their metabolism in tissues. The following abbreviations are used: 2E5, volatile fatty acids; EA, fatty acids; A-FA, free fatty acids in albumin complex; LDLP, low-density lip0proteins; ELK, glycerol; 9;, a-glycero- phosphate; QLQ, glucose. 45 mobilized from adipose tissues are re-esterified to glycerol to form new triglycerides (Olivecrons, 1962). The majority of the triglycerides syn- thesized by the liver are combined with protein to form lipoproteins, and they re-enter the plasma and are transported as low-density lipoproteins (Robinson, 1963). Triglycerides of chylomicrons and low density lipopro- teins are hydrolyzed for removal from the circulating blood lipids by the action of the enzyme lipoprotein lipase (LPL). Fatty Acid Uptake and Mobilization In adipose tissue the processes of circulating triglyceride uptake and of intracellular triglyceride release both require hydrolysis of the triglyceride molecule and these hydrolytic reactions are mediated by two distinct lipase systems. Lipoprotein lipase or clearing factor lipase first described by Korn (19553, b) plays an important role in lipoprotein metabolism as it is the enzyme responsible for the hydrolysis of the triglyceride moiety of chy- lomicrons and low density proteins. It is present in various tissues that utilize plasma triglycerides and several investigators have presented evidence to show that plasma triglycerides are hydrolyzed to free fatty acids before their removal from the blood (Robinson, 1960, 1963; Bezman e a1 , 1962a). The enzyme has been identified in heart and lung (Anfinsen ‘ C e a]_ , 1952), adipose tissue (Hollenberg, 1959; Rodbell, 1964; Pokrajac ‘ O ‘EE£31., 1967), muscle (Hollenberg, 1960) and in lactating mammary tissue (M93171de and Kern, 1963), but not in liver (Olson and Alaupovic, 1966). 46 Mayes and Felts (1968) reported that LPL was present in liver, but in an inactive state. The activity of lipoprotein lipase in adipose tissue is proportional to the rate of triglyceride uptake, therefore the enzyme may play a significant role in controlling lipid deposition in this tissue (Garfinkel st 21,, 1967; Bezman t 1., 1962b). There is considerable _-— evidence (Pav and Wenkeova, 1960; Salaman and Robinson, 1966; Wing _g‘gl., 1966; Reichl, 1970, 1972; Scow E£.il°: 1972; Schotz and Garfinkel, 1972) that LPL activity in adipose tissue is decreased by fasting and diabetes and increased by refeeding. Reichl (1972) found a progressive increase in LPL activity in adipose tissues that coincided with an increase in the plasma insulin concentration during the first 6 hours after feeding. This finding was consistent with the view of Robinson and Wing (1970) who earlier proposed that the increase in LPL activity that occurs during the transi- tion from the starved to the fed state is, at least in part, due to an increased secretion of insulin. Several investigators (Robinson, 1963; Patton and Hollenberg, 1969) have demonstrated that LPL is activated or released by heparin when in both in xitgg and in 3123 studies. Scow _£__l, (1972) observed that LPL is rapidly released into the blood stream when heparin is injected intravascularly and suggested that the enzyme is pre- sent in or near the vascular wall. This is in agreement with the earlier suggestions that the action of LPL takes place at the luminal surface of the capillary endothelium. These suggestions were based on the findings that the enzyme is released into the circulation of organs involved in triglyceride uptake, such as the perfused rabbit hindlimb (Robinson and French, 1960) and the lactating goat mammary gland (Barry t al., 1963). 47 Additional evidence to support the suggestions has been revealed by several researchers using the electron microscope. Electron microscopy of adipose tissues by Moskowitz and Moskowitz (1965), mammary gland by Schoefl and French (1968) and of adipose tissue and heart by Blanchette-Markie and Scow (1971) from normal animals have shown that chylomicrons and other lipid particles are attached to the luminal surface of the capillary endo- thelium, but none was seen inside the cells or in extracellular space. Furthermore, West _£ 31. (1972) concluded that hydrolysis occurs at the luminal surface of the capillary endothelial cells and the liberated gly- cerol and free fatty acids are then taken up by the tissue. It appears that intact chylomicrons and other lipid particles do not cross the cap- illary endothelium as intact particles in most tissues. The mobilization of adipose depot triglycerides as glycerol and free fatty acids requires the action of a lipase that is activated by the cate- cholamines and a variety of other hormones. The enzyme responsible for initiating the hydrolysis of the triglycerides stored in adipose tissues has been referred to as hormone sensitive lipase (Rizack, 1961). In an experiment designed to determine the intracellular location of hormone sensitive lipase, Khoo _£_al, (1972) found that a large fraction of this enzyme could be recovered in a large, phospholipid-rich particle present in rat adipose tissue. They suggested that this indicates that the enzyme in the intact cell might be associated with a lipid-rich matrix and their results strongly suggest that hormone sensitive lipase is predominantly a cytoplasmic enzyme. The regulation of this lipase is attributed to intra- cellular levels of cyclic AMP (CAMP) (Butcher t 1., 1965; Butcher and Sutherland, 1967; Meng and Ho, 1967; Davies, 1968; Patton, 1970) and in 48 this mechanism, cyclic AMP has been likened to a second messenger which carries the message from the hormone to its cellular action site (Robinson .E£.21“3 1967; 1968). Butcher (1969) indicated that hormone sensitive lipase regulation is mediated by a cyclic AMP dependent phosphorylation and dephosphorylation mechanism and that cyclic AMP effects the conversion of the enzyme from an inactive to an active form. Recent evidence accumu- lated by Huttunen t 1. (1970a, b), HutUmnen and Steinberg (1971), Mayer _£.§l, (1972) and Sidhu _5 a1. (1972) provide confirmation that hormonal activation of hormone sensitive lipase is mediated by cyclic AMP through a protein kinase and involves a phosphorylation activation of the enzyme. Thus, any condition which increases intracellular cyclic AMP would increase the activation of this enzyme. Bauman t al. (1972), Patton (1970) and Thorton gt _1. (1972) reported that lipolysis due to hormone sensitive lipase was elevated in both fasted lambs, and steers and decreased with refeeding. Beitz (1972) found that as steers matured and fat accumulated the maximwm rates of lipolysis generally declined; whereas, Sidhu _£‘_l. (1972) concluded that lipolysis seemed to increase with age and fatness in lambs. 0n the basis of the limited information to date it is difficult to draw conclusions as to the physiological interrelationship of hormone sensitive lipase, lipoprotein lipase and lipid metabolism generally. Hormonal Influences of Lipid Metabolism The major portion of the research efforts concerning the mechanisms of adipose tissue regulation has been conducted with non-ruminants, es- PECially the rat. In the ruminant the regulatory mechanisms and metabolic .49 pathways are somewhat different, although these differences may be quanti- tative rather than qualitative. Most of the hormones produced in an animal can affect some aspect of fat metabolism either by stimulation and/or inhibition of lipogenesis or lipolysis. The major lipogenic hormone in the non-ruminant is insulin which is ‘essentially the anabolic hormone of adipose tissue (Renold and Cahill, 1965). Insulin has two major effects on fat cells: 1) it stimulates the entry of glucose into the cell, and this is correlated with an accelerated metabolism of glucose and the synthesis and esterification of fatty acids (Crofford and Renold, 1965a, b; Turner and Bagnara, 1971); and 2) it exerts a potent antilipolytic effect, opposing the lipolytic actions of certain pituitary hormones and the catecholamines (Fain e£_§1., 1966; Rudman and Shank, 1966). Salans _£__l. (1968) and Nestel _£._l, (1969) reported that large adipocytes were less sensitive to the stimulatory effect of insulin on the oxidation of glucose and on the uptake of labeled triglyceride fatty acids than small cells. The former authors also found that biopsy samples of adipose tissues with different mean cell size incorporate glucose at similar rates when calculated on a per cell basis. However, Bjorntorp (1966) reported that on a per cell basis biopsy specimens from obese patients, which had enlarged adipocytes had a greater rate of lipid syn- thesis than specimens from nonobese patients. Additionally, Zinder _£ 21. (1967) and Smith (1971a) demonstrated that incorporation of glucose into lipids in human adipose tissue is dependent on cell size since the larger cells of a given fat depot had a greater rate of glucose incorporation 50 than smaller cells of the same fat specimen. Furthermore, the latter investigators observed that the larger adipocytes were less sensitive than smaller adipocytes to the stimulatory effect of insulin on glucose incorporation into lipids. The manner in which insulin is involved in carbohydrate and fat meta- bolism is readily apparent, since lipogenesis in adipose tissue is a function of the concentration of circulating insulin, and the circulating insulin concentration depends on blood glucose level (Adler and Wertheimer, 1968). Thus, as long as adipose tissue is responsive to insulin, the plasma level of insulin can regulate the lipogenic rate in adipose tissues. The major quantitative role of insulin in non-ruminants is the conversion of glucose to triglycerides in adipose tissue; whereas, this does not appear to be the case in ruminants because of the relatively low glucose levels and the presence of other substrates (VFA) for lipogenesis. The early work of Reid (1951) indicated that neonatal lambs and calves like non-ruminants are insulin sensitive, however, this insulin sensitivity decreases with increasing age. In more recent investigations, Manns t l. (1967); Horino _£._l° (1968) and Trenkle (1970) showed an increase in circulating insulin following the administration of butyric or propionic acid and they concluded that these two acids stimulated insulin secretion in ruminants. The actual physiological role of insulin in ruminant lipo- genesis has not been thoroughly elucidated. According to Ball (1970) the vast amount of research work in recent years on insulin has revealed a multiplicity of insulin action on adipose tissues. Perry and Bowen (1962) and Jungas and Ball (1962) reported that 51 insulin inhibited the customary production of free fatty acids and glycerol from rat adipose tissue following the administration of epinephrine, corticotropin, growth hormone, g1ucagon or thyroid stimulating hormone. Since the production of glycerol and free fatty acids from triglyceride is inhibited, this action has been termed the antilipolytic action of insulin. Butcher _£ _1. (1965) first showed that the induction of lipolysis in adipose tissue by epinephrine was accompanied by an increase in tissue cyclic AMP. Butcher _£__l, (1966, 1968) also indicated that the increase in cyclic AMP induced by epinephrine was lowered by the addition of insulin and Jungas (1966) demonstrated that adenyl cyclase activity of adipose tissue was lowered by pretreatment with insulin. Thus, Ball (1970) pro- posed that the effects of insulin on lipolysis are due to its lowering cyclic AMP levels, which in turn results in diminished activities of lipolytic enzymes. The cellular effects of insulin on lipid metabolism have been studied more intensively than any of the other hormones; none- theless, the mechanisms of insulin action on adipose tissues is still not completely resolved. In recent years it has become increasingly apparent that the lipolytic process in adipose tissues plays a key role in the energy metabolism of the animal. By regulating the extent of lipolysis in adipose tissues a variety of hormones exert important influences on the supply of free fatty acids made available as fuel for muscle and other tissues of the body. The catecholamines, epinephrine and norepinephrine, secreted by the adrenal medulla and the endings of the sympathetic nerve fibers, vigorously 52 promote hydrolysis of triglycerides to free fatty acids and glycerol in adipose tissues (Rudman, 1965; Steinberg, 1966; Fain, 1968; Corbin and Park, 1969; Exton g£_al., 1972). The catecholamines apparently do so by acti- vating the rate limiting enzymatic step in adipose tissue lipolysis, i.e., 1., 1972; Schwartz and Jungas, 1971; hormone sensitive lipase (Khoo SE Corbin gt al., 1970). They activate this enzyme by increasing 3',5'-cyclic AMP in adipocytes, which then activates the hormone sensitive lipase (Fain, 1968; Corbin £3 31,, 1970; Exton _£__l., 1972). Therriault _£‘_l. (1969) found that the lipolytic response of isolated adipocytes to norepinephrine was related to cell size since the larger adipocytes were less responsive to norepinephrine than the smaller cellS. In an experiment designed to study the influence of cell size on the release of free fatty acids from isolated adipocytes incubated with epinephrine and adrenocorticotropin (ACTH) Zinder and Shapiro (1971) concluded that free fatty acid release stimulated by either epinephrine or ACTH was a function of cell surface area as well as a function.of cell number. Hartman _£H_l. (1971) studied the lipolytic response of rat adipocytes to norepinephrine as related to cell size and reported that when the rate of lipolysis was based either on the amount of triglyceride in the incubation medium or on the cell surface area, lipolysis was inversely related to cell size. The pituitary peptide hormones, adrenocorticotrophin (ACTH), thyroid stimulating hormone (TSH), melanophore stimulating hormone (MSH) and vasopressin, are capable of promoting fat mobilization in a manner identical of the catecholamines. However, the lipolytic activity of each of these peptide hormones is mammalian species specific (Rudman, 1965). 