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Plasticy and control of mitochondrial metabolism in fish muscle Moyes, Christopher Douglas 1991

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PLASTICITY AND CONTROL OF MITOCHONDRIAL METABOLISM IN FISH MUSCLE By CHRISTOPHER DOUGLAS MOYES B.Sc, Zoology, University of Guelph, 1983 M.Sc, Biology, University of Ottawa, 1985 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE STUDIES (Department of Zoology) We accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH COLUMBIA June 1991 (5) Christopher Douglas Moyes In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholarly purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission. Department The University of British Columbia Vancouver, Canada DE-6 (2/88) ii ABSTRACT Problems associated with fuel preference, metabolic capacity and regulation were addressed using isolated mitochondria and enzyme analyses of cardiac and skeletal muscles of spiny dogfish (Squalus acanthias), common carp (Cyprinus carpio), skipjack tuna (Katsuwonus pelamis) and rainbow trout (Oncorhynchus my kiss). Chondrichthian muscle is unusual because it possesses no detectible activities of carnitine palmitoyl transferase (CPT), which is involved in transport of fatty acids into mitochondria, yet demonstrates activity of B-hydroxyacyl CoA dehydrogenase, a fatty acid B-oxidation enzyme. This paradox was addressed by searching for alternate pathways for mitochondrial fatty acid utilization possibly involving peroxisomal fatty acid processing. Dogfish red muscle and ventricle mitochondria did not utilize long, medium or short chain free fatty acids or fatty acyl carnitines under a variety of conditions. These substrates include the putative products of peroxisomal B-oxidation, which was not detectible in muscles of 2 chondrichthians. It was concluded that dogfish muscles do not utilize fatty acids by direct or indirect muscle pathways, but may rely on ketone bodies as "lipid" fuel. Carp and tuna red and white skeletal muscle, compact and spongy myocardium and atrium were compared to examine the importance of mitochondrial differences in determining tissue aerobic capacity. The spectrum of tissues examined possess a 26-fold difference in mitochondrial content, as indicated by citrate synthase activity. Higher aerobic capacities (tuna vs. carp) are attributed to a combination of three factors: i. greater recruitable muscle mass/kg body mass, ii. greater mitochondrial volume density/g tissue iii. increased mitochondrial specific activity. The relative importance of each factor varied between tissues. In general, the more aerobic tissues (ventricle, red muscle) differ primarily in recruitable mass/kg body mass. There was less than a 2-fold difference in mitochondrial content/g or mitochondrial specific activity between tuna and carp tissues. In less aerobic tissues (white muscle, atrium), a several fold greater mitochondrial content in tuna contributed to greater aerobic capacity. Coupled with the greater aerobic capacities of tuna muscles, was a shift in fuel preference toward fatty acid utilization, as indicated iii by CPT/mg mitochondrial protein and CPT/hexokinase ratios. Tuna ventricle mitochondria may operate close to their maximal in vitro rates (State 3) in situ at cardiac V02MAX. In contrast, trout white muscle mitochondria in vivo operate at a small fraction of their in vitro maximum. The maximal aerobic demands of white muscle probably occur during recovery from burst exercise, when a high lactate load is converted to glycogen in situ: This added cost of recovery represents an ATP demand that is only 3.5% of the maximal mitochondrial capacity. This capacity is suppressed in vivo by highly inhibitory ATP/ADP ratios and limiting phosphate. The primary signal for increased ATP synthesis associated with recovery is not ATP/ADP but probably phosphate, elevated due to phosphocreatine hydrolysis and adenylate catabolism in the purine nucleotide cycle. At low ADP availability and sub-optimal phosphate (<5mM), acidosis enhances respiration. At high respiratory rates mitochondrial pyruvate oxidation is sensitive to pyruvate concentration over the physiological range (apparent Km= 35-40 uM). The sensitivity is lost at the low rates that approximate in vivo respiration. Changes in lactate do not affect the kinetics of pyruvate oxidation. Fatty acid oxidation may spare pyruvate and lactate for use in glyconeogenesis, primarily through allosteric inhibition of pyruvate dehydrogenase, rather than covalent modification. iv T A B L E O F C O N T E N T S A B S T R A C T ii LIST O F T A B L E S viii LIST O F F I G U R E S ix LIST O F A B B R E V I A T I O N S x A C K N O W L E D G E M E N T S xi C H A P T E R 1: A C O M P A R I S O N O F F U E L P R E F E R E N C E S O F M I T O C H O N D R I A F R O M V E R T E B R A T E S A N D I N V E R T E B R A T E S 1 P R E F A C E 1 INTRODUCTION 1 T E R M I N O L O G Y 2 F A T T Y ACIDS 3 Catabolic pathway 3 Assay 3 Carnitine-dependent vs. carnitine-independent pathway 5 Chain length preference 6 Comparative aspects of fatty acid oxidation 7 K E T O N E BODIES 10 Catabolic pathway and assay conditions 10 Comparative aspects of ketone body catabolism 11 Ketone body vs. fatty acid or metabolic fuels 12 C A R B O H Y D R A T E S 13 Catabolic pathway 13 Comparative aspects of pyruvate oxidation 15 A M I N O ACIDS 17 V Catabolic pathway I7 Comparative aspects of amino acid oxidation 19 Amino acid metabolism in osmoconformers . 20 C H A P T E R 2: M I T O C H O N D R I A L A N D P E R O X I S O M A L B-OXIDATION IN E L A S M O B R A N C H S 23 P R E F A C E 23 INTRODUCTION 23 M A T E R I A L S A N D METHODS 24 Animals 24 Preparation of tissues for enzyme assays 25 Enzyme assays 26 Mitochondrial studies on dogfish red muscle and heart 27 RESULTS 28 CPT and HO AD 28 Mitochondrial studies 28 Fatty acids 32 Fatty acylcarnitines 32 Carbohydrate and ketone bodies 32 Peroxisomal fi-oxidation 33 DISCUSSION 33 C H A P T E R 3: D O F A S T FISH H A V E F A S T M U S C L E M I T O C H O N D R I A ? 39 P R E F A C E 39 INTRODUCTION 39 M A T E R I A L S A N D METHODS 41 Animals 41 vi Mitochondrial isolation 41 Mitochondrial oxygen consumption 42 Enzyme assays 42 RESULTS 43 Mitochondrial oxidation 43 Tissue enzymes 45 Mitochondrial enzymes 48 DISCUSSION 49 Aerobic capacity 49 Fuel preference 53 CHAPTER 4:RECOVERY METABOLISM IN TROUT WHITE MUSCLE: THE ROLE OF THE MITOCHONDRIA 57 INTRODUCTION 57 M A T E R I A L S A N D METHODS 59 Animals 59 Exercise protocol 59 Enzyme activities 59 Isolated mitochondrial studies 60 Mitochondrial oxygen consumption 61 Mitochondrial pyruvate oxidation 62 Statistics 63 RESULTS 64 Mitochondrial pyruvate oxidation i:kinetics 63 Mitochondrial pyruvate oxidation ii.effects of fatty acid oxidation 66 PDH activity during recovery 69 vii PDH in vitro vs in vivo 69 Mitochondrial oxygen consumption 69 Effects of ATP/ADP ratio 73 Effects of phosphate 76 DISCUSSION 80 Carbon metabolism 80 Metabolic rate 82 G E N E R A L DISCUSSION 88 L I T E R A T U R E C I T E D 91 vi i i LIST O F T A B L E S 1. Maximal activities of the mitochondrial enzymes B-hydroxyacyl CoA dehydrogenase (HOAD) and carnitine palmitoyl transferase (CPT) 29 2. Mitochondrial oxidation of fatty acids, ketone bodies and pyruvate by mitochondria from dogfish red muscle 30 3. Mitochondrial oxidation of fatty acids, ketone bodies and pyruvate by mitochondria from dogfish heart 31 4. Maximal activities of the peroxisomal enzymes catalase and peroxisomal fl-oxidation (PBO) . . 