53 Growth hormone and the adrenal glucocorticoids promote fat mobiliza- tion, but their stimulation of lipolysis in isolated adipocytes occurs by a mechanism quite different from that of the other lipolytic hormones (Fain 1., 1965). The lipolytic effect of either growth hormone or the adrenal _e__t_ glucocorticoids, unlike other lipolytic agents, requires a lag period of at least one hour (Fain, 1968; Moskowitz and Fain, 1970). The results of Fain and Saperstein (1970) indicate that the lipolytic action of growth hormone and the glucocorticoids is due to a direct activation of white adipocyte lipolysis. The mechanism by which these hormones stimulate lipo- lysis is different from the rapid activation of adenyl cyclase seen with other peptide hormones and catecholamines. The lipolytic effect of growth hormone and glucocorticoids is blocked by protein inhibitors and this suggests that the action of these lipolytic agents is to increase the synthesis of a protein or proteins which appears to increase the amount of cyclic AMP that accumulates in white adipose tissue cells (Fain, 1967; Moskowitz and Fain, 1970). The actions of glucagon in most instances, are antagonistic to those of insulin (Foa, 1964). Glucagon elevates cyclic AMP in liver and adipose tissue and activates hormone sensitive lipase, thereby stimulating lipo- lysis (Fain, 1968; Manganiello and Vaughan, 1972; Exton 35 al., 1972). However, the actual physiological role of glucagon in the regulation of fat mobilization has not been fully established. It is evident that many hormones influence fat mobilization, but the difficult question to answer in regard to these agents is whether their efforts are merely pharmacological or whether these agents function in Physiological control mechanisms. EXPERIMENTAL PROCEDURE An experiment involving 49 lambs was conducted to study the influence of breed type, sex and age on enzymatic activity and lipid biosynthesis. A commercial flock of 39 Western crossbred ewes was divided into two groups based upon their body length (anterior edge of scapula to tuber ischii) and body weight. The small ewes averaged 76.2 cm in length and weighed 70.9 kg while the larger ewes averaged 79.2 cm in length and 76.6 kg in weight. The smaller bodied ewes were mated to a Southdown ram and the larger bodied ewes were mated to a Suffolk ram in order to obtain two strains of lambs differing in propensity to fattening. Because of the differences in body type of the ewes and ram breed, the term breed type will be used to denote the matings employed in this study. Reference to the lambs resulting from these matings will be identified hereafter by their sire breed. Table I shows the design of the experiment and the number of lambs in each breed type, sex and age group. The ewes were maintained on pasture or hay until approximately one month before the first ewe lambed. The ewes were then moved to dry-lot where they were maintained on hay supplemented with a grain ration. The grain rations for the bred and lactating ewes are presented in tables A and B. in Appendix I. The lambs of the 8, 16 and 32 week age groups were exposed to gd’ libitum creep feed after 2 weeks of age and green, leafy alfalfa hay was fed twice daily. The composition of the creep feed and grain rations of the subsequent feeding periods are shown in table 2. The lambs were weaned at 2 months of age and remained in dry-lot on full feed until 54 55 Table 1. EXPERIMENTAL DESIGN Sex and breed type Ram lambs Ewe lambs Wether lambsa South- South- South- Age group Age downb Suffolkb downb Suffolkb downb Suffolkb total BirthC l 1 l 1 4 2a hourd 2 2 2 3 9 8 weeks 2 2 2 2 2 2 12 16 weeks 2 2 2 2 2 2 12 32 weeks 2 2 2 2 2 2 12 Sub-total 9 9 9 10 6 6 Total 18 19 12 49 aThe lambs of this sex group were castrated at 4 weeks of age. bRefers to the breed of the sire. cThe lambs of this age group were slaughtered immediately after birth and had not nursed. dThe lambs of this age group nursed and were slaughtered at 24 hours of age. as Table 2. COMPOSITION OF CONCICN'I‘RATI'I MIXTURES Age of lambs Ingredients 2 weeks 6 weeks 12 weeks 16 weeks onwards °/. °/. °/. °/. Crimped oats 24.92 14.95 ---------- Whole oats ----- 19.94 46.85 56.89 Crimped corn 51.84 23.93 ---------- Shelled corn ----- 19.94 34.90 22.95 Wheat bran 7.98 6.98 5.98 9.98 Soybean oil meal (497.) 9.97 9.97 9.97 9.97 Wet molasses 2.99 1.99 ---- ---_ Dicalcium phosphate 1.00 1.00 1.00 ---- Trace mineral salta 1.00 1.00 1.00 --.... Aurofacb-lo 0.30 0.30 0.30 0.20 Total 100.00 100.00 100.00 100.00 aGuaranteed analysis, 7.: Co, 0.012; Mn, 0.228; Cu, 0.040; Fe, 0.160; I, 0.007; Zn, 0.011. Product obtained from Michigan Salt Co., St. Louis, Mich. bChlortetracycline, 22 g per kilogram. Product obtained from American Cyanamid Co., Princeton, New Jersey. 57 slaughtered. Full feed of grain was defined as the amount of grain con- sumed in 20 min twice daily. Slaughter Procedure Lambs were slaughtered at the designated ages by bleeding without prior stunning. Blood was collected at the time of slaughter in a centri- fuge tube coated with heparin to prevent clotting and then centrifuged at 2000 x g for 10 min to obtain the plasma. The plasma was removed via pipette, frozen and stored at -30°C. Immediately following bleeding, samples of liver, longissimus muscle (LM), perirenal (PRF), and sub- cutaneous fat (SCF) were removed from the carcass. The longissimus muscle was excised from the 12th to 13th thoracic vertebrae region, except in the at birth and 24 hour age groups in which both intact longissimus muscles were excised in order to obtain sufficient sample. Subcutaneous fat samples were removed from the dorsal thoracic and lumbar regions of the carcass except in the at birth and 24 hour age groups where the absence of external fat made it impossible to obtain subcutaneous fat samples. The following enzymes were assayed on all four tissues: glucose-6- phosphate dehydrogenase (G-6PDH)‘, 6-phosphogluconate dehydrogenase (6- PGDH), NADP-malate dehydrogenase (ME), NADP-isocitrate dehydrogenase (ICDH), acetyl COA synthetase (Syn), acetyl COA carb0xylase (CBX) and ATP-citrate lyase (CCE). The adipose tissue and longissimus samples were assayed for 11p0protein lipase (LPL) and free fatty acids (FFA) were determined on blood plasma. o. be ' u '58 Extractions of acetyl COA synthetase and acetyl COA carb0xylase were made on all tissues immediately after sampling and the cytosol and mitochondrial fractions were obtained by centrifugation. These fractions were frozen and stored at -30°C. The remaining portions of the fresh tissue samples were frozen immediately in liquid nitrogen and stored at -30°C until powdered. The frozen tissue samples were powdered in a Waring blendor and the connective tissue particles were removed by sifting. All powdering procedures were carried out in a ~30°C freezer and following powdering the samples were stored at -30°C until assayed. Enzyme Assays Acetyl COA Synthetase Acetyl COA synthetase catalyzes the reaction of acetate with COASH to form acetyl COA. The assay measured the disappearance of the (-SH) group of COASH which was accomplished by breaking the 8-H bond shown in the following reaction. acetate + COA - SH + ATP Egg: acetyl COA + H20 + AMP + PPi The procedure followed was that described by Grunert and Phillips (1951). Enzyme extraction. Approximately 5 g of fresh tissue were homogenized 0° in 20 ml of 0.13M KCl at 2°C with a Virtis homogenizer for 30 seconds. The homogenate was centrifuged in a Sorvall RC2-B refrigerated centrifuge (0°C) for 10 min at 1000 X gravity.' The pellet was discarded and the 59 supernate was centrifuged at 20,000x g for 20 minutes. The cytosol frac- tion was decanted into a storage vial and the mitochondrial pellet was resuspended in 10 m1 of 0.13M KCl and both fractions were frozen and stored at -30°C until assayed. All preparatory work was carried out in a 2°C coldroom. Assay procedure. The assay reagents and the assay design are presented in table 3. All reagents, except the enzyme extract, were added to 15 ml conical centrifuge tubes. The reaction was started by the addition of 0.05 ml of enzyme extract diluted with buffer to a protein concentration of l to 2 mg protein/milliliter. The tubes were incubated in a gently shaking waterbath at 37°C for 10 minutes. The reaction was stopped by the addition of 2.8 ml of the color reagent (footnote e table 3) and the optical density was read exactly 30 see after the addition of the color reagent at 520 nm on a Bausch and Lomb Spectronic 20 Colorimeter. The calorimeter was set to zero with the blank. Calculations. O.D. (standard) - O.D. (assay) = A O.D. (A) A x 3.243 (conversion factor for 10 min assay) = umoles M COASH oxidized [tube (B) B x extract dilution factor X extract aliquot factor = mg protein 1 ml enzyme extract enzyme units/mg protein = Specific Activity enzyme unit - 1.0 ”moles M COASH reacting in 60 minutes A O.D. of 0.185 = 0.10 ”moles COASH oxidized Table 3. REAGENTS AND ASSAY DESIGN FOR ACETYL COA SYNTHETASE 60 Tubes Blank Standardb AssayC Assay Reagentsa vol (ml) vol (ml) vol (ml) conc (uM) 0.25M Mg CL2 0.01 0.01 0.01 2.5 0.08M ATP (dipotassium salt)d 0.01 0.01 0.01 0.8 0.5M acetate (K) 0.01 -- 0.01 5.0 0.05M tris pH 8.6 0.07 0.07 0.06 (35.0)(30.0) 0.017M COASH (tri-lithium salt) -- 0.01 0.01 0.17 Enzyme extract 0.05 0.05 0.05 -- Total 0.15 0.15 0.15 -- e after incubation Color reagent 2.8 2.8 2.8 -- 8Prepared in 0.05M tris pH 8.6. bDuplicated. CDuplicated. dPrepared in 0.05M tris pH 6.8 and diluted 1:3 with 0.05M tris pH 8.6 for each assay. eMixed fresh for each assay in the following proportions; 1 (2.0M Na2C03 in 0.09M NaCN) : S (6.8M.NaCl) stored in an amber bottle. : l (0.09M Nag [Fe (N0)(CN)5] "H20) and 61 Acetyl COA Carb0xylase Acetyl COA carb0xylase catalyzes the reaction of acetyl COA with 14C- bicarbonate to form radioactive malonyl COA as shown in the following reaction. acetyl COA + H14CO3 + ATP —E§%§f-> 14C-malonyl COA + ADP + Pi The 14c-malonyl COA was not utilized for fatty acid synthesis as NADPH was not present in the assay medium. The reaction was stopped by acidifi- cation and the radioactivity of the 14c-malonyl COA was measured by liquid scintillation counting. The procedure followed was a modified method of Dakshinanurti and Desjardins (1969). Enzyme extraction. Approximately 5 g of fresh tissue were homogenized in 25 ml of 0.4M glycylglycine in 0.25M sucrose pH 7.0 with a Virtis homo- genizer for 30 seconds at 2° centigrade. The homogenate was filtered through a thin layer of pyrex glass wool and centrifuged at lOOOXig for 10 minutes. The pellet was discarded and the supernate centrifuged at 20,000X.g for 20 minutes. The cytosol was decanted into a storage vial and the mitochondral pellet was discarded. All work was carried out in a 2°C cold room and the cytosol fraction was frozen and stored at -30°C until assayed. Assay procedure. The assay reagents and the assay design are pre- sented in table 4. All the reagents except acetyl COA (AcCOA) were added to an 12 mm by 75 mm disposable culture tube and incubated at 37°C for 30 min in a gently Table 4. REAGENTS AND ASSAY DESIGN FOR ACETYL COA CARBOXYLASE 62 - Tube Blank“, Assayc Assay cone Reagentsa vol m1) vol (m1) an 0.25M tris pH 7.6 0.32 0.25 (76.8)(60.0) 0.32M MgCl2 0.02 0.02 6.4 0.80M citrate (K+) 0.02 0.02 16.0 0.12M Glutathione (GSH) 0.02 0.02 2.4 60 mg/ml BSA (fraction V) 0.01 0.01 0.6 mg 0.01M EDTA 0.01 0.01 0.1 H20 0.20 0.20 ~- 0.08M ATPd -- 0.05 4.0 0.10M NaH 14(:03 (902, 910 cm/ 0.10 0.10 10.0 umole) 0.01M ACCOA -- 0.02 0.2 6N HCl 0.20 0.20 -- Enzyme extract .QLLQ Q;lQ _;;__ Total 1.0 1.0 -- After incubation Scintillation solutione 10.0 10.0 -- aPrepared in 0.05 tris pH 7.6. bDuplicated. CTriplicated. dPrepared in 0.05M tris pH 6.8 and frozen for stability. 8Composed of naphthalene (60 g): PPO (4g), POPOP (0.2 g), absolute methanal (100 m1), ethylene glycol (20 ml) and p-dioxane up to 1 liter. 63 shaking waterbath. The preincubation of the enzyme extract with citrate was necessary to attain maximum CBX activation. The reaction was started by the addition of ACCOA and incubated at 37°C for 2 minutes. After the 2 min incubation, 6N HCl was added to stop the reaction and the protein precipitate was sedimented by centrifugation at 1000 rpm for 3 minutes. A 0.2 m1 aliquot was removed via repipet and delivered into a scintilla- tion vial. The vials were placed in a 55°C sandbath to remove excess water and 14C02. After drying, 10 ml of scintillation solution were added and the radioactivity measured with a Chicago-Nuclear scintillation spectrophotometer. The micromoles of 14C02 incorporated were determined from the radioactivity of the malonyl COA and the specific activity of NaH 14CO3. Calculations. cpm (test) - cpm (blank) = Acpm (A) (A) x efficiency coefficient ‘ umoles 14C0 CC 2 incorporated/tube (B) 14 where CC = count coefficient CHM/umoles C02 = 902, 910 CPM/umole . l B x assay aliquot factor x extract aliquot factor = umoles 4C0 . . 2 2 minute assay x mg protein/ml enzyme extract incorporated/min/mg protein ATP - Citrate lyase Citrate cleavage enzyme catalyzes the cleavage of citrate to yield ACCOA and oxaloacetate (0AA). The reaction was measured by coupling it with the NAB-malate dehydrogenase (MDH) reaction. Malate dehydrogenase 64 catalyzes the conversion of 0AA to malate using NADH as the reducing agent. The oxidation of NADH as shown in the reaction below was followed spectro- photometrically at 340 nm. citrate + ATP + COA 99§—> Ac COA + ADP + Pi + 0AA . MD" *~ malate Mg++ NADH NAD The enzyme was assayed using the procedure described by Kornacker and Ball (1965). Enzyme extraction. Approximately 2 g of powdered tissue were homo- genized in 20 m1 of 0.15M KCl in 0.05M tris (CL-) pH 7.4 with a Virtis homogenizer at 2° centigrade. The homogenate was filtered through a thin layer of pyrex glass wool and centrifuged at 1000 x g for 10 minutes. The pellet was discarded and the supernate was centrifuged at 20,000 x g for 20 minutes. The cytosol fraction was decanted over glass wool to remove the fat particles and then frozen and stored at -30°C until assayed. Assay procedure. The assay reagents and design are presented in table 5. All reagents except ATP were added to an incubation tube and incubated at 25°C for 5 minutes. The activity of the blank was measured at 340 nm on a Beckman/DU spectrophotometer. The reaction was started by the addition of ATP and the change in optical density was recorded at one minute. Calculations. O.D. (blank) - O.D. (test) = A O.D. 65 Table 5. REAGENTS AND ASSAY DESIGN FOR CITRATE CLEAVAGE ENZYME. Blank AssayPI Assay conc Reagentsa vol (m1) vol (ml) QM 0.05M tris (CL‘) pH 7.3 1.40 1.30 (70.0)(60.0) 1.0M MgCl2 0.02 0.02 20.0 0.1M Dithiothreitol (DTT) 0.04 0.04 4.0 0.1M.KCN 0.02 0.02 2.0 0.4M citrate (K+) 0.10 0.10 40.0 2.5 mg/ml MDH 0.02 0.02 0.05 mg 1.67 mg/ml NADH 0.10 0.10 0.167 mg 0.008M COA 0.10 0.10 0.8 0.1M ATP - 0.10 10.0 Enzyme extract IQ;2Q Q;2Q -- Total 2.0 2.0 aPrepared in 0.05M tris (Cl‘) pH 7.3. bDuplicated. 66 A O.D. . . . "'—'- :. 13‘ l . E.C. (standard curve) umo £7 NADH OX1dlzed/tch/mln where E.C. = 0“D' conc ggmoles NADH oxidized/tubelmin X aliquot factor mg protein/ml enzyme extract = UmOICS NADH OXidiZEd/min/ mg protein Glucose-6-phosphate dehydrogenase and 6-Phosphogluconate dehydrogenase Glucose-6-phosphate dehydrogenase and 6-ph05phogluconate dehydrogenase catalyze the following reactions and their activities were determined collectively by measuring the generation of NADPH at 340 nm upon the addi- tion of both substrates (G-6-P and 6-PG). 6-Phosphogluconate dehydrogenase G-6PDH 6 PGDH - - ..EZQ... ._ - - _ G 6 P ++ > 6 PG ++ > Rlbulose 5 P NADP NADPH NADP NADPH was determined by the addition of only 6-PG and G-6-PDH was determined by difference. The enzyme was assayed using the procedure described by Lohr and Waller (1963). Enzyme.gxtraction. Approximately 2 g of powdered tissue were homo- genized in 25 ml of 0.25M sucrose with a Virtis homogenizer at 2° centi- grade. The homogenate was filtered through a thin layer of pyrex glass wool and centrifuged at 1000 x g for 10 minutes. The pellet was discarded and the supernate was centrifuged at 20,000 x g for 20 minutes. The cytosol fraction was decanted over glass wool to remove fat particles and frozen 67 and stored at -30°C until assayed. The mitochondrial pellet was resuspended in 5 ml of 0.25M sucrose and frozen and stored at -30°C until assayed for isocitrate dehydrogenase activity. Assay procedure. The assay reagents and design are presented in table 6. All reagents, except the substrate(s), were added to an incuba- tion tube and preincubated at 25°C for 5 minutes. The reaction was started by the addition of the substrate(s) and the change in O.D. at 340 nm after 1 min was recorded. Table 6. REAGENTS AND ASSAY DESIGN FOR GLUCOSE-6-PHOSPHATE DEHYDROGENASE AND 6-PHOSPHOGLUCONATE DEHYDROGENASE. Tubes G-6-Db Blank 6-PG 6-PGC Assay conc Reagentsa vol (m1) vol (ml) vol (m1) 7gp 0.05M glycylglycine pH 7.6 1.6 1.4 1.4 (80.0)(70.0) 0.40 Mg Cl2 0.1 0.1 0.1 40.0 0.0015M NADP - 0.2 0.2 0.3 0.01M G-6-P 0.01M 6-PG 0.2 0.2 - 2.0 0.01M 6-PG - - 0.2 2.0 Enzyme extract 0.1 0.1 0.1 - Total 2.0 2.0 2 0 8Prepared in 0.05M tris (Cl') pH 7.6. bDuplicated. cDuplicated. 68 Calculgtions. O.D. (both) - O.D. (blank) = total A O.D. (G-6PDH + 6-PGDH) O.D. (6 - PG) - O.D. (blank) = A O.D. (6-PGDH) Total A O.D. - A O.D. (6-PGDH) = A O.D. (G-6PDH) .Q_QLQ; = umoles NADPH generated/min/tube E.C. (standard curve O.D. conc where E.C. nmoles NADPH generatedjmin/tube X extract gliquot factor = mg protein/ml enzyme extract “moles NADPH generated/min/mg protein NADP - Malate dehydrogenase NADP-malate dehydrogenase (ME) catalyzes the decarboxylation of L- malate to pyruvate and the activity was determined by measuring the NADPH generated by the reaction shown below. ME L-malate Mn ++ > pyruvate + C02 NADP NADPH The enzyme was assayed using the procedure described by Ochoa (1955) and the generation of NADPH was followed spectrophotometrically at 340 nm. The enzyme extraction procedure was identical to that previously described for the extraction of G-6PDH and 6-PGDH. Assay procedure. The reagents and assay design are presented in table 7. All reagents except L-malate were added to an incubation tube and preincubated at 25°C for 5 minutes. The blank contained all reagents except NADP. The reaction was started by the addition of L-malate and the change in optical density at 340 nm was recorded at 1 minute. 69. Table 7. REAGENTS AND ASSAY DESIGN FOR MALIC ENZYME. Tubes Blank Assay“ Assay conc Reagentsc vol (.nl) vol @L pm 0.05M tris (Cl') pH 7.4 1.6 1.4 (80.0)(70.0) 0.02M Mn Cl2 0.1 0.1 2.0 0.0015M NADP - 0.2 0.3 0.015M L-malate 0.2 0.2 3.0 Enzyme extract 0.1 0.1 - Total 2.0 2.0 aPrepared in 0.05 tris (Cl') pH 7.4. bDuplicates. Calculations. O.D. (test) - O.D. (blank) = A O-D. A—QLQL = umoles NADPH generated/min/tube E.C. (standard curve) where: E.C. = O.D. conc Emoles NADPH generated/minjtube X extract aliquot factor 3 mg protein/ml enzyme extract pmoles NADPH gerierated/min/mg protein NADP - Isocitrate Dehydrogenase NADP-isocitrate dehydrOgenase (ICDH) catalyzes the decarboxylation of isocitrate to a-ketoglutarate both intra- and extra-mitochondrially. The activity of this enzyme was determined by following the generation of NADPH (as shown in the reaction below) spectrophotometrically at 340 nm. 70 . ICDH - Isoc1trate Mn ++ > a ketoglutarate NADP NADPH The enzyme was assayed according to the method of Plaut (1962) for both the cytosol and mitochondrial fractions. The enzyme extraction procedure was identical to that used for the extraction of G-flflfll: and 6-PGDH as previously described. Assay procedure. The reagents and assay design are presented in table 8. In the assay of the cytosol fraction all the reagents except isocitrate were added to an incubation tube and preincubated at 30°C for 5 minutes. The reaction was initiated by the addition of isocitrate and the change in optical density at 340 nm at 1 min was recorded. In the assay of the mitochondrial fraction all the reagents except NADP were added to an incubation tube and preincubated for 5 min at 30° centigrade. The reaction was started by the addition of NADP and the change in optical density at 340 nm was recorded at 1 minute.. The blanks in both cases con- tained all of the reagents except NADP. The calculations for ICDH were identical to those for the calculation of malate enzyme already described. 71 Table 8. REAGENTS AND ASSAY DESIGN FOR ISOCITRATE DEHYDROGENASE. Tubes Blank Assay Assay conc Reagentsa vol (ml), vol (ml)g HM 0.05M tris (CL') pH 7.4 1.2 1.0 (60 0)(50.0) 0.0034M EDTA 0.2 0.2 0.68 0.03M MnCl2 0.1 0.1 3.0 0.0015M NADP - 0.2 0.30 0.3% gelatin 0.2 0.2 - 0.015M dl-isocitrate 0.2 0.2 3.0 Enzyme extract 0.1 0.1 Total 2.0 2.0 aPrepared in 0.05M tris (Cl') pH 7.4. bDuplicated. Lipoprotein Lipase Lipoprotein lipase is a specific lipase which preferentially hydro- lyzes triglycerides when they are components of lipoproteins. The en- zyme was assayed by a modification of the method of McBride and Korn (1963). Enzyme extraction. Approximately 3 g of powdered tissue was homogen- ized in 12 ml of 0.15M KCl pH 8.5 with a Virtis homogenizer for 30 seconds. The homogenate was centrifuged at 2750 rpm for 10 min and the supernate decanted over a thin layer of pyrex glass wool. Assay procedure. The reagents and assay design are presented in table 9. The substrate employed in this assay was a mixture of (1:1) fresh 72 Table 9. REAGENTS AND ASSAY DESIGN FOR LIPOPROTEIN LIPASE. Tubesa Boiled extractb blank Assay Reagents vol (m1) vol (ml)g 107. BSA (fraction v>° 1.0 1.0 0.15M KCl pH 8.5 1.0 1.0 1:1 serumzediol 0.5 0.5 Enzyme extract 0.5 0.5 After 45 minute incubation d LPL extraction fluid 5.0 5.0 Hexane 3.0 3.0 H20 2.0 2.0 aDuplicated. bThe extract was placed in boiling waterbath for 3 min to inactivate the enzyme. cPrepared in 0.15M KCl pH 8.5. dComposed of isopropanol:heptane:lN H 2804 (40:10:l). ovine serum and ediol, a coconut oil emulsion (Calbiochem) mixed 1:6 with distilled water. The appropriate amount of serum and ediol were combined prior to assay and incubated for 30 min at 30°C in a gently shaking water- bath. The KC], BSA and preincubated substrate (serumzediol) were added to a 25 ml screw cap vial. The specified amounts of the enzyme extracts were added to start the reaction and the reaction mixtures were incubated at 37°C for 45 minutes in a gently shaking water bath. After the 45 min incubation period 5 m1 of the LPL extraction solution was added to each vial to stop the reaction and the vials were allowed to stand at room 73 temperature for 5 minutes. Following the 5 min equilibrium period, 3 ml of heptane and 2 ml of H 0 were added and the mixture was allowed to stand 2 until the layers separated. Two m1 of the top layer were transferred to another test tube via repipet and dried in a 55°C sandbath for the subse- quent colorimetric determination. After drying 4 ml of toluene were added to the tubes and the tubes vortexed at low speed. Then 3 ml of a 1:1 mixture of Rhodamine B (1 mg/ ml H20) and uranyl acetate (1% aqueous solution) were added and the tubes vortexed three times at 10 min intervals. Each time the mixture turns from purple to pink and becomes turbid. The tubes were centrifuged at 2000 rpm for 5 min and the top layer transferred via Pasteur disposable capillary pipette into a spectrophotometer cuvette. The optical density at 545 nm was recorded and compared to that of a palmitic acid standard curve. Calculations. O.D. (assay) - O.D. (blank) = A O.D. A O.D. compared to the palmitic standard curve to obtain the umoles FFA released/m1 toluene umoles FFA/m1 toluene X dilution factor X aliquot factor X extrgct_§liquot mg protein/ml enzyme extract factor X time factor = umoles FFA released/hr/mg protein Free Fatty Acids Assay procedure. A colorimetric method described by Mackenzie g; l. (1967) was used to determine the free fatty acid (FFA) concentration in 741 blood plasma. The reagents and assay design are presented in‘table 10. One gm of zeolite was added to a 25 ml screw cap test tube followed by 1 ml of the plasma sample and 2 drops of 85% phosphoric acid. With constant mixing 3 m1 of methylal:methanol (4:1) were added followed by 4 ml methanol, 1 ml H20 and 3 ml petroleum ether. The test tubes were capped and vortexed twice at 5 min intervals for 30 sec at low speed. The reaction mixture was centrifuged at 2000 rpm for 5 min and a 1 ml aliquot from the top layer was transferred via capillary pipette to another test tube. The tubes were placed in a 55°C sandbath to evaporate the petroleum ether. After drying the same colorimetric method described for the lipoprotein lipase assay was employed to determine the FFA. The blank was used to set the spectrophotometer to zero. Table 10. REAGENTS AND ASSAY DESIGN FOR FREE FATTY ACIDS. Tubes Blank Assaya ‘_ Reagents vol (ml) vol (ml) Zeoliteb 1 g 1 8 Plasma - 1.0 85% phosphoric acid 2 drops 2 drops Methylal:Methanol (4:1) 3.0 3.0 Methanol (reagent grade) 4.0 4.0 Distilled H20 2.0 1.0 Petroleum ether (redistilled) 3.0 3.0 aDuplicated. Ground to 100 mesh and dried at 100°C. 75 Calculations. The test optical density was compared to a palmitic standard curve to obtain the “M FFA/m1 toluene. _gmoles FFA/ml toluene X dilution factor X aliquot factor = umoles FFA/mg mg protein/ml plasma protein Protein Determination Protein was determined by the method of Lowry £3 11. (1951) and ex- pressed on a mg/ml of tissue extract basis. Statistical Analysis Data were anlyzed on the C.D.C. 3600 computer at Michigan State University Computer Laboratory. A least squares analysis was used to determine differences between treatment means. The Duncan's Multiple Range Test was employed when significant differences were observed (Snede- cor and Cochran, 1969). Simple correlation coefficients were also determined (Snedecor and Cochran, 1969). RESULTS AND DISCUSSION The results and discussion of the enzyme analyses will be presented in groups according to the phase of lipogenesis in which they are involved. The first group includes acetyl COA synthetase and ATP-citrate lyase which are involved in the formation of the cytoplasmic acetyl COA necessary for fatty acid synthesis. Acetyl COA carb0xylase which is considered to be a key enzyme in d2 ngyg_fatty acid synthesis from acetyl COA will also be included with the first group. The second group includes glucose-6- phosphate dehydrogenase, 6-phosphog1uconate dehydrogenase, NADP-isocitrate dehydrogenase and NADP-malate dehydrogenase which have been regarded as being functional for the generation of the reducing equivalents (NADPH) necessary for lipogenesis. Thcfinal. group will include the enzyme lipo- protein lipase which plays an important role in lipoprotein metabolism since it is responsible for hydrolysis and uptake of the triglyceride moiety of circulating chylomicrons and low density proteins from plasma. In the discussion of the results several terms are used which require definition or clarification. In the discussion of the level of lipogenic activity, its use in this dissertation refers to the rate of fatty acid synthesis and it will be used synonymously with the rate of fat deposi- tion. A decrease in lipogenic activity or rate of fat deposition indi- cates a decreased rate of fatty acid synthesis, but it does not imply that fatty acids are not being synthesized or that fat is not being de- posited. An increase in lipogenic activity refers to an increase in the rate of fatty acid synthesis and/or rate of fat deposition. Under normal nutritional regimes, fatty acids are being synthesized and fat is deposited 76 77 during the entire growth and development of the lamb and continues, al- though to a more limited extent, even after maturity has been reached. However, the rate of fatty acid synthesis and fat deposition varies throughout the life of the sheep. Reference to adipose tissue depots in this dissertation will include the subcutaneous (SCF) and perirenal (PRF) fat. While intramuscular fat (marbling) is generally considered as a fat depot, because of the low quantity of fat observed in the longissimus muscle (LM) of the lambs included in this study, the LM will not be included in the discussion of the adipose tissue depots (SCF AND PRF),but will be discussed separately. Acetyl COA Generation Acetyl COA Synthetase The enzyme acetyl COA synthetase (SYN) allows the ruminant to form acetyl COA directly in the cytoplasm; thus, the conversion of acetate to acetyl COA for gg_flgyg fatty acid synthesis is of considerable physiolo- gical importance in the ruminant. The means and standard error of the means of acetyl COA synthetase activity are presented in figures 7, 8 and 9. Cytoplasmic SYN (figure 7) was significantly (P <.05) higher in perirenal fat (PRF) than in subcutaneous fat (SCF) with both PRF and SCF possessing greater enzyme activities than longissimus muscle (HM) and liver. Mitochondrial SYN activity (figure 8) followed a similar pattern with the adipose tissue depots showing significantly (P <.05) higher activity than muscle and liver. A pattern similar to that of both the \1 co enzyme units A I.4o- § ° \ A \ N \ (a \ a \ \ N \ . M S 0.20? a \ As a s Liver L M .0 :U 11 (0 (5 1n Figure 7. Means and standard error of the means of cytoplasmic acetyl COA synthetase activity in var ous tissues. a’b":Means with different superscripts differ significantly (P <.05). p. 79 enzyme units mg protein A s s A \ W \ \ a \ _ \ 0... § § 0.40 ~ § § a as 0.20 - g \ x . , \ A Figure 8. Means and standard error of the means of mitochondrial acetyl COA synthetase activity in various tissues. 