34 5. Oxidation of various substrates by mitochondria from heart and skeletal muscle of carp and tuna 44 6. Tissue activities of citrate synthase (CS), carnitine palmitoyl transferase (CPT), and hexokinase (HK) in tuna and carp 46 7. Mitochondrial enzymes assayed in isolated mitochondria of carp and tuna 47 8. Tuna muscle mitochondrial ultrastructure 51 9. Effect of palmitoyl carnitine on P D H activation state in isolated trout white muscle mitochondria 68 10. Oxidation of physiological fuels by trout white muscle mitochondria (15°C) 71 11. Activities of citrate synthase and pyruvate dehydrogenase in trout white muscle and isolated mitochondria at 15°C and 25°C 72 12. M g 2 + effects on respiratory properties of isolated trout white muscle mitochondria 75 13. Influence of pH on the respiratory rates of isolated trout white muscle mitochondria 79 ix LIST OF FIGURES 1. Carnitine-dependent and carnitine-independent fatty acid oxidation by mitochondria 4 2. Relationship between mitochondrial content (CS/g) and CPT activity/mg mitochondrial protein 56 3. Kinetics of pyruvate oxidation by trout white muscle mitochondria 64 4. Kinetics of carp red muscle mitochondrial pyruvate oxidation 65 5. Effects of 50 u M palmitoyl carnitine on pyruvate dehydrogenase flux in trout white muscle mitochondria 67 6. Pyruvate dehydrogenase (PDH) activation state in white muscle of burst exercised trout 70 7. Influence of A T P / A D P ratios on trout white muscle mitochondrial respiration 74 8. Effects of phosphate concentration on mitochondrial respiration rate and A D P / O ratio 77 9. Influence of pH on phosphate dependency of mitochondrial respiration 78 X ABBREVIATIONS A M P adenosine monophosphate A D P adenosine diphosphate A T P adenosine triphosphate B C A A branched chain a-amino acids B C K A branched chain a-keto acids BHB 6-hydroxybutyrate B H B D H 8-hydroxybutyrate dehydrogenase BSA bovine serum albumin C A T carnitine acetyl transferase CoA coenzyme A CPT carnitine palmitoyl transferase CS citrate synthase F A D (FADHj) oxidized (reduced) flavin adenine dinucleotide G D H glutamate dehydrogenase HEPES N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid H K hexokinase H O A D 6-hydroxyacyl CoA dehydrogenase H P L C high performance liquid chromatography L C F A long chain fatty acids (C16-C22) M C F A medium chain fatty acids (C8-C14) N A D + (NADH) oxidized (reduced) nicotinamide adenine dinucleotide N A D P + oxidized nicotinamide adenine dinucleotide phosphate PBO peroxisomal 8-oxidation P D H pyruvate dehydrogenase PDHa active pyruvate dehydrogenase PEPCK phosphoenolpyruvate carboxykinase RCR respiratory control ratio SCFA short chain fatty acids (<C8) T C A tricarboxylic acid V v(m,f) volume/volume of mitochondria/fiber V v(ms,f) as above, for subsarcolemmal mitochondria Vv(mt,f) as above, for total mitochondria Sv(im,m) surface area/volume of inner membrane/mitochondria Sv(im,ms) as above, for subsarcolemmal mitochondrondria Sv(im,mi) as above, for interfibrillar mitochondria [X] concentration of metabolite X xi A C K N O W L E D G E M E N T S I would like to thank my supervisor, Peter Hochachka, for his encouragement, enthusiasm, imagination and vats of money. Discussions and interactions with friends and colleagues, particularly Raul Suarez, Claudia Kasserra, Tim West, Trish Schulte, Les Buck and Peter Arthur, were constant sources of intellectual challenge, and occasionally confusion and infuriation. They may decide amongst themselves who was most responsible for which. Financial support was provided by a Natural Sciences and Engineering Research Council Post-Graduate Fellowship, U B C Graduate Fellowship, Dominos Pizza and Bank of Mom and Dad. 1 C H A P T E R 1: A C O M P A R I S O N O F F U E L P R E F E R E N C E S O F M I T O C H O N D R I A F R O M V E R T E B R A T E S A N D I N V E R T E B R A T E S P R E F A C E The chapter is excerpted from a review published by Moyes, R.K Suarez, J.S.Ballantyne and P.W.Hochachka (Can.J.Zool. 68:1337-1349,1990). The text in this thesis chapter was written by myself, with critical reviews by the other authors gratefully acknowledged. Sections of the published review written by my co-authors are not included in this thesis. I N T R O D U C T I O N Cellular energy production via the oxidation of glucose, fatty acids, ketone bodies and amino acids requires participation of mitochondrial transporters, mitochondrial matrix and membrane-bound enzymes and oxidative phosphorylation. Consequently, studies of metabolite oxidation by isolated mitochondria provide insights into the fuel preferences of various tissues. Because studies which make use of isolated mitochondria minimize disruption of enzyme-enzyme, enzyme-membrane and enzyme-transporter interactions, they are useful in assessing transport capacities, metabolic flux rates, enzyme activities in situ and regulation of mitochondrial pathways. Such studies may also yield important information about anabolic capacities since mitochondria participate in biosynthetic processes such as gluconeogenesis and fatty acid synthesis. Fundamental differences are evident when comparing the mitochondrial properties of various tissues of an organism or homologous tissues of different organisms. These variations may reflect differences in metabolic organization of the tissue. A greater awareness of the spectrum of mitochondrial adaptations employed by animals may provide models which are better suited for the study of various aspects of mitochondrial metabolism. In this review I discuss the conditions under which mitochondrial substrate oxidation rates are determined, the classes of substrates utilized, and the metabolic significance of various patterns of substrate preference observed. The properties of mitochondria from homologous vertebrate and invertebrate tissues are compared to determine conserved and malleable components of mitochondrial 2 oxidative metabolism. I have restricted the comparisons to properties of mitochondria from liver, skeletal muscle and heart, tissues for which comparable studies are available for non-mammalian vertebrates and invertebrates. TERMINOLOGY The terminology for mitochondrial respiratory states is that of Chance and Williams (1956). Mitochondria are in State 3 when given saturating amounts of A D P , in the presence of oxidizable carbon substrates and phosphate. When translated to the whole tissue level, the mitochondrial State 3 rate typically corresponds very closely to the maximal rate of oxygen consumption by the whole tissue, athough they tend to underestimate whole tissue rates (Schwerzmann et al. 1989, Suarez et al. 1990, Suarez and Moyes submitted). Several of the studies in this thesis support this observation, but a markedly different situation occurs with trout white muscle, where mitochondrial State 3 is probably never reached in vivo (see Chapter 4). Once the A D P is phosphorylated to A T P , oxygen consumption decreases to a low rate (State 4). Mitochondrial preparations are considered coupled, (i.e., respiration or the consumption of oxygen is dependent upon phosphorylation of ADP to ATP) when the ratio of the State 3/State 4 (= respiratory control ratio or RCR) is in excess of 4-6 when tested using a preferred substrate (Chappell and Hansford 1972). Many of the interspecific and intertissue differences observed in RCR can be attributed to variations in methodology; however, inherent loose coupling in vivo, as reflected by low RCR in vitro, has been suggested as a mechanism for generation of heat. This is exemplified by brown adipose tissue, which possesses mitochondria which are uncoupled by a specific protein, a process under hormonal control (Nicholls 1979). Inherent loose coupling for heat production has been proposed for skeletal muscle of mammals (mice, Skulachev et al. 1963; seals, Grav and Blix 1979) and birds (Barre et al. 1989). Akhmerov (1986) has suggested that, in general, homeotherm mitochondria are more loosely coupled than poikilotherm mitochondria as a mechanism to generate heat. 3 F A T T Y ACIDS Catabolic pathway Although there are several pathways in various intracellular compartments involved in breakdown of fatty acids (a-oxidation, mitochondrial and peroxisomal 8-oxidation, omega-oxidation, see Bremer and Omundsen 1984), the most important pathway in production of energy is mitochondrial fi-oxidation. Mitochondrial fatty acyl CoA enters the 8-oxidation spiral resulting in production of reducing equivalents ( F A D H 2 , NADH) and acetyl CoA. Two general pathways, differing in the intracellular location of the fatty acyl CoA synthase step, exist for production of mitochondrial fatty acyl CoA from cytoplasmic fatty acids (Figure 1). If the appropriate fatty acyl CoA synthase occurs in the matrix, fatty acids can be activated direcdy inside the mitochondrion. If the mitochondrial matrix lacks the capacity for fatty acyl CoA synthesis, fatty acids are activated extramitochondrially. As fatty acyl CoA does not penetrate the inner mitochondrial membrane, a carnitine shuttle is neccessary for the transfer. The relative importance of the carnitine-dependent and carnitine-independent pathways is highly dependent on fatty acid chain length, and varies between tissues and species. Assay Fatty acids are a group of substrates with a wide range of chain lengths oxidized by mitochondria by 2 general pathways involving several chain length-specific families of enzymes. Despite this, the fatty acid oxidizing capacity of mitochondria is usually estimated from the rate of oxidation of optimal concentrations of palmitoyl carnitine (0.01-0.1 mM) or palmitoyl CoA (0.01-0.1 mM) + L-carnitine (1-5 mM). Many of the problems associated with the use of acyl CoA (binding to soluble and membrane proteins, detergent effects, micelle formation, substrate for 8-oxidation by contaminating peroxisomes) can be reduced by using palmitoyl carnitine instead of palmitoyl CoA (+carnitine). However, oxidation of fatty acyl carnitine bypasses the regulatory influence of the outer carnitine palmitoyl transferase (CPT) (Bremer 1983). The capacity of mitochondria to utilize fatty acids should be determined with fatty acids and fatty acyl carnitines of various chain lengths. 4 Figure 1. Carnitine-dependent and carnitine independent fatty acid oxidation by mitochondria. The upper pathway is the carnitine-dependent route. The fatty acid is activated into its CoA ester by fatty acylCoA synthetase located on the outer membrane (1). The outer carnitine palmitoyl transferase (CPT) (3) catalyzes production of fatty acyl carnitine. It has been generally accepted that the outer CPT is located on the outer side of the inner mitochondrial membrane (solid line), but more recent evidence suggests it is present on the inner side of the outer mitochondria] membrane (broken line, Murthy and Pande 1987). After transport into the mitochondrion by acylcarnitine-carnitine exchanger (4), the inner CPT (5) regenerates the acyl Co A , which enters the B-oxidation pathway. The lower pathway is the carnitine-independent route. Fatty acids enter the matrix where they are activated into acylCoA esters by fatty acylCoA synthetase (2). (Modified from Bremer and Osmundsen 1984). 5 Carnitine-dependent vs. carnitine-independent pathway As fatty acid oxidation by the carnitine-dependent pathway is primarily regulated at CPT, use of the carnitine-independent pathway may alter the interaction between cytoplasmic parameters and mitochondrial fatty acid oxidation (e.g. malonyl CoA, carnitine, CoA, phosphorylating hormones; Bremer 1983, Harano et al. 1985, Lund 1987, Murthy and Pande 1987). Furthermore, each pathway for fatty acyl CoA synthesis draws on different intracellular pools of A T P and CoA and it is difficult to ascribe regulatory significance to kinetic differences between the enzymes in relation to these separate pools. Several environmental factors have the potential to perturb either fatty acid oxidation or the balance between the carnitine-dependent and carnitine-independent pathways. Fatty acid oxidation is strongly affected by assay medium composition, especially ionic strength. Oxidation of palmitoyl CoA, but not palmitoyl carnitine, is markedly enhanced by elevating assay medium ionic strength to physiological levels (Brosnan and Fritz 1971), possibly due to the sensitivity of CPT to this parameter (Saggerson 1982). This sensitivity may have physiological relevance in tissues of animals which experience environmentally-induced changes in ionic strength. In mitochondria from various osmoconformers, fatty acyl carnitine oxidation is enhanced by osmotic swelling, possibly due to changes in matrix ionic strength (Ballantyne and Moon 1986, Ballantyne and Moyes 1987a). Freeze-tolerant tissues would also be expected to experience changes in intracellular ionic strength associated with removal of free water (Liu 1979); fatty acid oxidation by mitochondria from freeze tolerant insects also demonstrates sensitivity to ionic strength (Ballantyne and Storey 1985). Urea and methylamines, important solutes in elasmobranch tissues, also affect mitochondrial fatty acid oxidation (Ballantyne and Moon 1986) and may be the reason extrahepatic tissues rely less on lipid than other substrates (Ballantyne et al. 1987, Ballantyne and Chamberlin 1988). Temperature may also influence the balance between the carnitine-dependent and -independent pathways. For example, the capacity of lake charr (Salvelinus namaycush) liver mitochondria to utilize fatty acids increases, relative to fatty acyl carnitines, with decreasing assay temperature (Ballantyne et al. 1989). This may be related to the very high enthalpy of the carnitine-acyl 6 carnitine exchanger (Ramsay and Tubbs 1976) (Q 1 0 = 8.5) which suggests the rate of this reaction would be markedly decreased at lower temperature. Chain length preference Fatty acid chain length is one variable often overlooked in studies assessing the fatty acid oxidizing capacity of mitochondria. Diet-dependent differences in fatty acid chain length profile may be expected to lead to differences in mitochondrial properties. The chain length of fatty acids available to mitochondria varies between tissues and species. In monogastric animals, liver processes the mixed chain length fatty acids obtained in the diet, but L C F A are the predominant chain lengths available to the extrahepatic tissues due to chain length-specific differences in digestive physiology (site of absorption) and differential utilization by liver (see Frost and Wells 1981). Because of the product specificity of fatty acid synthase (see Wakil et al. 1983), palmitate and oleate are "typical" substrates in species where fatty acids synthesized de novo are an important contribution to lipid stores (e.g., hummingbirds) . In ruminants, SCFA produced in bacterial fermentations are a major carbon source. M C F A are the predominant fatty acids in the milk diet of many mammalian neonates (elephant, rat, rabbit; Jenness 1974). Polyunsaturated L C F A are important in the diet of marine species such as seals, seabirds and fish (Sargent et al. 1976). As families of enzymes are responsible for transport (CAT, CPT), activation (short, medium, long, very long chain fatty acyl CoA synthases), and oxidation (short, medium and long chain fatty acyl CoA dehydrogenases) of fatty acids (see Bremer and Osmundsen 1984), interspecific differences in chain length specificity are not surprising. Chain length-specific differences in the balance between the