8’ Means with different superscripts differ significantly (P <.05). 8 O 4.00- §% 3.00- k \ A A 2...- §§ ”s AA A a A A ..o_\ A \ -A A A 0.20-§ § § 0 S a §t§ Liver L.M PRF. 3.0.5 Figure 9. Means and standard error of the means of total acetyl COA synthetase activity in various tissues. a, eans with different superscripts differ significantly (P <.05). 8| cytoplasmic and mitochondrial SYN activity was observed for total SYN activity in the four tissues. These results are in contrast to the find- ings of Hanson and Ballard (1967) and Ballard _£__l, (1969) who reported much higher acetyl COA synthetase activity in liver of adult ruminants than in adipose tissue. However, these results are in agreement with the more recent observations of Cramer (1972) who found very low SYN activity in the liver of steers and an especially higher level of enzyme activity in the adipose tissue depots. The data indicate that SYN is located both in the cytoplasm and mitochondria and agree with the earlier findings of Kornacker and Lowenstein (1965) and Hanson and Ballard (1967). However, these results are in contrast to the findings of Martin and Denton (197Qfl and Barth _£._l. (1971) who concluded that this enzyme was preferentially located in the mitochondria. The results showed that 35 to 40 percent of the SYN activity in ovine adipose tissue was of cytoplasmic origin and 60 to 65 percent of mitochondrial origin. The means and standard error of the means of cytoplasmic SYN in var- ious tissues are presented by breed, sex and age in table 11. All of the tissues of the Southdown lambs had greater SYN activities than those of Suffolks, but only in PRF were the differences significant (P = .01). These data indicate that Southdown lambs which have a greater propensity to fatten, possess a higher potential to utilize acetate for acetyl COA synthesis and possibly for lipogenesis. The influence of sex on cytoplasmic SYN activities in the various tissues was not significant and there was no apparent or consistent pattern of enzyme activity among the various sexes. SYN activity increased with age in the liver. The livers at 32 82 .Amo. V mv maucmeamacmam nemwwv menace was manmwum> oweuws munfiuomumaom uoouewwwv saws mamQZm.e.p .mmmesuaoumm :H asoem momma“ was manmwum> some now oocsoawacwwm mo Ho>eAu .mcowum>ummno mo noneszn .cHa co aw wawuomou mmua>auum uwwwommm mm vowmouaxe mesam>m amen. A Nmm.o Amsv uses. a mNH.H Away Nmo. a mma.o Amav ammo. a oH¢.o a-v ems: «m mass. A cam.a Away uses. a was.o a~av Nmo. a Hoo.o Amav mumNO. A ~mm.o Aafiv sums on «was. a omm.a AHHV uses. A sma.a Amos «no. a maa.o Awav «ammo. A mm~.o a~av sums w macs. a maH.~ Ase one. A mnH.o Ame memmo. a mo~.o Ame use: an ean. a mm~.m Ase «no. A emo.o Aev some. a naa.o Ase sense Aao.umv Aao.umv Ame.umv Aao.mmv awe was. a owo.a AHHV Has. a Hms.a a~av Nmo. a “no.0 a~Hv mmo. n mm~.o Amav amass: mom. a nmm.a Amav NEH. A aoe.~ Aaav mac. A «NH.o Amos mmo. a omm.o Aauv was man. a Ham.a Amos was. a HHN.H Awav owe. A oma.o Ammo mac. a mmm.o Amav sum am A: 99“.: sent sense .3 mmu. A ooe.a Ammo woes. A Nas.a flame Nuo. a saa.o Aemv omo. A mim.o Asmv asoeeusom mma. + HHN.H Asav wwso. A Nos.a Amwv NNO. A ecu.o AmNV owe. a mm~.o Ammo xuommam Amm.umv afio.umv Aas.tmv uAso.umv swans umm G umw fi gnome: G H0>w4 fl man—mwumxr msoesmuaonsm Hmaeuwumm maawmmawaoq a Nmfimwz map No Houum vhmvcmum Bum mfimflz .me< az< xmm .ommem em mmsmmHs mson<> zH ssH>Heo< mmoa .cHE oo cw wowuemou mmwuom uwmaumam mm commepmxo monHm> .wo>ummno ouo3 mcoauemueuca unwowmwcwam aes3 mace meuoemmua mum momma one mo muouue vumecmum was masons Aom.umv Amm.umv Ams.umv use moomcmuaonom ueeN. a mmo.u oesw. a NHN.H sums um seem. a Hmm.o ueem. a mms.o ems: SH seem. a swa.a oesm. a Hoo.a sum; m seew. a wa¢.~ euaw. n mam.H “no; em s.moem. a moa.s x.eoem. A Noe.~ euuam caoseasom xaommsm Asm.umv Amo.umv AoH.umv use HNGGHHHmm Aam.uav Asm.umv Aoo.umv waomaa msawmmwwaoq Aso.umv Amm.umv naeo.uav um>aq mw< x xom ew< x mooum xmm x peeum oommwa mqmmbmmHH mDOHm<> zH WHH>HHU< mm Casuas muawuemummom uceuewmwv Lugs mameZe.e .memmnucmuma CH macaw esmmwu was oHnmwum> nose now eucmowwwcwwm mo He>eqo .mcowum>uemno mo usefide mmfluem oamwoomm mm pemmeumxe wosHm>m «mm. A sqm.m Amav soc. a mes.m Amos «so. A Noe.o a~HV use. a ssm.o Awav ems: mm mmm. A oma.u Ammo ems. a oo~.~ AHHV smo. a so~.o Awav use. A o-.o Amos saws so Nmm. A ao~.~ Amie soc. A ems.a Away see. a oma.o a~av use. a mma.o Amfiv ewes m use. a emH.~ ass was. A osm.o Ame smo. a mso.o Ame use; «N sso.H a om~.s Ase Nos. a mom.o Ase «me. A mmH.o Ase cream Ame.uav Ama.uav Aom.umv Aao.umv mw< «Nam. a Ham.m Amos ems. A moo.~ Anne emo. a mmm.o a~uv see. a moH.o Ammo assume eNmm. a sam.s Awav owe. a awn.N Amie sue. a ma~.o Ammo wmo. a mm~.o Amav mam smmm. a Nos.a Amav mas. A ao~.m Amie one. a ma~.o Amav mmo. a sm~.o Away awe Ame." V Amsnumv Awm.umv Aem.umv xmm ass. a amm.a Ammo one. a mNm.~ Ammo see. A osm.o Aemv mmo. a eo~.o Acme czoseusom ass. A aom.~ Amos was. a oms.~ Ammo «so. a oom.o Ammo Nmo. A NwH.o Ammo xsomunm STA: Gene SEA: segue 385 own a umm c mHomse : uo>wq c mflnmwum> msomcmu=on=m Hmceuwuom mafiwmmawcoq n mmcmmz onuuo uouum pumvcmum mam mammz .mu< aza xmm .mmmem we mesmmus mson<> zH ssH>Hse< mmm «ou ussmo< aaHeozomoost .Na mumqu .awa 00 cu wawuommu =m¢oo 81 o.H u awn: mahuam mums: ucflmuoua wE\muHa= mamucm u zuw>wuom owmwomam m vmmmmuaxm mosam> .vm>ummno wuw3 macfiuvmumucw ucmofimwcwfim coca haco pwucowwua mum momma «no mo nouns cumvcmuw cam mcmoZm wona.fi A qmm.¢ omH.H A nmo.m Hmaum3 oomH.H A n¢N.o onH.H A osw.m mam momma.a A «mo.a omfi.H A moH.N sum :3ovsusom xaowmsm Ammoumv Ammonmv Amo.umv amt msomcmununsm mmmm. H n¢H.¢ x.o¢mm. A moo.H umsumz now. A NHH.N vumoo. A «No.~ mam moo. A o~m.~ sumo. A mom.m 5mm czocnusom xHowmsm Asm.umv Amm.nmv Aqo.nmv “mm Hmcmuwumm Amw.nmv Aqw.umv Aoo.umv maumsE mafiwmmwwcoq Aoo.umv Amm.umv nAmm.umV um>AA mm< x xmm mw< x vmmum xmm x vmmum commas MJmMDmmHH mDon<> zH wHH>HHO< mm cwnuwz muaguumumadm ucmummwwc sums mamm2m.o.w .mommcucmumd CA :3ocm mammfiu cam manmwum> comm pow cosmoawwcmwm mo Hm>oqu .mSOAum>ummno mo uwnadzn .cwa 00 Ca wcfluumwu mmwuom owwfiumam mm vmmmmuaxm mosam>m max. A aaa.m Ammv ccam. A ana.¢ Amcc mac. A mam.c Amcv mmmc. A mma.c Ammc moms mm max. A aa¢.m Amsc ccam. A Hma.~ AHAV mac. A mcm.c Amac mammc. A Nam.c ammv xmas mm «mm. A caH.c AAAV macm. A man." AmAc mac. A mAm.c AmAV acmmc. A ass.c AmAc mama m coca. A c~m.s Aac NAA. A Amm.c Aac cmcc. A mmm.c Aac Ago; cm mcHA.H A mmm.a Aac mmc. A Hms.c Asv acmac. A Nam.c Asc cuAAm aqm. cc asc.umc amm.umc Aac.ucc mmm mcam. A cam.m AHAV cam. A acm.q Amsc mac. A aAc.c Amfic mac. A amm.c Amfic Ameumz cmaa. A acc.m Amsc ccm. A Amm.s AaAc mac. A ass.c aaAv mac. A «mm.c aaAc mam cmaa. A mma.~ ANAV «mm. A cma.¢ Amsc cmc. A mAc.c amcv sac. A mcm.c Amac 5mm 39".: ASN.5: aaa.umv GAE: xmm mac. A cam.a Amsc acq. A asm.q Ammc mmc. A mas.c Asmv mac. A mam.c Asmc csoccucom cam. A amm.q Aamc «as. A cac.c Ammc mmc. A mcc.c Ammv nmc. A aa¢.c ammc scammsm Cmfmv Ammfmv afiumv ufiquv wowum umw c umm c wHomDE c uw>wq c manmwum> mnemcmuoonsm Hmcmuwpwm msafimmwwcoq n Mmfimmz mfiu .MOHOHhm flhmvcmum Ucm wfimwz .mc< cz< xmm .cmmmm mm mmcmmHa mcch<> zH meH>Hac< mmmammszmm mAn .oAE 00 :A wowuommu mm .vm>ummno mums mcvoomumu:A uamoAmAawAm coca hflco voodommuo ncAmuoum wE\muA:= mahuom u qu>Auom koAooom mm mum momma on» mo uouuo vuwvcmum cam mcmozm mwm.m uwNH.H A moo.m omN.H A panama owNH.H A amo.H wNH.H A m¢¢.¢ m3m vomNH.H A nm¢.~ wNH.H A mmm.m 5mm c3ovnusom xaommom an.umc aaa.uac amc.umc umm msoocmuoooSm momm. A Ham.o x.oooo. A nom.u umnuoz ooh. A w¢m.m women. A qu.¢ mam ooh. A ww~.¢ noon. A Nmm.m 8mm coconusom xHommsm Amm.umc aam.umc Amc.umv gum HmoonAuom Asa.umc amm.umc Asm.umc mAuasa msaAmwAwooA Ammo. A owo.o %.vmmo. A me.H x.vommo. A oo¢.o Mama mm «me. A mm¢.o ammo. A omm.o wmmo. A swo.o xoma ca mmo. A mw¢.o ammo. A 0mm.o comma. A «om.o xwo3 w ammo. A ~m¢.o ammo. A onm.o use; AN N.UOMH. H me.O %.wUOMH. H mmhco Suuwm Assam: mam Ema . AAc.nmc Aac.ncc mamc.umc Aa>Ac mm< x xmm ow< x vooum xom x vooum oommAH «.mmammHH mDOHM<> zH NHH>HHU< mm casuAB muawuomumoom ucmumMMAn £uw3 mam02m.m.v .mmmmnuaoumm 5A czosm mommAu was oHnmAum> some now wocmowwwchm mo Hm>oqo .mcoAum>uomno mo umcsdzn .cAououa wa\cAE\woNAcho mam c---- A ccc.c Ammo c---- A cccnc Ammo c---- A ccc.c ammo mama mm c---- A ccc.c Ammo c---- A ccc.c Amsc c---- A ccc.c Ammc saws mm m---- A ccc.c AmAV c---- A ccc.c Amsc c---- A ccc.c Ammo mama m mesa. A asa.m Ace ommm. A asa.A Aac scam. A ma¢.A aac Anon am mesa. A smm.m Ame macs. A amm.c Asc momma. A «Na.c ch muAAm Amcdumc amc.wav aqc.umv .mmq ---- A ccc.c aNAV --u- A ccc.c Ammc ---- A ccc.c Ammo Assam: «as. A cmN.A AAAV mmA. A mmm.c AaAc amA. A ams.c Aamv mam asA. A mam.c Amsv cam. A cmm.c Amsv «as. A Ham.c Ammo sum Amc.wac Aca.nac Ama.umv .mmw comm. A Nmm.c Aamc mma. A mcs.c Acme mmA. A cmH.c Aamc csocnoaom mama. A Ama.c Asmc mma. A acm.c Ammc smA. A mam.c Ammo xaommcm Asc.umc amm.umc uaac.umv cmoAm umm C wfiumg d Hm>HA fi anmem> HmcmuAumm moaAmmchoq n Madam: ~30 mo Houum Unmvcmuw @Gm mammz .mo< nz< xmm .QMMMm Mm mmDmmHH mDOHm<>~fiH MHH>HHU< mmog .awmuoum wE\cAB\0mNA0Axo mm n .0o>ummno mueB mcoAuomumucH osmoAMAomAm cos? xaco cmucmmwuo mum momma use 00 uouum wumwcmum 0cm annex” -- ccc.c o---- ccc.c o-- ccc.c mama mm nun 000.0 ouuuu 000.0 ounnu 000.0 x003 0H nun 000.0 onus: 000.0 onus: 000.0 xoma w a.assm. A amm.a x.csaa. A mmm.~ use; am 0mmd. A Hmm.H mmmm. A HH¢.m Luuwm umnuwz mam 6mm AAc.uav Amc.umc Aam.umc cam HmcmuAumm Amm.umv Amm.umv Amq.umv saunas mDEAmmAmaoa Amm.umc AsA.ucc maam.nmc Am>Au ow< x sow mw< x woman xmm x woman osmmAH ZH wHH>HHU< mm .msH mamas 98 in this study. At low levels of CCE activity, the homogenate must be in a fairly pure state in order to obtain a significant enzyme response. Although CCE activity was highest in adipose tissues as was SYN activity, it was not closely related to mitochondrial acetyl COA synthe- tase activity. Simple correlations between CCE and the other enzymes measured are presented in Appendixes IV, V and VI. In summarizing the results of this study, perirenal fat showed sig- nificantly (P<;05) greater CCE activities than muscle and liver. There were no significant trends for breed and sex effects on CCE activity in the various tissues studied. Age had the greatest influence with PRF from the neonate and 24 hour age groups possessing significantly (P <.05) higher levels of CCE than that at the 8, l6 and 32 weeks. The level of CCE activity was not highly associated with the level of mitochondrial acetyl COA synthetase activity. Malonyl COA Formation The immediate precursor of fatty acids, malonyl COA, is formed from cytoplasmic acetyl COA by the action of the enzyme acetyl COA carb0xylase (CBX). CBX is the first enzyme involved in the sequence of the cytoplasmic pathway of fatty acid synthesis and has been implicated by many researchers (Vagelos, 1964; Numa g5 al,, 1965; Lane and Moss, 1971) as the key enzyme in the regulation of gg_gggg fatty acid synthesis in animal tissues. Bau- man _£ _1. (1972) reported that in sheep adipose tissue CBX was the most closely related to lipogenesis of all the enzymes measured. However, the specific involvement of CBX in affecting the rate of lipogenesis has not been thoroughly elucidated. 99 Acetyl COA Carb0xylase The means and standard error of the means of CBX activity in various tissues are presented in figure 11. SCF showed significantly (P <.05) greater CBX activity than liver, LM and PRF. PRF and liver possessed significantly (P <.05) higher enzyme activity than LM. These results are in partial agreement with the findings of Chakrabarty and Leveille (1969) who also reported greater CBX in rat adipose tissue than in liver; however, the observation that CBX activity in sheep liver was essentially equal to that found in PRF is in disagreement with their results. Lower levels of CBX activity were found in sheep liver, PRF and SCF in this study compared to that reported by the latter authors in rat liver and adipose tissue. These results may be due to the crude homogenate (high protein concentration) used in this study and/or the effect of frozen storage on enzyme activity. However, the possibility of a species difference cannot be excluded since this comparison is made between non-ruminants (rat) and ruminants (sheep). Means and standard error of the means of CBX activity in various tissues by breed, sex and age are presented in table 15. Except for muscle, the tissues of Southdowns showed greater CBX activity than those of Suffolks, but none of these differences was significant. However, the breed differ- ences in CBX activities in muscle and SCF approached statistical signifi- cance (P = .09 and P = .11, respectively). CBX activities due to sex differences in liver, LM and PRF were nonsignificant, however, those of SCF approached significance (P = .09). The SCF and PRF of ram and ewe lambs had considerably greater CBX activities than those from wethers. 100 nM C02 incorporated/min/mg protein V |.20 0.80 - 0.40 ///////////////////////%}—*° W” Liver LM PRF SCF 0 Figure 11. Means and standard error of the means of acetyl COA carb0xy- lase activity in various tissues. a’b’cMeans with different superscripts differ significantly (P <.05). lOl .Amo. V my kHqukoAcmAm noMMAc canaoo can manmAum> awnuws muawuomuoaom uchmMMAw nuw3 mammz .o.v .mommnucoumo GA :3osm mommuu 0cm oHnmAum> some you ooomkoAawwm mo HW>QH0 .mCOququmQO NO hmnszn .cAwuouo wa\cA8\coumuoouoocA N00 00 mmHoaocmc mm commouoxo monfim>m AAAA. A aqm.A ANAV acmA. A amm.A ANAc cAmc. A mmA.c AAAV cac. A mca.c AmAc Aces Na cssA. A mam.c ANAc AVcmA. A cma.c ANAV cANc. A msA.c AmAc cac. A maa.c ANAV Amma mA AAA. A aca.c ANAc ccmA. A mmm.c ANAV «Amc. A mm~.c AmAc cac. A Ama.c AmAc Am»; m acmmA. A sma.c Aac mcmc. A Amm.c Aav acA. A mac.c Aac use; AN acccm. A mcm.c Ase mmmc. 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A A0a.m xoms mm MON. A mmm.0 umom. A doq.o omom. A omH.H xmos 0A now. A now.o umom. A mow.o omom. A 000.0 xmoa w nosuoz 03m 5mm AAc.nmv Amn.umv AcA.nmc cam msoocmuSoQSm mw< x xom ow< x vwonm xmm x vomnm osmmAH Acmscnuaoov «.mmcmmAe mccAmm> zA mnA>AHc< mchmxcmm :AnuA3 moaAnomnooom ucmanMAw nuA3 mammZm.e .momwcucenmo aA osonm mommAu was oAnmAnm> some now mocmoAWAchm mo Hm>mqo .mcoAum>nmmno mo nwnaozb .cwmuono wE\cAE\emo:wono mmam Am.m A cm.ss AmAc m.cmAa A ms.mm ANAV AA. A sm.c ANAV men. A ac.s AmAc Ammz mm Am.c A ma.mm ANAV mmA.A A sm.ma ANAV mm. A ma.c AAAV can. A mA.m ANAV Aces mA Am.m A mA.mm ANAV cmA.s A Ac.mm ANAV mm. A ma.c ANAV use. A mA.A ANAV Ame; m cam.m A cs.mm Ame mm. A ma.c mac mmm. A am.cA Aac Ago; Am cmA.a A mn.mm Ase mm. A mm.c Asc ca~.A A ms.m Ase euAAm Anc.umv AAc.nmv Amm.nmc AAc.umc mm< msm.m A cm.Am AmAc AA.A A mA.Am ANAc mm. A mm.c AmAv An. A ~m.m ANAV Ameumz Asm.m A Am.mm AAAV us.m A aa.a~ AAAV mA. A mn.c AaAc am. 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A Amo.o :nnAm nonnoz osm Eom AAc.nmc Amm.umc mAaA.umc mAnass mDEAmmAwooq ow< x xom ow¢ x woonm xom x woonm oommAH ommbmmHH mDOHm¢> ZH m0H0¢ wHH¢m 0000 02¢ wHH>HHO¢ mm¢0HA ZHMHommoqu mo mZOHHU¢000HZH 00¢ x xmm 02¢ 00¢ x 00000 .xmm x 00000 .¢mm 000¢H 151 PRF of lambs at 8 weeks of age exhibited significantly (P <.05) higher LPL activities than that at birth, 24 hours and 32 weeks of age. The high level of LPL activities in PRF appears to coincide with the high level of lipogenic activity usually found in this depot at 8 weeks of age. These observations suggest that during periods of high rates of fat deposi- tion in PRF, deposition of fat may be the result of both lipogenic activity and circulating triglyceride uptake. Also, these results indi- cate that the maximum rate of fat deposition in PRF probably occurs before the lambs reach 16 weeks of age. The influence of age on LPL activity in SCF was not significant and does not appear to be related to the level of lipogenic activity or rate of fat deposition. The influence of breed and sex on free fatty acid levels in blood plasma were not significant (table 23). Free fatty acid concentrations in blood plasma at birth were significantly (P <.05) higher than those at 24 hours, 8, l6 and 32 weeks of age. These high FFA levels no doubt reflect the fasted state of the neonatal lambs and probably resulted from mobilization of body fat to supply energy for vital functions. After milk ingestion the FFA concentration in blood decreased to a low level as indicated by the plasma FFA at 24 hours. This low level probably resulted from increased blood glucose which stimulated insulin secretion. The effect of insulin is two-fold as it exerts an antilipolytic effect which decreases fatty acid mobilization and it simultaneously stimulates triglyceride uptake. These two effects could account for the low FFA observed in plasma at 24 hours. FFA concentrations in plasma at 16 weeks were significantly (P <.05) higher than those at 24 hours. The plasma FFA 152 increased between 24 hours and 8 weeks, and then essentially stabilized. These results probably reflect increased rumen function which is associated with reduced blood glucose. Reduced blood glucose generally results in decreased insulin secretion which would tend to diminish triglyceride up- take and this coupled with the increased insulin insensitivity of maturing ruminants could possibly account for the higher FFA levels in blood of the older lambs. Moreover, the increase in rumen function generally results in increased plasma FFA. The simple correlations for lipoprotein lipase activity and free fatty acids with the activity of the other enzymes measured are presented in Appendixes IV, V, VI and VII. In summarization, lipoprotein lipase activities were greater in adipose tissues than in muscle. The level of LPL activity in adipose tissues does not appear to be related to propensity to fatten. LPL acti- vity seems to be related to the level of lipogenic activity only in PRF of lambs. Plasma FFA appeared to be associated with the energy state of lambs as well as rumen development. SUMMARY Forty-nine lambs were included in an experiment designed to study the influence of breed, sex and age on lipogenic enzyme activities in some ovine tissues. Thirty-nine western crossbred ewes were divided into two groups based upon body length and body weight. The smaller bodied ewes were mated to a Southdown ram and the larger bodied ewes were mated to a Suffolk ram in order to obtain two strains of lambs differing in propen- sity to fatten. Lambs were slaughtered at birth, 24 hours, 8, l6 and 32 weeks of age and blood, liver, longissimus muscle (LM), perirenal (PRF) and subcutaneous fat (SCF) samples were collected from the carcass. En- zymes assayed were glucose-6-phosphate dehydrogenase (G-6PDH), 6-phospho- gluconate dehydrogenase (6-PCDH), NADP-malate dehydrogenase (ME), NADP- isocitrate dehydrogenase (ICDH), acetyl COA carb0xylase (CBX), acetyl COA synthetase (SYN), lipoprotein lipase (LPL) and ATP-citrate lyase (CCE). Plasma free fatty acids (FFA) were also determined. Acetyl COA synthetase activities were significantly (P <.05) higher in adipose tissues than in liver and LM. The results showed that 35 to 40 percent of the SYN activity in adipose tissue was of cytoplasmic origin and 60 to 65 percent was located intramitochondrially. Tissues of South- down lambs had higher SYN activities than those of Suffolks. These data indicated that Southdowns, which have a greater propensity to fatten, possessed a greater potential to utilize acetate for acetyl COA synthesis and possibly for lipogenesis. There were no consistent or apparent patterns of SYN activities due to sex in the various tissues. The influ- ence of age on the activities of SYN was significant, but the pattern 153 154 observed in each instance was tissue specific. The SYN activities appeared to be related to the level of lipogenic activity only in PRF. PRF showed significantly greater CCE activities than LM and liver. The influence of breed and sex on CCE activities in the various tissues studied were not significant. Neonate and 24 hour lambs possessed greater CCE activities than those at 8, 16 and 32 weeks. CCE did not appear to be related to the predisposition to fatten and the level of lipogenic activity. SCF had the greatest CBX activities followed in order by liver, PRF and LM. Adipose tissues of Southdowns showed higher CBX activities than those of Suffolks. There appeared to be a relationship between CBX activity in ovine adipose tissues and the propensity to fatten. PRF and SCF of 32 week ewe and ram lambs had greater CBX activities than those of wethers. CBX activities in the adipose tissues of 32 week rams and ewes were greater than those at all other ages; whereas, the 32 week wethers possessed lower CBX activities than at 16 weeks. The influence of sex on predisposition to fatten tended to be related to CBX activities in adipose tissues, but this relationship did not apply to ewes. CBX activities in PRF and SCF appeared to be related to the level of lipo- genic activity. CBX appeared to be an important enzyme in fatty acid synthesis in ovine adipose tissues. PRF and SCF had significantly (P <.05) higher G-6PDH activities than liver and LM. 6-PGDH activity was greatest in SCF followed in order by liver, PRF and LM. 6-PGDH activities were greater than those of G-6PDH in ovine adipose tissues. The effect of breed on G-6PDH activities was 155 not significant and the general pattern of G-6PDH activities among differ- ent sexes did not appear to be related to the predisposition to fatten. G-6PDH activity tended to be related to lipogenic activity in lambs as it adapted to changes in the level of lipogenic activity due to age. Tissues of Southdowns exhibited greater 6—PGDH activities than those of Suffolks which indicated that this enzyme was related to the propensity to fatten. There were no consistent or apparent patterns due to sex on 6-PGDH activities in the various tissues studied. 6-PGDH activities in- creased considerably with age in PRF and SCF and it appeared that this enzyme was related to the level of lipogenic activity. 6-PGDH activities tended to be regulated by the level of lipogenic activity and this indi- cated that this enzyme was adaptable as well as not rate limiting. The pentose shunt dehydrogenases, G-6PDH ande-PGDH, are capable of generating NADPH for lipogenesis as indicated by the fact that they contributed 33 to 50 percent of total NADPH generating potential in SCF and PRF of lambs from 8 to 32 weeks of age. ME activities in the adipose tissues were greater than those in liver and LM. The effect of breed on ME activities in the various tissues was not significant. Liver and LM of rams possessed the highest ME activities followed in order by ewes and wethers, but the differences due to sex in PRF and SCF were nonsignificant. ME activities decreased with age in liver and LM which appeared to reflect the decreased citrate cycle activity usually associated with maturing lambs. However, ME activities increased with age in PRF and SCF and this trend followed the general pattern ob- served for lipogenic activity in growing lambs. ME appeared to play a 156 limited role in NADPH generation in ovine adipose tissues as indicated by its slow adaptation and the fact that its contribution to the total NADPH generating potential was minor (13 to 19 percent) in PRF and SCF. Cytoplasmic ICDH activities in PRF and SCF were greater than those in liver and LM. Mitochondrial ICDH activities were highest in LM followed in order by SCF, PRF and liver. The effect of breed on cytoplasmic ICDH in the adipose tissue depots was not significant. PRF of wethers possessed greater cytoplasmic ICDH activities than those of ewes and both sexes had greater enzyme activities than rams. However, SCF of rams possessed the highest cytoplasmic ICDH followed in order by wethers and ewes. The effects of breed and sex on mitochondrial ICDH in the various tissues stu- died were not significant. The response of cytoplasmic ICDH to changes in the level of lipogenic activity was not as great as that observed for the pentose shunt dehydrogenases. Mitochondrial ICDH activities in the adipose tissues gradually increased with age and appeared to be related to changes in the level of lipogenic activity usually associated with maturing lambs. The presence of significant ICDH activities in ovine adipose tissues and the finding that ICDH contributed 35 to 54 percent of the total NADPH generating potential in PRF and SCF indicates that ICDH plays a major role in the generation of NADPH for lipogenesis in the rumin- ant. LPL activities were highest in SCF followed in order by PRF and LM which indicated that adipose tissues were more actively involved in tri- glyceride uptake than LM and this probably reflected the larger quantities of triglycerides being deposited in those depots at the ages studied. The 157 influence of breed on LPL activities in the various tissues studied was not significant. LPL appeared to be related to the propensity to fatten in LM since lambs with the greatest predisposition to fatten (wethers and ewes) had the greatest activities. LPL activities in PRF appeared to coincide with the changes in rate of fat deposition usually found in lambs between birth and 32 weeks of age. These data suggested that the maximum rate of fat deposition in PRF probably occurred before the lambs reached 16 weeks of age. The results of this study indicated that during periods of high rates of fat deposition in PRF, fat deposition may be the result of both high lipogenic activity and uptake of circulating triglycerides. The influence of age on LPL activities was not significant and this enzyme did not appear to be related to the rate of fatty deposition in SCF. The influence of breed and sex on plasma FFA levels was nonsignifi- cant. The energy states of the lambs as well as the development of rumen function appeared to be the major factors affecting plasma FFA. LITERATURE CITED Aas, M. 1971. Organ and subcellular distribution of fatty acid activating enzymes in the rat. Biochem. Biophys. Acta. 231:32. Aas, M. and J. Bremer. 1968. Short-chain fatty acid activation in rat liver. A new assay procedure for the enzymes and studies on their intracellular localization. Biochem. Biophys. Acta. 164:157. Abraham, 8., K. J. Matthes and I. L. Chaikoff. 1961. Factors involved in synthesis of fatty acids from acetate by a soluble fraction obtained from lactating rat mammary gland. Biochem. Biophys. Acta. 49:268. Adler, J. H. and H. E. Wertheimer. 1968. Aspects of fat deposition and mobilization in adipose tissue. In G. A. Lodge and G. E. Lamming (Ed.) Growth and Development of Mammals. New York Plenum Press, London. Allee, G. L., D. Romsos, G. A. Leveille and D. H. Baker. 1971. Influence of age on ig_vitro lipogenesis and enzymatic activity in pig adipose tissue. Proc. Soc. Exp. Biol. Med. 137:449. Anderson, D. B. and R. G. Kauffman. 1972. Cellular and enzymatic changes in porcine adipose tissue growth. J. Lipid Res. (Submitted). Anderson, D. B., R. G. Kauffman and L. L. Kastenschmidt. 1972. Lipogenic enzyme activities and cellularity of porcine adipose tissue from various anatomical locations. J. Lipid Res. (In Press). Anfinsen, C. B., E. Boyle and R. K. Brown. 1952. The role of heparin in lipoprotein metabolism. Science 115:583. Annison, E. F., J. L. Linzell, S. Fazakerley and B. W. Nichols. 1967. The oxidation and utilization of palmitate, stearate, oleate and acetate by the mammary gland of the fed goat in relation to their overall metabolism, and the role of plasma phospholipids and neutral lipids in milk fat synthesis. Biochem. J. 102:637. Armstrong, D. G. 1965. Carbohydrate metabolism in ruminants and energy supply. In R. W. Daugherty (Ed.) Physiology of Digestion in the Ruminant. Butterworth, Washington. Armstrong, D. G. and K. L. Blaxter. 1957. The utilization of acetic, propionic and butyric acids by fattening sheep. Brit. J. Nutr. 11:413. Armstrong, D. G., K. L. Blaxter and N. McC. Graham. 1957. The heat in- crements of mixtures of steam-volatile fatty acids in fasting sheep. Brit. J. Nutr. 11:392. 158 159 Armstrong, D. G., K. L. Blaxter, N. McC. Graham and F. W. Wainman. 1958. The utilization of energy of two mixtures of steam-volatile fatty acids by fattening sheep. Brit. J. Nutr. 12:177. Babatunde, G. M., W. G. Pond, E. F. Walker, Jr. and P. Chapman. 1968. Effects of dietary safflower oil or hydrogenated coconut oil on heart, liver and adipose tissue fatty acid levels and physical carcass measurements of pigs fed a fat-free diet. J. Anim. Sci. 27:1290. Baldwin, R. L. and L. P. Milligan. 