carnitine-dependent and -independent pathways are clear from mammalian studies (see Groot et al. 1976). This complexity makes it difficult to determine whether species-specific differences in fatty acyl carnitine chain length preference (e.g. Osmundsen and Bremer 1978) are due to the properties of individual enzymes, or the relative activities of a member of the family of enzymes catalyzing a single step. An additional consideration is the influence of intracellular fatty acyl-binding protein in metabolism of fatty acids of different chain lengths. The fate of L C F A may be controlled by concentration-dependent 7 interactions with fatty acyl-binding protein but M C F A do not bind (Wu-Rideout et al. 1976). Another factor which may influence the mitochondrial metabolism of fatty acids of different chain lengths is peroxisomal B-oxidation. This pathway shortens L C F A , producing M C F A for oxidation by mitochondria (Lazarow and De Duve 1976). While its quantitative importance is controversial (Mannaerts et al. 1979, Kondrup and Lazarow 1985, Christensen et al. 1986), it is thought to be important in processing fatty acids which are poor substrates for mitochondria (Osmundsen et al. 1987). Increasing the dietary content of these fatty acids increases the capacities for both peroxisomal and mitochondrial B-oxidation in rat liver, but the peroxisomal response is more pronounced (Berge et al. 1987). The peroxisomal response persists as long as the fatty acids remain in the diet (Thomassen et al. 1985). Increasing the degree of unsaturation of L C F A decreases mitochondrial oxidation (Osmundsen and Bjornstad 1985) but increases peroxisomal oxidation (Hovik and Osmundsen 1987). Marine animals have naturally high levels of polyunsaturated L C F A in their diet, originating from zooplankton (Sargent et al. 1976). Studies with liver mitochondria from seals (Osmundsen 1981) and salmonids (Henderson and Sargent 1985) show that such animals possess mitochondria which are better able to oxidize these fatty acids than are rat liver mitochondria. These data suggest that mitochondrial adaptations in response to dietary fatty acids may arise over evolutionary time, or possibly even during chronic exposure, whereas the peroxisomal pathway may be most important in responding to acute changes in dietary fatty acids. Comparative aspects of fatty add oxidation In mammalian liver, when carbohydrate status is high, hormonal and allosteric effectors direct the tissue toward net fatty acid conservation (inhibition of fatty acid oxidation, stimulation of fatty acid synthesis from carbohydrate, stimulation of triglyceride formation). When carbohydrate status is low, mitochondrial B-oxidation of fatty acids increases, largely due to a decrease in malonyl CoA inhibition of CPT (reviewed by McGarry and Foster 1980, Robinson and Williamson 1980). High rates of oxygen consumption are observed with isolated mammalian liver mitochondria given palmitoyl carnitine due to production of reducing equivalents in the B-oxidation spiral, but most of the acetyl CoA generated is used 8 in production of ketone bodies (e.g., Chatzidakis and Otto 1987). The fate of acetyl CoA in the mitochondria (fatty acids synthesis vs. ketogenesis) is regulated by oxaloacetate availability (Lane and Mooney 1981). Mammalian liver typically oxidizes L C F A , M C F A and S C F A by both the carnitine-dependent and carnitine-independent pathways (see Groot et al. 1976). Palmitoyl carnitine is generally oxidized at high rates by vertebrate liver mitochondria (mammals, Bode and Klingenberg 1965; birds, Ballantyne et al. 1988; teleosts, Suarez and Hochachka 1981a, Ballantyne et al. 1989; elasmobranchs, Moyes et al. 1986). Liver mitochondria of mammals (Bode and Klingenberg 1965, de Jong 1971) and lower vertebrates (teleosts, Ballantyne et al. 1989) can oxidize palmitate and M C F A in the absence of carnitine at moderate rates, however carnitine-dependent oxidation usually occurs at greater rates than carnitine-independent fatty acid oxidation. The invertebrate tissues which apparently serve the function of the vertebrate liver in fatty acid processing and storage, also oxidize palmitoyl carnitine at high rates (hepatopancreas of bivalve molluscs (Ballantyne and Storey 1984, Ballantyne and Moon 1985) and crustaceans (Panulirus argus, Tsokos et al. 1983), insect fat body (locust and blowfly; Ballantyne and Storey 1983a)). The ability of invertebrate "livers" to oxidize fatty acids by the carnitine-independent pathway has not been assessed. The fate of hepatic acetyl CoA in species with low reliance on ketone bodies has not been examined (e.g., teleost fish, Zammit and Newsholme 1979). Unlike liver, cardiac and skeletal muscle mitochondria are specialized for oxidation of acetyl CoA produced in 8-oxidation. Red muscle and heart mitochondria from non-mammalian vertebrates typically oxidize long chain fatty acyl carnitine as a preferred substrate (e.g., pigeon breast muscle, Tyler and Gonze 1967, Ernster and Nordenbrand 1967; hummingbird flight muscle, Suarez et al. 1986; frog sartorius, Skoog et al. 1978; carp red muscle, Moyes et al. 1989). Mammalian skeletal and cardiac muscle mitochondria have little capacity to oxidize L C F A by the carnitine-independent pathway (Bode and Klingenberg 1965, Huxtable and Wakil 1971). Mammalian cardiac, but not skeletal, muscle mitochondria can oxidize M C F A in the absence of carnitine (Bode and Klingenberg 1965, Hansford 9 1978) . These differences in capacity to oxidize fatty acids are probably due to differences in chain length preferences of the matrical fatty acyl CoA synthetases. Not all skeletal and cardiac muscles are capable of oxidizing fatty acyl carnitines at high rates. Red muscle mitochondria oxidize fatty acyl carnitines at higher rates than do white muscle mitochondria (rabbit-Pande and Blanchaer 1971, carp-Moyes et al. 1989). Turtle heart mitochondria oxidize acyl carnitines at less than 50% the rate of pyruvate (Kane and Privitera 1970, L.T.Buck and P.W.Hochachka unpubl.). Skeletal muscle and heart mitochondria of elasmobranchs are also unusual in their capacity to oxidize fatty acids. Although elasmobranch heart and skeletal muscle mitochondria possess HO A D , suggesting a capacity for 8-oxidation, they lack detectable CPT activity (Zammit and Newsholme 1979, Sidell et al. 1987, Moon and Mommsen 1987, Moyes et al. 1990). Dogfish red muscle and heart mitochondria oxidize palmitoyl carnitine at less than 10% the rate of pyruvate, although they oxidize medium chain fatty acyl carnitine at greater rates than long chain fatty acyl carnitines. Fatty acids (octanoate, palmitate) are not oxidized at detectible rates even in the presence of 5 m M A T P (Moyes et al. 1990). Mitochondria from mollusc hearts are similar to elasmobranch hearts as they cannot oxidize palmitoyl carnitine (squid, Ballantyne et al. 1981; clam, Ballantyne and Storey 1984). The extent to which mollusc hearts can utilize medium chain fatty acyl carnitine or M C F A is not known. The biochemical strategies which have evolved to fuel flight in insects are remarkably diverse (see reviews by Beenakkers et al. 1984, Storey 1985). Trends regarding fuel preference in insect species generally reflect similar diets or lifestyles, rather than strict phylogenetic relationships (Martin and Lieb 1979) . The insect species which are well-known for prolonged periods of flight are thought to use fatty acids as fuel for long-term flight (e.g. locust, Krogh and Weis-Fogh 1951; Van Der Horst et al. 1978); their mitochondria oxidize palmitoyl carnitine and pyruvate at high rates (e.g. locusts, Van Den Bergh 1967; moths, Hansford and Johnson 1976). Conversely, insects which make relatively short-duration flights generally rely on carbohydrate and/or proline as fuel (Sacktor 1975): Flight muscle mitochondria from such species oxidize palmitoyl carnitine at only 10% the rate of pyruvate (blowfly, Childress and 10 Sacktor 1966; cicada, Hansford 1971), which is similar to the situation with elasmobranchs and molluscs. Mitochondria of some moth species (e.g., Prodenia eridania) are capable of oxidizing fatty acids (palmitate, hexanoate) in the absence of carnitine (Stevenson 1968). This unusual capacity is probably due the presence of a matrical L C F A CoA synthetase, although Beenakkers et al. (1985) have suggested it may be related to elevated thoracic temperatures. Other moth species oxidize palmitoyl carnitine but not palmitate (in the absence of carnitine) (e.g., Manduca sexta, Hansford and Johnson 1976). The energetic benefits of fatty acid-based energy metabolism are well established. One explanation why many species and tissues possess mitochondria which rely less on fatty acids as metabolic fuels may be related to toxic effects of fatty acids and fatty acid oxidation intermediates under hypoxic conditions (Moore 1985). Greater reliance on non-fatty acid substrates occurs in several tissues and species which frequently experience hypoxic conditions. Turtles and bivalve molluscs commonly experience extended periods of environmental hypoxia; their extrahepatic tissues oxidize fatty acyl carnitines at low rates. Vertebrate white muscle also experiences tissue hypoxia and oxidizes fatty acids at low rates. K E T O N E BODIES Catabolic pathway and assay conditions Ketone bodies (acetoacetate and BHB) and pyruvate are transported into mitochondria on the monocarboxylate carrier in exchange for an hydroxyl or another monocarboxylate (reviewed by LaNoue and Schoolwerth 1979). Unlike pyruvate, non-carrier mediated diffusion may be important at physiologically relevant [ketone body] (Halestrap 1978). Once inside the mitochondrion, BHB is oxidized by B H B D H to form acetoacetate. As this enzyme is involved in both synthesis and catabolism of B H B , it is present in both ketogenic and ketolytic tissues. In ketogenic tissues, such as mammalian liver, mitochondria synthesize acetoacetate and BHB from fatty acids and some amino acids. As liver mitochondria cannot catabolize ketone bodies, they are expelled from the mitochondria, in exchange for pyruvate (Patel et al. 1982, Zweibel et al. 1982). 11 Isolated liver mitochondria given BHB consume oxygen at high rates due to N A D H production in the B H B D H reaction, but acetoacetate is not oxidized (e.g., elasmobranch liver, Moyes et al. 1986). This pattern of ketone body oxidation (high rates of oxygen consumption with BHB but no oxidation of acetoacetate) reflects a ketogenic tissue. In extrahepatic tissues, ketone bodies are converted to acetyl CoA which is oxidized in the TCA cycle. In extrahepatic tissues 3-oxo acid CoA-transferase activates acetoacetate into its CoA ester. If the mitochondria are depleted of the co-substrate succinyl CoA, this reaction may limit acetoacetate oxidation. Thiolase cleaves acetoacetyl CoA into two acetyl CoA units which enter the T C A cycle. Addition of malate as a sparker may enhance mitochondrial ketone body oxidation by providing a source of oxaloacetate. Inhibition of mitochondrial BHB oxidation in response to T C A cycle inhibitors suggests a ketolytic tissue. Malonate cannot be used as it directly inhibits B H B D H as well as succinate dehydrogenase (Moyes et al. 1986). Oxidation of acetoacetate, but not B H B , suggests a ketolytic tissue which is deficient in B H B D H (e.g., squid heart, Ballantyne et al. 1981). Comparative aspects of ketone body catabotism Vertebrate liver is a ketogenic, non-ketolytic tissue responsible for ketone body export. As mentioned previously, vertebrate liver mitochondria typically oxidize BHB but not acetoacetate. One exception amongst vertebrates is teleost liver which lacks detectable B H B D H (Zammit and Newsholme 1979). Mitochondria from this tissue have a poor capacity for BHB oxidation (Ballantyne et al. 1989), and therefore synthesis. This is reflected in low circulating BHB levels in vivo (Zammit and Newsholme 1979). Teleost livers can synthesize acetoacetate, but even after extended periods of food deprivation plasma [acetoacetate] remains low (Zammit and Newsholme 1979). Another unusual feature of teleost ketone metabolism is the presence of hepatic 3-oxoacid CoA-transferase (Philips and Hird 1977, Zammit and Newsholme 1979). In mammals this enzyme is involved in ketolysis, but in teleosts it has been suggested to be involved in ketogenesis (Philips and Hird 1977). As in vertebrate liver, hepatopancreas mitochondria of the spiny lobster (Panulirus argus) oxidize BHB at high rates (Tsokos et al. 1983) 12 although hepatopancreas mitochondria of crab (Callinectes sapidus) do not (Chen and Lehninger 1972), suggesting a lack of active hepatic B H B D H in this tissue or preparation. In mammals, extrahepatic ketone utilization appears dependent on substrate availability (e.g., rat hindlimb, Ruderman et al. 1971, Robinson and Williamson 1980). Mitochondria from heart of rat, pigeon (Tyler and Gonze 1967) and turtle (Kane and Privitera 1970, L.T.Buck and P.W.Hochachka unpubl.) oxidize BHB at high rates. Elasmobranch skeletal muscle and heart mitochondria oxidize B H B , acetoacetate and pyruvate at similar rates (dogfish, Moyes et al. 1990) and isolated, perfused elasmobranch hearts treated with iodoacetate perform well when given acetoacetate as fuel (skate, Driedzic and Hart 1984). Apparently unique amongst vertebrates, teleost heart and skeletal muscle do not rely on ketone bodies. Isolated perfused teleost hearts do not use acetoacetate effectively (Driedzic and Hart 1984) and red muscle mitochondria oxidize ketone bodies at very low rates (Moyes et al. 1989). Ketone body oxidation in invertebrates has not been extensively studied. Although crustacean (Orconectes limosus) abdomen muscle mitochondria do not oxidize BHB (Skorkowski et al. 1976), this may be due to low B H B D H activity rather than lack of ketolytic enzymes, as acetoacetate oxidation was not examined. In general, mollusc extrahepatic tissues possess negligible B H B D H activities resulting in low rates of mitochondrial BHB oxidation, but acetoacetate is oxidized at high rates, suggesting that these tissues are ketolytic (Ballantyne et al. 1981, Ballantyne and Moyes 1987a). Ketone bodies vs. fatty acids as metabolic fuels Species which use ketones as the primary lipid-based fuel for extrahepatic tissues (e.g., elasmobranchs, molluscs) must differ from ketotic mammals in several aspects of regulation of ketone metabolism. In mammals, when hepatic carbohydrate status is low, ketogenesis converts the stored fatty acids into substrates useable by tissues which normally rely on glucose, such as brain (McGarry and Foster 1980, Robinson and Williamson 1980). Thus, in mammals, ketone body utilization occurs when carbohydrate status is low but, in species such as elasmobranchs where ketones functionally replace fatty acids, the regulatory dependency of ketogenesis on hepatic carbohydrate status must be altered. 