1966. Enzymatic changes associated with the initiation and maintenance of lactation in the rat. J. Biol. Chem. 241:2058. Baldwin, R. L. and M. Ronning. 1966. Dietary fat levels and carbohydrate and fat metabolism in calves. J. Dairy Sci. 49:688. Baldwin, R. L., M. Ronning, C. Radanovics and G. Plange. 1966. Effect of carbohydrate and fat intakes upon the activities of several liver enzymes in rats, guinea piglets, piglets and calves. J. Nutr. 90:47. Ball, E. G. 1966. Regulation of fatty acid synthesis in adipose tissue. In G. Weber (Ed.) Advances in Enzyme Regulation. Vol. 4. Pergamon Press, New York. Ball, E. G. 1970. Some considerations of the multiplicity of insulin action on adipose tissue. In B. Jeanrenaud and D. Hepp (Ed.) Adipose tissue. Regulation and Metabolic Functions. Academic Press, New York. Ballard, F. J. and I. T. Oliver. 1964. The effect of concentration on glucose phosphorylation and incorporation into glycogen in the livers of foetal and adult rats and sheep. Biochem. J. 92:131. Ballard, F. J. and R. W. Hanson. 1967. The citrate cleavage pathway and lipogenesis in rat adipose tissue: replenishment of oxaloacetate. J. Lipid Res. 8:73. Ballard, F. J., R. W. Hanson and D. S. Kornfeld. 1969. Gluconeogenesis and lipogenesis in tissue from ruminant and non-ruminant animals. Fed. Proc. 28:218. Ballard, F. G., O. H. Filsell and T. G. Jarrett. 1972. Effects of carbo- hydrate availability on lipogenesis in sheep. Biochem. J. 126:193. Barcroft, J., R. A. Mc Anally and A. T. Phillipson. 1944. Absorption of volatile fatty acids from the alimentary tract of sheep and other animals. J. Exp. Biol. 20:120. 160 Barrick, E. R., T. N. Blumer, W. L. Brown, F. H. Smith, S. B. Tove, H. L. Lucas and H. A. Stewart. 1953. The effect of feeding several kinds of fat on feed lot performance and carcass characteristics of swine. J. Anim. Sci. 12:899. Barry, J. M., W. Bartley, J. L. Linzell and D. S. Robinson. 1963. The uptake from the blood of triglyceride fatty acids of chylomicra and low-density lipoproteins by the mammary gland of the goat. Biochem. J. 89:6. Barth, C., M. Sladek and K. Decker. 1971. The subcellular distribution of short-chained fatty acyl-COA synthetase activity in rat tissues. Biochem. Biophys. Acta. 248:24. Bartley, J. C., R. A. Freedland and A. L. Black. 1966. Effect of aging and glucose loading on the activities of glucose-6-phosphatase and phosphorylase of livers of cows and calves. Amer. J. Vet. Res. 27:1243. Bauman, D. E., R. E. Brown and C. L. Davis. 1970. Pathways of fatty acid synthesis and reducing equivalent generation in mammary gland of rat, sow and cow. Arch. Biochem. Biophys. 140:237. Bauman, D. E., D. E. Johnson and R. W. Mellenberger. 1972. Lipogenesis and lipolysis of sheep adipose tissue during fattening. J. Anim. Sci. 35:203. (Abstr.). Beinhert, H., D. E. Green, P. Hele, H. Heft, R. W. Von Korff and C. V. Ramakrishnan. 1953. The acetate activating enzyme of heart muscle. J. Biol. Chem. 203:35. Beitz, D. 1972. Personal communication. Department of Animal Science. Iowa State University. Ames, Iowa. Bergman, E. N., R. S. Reid, M. G. Murray, J. M. Brockway and F. G. Whitelow. 1965. Interconversions and production of volatile fatty acids in the sheep rumen. Biochem. J. 97:53. Bergman, E. N., W. E. Roe and K. Kon. 1966. Quantitative aspects of pro- pionate metabolism and g1uconeogenesis in sheep. Amer. J. Physiol. 211:793. Bergman, E. N., M. L. Katz and C. F. Kaufman. 1970. Quantitative aspects of hepatic and portal glucose metabolism and turnover in sheep. Amer. J. Physiol. 219:785. Bezman, A., J. M. Felts and R. J. Havel. 1962a. The incorporation of tri- glyceride fatty acid into adipose tissue as a function of lipoprotein lipase activity. Clin. Res. 10:84. 161 Bezman, A., J. M. Felts and R. J. Havel. 1962b. Relation between incor- poration of triglyceride fatty acids and heparin-released lipoprotein lipase from adipose tissue slices. J. Lipid Res. 3:427. Bickerstaffe, R. 1971. Uptake and metabolism in lactating mammary gland. In I. R. Falconer (Ed.) Lactation. The Pennsylvania State University Press, University Park. Bishop, C., T. Davies, R. F. Glascock and V. A. Welch. 1969. Studios on the origin of milk fat. Biochem. J. 113:629. Bjorntorp, P. 1966. Studies on adipose tissue from obese patients with or without diabetes mellitus. Acta. Med. Scand. 179:229. Bjorntorp, P. and L. Sjostrom. 1971. Number and size of adipose tissue fat cells in relation to metabolism in human obesity. Metab. Clin. Exp. 20:703. Blanchette-Mackie, E. J. and R. 0. Scow. 1971. Sties of lipoprotein lipase activity in adipose tissue perfused with chylomicrons. Cell Biol. 51:1. Blaxter, K. L. 1962. In The Energy Metabolism of Ruminants. Hutchinson Scientific and Technical, London. Blaxter, K. L. 1966. In The Energy Metabolism of Ruminants. Charles C. Thomas Publishing, Springfield, Illinois. Bortz, W. M. and F. Lynen. 1963a. Elevation of long chain acyl-COA de- irivatives in liver of fasted rats. Biochem. Z. 339:77. Bortz, W. M. and F. Lynen. 1963b. The inhibition of acetyl COA carb0xylase by long chain acyl-COA derivatives. Biochem. Z. 337:505. Brady, R. 0. 1958. The enzymatic synthesis of fatty acids by aldol con- densation. Proc. Nat. Acad. Sci. U.S.A. 44:993. Brown, J. and P. McLean. 1965. Effect of alloxan-diabetes on the activity of citrate enzyme in adipose tissue. Nature 207:407. Brown, J., P. McLean and A. L. Greenbaum. 1966. Influence of thyroxine and luteinizing hormone on some enzymes concerned with lipogenesis in adipose tissue, testis and adrenal gland. Biochem. J. 101:197. Butcher, R. W. 1969. Cyclic 3', 5'-AMP and the lipolytic affects of hormones on adipose tissue. Pharm. Rev. 18:237. Butcher, R. W. and E. W. Sutherland. 1967. The effects of the catechola- mines, adrenergic blocking agents, prostaglandin E, and insulin on cyclic AMP levels in the rat epididymal fat pad in vitro. Ann. N.Y. Acad. Sci. 139:849. 162 Butcher, R. W., R. J. Ho, H. C. Meng and E. W. Sutherland. 1965. Adenosine 3', 5'-monophosphate in biological materials. II. The measurement of adenosine 3', 5'-monophosphate in tissues and the role of the nucleo- tide in the lipolytic response of fat to epinephrine. J. Biol. Chem. 240:4515. Butcher, R. W., J. T. Sneyd, C. R. Park and E. W. Sutherland. 1966. Effect of insulin on adenosine 3', 5'-monophosphate in the rat epididymal fat pad. J. Biol. Chem. 241:1651. Butcher, R. W., C. E. Baird and E. W. Sutherland. 1968. Effects of lipo- lytic and antilipolytic substances on adenosine 3', 5'-monophosphate levels in isolated fat cells. J. Biol. Chem. 243:1705. Cahill, G. F., B. Leboeuf and R. B. Flinn. 1960. Studies on rat adipose tissue in vitro. VI. Effect of epinephrine on glucose metabolism. J. Biol. Chem. 235:1246. Chakrabarty, K. and G. A. Leveille. 1969. Acetyl COA carb0xylase and fatty acid synthetase activities in liver and adipose tissue of meal fed rats. Proc. Soc. Exp. Biol. Med. 131:1051. Chakrabarty, K. and J. R. Romans. 1972. Lipogenesis in the adipose cells of the bovine (Bos Taurus) as related to their intramuscular fat content. Comp. Biochem. Physiol. 418:603. Chang, H. C., I. Seidman, G. Teebor and M. D. Lane. 1967. Liver acetyl COA carb0xylase and fatty acid synthetase. Relative activities in the normal state and in hereditary obesity. Biochem. Biophys. Res. Comm. 28:682. Church, D. C. 1969. In Digestive Physiology and Nutrition of Ruminants. Vol. I. Oregon State University Bookstore Inc., Corvallis, Oregon. Cook, R. M. and L. D. Miller. 1965. Utilization of volatile fatty acids in ruminants. I. Removal of them from portal blood by the liver. J. Dairy Sci. 48:1339. Corbin, J. D. and C. R. Park. 1969. Permissive effects of glucocorticoids on lipolysis in adipose tissue. Fed. Proc.28:702. Corbin, J. D., E. M. Reimann, D. A. Walsh and E. G. Krebs. 1970. Activa- tion of adipose tissue lipase by skeletal muscle cyclic adenosine 3', 5'-monophosphate-stimulated protein kinase. J. Biol. Chem. 245: 4849. Cramer, D. A. 1972. Personal communications. Department of Animal Science. Colorado State University. Fort Collins, Colorado. 163 Crofford, O. B. and E. A. Renold. 1965a. Glucose uptake by incubated rat epididymal adipose tissue. Rate limiting steps and site of insulin action. J. Biol. Chem. 240:14. Crofford, 0. B. and E. A. Renold. 1965b. Glucose uptake by incubated rat epididymal adipose tissue. Characteristics of the glucose trans- port system and action of insulin. J. Biol. Chem. 240:3237. Dakshinamurti, K. and P. R. Desjardins. 1969. Acetyl COA carb0xylase from rat adipose tissue. Biochem. Biophys. Acta. 176:221. Davies, J. L. 1968. In vitro regulation of the lipolysis of adipose tissue. Nature 218:349. Davis, C. L. 1967. Acetate production in the rumen of cows fed either control or low fiber, high grain diets. J. Dairy Sci. 50: 1621. Davis, C. L., R. E. Brown, J. R. Staubus and W. 0. Nelson. 1960a. Avail- ability and metabolism of various substrates in ruminants. I. Absorp- tion and metabolism of acetate. J. Dairy Sci. 43:231. Davis, C. L., R. E. Brown and J. R. Staubus. 1960b. Availability and metabolism of various substrates in ruminants. III. The effect of propionate on acetate oxidation in_vivo. J. Dairy Sci. 43:1783. Del Boca, J. and J. P. Flatt. 1969. Fatty acid synthesis from glucose and acetate and the control of lipogenesis in adipose tissue. Europ. J. Biochem. 11:127. Denton, R. M. and M. L. Halperin. 1968. The control of fatty acid tri- glyceride synthesis in rat epididymal adipose tissue. Roles of coenzyme A derivatives, citrate and L-glycerol-B-phosphate. Biochem. J. 110:27. Di Girolamo, M. and S. Mendlinger. 1971. Role of fat cell size and number in enlargement of epididymal fat pads in three species. Amer. J. Physiol. 221:859. Di Girolamo, M., S. Mendlinger and J. W. Fertig. 1971. A simple method to determine fat cell size and number in four mammalian species. Amer. J. Physiol. 221:850. Di Luzio, N. R. 1960. Hepatic participation in lipid metabolism. J. Amer. Oil Chem. Soc. 37:163. Donaldson, W. E., E. M. Wit-Peeters and H. R. Scholte. 1970. Fatty acid synthesis in rat liver: Relative contributions of the mitochondrial, microsomal and non-particulate systems. Biochem. Biophys. Acta. 202:35. 164 Elsden, S. R., M. S. Hitchcock, R. A. Marshall and A. T. Phillipson. 1946. Volatile acids in the digesta of ruminants and other animals. J. Exp. Biol. 22:191. Exton, J. H., N. Friedmann, E. H. Wong, J. P. Breneaux, J. D.Corbin- and C. R. Park. 1972. Interaction of glucocorticoids with glucagon and epinephrine in the control of g1uconeogenesis, glycogenolysis in the liver and of lipolysis in adipose tissue. J. Biol. Chem. 247:3579. Fain, J. N. 1967. Andrenergic blockage of hormone induced lipolysis in isolated fat cells. Ann. N. Y. Acad. Sci. 139:879. Fain, J. N. 1968. Effect of dibutyryl 3', 5'-AMP, theophylline and norepinephrine on lipolytic action of growth hormone and glucocorti- coid in white fat cells. Endocrinol. 82:825. Fain, J. N. and R. Saperstein. 1970. The involvement of RNA synthesis and cyclic AMP in the activation of fat cell lipolysis by growth hormone and glucocorticoids. In B. Jeanrenaud and D. Hepp (Ed.) Adipose Tissue: Regulation and Metabolic Functions. Academic Press, New York. Fain, J. N., V. P. Kovacev and R. 0. Scow. 1965. Effect of growth hormone and dexamethasone on lipolysis and metabolism in isolated fat cells of the rat. J. Biol. Chem. 240:3522. Fain, J. N., V. P. Kovacev and R. O. Scow. 1966. Antilipolytic effect of insulin in isolated fat cells of the rat. Endocrinol. 78:773. Felinski, L., G. A. Carton, A. K. Lough and A. T. Phillipson. 1964. Lipids of sheep lymph: transport from the intestine. Biochem. J. 90:154. Feller, D. D. 1954. Metabolism of adipose tissue. J. Biol. Chem. 206:171. Filsell, O. H., I. G. Jarrett, M. R. Atkinson, P. Caiger and R. K. Morton. 1963. Nicotinamide nucleotide coenzymes and glucose metabolism in the livers of foetal and newborn lambs. Biochem. J. 89:92. Flatt, J. P. and E. G. Ball. 1964. Studies on the metabolism of adipose tissue. XV. An evaluation of the major pathways of glucose catabo- lism as influenced by insulin and epinephrine. J. Biol. Chem. 239:675. Foa, P. P. 1964. Glucagon. In G. Pincus, K. V. Thimann and E. B. Astwood (Ed.) The Hormones. Vol. 4. Academic Press, New York. Ford, E. H. 1965. The importance of glucose in ruminant metabolism. In K. L. Blaxter (Ed.) Energy Metabolism. Academic Press, New York. 165 Ford, E. H. and P. B. Reilly. 1969. Amino acid utilization in the ruminant. Res. Vet. Sci. 10:96. Formica, J. V. 1962. Utilization of citric acid for the biosynthesis of long-chained fatty acids. Biochem. Biophys. Acta. 59:739. Foster, D. W. and P. A. Srere. 1968. Citrate cleavage enzyme and fatty acid synthesis. J. Biol. Chem. 243:1926. Friend, D. W., H. M. Cunningham and J. G. Nicholson. 1962. The production of organic acids in the pig. 1. The effect of diet on the proportions of volatile fatty acids in pig feces. Can. J. Anim. Sci. 42:55. Friend, D. W., H. M. Cunningham and J. G. Nicholson. 1963a. The pro- duction of organic acids in the pig. II. The effect of diet on the levels of volatile fatty acids and lactic acid in sections of the alimentary tract. Can. J. Anim. Sci. 43:156. Friend, D. W., H. M. Cunningham and J. G. Nicholson. 1963b. Volatile fatty acids and lactic acid in the alimentary tract of the young pig. Can. J. Anim. Sci. 43:174. Friend, D. W., J. G. Nicholson and H. M. Cunningham. 1964. Volatile fatty acid and lactic acid content of pig blood. Can. J. Anim. Sci. 44:303. Garfinkel, A. S., N. Baker and M. C. Schotz. 1967. Relationship of lipo- protein lipase activity to triglyceride uptake in adipose tissue. J. Lipid Res. 8:274. Garton, G. A. 1965. The digestion and assimilation of lipids. In R. W. Dougherty (Ed.) Physiology of Digestion in the Ruminant. Butterworth, Washington. Carton, G. A. and W. H. Duncan. 1954. Dietary fat and body fat: The composition of the backfat of pigs fed on diets rich in cod-liver oil and lard. Biochem. J. 57:125. Gellhorn, A. and W. Benjamin. 1965. Effects of aging on the composition and metabolism in adipose tissue in the rat. In A. E. Renold and G. F. Cahill (Ed.) Handbook of Physiology. Section 5. Adipose Tissue. Amer. Physiol. Soc. Washington, D. C. Goetsch, D. D. 1966. Liver enzymes changes during rumen development in calves. Amer. J. Vet. Res. 27:1187. Goodridge, A. G. and E. G. Ball. 1967. Lipogenesis in the pigeon: in vivo studies. Amer. J. Physiol. 