13 Furthermore, the use of ketones instead of fatty acids necessitates differences in whole body triglyceride metabolism. Unlike fatty acids, intramuscular stores of ketones do not exist, so that muscle demand for lipid-derived acetyl CoA must be transmitted to ketogenic pathways. In elasmobranchs, which lack adipose tissue, the liver is both the ketogenic tissue and the major lipid store. At the target tissues the communication between pyruvate and lipid (ketone) oxidation must also be different. The metabolite ratios which allow communcation between fatty acid oxidation and P D H (acetyl CoA /CoA, N A D H / N A D + ) would be expected to be affected in a subdy different manner by ketone body oxidation. Consider, for example, production of 8 moles acetyl CoA from different fuels. From palmitate, 7 cycles of^-oxidation yields 7 moles N A D H (and 7 moles FADHj) . If 4 moles of ketone bodies are used to generate the 8 moles acetyl CoA, 4 moles N A D H result from BHB but no N A D H from acetoacetate. Interactions of pyruvate and ketone bodies at the level of mitochondrial transport occur in mammalian liver. Ketones body expulsion from liver mitochondria, in exchange for pyruvate, stimulates pyruvate oxidation (Patel et al. 1982, Zweibel et al. 1982). Markedly different effects may occur in ketolytic tissues where ketone bodies and pyruvate compete for the monocarboxylate transporter, a competition which may affect the kinetics of oxidation of both fuels. CARBOHYDRATE Aerobic utilization of glucose, lactate or alanine involves mitochondrial oxidation of pyruvate. Mitochondria are also involved in oxidation of reducing equivalents, generated in the cytoplasm during glycolysis, through N A D H shuttles such as the a-glycerophosphate and malate/aspartate shuttles (see review by Dawson 1979). Catabolic pathway Pyruvate enters mitochondria by electroneutral exchange with an hydroxyl or another monocarboxylate on the monocarboxylate transporter (see LaNoue and Schoolwerth 1979). Unfacilitated diffusion becomes an increasingly important mechanism of entry into the mitochondria at [pyruvate] greater than 5 m M , a situation probably never occurring in vivo (Pande and Parvin 1978). As pyruvate 14 transport is dependent on the magnitude of the pH gradient across the mitochondrial membrane, it may be important to consider factors which may afreet this gradient such as the concentration of weak acids and metabolites which are transported on proton symports (pyruvate, phosphate), as well as temperature (Moyes et al. 1988). Once inside the mitochondrion, pyruvate is oxidized by P D H , forming acetyl CoA. This enzyme is under complex allosteric and covalent regulation (Wieland 1983, Reed 1981). In mammals, most of the control of pyruvate oxidation has been attributed to regulation of P D H . The rate of pyruvate oxidation may also be affected by changes in the rate of pyruvate transport (Halestrap et al. 1980, Patel et al. 1982, Zweibel et al. 1982). The influence of concentration of other monocarboxylates (ketones, lactate) on pyruvate transport is not known in non-mammalian vertebrates. Lactate competes with pyruvate for the monocarboxylate transporter of rat heart with lower affinity (Halestrap 1978). This competition may be relevant in species where tissue [lactate] can reach high levels (80 mM in tuna, Guppy et al. 1979; 150 m M in turtles, Jackson and Heisler 1983). Later in this thesis (Chapter 4) I provide evidence that lactate does not compete with pyruvate for the trout white muscle mitochondria. The potential significance of ketone body/pyruvate competition in species which use ketones as the primary lipid source has been discussed previously. In liver several anabolic and catabolic pathways involve mitochondrial transport and metabolism of pyruvate. Consequently, high rates of oxygen consumption in response to pyruvate tell relatively little about pyruvate metabolism. However, if pyruvate is poorly oxidized by mitochondria, it is important to resolve whether the low rate of oxidation is due to low activity of the monocarboxylate carrier or P D H . Low monocarboxylate carrier activity would limit the capacity of the tissue for aerobic glycolysis, gluconeogenesis, lipogenesis and ketolysis; low P D H activity would have similar implications for glycolysis and lipogenesis but may favour gluconeogenesis from lactate or alanine, as these pathways require pyruvate carboxylase-mediated conversion to oxaloacetate. A comparison of the rates of oxidation of alanine, ketone bodies and pyruvate can be used to distinguish between these situations. Alanine can enter mitochondria and give rise to pyruvate via intramitochondrial alanine aminotransferase, precluding 15 the influence of the pyruvate carrier. High rates of alanine oxidation in mitochondria which oxidize pyruvate at low rates suggests a low capacity for monocarboxylate transport. As each of the monocarboxylates are transported with similar maximal velocities (Halestxap 1978), high rates of ketone body oxidation in mitochondria which oxidize pyruvate at low rates suggests low P D H activity rather than monocarboxylate transporter capacity. If P D H is found to be limiting pyruvate oxidation, this could be due to chronically low total enzyme activity which may be related to developmental (Malloch et al. 1986), seasonal or nutritional (Wieland et cd. 1972) factors. Alternately, P D H activity may be subject to inhibition by covalent modification or allosteric effectors (Reed 1981, Wieland 1983). Comparative aspects of pyruvate oxidation Several factors make it difficult to interpret the metabolic significance of either high or low rates of pyruvate oxidation in liver mitochondria. Liver mitochondrial pyruvate metabolism is required in several metabolic pathways, both catabolic (glucose and lactate oxidation) and anabolic (l'pogenesis from glucose, gluconeogenesis from lactate and some amino acids). Pyruvate oxidation may be particularly sensitive to isolation artifacts such as divalent cation depletion and dissipation of the pH gradient. In some cases, liver tissue of closely related species oxidize pyruvate at markedly different rates. For example, Mercenaria hepatopancreas mitochondria oxidize pyruvate at about 50% of the rate of palmitoyl carnitine (Ballantyne and Storey 1984) but no pyruvate oxidation is detectable with Mytilus edulis hepatopancreas mitochondria (Ballantyne and Moon 1985). It is possible that differences in the rate of pyruvate oxidation in some bivalve tissues (e.g. oyster gi l l , Burcham et al. 1983 vs. Ballantyne and Moyes 1987a) are artifacts due to inactivation of P D H by divalent cation depletion during isolation (Ballantyne and Moon 1985). In tissues which are less biosynthetically active, mitochondrial rates of pyruvate oxidation reflect the capacity of the tissue for carbohydrate oxidation. Pyruvate is typically oxidized at rates as great as any other substrate in vertebrate heart and muscle mitochondria (heart mitochondria of mammals (Bremer 1965), turtles (Mersmann and Privitera 1964), elasmobranchs (Moyes et al. 1990); muscle mitochondria 16 of mammals (rabbit red muscle, Pande and Blanchaer 1971), birds (hummingbird flight muscle, Suarez et al. 1986), frog (gastrocnemius, Skoog et al. 1978), teleosts (carp red and white muscle, Moyes et al. 1989 but not goldfish red muscle, Mourik 1983), elasmobranch (dogfish red muscle (Moyes et al. 1990). Mitochondria from invertebrate heart (squid, Ballantyne et al. 1981; bivalve, Ballantyne and Storey 1983b) and muscle (insect flight muscle, Hansford 1971, Bursell 1975, Hansford and Johnson 1976, Weeda et al. 1980) also oxidize pyruvate as preferred substrates. There are several noteworthy exceptions amongst invertebrate tissues. Several bivalve tissues have been reported to oxidize pyruvate at low rates. While isolation artifacts cannot be ruled out, it is possible that the often contradictory results may be due to peculiarities associated with osmolyte metabolism. Depressed pyruvate oxidation may be an adaptation which allows accumulation of alanine, an important osmolyte in these species (Paynter et al. 1985). The stimulation of pyruvate oxidation with hypoosmotic acclimation or hypotonic incubation (Ballantyne and Moyes 1987a) may be related to the sensitivity of P D H to ionic environment (e.g. rat kidey, Pawekzyk et al. 1988). Tsetsefly flight muscle mitochondria are unusual amongst insects as they appear to rely primarily on proline oxidation to fuel flight, with pyruvate oxidized at low rates relative to proline (Bursell 1975). Although mitochondria from virtually every muscle examined have the capacity to oxidize pyruvate at high rates, it is likely that interspecific differences in pyruvate metabolism have been masked because most studies employed optimal conditions (single substrate, saturating concentrations). Isolated mitochondria oxidize pyruvate and fatty acyl carnitines at similar rates, but when presented together oxidation of fatty acids inhibits pyruvate oxidation (e.g., Hansford and Cohen 1978). Thus, in perfused mammalian hearts given glucose and fatty acids, about 80% of C 0 2 produced originates from the fatty acids (Bremer et al. 1984). Even those insect species which appear to fly using lipid fuels, carbohydrate oxidation fuels the pre-flight warm-up (reviewed by Storey 1985). Oxidation of fatty acids causes changes in mitochondrial metabolites ( N A D H / N A D + , acetyl CoA/CoA and ATP/ADP) which result in inactivation (covalent) and inhibition (allosteric) of P D H (e.g., Hansford and Cohen 1978). A similar interaction 17 between oxidation of fatty acids and pyruvate is evident in mitochondria from non-mammalian vertebrates (teleost liver, Suarez and Hochachka 1981b; hummingbird flight muscle, Suarez et al. 1986). It is not known if fundamentally similar interactions are involved in P D H regulation in mitochondria from species which utilize metabolites other than fatty acids to fuel exercise (e.g., ketones in elasmobranchs, proline in squid and tsetsefly). Another important regulator of P D H in mammals is mitochondrial [Ca 2 + ] , as affected by cytoplasmic [Ca 2 + ] . Changes in cytosolic [Ca 2 +] arising through the effects of Ca2+-dependent hormones modulate P D H activity (Oviasu and Whitton 1984, McCormack and Denton 1985, Staddon and Hansford 1987). An increase in work of heart and skeletal muscle stimulates P D H (Hagg et al. 1976, Illingworth and Mullings 1976, Pearce et al. 1980) possibly through the effects of increased cytoplasmic [Ca 2 +] (McCormack and England 1983, Hansford 1987). The relationship between P D H , cytoplasmic and mitochondrial [Ca 2 + ] is unknown in animals which may experience high or fluctuating intracellular [Ca 2 +] associated with osmotic changes (salinity stress in osmoconformers, extracellular freezing) or acid-base changes (mobilization of calcium salts in shells during hypoxia). AMINO ACIDS Catabolic pathway It has been suggested that neutral amino acids enter mitochondria by non-facilitated diffusion (Hailing et al. 1973). However rat mitochondria accumulate neutral amino acids in a stereospecific manner, indicative of a specific carrier (Cybulski and Fisher 1977). Glycine is transported at much lower maximal velocities in several species and is thought to enter rat brain and liver mitochondria by a separate carrier (Benavides et al. 1980). Mitochondrial glycine oxidation may occur through serine after conversion and reversion by cytoplasmic and mitochondrial forms of serine hydroxymethyl transferase. This is thought to aid in the transfer of one carbon units into the mitochondria (Cybulski and Fisher 1976). The proline transporter is separate from the neutral amino acid transporter as suggested by differential sensitivity to thiol blocking agents and sensitivity to uncouplers (Meyer 1977, Moyes and Ballantyne 1987). Maximal rates of neutral amino acid and proline oxidation are obtained with less than 18 10 mM concentrations. Two general pathways exist for glutamate catabolism in mammalian tissues. Glutamate oxidation occurs through glutamate dehydrogenase (GDH) following transport by the glutamate/OH" exchanger (bidirectional). In the second pathway glutamate enters the mitochondria by the glutamate/aspartate (unidirectional) exchanger and is transaminated by aspartate aminotransferase to form a-ketoglutarate (see LaNoue and Schoolwerth 1979). The latter pathway is required in the malate-aspartate shuttle for transport of reducing equivalents into mitochondria (see Dawson 1979). Relative fluxes of glutamate through these pathways may be dependent on [a-ketoglutarate], regulation of G D H and pH (Tannen and Ross 1979, LaNoue et al. 1983, Moyes et al. 1985). As with BHB, high rates of oxygen consumption do not necessarily indicate entrance of glutamate carbon into the T C A cycle or the importance of the G D H route relative to the aminotransferase pathway. As many amino acids can be catabolized through aminotransferases, it may be necessary to provide mitochondria with a-ketoglutarate as the amino acceptor to achieve maximal rates of amino acid oxidation. B C A A (branched-chain amino acids; valine, leucine, isoleucine) catabolism is initiated cytoplasmically by B C A A transaminase, which is present in relatively high activity in muscle but also occurs in liver (Harper et al. 1984). Transport of the resulting B C K A (branched-chain keto acids) into the mitochondrion occurs by a specific transporter which is sensitive to the mitochondrial pH gradient (Hutson 1986, Hutson and Rannels 1985). B C K A dehydrogenase is the first step of the mitochondrial pathway which leads to production of succinyl CoA (isoleucine, valine) or acetoacetyl CoA (leucine). It is under covalent control with the active form (dephosphorylated) predominant in the liver and the inactive form (phosphorylated) predominant in the muscle (Harper et al. 1984, Wagenmakers et al. 1984). These observations have lead to the suggestion that B C A A are oxidized primarily in liver, after transamination in the muscle (see Harper et al. 1984). However, under exercise conditions there is an activation of muscle B C K A D H (Kasperek et al. 1985, Kasperek and Snider 1987) which coincides with an increase in B C A A uptake by muscle, suggesting that these amino acids may be relevant fuels for 19 muscle work (Ahlborg et al. 1974). Comparative aspects of amino acid oxidation Amino acids are derived from the diet, routine protein turnover, and other pathways such as the giucose-alanine cycle. Amino acid catabolism is centralized in the liver, which possesses the pathways for both conversion of carbon skeletons to storage products (lipid, carbohydrate) and detoxification of the resulting ammonia. The changes in mitochondrial ammonia metabolism associated with the evolution of uricogenesis and ureagenesis are discussed in reviews by Campbell et al. (1987) and Mommsen and Walsh (1989). Hepatic amino acid oxidation spares dietary carbohydrate and lipid. Consequently, vertebrate liver mitochondria are generally capable of transporting and oxidizing most amino acids or their cytoplasmically-generated metabolites (e.g. Moyes et al. 1986). Direct utilization of amino acids by mammalian muscle is limited. Most of the amino acids produced during starvation are released from the tissue or transaminated to form alanine or glutamine (Goldberg and Chang 1978). One reason for restricting amino acid oxidation to liver in tetrapods is that ammonia release occurs in the same tissue as the pathway for ammonia detoxification. Although this would not apply to ammoniotelic animals, migrating salmon, which catabolize white muscle protein, also release alanine into the plasma (Mommsen et al. 1980, French et al. 1983). It is not known if alanine can be utilized directly by oxidative tissues (red