213:245. Gregolin, C., E. Ryder, A. K. Kleinschmidt, R. C. Warner and M. D. Lane. 1966. Molecular characteristics of liver acetyl COA carb0xylase. Proc. Nat. Acad. Sci. U.S.A. 56:148. 166 Gregolin, C., E. Ryder and M. D. Lane. 1968a. Liver acetyl coenzyme A carb0xylase. J. Biol. Chem. 243:4227. Gregolin, C., E. Ryder, R. C. Warner, A. K. Kleinschmidt, H. Chang and M. D. Lane. 1968b. Liver acetyl coenzyme A carb0xylase. J. Biol. Chem. 243:4236. Grunert, R. R. and P. H. Phillips. 1951. A modification of the nitro- prusside method of analysis for glutathione. Arch. Biochem. 30:217. Gul, B. and R. Dils. 1969. Enzymatic changes in rabbit and rat mammary gland during the lactation cycle. Biochem. J. 112:293. Hanson, R. W. and F. J. Ballard. 1967. The relative significance of acetate and glucose as precursors for lipid synthesis in liver and adipose tissues from ruminants. Biochem. J. 105:529. Hanson, R. W. and F. J. Ballard. 1968. The metabolic fate of the pro- ducts of citrate cleavage. Biochem. J. 108:705. Hardwick, D. C. 1966. The fate of acetyl groups derived from glucose in the isolated perfused goat liver. Biochem. J. 99:228. Hardwick, D. C., J. L. Linzell and T. B. Mepham. 1963. The metabolism of acetate and glucose by the isolated perfused udder. Biochem. J. 88:213. Harlen, W. R. and S. J. Wakil. 1963. Synthesis of fatty acids in animal tissues. J. Biol. Chem. 238:3216. Hartman, A. D., A. I. Cohen, C. J. Richane and T. Hsu. 1971. Lipolytic response and adenyl cyclase activity of rat adipocytes as related to cell size. J. Lipid Res. 12:498. Hele, P. 1954. The acetate activating enzymes of beef heart. J. Biol. Chem. 206:671. Hirsch, J. and P. W. Han. 1969. Cellularity of rat adipose tissue of growth, starvation and obesity. J. Lipid Res. 10:77. Hirsch, J. and J. L. Knittle. 1970. Cellularity of obese and nonobese human adipose tissue. Fed. Proc. 29:1516. Hobson, P. N. and R. Summers. 1966. Effect of growth rate on the lipase activity of a rumen bacterium. Nature 209:736. Hollenberg, C. H. 1959. Effect of nutrition on activity and release of lipase from rat adipose tissue. Amer. J. Physiol. 197:667. 167 Hollenberg, C. H. 1960. The effect of fasting on the lipoprotein lipase activity of rat heart and diaphragm. J. Clin. Invest. 39:1282. Holter, J. B., R. D. McCarthy and E. M. Kesler. 1963. Butyrate metabolism in the isolated, perfused goat liver. J. Dairy Sci. 46:1256. Hood, R. L. 1972. Adipose Tissue Cellularity and Lipogenic Activity in Porcine and Bovine Animals. Ph.D. Thesis. University of Minnesota, St. Paul. Horino, M., L. J. Machlin, F. Hertelendy and D. M. Kipnis. 1968. Effect of short-chain fatty acids on plasma insulin in ruminant and non- ruminant species. Endocrinol. 83:118. House, W. A. and R. W. Phillips. 1968. Changes in glucose metabolism during postnatal development of the lamb. Fed. Proc. 27:557. Howanitz, T. J. and H. R. Levy. 1965. Acetyl COA carb0xylase and citrate cleavage enzyme in the rat mammary gland. Biochem. Biophys. Acta. 106:430. Huttunen, J. K. and D. Steinberg. 1971. Activation of phosphorylation of purified adipose tissue hormone-sensitive lipase by cyclic AMP- dependent protein kinase. Biochem. Biophys. Acta. 239:411. Huttunen, J. K., D. Steinberg and S. E. Mayer. 1970a. ATP-dependent and cyclic AMP-dependent activation of rat adipose tissue lipase by protein kinase from rabbit skeletal muscle. Proc. Nat. Acad. Sci. U.S.A. 67:290. Huttunen, J. K., D. Steinberg and S. E. Mayer. 1970b. Protein kinase activation and phosphorylation of a purified hormone sensitive lipase. Biochem. Biophys. Res. Commun. 41:1350. Iliffe, J. and N. B. Myrant. 1970. The sensitivity of acetyl coenzyme A carb0xylase to citrate stimulation in a homogenate of rat liver. Biochem. J. 117:385. Jansen, G. R., C. F. Hutchison and M. E. Zanetti. 1966. Studies of lipo- genesis in vivo. Effect of dietary fat or starvation on conversion of 14C-glucose into fat and turnover of newly synthesized fat. Biochem. J. 99:333. Jarrett, I. G., G. B. Jones and B. J. Potter. 1964. Changes in glucose utilization during development of the lamb. Biochem. J. 90:189. Jungas, R. L. 1966. Role of cyclic 3', 5'-AMP in the response of adipose tissue to insulin. Proc. Nat. Acad. Sci. U.S.A. 56:757. Jungas, R. L. and E. G. Ball. An antilipolytic action of insulin on adipose tissue. Fed. Proc. 21:202. 168 Kallen, R. G. and J. M. Lowenstein. 1962. The stimulation of fatty acid synthesis by isocitrate and malonate. Arch. Biochem. Biophys. 96:188. Katz, J., B. R. Landau and G. E. Bartsch. 1966. The pentose cycle, triose phosphate isomerization and lipogenesis in rat adipose tissue. J. Biol. Chem. 241:727. Keech, D. B. and M. Futter. 1963. Pyruvate carb0xylase. II. Properties. J. Biol. Chem. 238:2609. Keeny, M. 1970. Lipid metabolism in the rumen. In A. T. Phillipson (Ed.) Physiology of Digestion and Metabolism in the Ruminant. Oriel Press Limited, Newcastle. Khoo, J. C., L. Jarrett, S. E. Mayer and D. Steinberg. 1972. Subcellular distribution of epinephrine induced changes in hormone sensitive lipase, phosphorylase and phosphorylase kinase. J. Biol. Chem. 247:4812. Kinsella, J. E. 1970. Biosynthesis of lipids from [2-14C] acetate and D(-)-B-hydroxy [1,3-14C] butyrate by mammary cells from bovine and rat. Biochem. Biophys. Acta. 210:28. Knittle, J. L. and J. Hirsch. 1968. Effect of early -nutrition on the development of rat epididymal fat pads: cellularity and metabolism. J. Clin. Invest. 47:2091. Korchak, H. M. and E. J. Masoro. 1962. Changes in the level of fatty acid synthesis enzymes during starvation. Biochem. Biophys. Acta. 58:354. Korchak, H. M. and E. J. Masoro. 1964. Fatty acids as lipogenic inhibi- tors. Biochem. Biophys. Acta. 84:750. Korn, E. D. 1955a. Clearing factor, heparin-activated lipase. I. Isola- tion and characterization of enzyme from normal rat heart. J. Biol. Chem. 215:1. Korn, E. D. 1955b. Clearing factor, heparin-activated lipase. II. Sub- strate specificity and activation of coconut oil. J. Biol. Chem. 215:15. Kornacker, M. S. and E. G. Ball. 1965. Citrate cleavage in adipose tissue. Proc. Nat. Acad. Sci. U.S.A. 54:899. Kornacker, M. S. and J. M. Lowenstein. 1964. Citrate cleavage and acetate activation in liver of normal and diabetic rats. Biochem. Biophys. Acta. 84:490. Kornacker, M. S. and J. M. Lowenstein. 1965. Citrate and the conversion of carbohydrate into fat. Biochem. J. 94:209. 169 Kronfeld,. D. S. 1968. Acetate kinetics in normal and ketotic cows. J. Dairy Sci. 51:397. Landau, B. R. and J. Katz. 1965. Pathways of glucose metabolism. In A. E. Renold and G. F. Cahill (Ed.) Handbook of Physiology. Section 5. Adipose Tissue. Amer. Physiol. Soc. Washington, D. C. Lane, M. D. and J. Moss. 1971. Regulation of fatty acid synthesis in animal tissues. In H. J. Vogel (Ed.) Metabolic Pathways: Metabolic Regulation. Vol. 5. Academic Press, New York. Lane, M. D., J. Moss, E. Ryder and E. Stoll. 1971. The activation of acetyl COA carb0xylase by tricarboxylic acids. In G. Weber (Ed.) Advances in Enzyme Regulation. Vol. 9. Pergamon Press, New York. Langdon, R. G. 1957. The biosynthesis of fatty acids in rat liver. J. Biol. Chem. 226:615. Leat, W. F. 1970. Carbohydrate and lipid metabolism in the ruminant during post-natal development. In A. T. Phillipson (Ed.) Physiology of Digestion and Metabolism in the Ruminant. Oriel Press Limited, Newcastle. Lee, Y. B. 1972. The Effect of Growth and Environment on the Biochemistry of Porcine Lens and Adipose Tissue. Ph.D. Thesis. University of Wisconsin, Madison. Lee, Y. B. and R. G. Kauffman. 1971. Lipogenic enzyme activities in intramuscular adipose tissue of the pig. J. Animal Sci. 33:1144. (Abstr.). Lee, Y. B. and R. G. Kauffman. 1972. Personal communications. Department of Meat and Animal Science. University of Wisconsin. Madison, Wisconsin. Lee, S. D. and W. F. Williams. 1962. Acetate turnover in the bovine. J. Dairy Sci. 45:893. Lehninger, A. L. 1970. Biochemistry. Worth Publishing Inc., New York. Leng, R. A. 1970. Formation and production of volatile fatty acids in the rumen. In A. T. Phillipson (Ed.) Physiology of Digestion and Metabolism in the Ruminant. Oriel Press Limited, Newcastle. Leng, R. A. and E. F. Annison. 1963. Metabolism of acetate, propionate and butyrate by sheep liver slices. Biochem. J. 86:319. Leng, R. A., J. W. Steel and J. R. Luick. 1967. Contribution of propionate to glucose synthesis in sheep. Biochem. J. 103:785. 170 Leveille, G. A. 1970. Adipose tissue metabolism: influence of periodicity of eating and diet composition. Fed. Proc. 29:1294. Leveille, G. A. and R. W. Hanson. 1966. Adaptive changes in enzyme activity and metabolic pathways in adipose tissue from meal-fed rats. J. Lipid Res. 7:46. Linzell, J. L. 1968. The magnitude and mechanisms of the uptake of milk precursors by the mammary gland. Proc. Nutr. Soc. 27:44. Linzell, J. L., T. B. Mepham, E. F. Annison and C. E. West. 1969. Mammary metabolism in lactating sows: arteriovenous differences of milk precursors and the mammary metabolism of [14C] glucose and [14C] acetate. Brit. J. Nutr. 23:319. Lohr, G. W. and H. D. Waller. 1963. Glucose-6-phosphate dehydrogenase (Zwischenferment). In H. V. Bergmeyer (Ed.) Methods of Enzymatic Analysis. Academic Press, New York. Lowenstein, J. M. 1961. The pathway of hydrogen biosynthesis. J. Biol. Chem. 236:1217. Lowry, O. H., N. O. Rosebrough, A. L. Farr and R. J. Randall. 1951. Protein measurement with the folin phenol reagent. J. Biol. Chem. 193:265. Lynen, F. 1967. The role of biotin-dependent carboxylations in biosyn- thetic reactions. Biochem. J. 102:381. Mackenzie, R. D., T. R. Blohm, E. M. Auxier and A. C. Luther. 1967. Rapid colorimetric micromethod for free fatty acids. J. Lipid Research. 8:589. Manganiello, V. and M. Vaughan. 1972. Selective loss of adipose cell responsiveness to glucagon with growth in the rat. J. Lipid Res. 13:12. Manns, J. G., J. M. Boda and R. F. Willes. 1967. Probable role of propionate and butyrate in control of insulin secretion in sheep. Amer. J. Physiol. 212:756. Martin, B. R. and R. M. Denton. 19703. The intracellular localization of enzymes in white adipose tissue fat cells and the permeability of fat cell mitochondria. Biochem. J. 117:861. Martin, B. R. and R. M. Denton. 1970b. Conversion of pyruvate into citrate by mitochondria isolated from rat epididymal fat cells. Biochem. J. 119:38p. 171 Martin, D. B. and R. R. Vagelos. 1962. The mechanism of tricarboxylic acid cycle regulation of fatty acid synthesis. J. Biol. Chem. 237: 1787. Mason, J. V. and R. F. Sewell. 1967. Influence of diet on the fattyuacid composition of swine tissues. J. Animal Sci. 26:1342. Masoro, E. J. 1961. Biochemical mechanisms related to the homeostatic regulation. J. Lipid Res. 3:149. Masoro, E. J. 1968. Physiological Chemistry of Lipids in Mammals. W. B. Sanders Company, Philadelphia. Masoro, E. J., I. L. Chaikoff and W. G. Dauben. 1949. Lipogenesis from glucose in the normal and liverless animal is studied with 14C- labeled glucose. J. Biol. Chem. 179:1117. Matsuhashi, M., S. Matsuhashi, S. Numa and F. Lynen. 1962. Citrate- dependent carboxylation of acetyl COA in rat liver. Fed. Proc. 21:288. Mayer, S. E., J. C. Khoo, D. Steinberg and L. Jarrett. 1972. Comparison of phosphorylase and lipase activation in adipocytes by epinephrine. Fed. Proc. 31:555. (Abstr.). Mayes, P. A. and J. M. Felts. 1968. The functional status of lipoprotein lipase in rat liver. Biochem. J. 108:483. McBride, 0. W. and E. D. Korn. 1963. The lipoprotein lipase from mammary gland and the correlation of its activity to lactation. J. Lipid Res. 4:17. McCarthy, R. D. 1962. Use of Radioisotopes in Animal Biology and the Medical Sciences. Vol. 2. Academic Press, New York. McClymont, G. L. 1951. Identification of the volatile fatty acids in the peripheral blood and rumen of cattle and the blood of other species. Australian J. Agr. Res. 2:103. Meng, H. C. and R. J.Ho. 1967. Quantitative relationship of some factors affecting fatty acid mobilization and the role of adenosine 3', 5'-monophosphate in the activation of epinephrine sensitive lipase in adipose tissue. Prog. Biochem. Pharm. 3:207. Moody, W. G. and R. G. Cassens. 1968. A quantitative and morphological study of bovine longissimus fat cells. J. Food Sci. 33:47. Moskowitz, J. and J. N. Fain. 1970. Stimulation by growth hormone and dexamethasone of labeled cyclic adenosine 3', 5'-monophosphate accu- mulation by white fat cells. J. Biol. Chem. 245:1101. 172 Moskowitz, M. S. and A. A. Moskowitz. 1965. Lipase: Localization in adipose tissue. Science 149:72. Moss, J. and M. D. Lane. 1971. Biotin-dependent enzymes. In A. Meister (Ed.) Advances in Enzymology. Vol. 35. Interscience Publishers, New York. Nestel, P. J., W. Austin and C. Foxman. 1969. Lipoprotein lipase con- tent and triglyceride fatty acid uptake in adipose tissue of rats of differing body weights. J. Lipid Res. 10:383. Nugteren, D. H. 1963. The conversion of y-linolenic acid into homo-v- linolenic acid. Biochem. J. 89:28P. Numa, S., W. M. Bortz and F. Lynen. 1965. Regulation of fatty acid synthesis at the acetyl COA carboxylation step. In G. Weber (Ed.) Advances in Enzyme Regulation. Vol. 3. Pergamon Press, New York. Numa, S., E. Ringelmann and B. Reidel. 1966. Further evidence for poly- meric structure of liver acetyl COA carb0xylase. Biochem. Biophys. Res. Commun. 24:750. Ochoa, S. 1955. Malic enzyme. In S. P. Colowick and N. 0. Kaplan (Ed.) Methods of Enzymology. Vol. 1. Academic Press, New York. O'Hea, E. K. and G. A. Leveille. 1968. Lipid metabolism in isolated adipose tissue of the domestic pig (Sus domesticus). Compt. Biochem. Physiol. 26:1081. O'Hea, E. K. and G. A. Leveille. 1969a. Influence of feeding frequency on lipogenesis and enzymatic activity in adipose tissue and on the performance of pigs. J. Animal Sci. 28:336. O'Hea, E. K. and G. A. Leveille. 1969b. Significance of adipose tissue and liver sites of fatty acid synthesis in the pig and the efficiency of utilization of various substrates for lipogenesis. J. Nutr. 99:338. O'Hea, E. K. and G. A. Leveille. 1969c. Lipid biosynthesis and trans- port in the domestic chick (Gallus domesticus). Compt. Biochem. Physiol. 30:149. O'Hea, E. K. and G. A. Leveille. 1969d. Influence of fasting and re- feeding on lipogenesis and enzymatic activity of pig adipose tissue. J. Nutr. 99:345. Olivecrona, T. 1962. Metabolism of chylomicrons labeled with 14C-glycerol -H3 palmitic acid in the rat. J. Lipid Res. 3:439. 173 Olson, A. C. and P. Alaupovic. 1966. Rat liver triglyceride synthetase. Biochem. Biophys. Acta. 125:185. Opstvedt, J., R. L. Baldwin and M. Ronning. 1967. Effect of diet upon activities of several enzymes in abdominal adipose and mammary tissues in the lactating dairy cow. J. Dairy Sci. 50:108. Pande, S. V., R. P. Khan and T. A. Venkitasubramanian. 1964. Nicotina- mide adenine dinucleotide phosphate-specific dehydrogenase in rela- tion to lipogenesis. Biochem. Biophys. Acta. 84:239. Patton, R. L. 1970. The reciprocal regulation of lipoprotein lipase activity and hormone sensitive lipase activity in rat adipocytes. J. Biol. Chem. 245:5577. Patton, R. L. and C. H. Hollenberg. 1969. The mechanism of heparin stimulation of rat adipocyte lipoprotein lipase. J. Lipid Res. 10:374. Pav, J. and J. Wenkeova. 1960. Significance of adipose tissue lipopro- tein lipase. Nature 185:926. Pennington, R. J. and T. M. Sutherland. 1956. The metabolism of short- chain fatty acids in the sheep. Biochem. J. 63:618. Perry, W. F. and H. F. Bowen. 1962. Factors affecting the in vitro production of non-esterified fatty acid from adipose tissue. Can. J. Biochem. Physiol. 40:749. Pihl, A. and K. Bloch. 1950. The relative rates of metabolism of neutral fat and phospholipids in various tissues of the rat. J. Biol. Chem. 183:431. Plaut, G. E. 1962. Isocitrate dehydrogenase (TPN-linked) from pig heart (revised procedure). In S. P. Colowick and N. 0. Kaplan (Ed.) Methods of Enzymology. Vol. V. Academic Press, New York. Pokrajac, N., W. J. Lossow and I. L. Chaikoff. 1967. The effect of nutritional state on lipoprotein lipase activity in isolated rat adipose tissue cells. Biochem. Biophys. Acta. 138:123. Polan, C. W., J. J. McNeill and S. B. Tove. 1964. Biohydrogenation of unsaturated fatty acids by rumen bacteria. J. Bact. 88:1056. Raggi, F., E. Hansson, M. G. Simesen, D. S. Kronfield and J. R. Luick. 1961. Pentose cycle activity in various bovine tissues. Res. Vet. Sci. 2:180. Randle, R. J., P. B. Garland, C. N. Hales, E. A. Newsholme, R. M. Denton and C. I. Pogson. 1966. Interactions of metabolism and physiological role of insulin. In G. Pincus (Ed.) Recent Progress in Hormone Research Vol. 22. Academic Press, New York. 174 Reichl, D. 1970. Adipose tissue lipoprotein lipase in rats adapted to controlled feeding schedules. Biochem. J. 119:49P. Reichl, D. 1972. Lipoprotein lipase activity in adipose tissue of rats adapted to controlled feeding schedules. Biochem. J. 128:79. Reid, R. L. 1950. Utilization of acetic and propionic acids in sheep. Nature 165:448. Reid, R. L. 1951. Studies on the carbohydrate metabolism of sheep. III. The blood glucose during insulin hypoglycemia. Australian J. Agr. Res. 2:132. Reid, R. L. 1953. Studies on the carbohydrate metabolism of sheep. Australian J. Agr. Res. 4:213. Renold, A. G. and J. F. Cahill. 1965. Metabolism of isolated adipose tissue (a summary). In Handbook of Physiology. Section 5. Adipose Tissue. Amer. Physiol. Soc. Washington, D. C. Rizack, M. S. 1961. An epinephrine sensitive lipolytic activity in adi- pose tissue. J. Biol. Chem. 236:657. Robinson, D. S. 1960. The effect of changes in nutritional state on the lipolytic activity of rat adipose tissue. J. Lipid Res. 1:332. Robinson, D. S. 1963. The clearing factor lipase and its action in the transport of fatty acids between the blood and the tissues. In R. Paoletti and D. Kritchevsky (Ed.) Advances in Lipid Research. Vol 1. Academic Press, New York. Robinson, D. S. and J. E. French. 1960. Heparin, the clearing factor lipase and fat transport. Pharm. Rev. 12:241. Robinson, D. S. and D. Wing. 1970. Regulation of adipose tissue clearing factor lipase. In B. Jeanrenaud and D. Hepp (Ed.) Adipose Tissue: Regulation and Metabolic Functions. Academic Press, New York. Robinson, G. A., R. W. Butcher and E. W. Sutherland. 1967. Adenyl cyclase as an adrenergic receptor. Ann. N. Y. Acad. Sci. 39:703. Robinson, G. A., R. w. Butcher and E. W. Sutherland. 1968. Cyclic AMP. Annu. Rev. Biochem. 37:149. Rodbell, M. 1964. Localization of lipoprotein lipase in fat cells of rat adipose tissue. J. Biol. Chem. 239:753. Roe, W. E., E. N. Bergman and K. Kon. 1966. Absorption of ketone bodies and other metabolites via the portal blood of sheep. Amer. J. Vet. Res. 27:729. 175 Rognstad, R. and J. Katz. 1966. The balance of pyridine nucleotides in adipose tissue. Proc. Nat. Acad. Sci. U.S.A. 55:1148. Rous Simonne. 1971. The origin of hydrogen in fatty acid synthesis. In R. Paoletti and D. Kritchevsky (Ed.) Advances in Lipid Research. Vol. 9. Academic Press, New York. Rudman, D. 1965. The mobilization of fatty acids from adipose tissue by pituitary peptides and catecholamines. Ann. N. Y. Acad. Sci. 131:102. Rudman, D. and P. W. Shank. 1966. Comparison of the responsiveness of perirenal adipose tissue of the rat, hamster, guinea pig and rabbit to the antilipolytic action of insulin. Endocrinol. 79:565. Saggerson, E. D. and A. L. Greenbaum. 1970. The regulation of trigly- ceride synthesis and fatty acid synthesis in rat epididymal adipose tissue. Biochem. J. 119:193. Salaman, M. R. and D. S. Robinson. 1966. Clearing factor lipase and adipose tissue. A medium in which the enzyme activity of tissue from starved rats increases in vitro. Biochem. J. 99:640. Salans, L. B., J. L. Knittle and J. Hirsch. 1968. The role of adipose cell size and adipose tissue insulin sensitivity in the carbohydrate intolerance of human obesity. J. Clin. Invest. 47:153. Salans, L. B., E. S. Horton and E. H. Sims. 1971. Experimental obesity in man: Cellular character of adipose tissue. J. Clin. Invest. 50:1005. Satter, L. D., J. W. Suttie and B. R. Baumgardt. 1964. Dietar induced changes in volatile fatty acid formation from¢y-cellulose- 4C and hemicellulose-IAC. J. Dairy Sci. 47:1365. Satter, L. D., J. W. Suttie and B. R. Baumgardt. 1967. Influence of accompanying substrate on 1g vitro volatile fatty acid formation from a-cellulose-lac. J. Dairy Sci. 50:1626. Schoefl, G. I. and J. E. French. 1968. Vascular permeability to parti- culate fat: Morphological observations on vessels of lactating mammary gland and of lung. Proc. Roy. Soc. Ser. B. 169:153. Schotz, M. C. and A. S. Garfinkel. 1972. Effect of nutrition on species of lipoprotein lipase. Biochem. Biophys. Acta. 270:472. Schwartz, J. P. and R. L. Jungas. 1971. Studies of the hormone sensitive lipase of adipose tissue. J. Lipid Res. 12:553. 176 Scow, R. 0., M. Hamosh, E. J. Blanchette-Mackie and A. J. Evans. 1972. Uptake of blood triglycerides by various tissues. Lipids 7:497. Shaw, J. C. and W. L. Ensor. 1959. Effect of feeding cod liver oil and unsaturated fatty acids on rumen volatile fatty acids and milk fat content. J. Dairy Sci. 42:1238. Sidhug K. S., R. S. Emery, R. A. Merkel and A. F. Parr. 1972. Relation of lipolysis to fatness in lambs. J. Anim. Sci. 35:207. (Abstr.). Smith, U. 1971a. Effect of cell size on lipid synthesis by human adipose tissue in vitro. J. Lipid Res. 12:65. Smith, R. M. 1971b. Interactions of acetate, propionate and butyrate in sheep liver mitochondria. Biochem. J. 124:877. Snedecor, G. W. and W. G. Cochran. 1969. Statistical Methods. Sixth Edition. The Iowa State University Press, Ames. Spencer, A. F. and J. M. Lowenstein. 1962. The supply of precursors for the synthesis of fatty acids. J. Biol. Chem. 237:3640. Spincer, J., J. F. Rook and K. G. Towers. 1969. The uptake of plasma constituents by the .mammary gland of the sow. Biochem. J. 111:727. Srere, P. A. 1959. The citrate cleavage enzyme. J. Biol. Chem. 234:2544. Srere, P. A. 1965. The molecular physiology of citrate. Nature 205:766. Srere, P. A. and A. Bhaduri. 1962. Incorporation of radioactive citrate into fatty acids. Biochem. Biophys. Acta. 59:487. Srere, P. A. and D. W. Foster. 1967. On the proposed relation of citrate enzymes to fatty acid synthesis and ketosis in starvation. Biochem. Biophys. Res. Commun. 26:556. Steinberg, D. 1966. Catecholamine stimulation of fat mobilization and its metabolic consequences. Pharm. Rev. 18:217. Tepperman, H. M. and J. Tepperman. 1958. The hexosemonophosphate shunt and adaptive hyperlipogenesis. Diabetes 7:478. Tepperman, H. M. and J. Tepperman. 1963. On the response of hepatic- glucose-é-phosphate dehydrogenase activity to changes in diet com- position and food intake pattern. In G. Weber (Ed.) Advances in Enzyme Regulation. vol. 1. Pergamon Press, New York. Therriault, D. G., R. W. Hubbard and D. B. Mellin. 1969. Endocrine control of fat mobilization in the isolated fat cells of cold-exposed rats. Lipids 4:413. L77 Thorton, J. H., D. C. Beitz and A. D. McGilliard. 1972. Lipolysis in bovine adipose tissue. J. Animal Sci. 35:208. Tove, S. B. 1965. Fat metabolism in ruminants. In R. W. Daugherty (Ed.) Physiology of Digestion in the Ruminant. Butterworth, Washington. Tove, S. B. and F. H. Smith. 1960. Changes in the fatty acid composition of the depot of mice induced by feeding oleate and linoleate. J. Nutr. 71:264. Trenkle, A. 1970. Effects of short-chain fatty acids, feeding, fasting and type of diet on plasma insulin levels in sheep. J. Nutr. 100:1323. Tubbs, P. K. and P. B. Garland. 1963. Fatty acyl thioesters of coenzyme A: Inhibition of fatty acid synthesis in liver of normal, fasted and fat or sugar fed rats. Biochem. J. 89:25. Turner, C. D. and J. T. Bagnara. 1971. Mechanisms of action of islet hormones. In General Endocrinology. W. B. Sanders Company, Phila- delphia. Vagelos, P. R. 1964. Lipid Metabolism. Annu. Rev. Biochem. 33:139. Wadsworth, J. C. 1968. Fatty acid composition of lipid in the thoracic duct lymph of grazing cows. J. Dairy Sci. 51:876. Waite, M. 1962. The carboxylation of acetyl COA by acetyl carb0xylase. Fed. Proc. 21:287. Wakil, S. J. 1961. Mechanism of fatty acid synthesis. J. Lipid Res. 2:1. Wakil, S. J., J. W. Porter and D. M. Gibson. 1957. Studies on the mech- anisms of fatty acid synthesis. Biochem. Biophys Acta 24:453. Warner, A. I. 1964. Production of volatile fatty acids in the rumen. Nutr. Abstr. Rev. 34:339. Watson, J. A. and J. M. Lowenstein. 1970. Citrate and the conversion of carbohydrate into fat. J. Biol. Chem. 245:5993. Weigand, E., J. W. Young and A. D. McGilliard. 1972. Extent of propionate metabolism during absorption from the bovine ruminoreticulum. Biochem. J. 126:201. West, C. E., R. Bickerstaffe, E. F. Annison and J. L. Linzell. 1972. Studies on the mode of uptake of blood triglycerides by the mammary gland of the lactating goat. Biochem. J. 126:477. ,1-78 Wing, D. R., M. R. Salaman and D. S. Robinson. 1966. Clearing factor lipase in adipose tissue. Biochem. J. 99:648. Wise, E. M. and E. G. Ball. 1964. Malic enzyme and lipogenesis. Proc. Nat. Acad. Sci. U.S.A. 52:1255. Wood, R. D., M. C. Bell, R. B. Grainger and R. A. Teekell. 1963. Metabo- lism of labeled linoleic-l-14C acid in sheep rumen. J. Nutr. 79:62. Wright, D. E. 1960. Hydrogenation of chloroplast lipids by rumen bacteria. Nature 185:546. Young, J. W., E. Shrago and H. A. Lardy. 1964. Metabolic control of enzymes involved in lipogenesis and g1uconeogenesis. Biochem. 3:1687. Young, J. W., S. L. Thorp and H. 2. de Lumen. 1969. Activity of selected gluconeogenic and lipogenic enzymes in bovine rumen mucosa, liver and adipose tissue. Biochem. J. 114:83. Zinder, O. and B. Shapiro. 1971. Effect of cell size on epinephrine and ACTH-induced fatty acid release from isolated fat cells. J. Lipid Res. 12:91. Zinder, 0., R. Arad and B. Shapiro. 1967. Effect of cell size on the metabolism of isolated fat cells. Israel J. Med. Sci. 3:787. APPENDIX 179 APPENDIX I. Ewe Grain Rations Table A. Composition of Grain Mixture for Bred Ewes Ingredients Percent Oats 55.00 Corn 13.75 Wheat 20.00 Wheat bran 10.00 Wet molasses 1.00 Aurofac .25 Total 100.00 Table B. Composition of Grain Mixture for Lactating Ewes Ingredients Percent Oats 53.00 Corn 25.00 Wheat bran 15.00 Soybean oil meal 5.00 Wet molasses 2.00 Total 100.00 180 APPENDIX 11. Raw Dataa a Values expressed nanomoles NADPH produced/min./mg protein b 1 Suffolk breed type, 2 = Southdown breed type c l = Ram; 2 = Ewe; 3 - Wether d l = Birth; 2 = 24 hour; 3 - 8 week; 4 = 16 weeks; 5 = 32 weeks e 1 = Liver; 2 = Longissimus muscle; 3 = Perirenal fat; 4 = Sub- cutaneous fat; 5 = Blood plasma G-6 PDH = glucose-6-phosphate dehydrogenase 6-PGDH = 6-phosphogluconate dehydrogenase NADP-ME = NADP-malate dehydrogenase ICDH-CYTO Cytoplasmic NADP-isocitrate dehydrogenase ICDH-MITO mitochondrial NADP=isocitrate dehydrogenase 181 mom.mNH «mm.Nm mm¢.wH qu.oH ccc.c N m H H HH me.¢q me.Nw Ho¢.a oem.mn NwN.¢ H m H H HH HNm.Nm me.o¢ ONo.mm mNo.om oom.o e m N N oH HHH.qo aNo.mm omH.mN mNN.no mN¢.qH m m N N oH NoN.ooH meo.me mNH.NN «mm.mH Nom.H N m N N oH mNe.No mq¢.mm omH.NH o¢¢.mc mmN.q H m N N oH mNe.mm m¢a.mo ooN.Nm NN¢.om oqo.mN a m N N m oom.o¢ Neo.HNH mam.m¢ mmm.¢m coo.H¢ m m N N m HHH.¢o qu.Nm moN.oH qu.¢H ¢m0.N N m N N m mmm.mN Ham.ma Ham.HH HmN.mm mNm.H H m N N m oao.me mmn.¢o mNN.¢N an.Hm mNm.o¢ e m H H m «mo.wq oH¢.mmH qu.mH «oN.oq Noa.¢H m m H H w Noq.N¢H me¢.mq mmo.aH mmN.mH ow¢.o N m H H w omo.qm moN.oHH HNo.0N ema.o¢ aao.q H m H H m mmo.we «mo.Ne «wo.mN HqN.N¢ mHN.mm e m m H N mmN.aoH mNH.ooH qu.mm mmm.mm Nw¢.NN m m m H N NNN.HoH me.N¢ Noo.HN q¢m.mH omn.o N m m H N woo.o¢ oHe.mw mom.N mHo.NN moN.m H m m H N omN.mN H¢9.HN Hom.H¢ mNH.oH «NH.Nq e m N H o moo.Ho aem.NmH mNa.mm mom.m¢ HHa.om m m N H o www.mo ooe.mm wNm.mN Nom.oH on.H N m N H o NNH.wm oqm.Hm mNm.NH mmm.mm qNN.m H m N H o Nnm.¢oH HoN.NN www.mN N¢¢.Hm ccc.c N N N N e wNm.Nm Noq.Nm qu.oH NHN.mn oHN.oH H N N N e eoH.Nm mo¢.mmm mH¢.NN HHN.mN moH.oq m N N H m wNm.mmH qu.Nm ONo.¢N omo.om 000.0 N N N H m mmN.¢¢ m¢m.¢o Nmo.mH mHm.Nn ONN.m H N N H m on.omH mqm.NN NNo.wm coo.mm mNm.H N N N H N mqm.¢w Hom.oN mNm.mH HNm.qm oqo.m H N N H N NNN.mq on.moq omo.HH NHH.N¢ m¢m.mH m N N H H Hm¢.m¢H me.om mNm.Nm eNN.om ONo.N N N N H H Nmo.qq ooH.Nm Hom.N¢ «om.Nm NNm.oH H N N H H OHqumooH oawoumaoH mzumaHH . NcoNumeuuoo quEHm .>H anzmmm< 197 NO0.0n mao.o om~.o o¢m.o .mm.0 NHN.O owo.0u onN.o qmn.o NH0.0u NN0.0 «no.0 wom.o .HH.o 000.0 uo~.o mmN.o mHo.0u NNH.O «00.0 nmm.o mom.o mHo.o m¢o.o 00H.Ou oHo.o mNo.0n muo.0n ¢m0.0u moo.0a Hmm.o N.0.0 «oo.o ONH.O ONH.O aNH.o Nom.o oom.0 om~.o HNH.O: o~m.o m~¢.o .ON.O NMH.O ONH.O wmo.o o~o.o- «00.0: qmo.o- ONH.O: Hmuoa-cNN- anzmmm< 198 .N. O NNN.O OO0.0- ..0.0 ONN.O NON.O NNN.O OON.O .N..O ON0.0- NNO.O NNH.O- HNOON-ONN- xHQZMmm< 199 000.0 wm~.o: mmm.o mmm.o .m¢.o .wN.o OO0.0 w~o.o mom.o mao.o «mm.o ¢o¢.o HmuOHuchma<00 u< .ON.O- NNN.O NNN.O NON.O NHN.O OOO O ON0.0 OON.O NNN.O OON.O NON.O Hmuoa-NOOH N.N.O- ..H.O- .O0.0- NNH.O- OO0.0 .NN.O- .HN.O- NOH.O- NOH.O- ONH.O- OooHN- xHszmm<