muscle, heart) under these conditions, but carp red muscle mitochondria do not oxidize alanine at detectable rates (Moyes et al. 1989). In invertebrate tissues, direct utilization of amino acids in energy metabolism may be more extended. In squid, proline is present at high tissue concentrations and oxidized at high rates by heart mitochondria (Ballantyne et al. 1981, Mommsen and Hochachka 1981, Mommsen et al. 1982). Proline serves several roles in insect flight muscle metabolism. Tsetse fly flight muscle mitochondria oxidize proline at rates 50- to 100-fold greater than pyruvate (Bursell 1975). Flight muscle mitochondria of the Colorado beede QLeptinotarsa decemlineata) are also able to oxidize proline at greater rates than pyruvate or fatty acyl carnitine (de Kort et al. 1973, Weeda et al. 1980). The peculiar pathway which allows 20 proline to be utilized as sole substrate is described by Storey (1985). Briefly, proline is converted to glutamate which enters the T C A cycle as a-ketoglutarate following transamination by alanine aminotransferase. This reaction requires equimolar amounts of pyruvate as amino group acceptor. Pyruvate is probably produced from T C A cycle malate via NAD+-dependent malic enzyme (Weeda et al. 1980). Ultimately proline carbon is the source of both glutamate and pyruvate, with the short circuit allowing a portion of the T C A cycle to proceed without a source of acetyl CoA. Other insects have mitochondria which oxidize proline at high rates and utilize proline as fuel for flight, but these mitochondria are also capable of utilizing pyruvate as an acetyl CoA source (e.g., Popillia japonica, Hansford and Johnson 1975). Flight muscle mitochondria of other species oxidize proline at low rates relative to pyruvate (e.g. Musca domestica, Bursell 1975) and may use proline primarily in an anaplerotic role. Chamberlin and Phillips (1983) found that locust midgut oxidizes proline to provide the energy for ion pumping. Amino acid metabolism in osmoconformers Although osmoconforming animals maintain high levels of plasma and intracellular amino acids (proline, alanine, 6-alanine, glycine, glutamate, sarcosine, betaine) little is known of the metabolic mechanisms which control osmolyte levels. Osmotically-active amino acids are oxidized primarily by mitochondrial pathways, and mitochondrial density appears to be an important determinant of the osmolyte profile. In mitochondria-rich tissues, oxidizable solutes are partially replaced with non-oxidizable solutes such as urea and T M A O in elasmobranchs and taurine in bivalve molluscs (see Ballantyne et al. 1987). In mammals, deamination of amino acids occurs primarily in the liver with ammonia released in the G D H reaction. Gi l l (Zurburg and de Zwaan 1981) and mantie (Bartberger and Pierce 1976) may be centralized tissues for deamination of amino acid osmolytes in hypo-osmotically stressed bivalve molluscs. Mitochondria from these tissues oxidize many of the important amino acid osmolytes at high rates. Glutamate is oxidized at high rates in bivalve gill (Burcham et al. 1983, 1984) and mantie (Moyes et al. 21 1985) mitochondria. Although it has been suggested that the levels of G D H are too low to be important in amino acid deamination (deZwaan and van Marrewijk 1973, Reiss et al. 1977), studies of glutamate oxidation by clam mantle mitochondria suggest that in situ G D H activity is sufficient to catalyze the high rate of glutamate oxidation (Moyes et al. 1985). Proline is another important osmotically active solute which is oxidized through G D H . Proline oxidation does not occur at detectable rates in hepatopancreas mitochondria (Mercenaria, Ballantyne and Storey 1984, Mytilus edulis, Ballantyne and Moon 1985). Proline is oxidized at moderate rates in gill mitochondria of mussels (Modiolus demissus, Burcham et al. 1984) and oysters (Crassostrea virginica, Moyes and Ballantyne 1987), contrary to a previous study on the same species (Burcham et al. 1983). The mechanisms which control osmolyte metabolism, particularly during osmotic stress, are not well established and may differ for each metabolite and species. Osmotic effects on mitochondrial oxidation of osmolytes may be important in cell volume regulation in osmoconforming animals (Ballantyne et al. 1987). Mitochondrial oxidation of various osmotically-active amino acids is sensitive to changes in mitochondrial volume (Ballantyne and Storey 1983, Ell is et al. 1985, Moyes et al. 1986). As the effects of hypotonic incubation can be mimicked by acclimation to dilute sea water (with incubation in isotonic medium), it is unlikely that mitochondrial oxidation responds to mitochondrial volume per se but rather some mitochondrial parameter altered by both mitochondrial volume changes and acclimation to dilute sea water (Ballantyne and Moyes 1987c). In mammals increased intracellular [Ca 2 + ] stimulates mitochondrial oxidation through direct effects on P D H and T C A cycle dehydrogenases (McCormack and Denton 1986, Rutter and Denton 1986, Johnston and Brand 1987). C a 2 + may also affect mitochondrial oxidation by altering matrix volume (Halestrap et al. 1986, Halestrap 1987, Davidson and Halestrap 1987, Halestrap and Dunlop 1987). In osmoconformers, mitochondrial oxidation of osmolytes is stimulated by hypoosmotic stress in hepatocytes in the same manner as observed with isolated mitochondria suggesting mitochondria swell in situ (Ballantyne et al. 1986). However, stimulation of mitochondrial oxidation with hypoosmotic incubation appears unrelated to changes in mitochondrial [Ca 2 +] 22 as the effects of osmotic stress persist in mitochondria treated with E D T A and the calcium ionophore A-23187 (Ballantyne and Moyes 1987b). Several of the enzymes and transporters involved in osmolyte oxidation are sensitive to changes in ionic environment (Moyes et al. 1986, Moyes and Moon 1987, Ballantyne and Moyes 1987b) but it is not known how matrix ionic stength changes with osmotic stress. Considerable information can be gained about how a muscle functions when one understands how the mitochondria function, because of their central role in aerobic metabolism. Fish are useful and interesting models because comparisons between different species and muscles demonstrate a much greater range in mitochondrial parameters than any other vertebrate group. In this thesis, I use fish muscle mitochondria to addressing questions of control of fuel preference, aerobic capacity and metabolic regulation. Dogfish muscles are unusual in that they lack CPT, the enzyme capable of transporting fatty acids into mitochondria but possess H O A D , an enzyme involved in fatty acid oxidation. In Chapter 2,1 address this paradox using dogfish mitochondria to establish whether elasmobranchs are able to utilize fatty acids by alternate pathways that bypass CPT. Chapter 3 examines the nature and importance of mitochondrial differences in muscles with dramatically different aerobic capacities. Muscles from carp (low aerobic potential) and tuna (high aerobic potential) are compared to determine the contributions of intrinsic and extrinsic factors in determining tissue aerobic capacity. Chapter 4 deals specifically with the role and regulation of mitochondria from white muscle. Burst exercise does not involve mitochondrial ATP producing pathways but recovery metabolism demands mitochondrial A T P production. I examine control of mitochondrial metabolism by extreme and prolonged changes in metabolite levels which are characteristic of recovery from burst exercise in fish. 23 C H A P T E R 2: M I T O C H O N D R I A L A N D P E R O X I S O M A L F A T T Y A C I D O X I D A T I O N IN E L A S M O B R A N C H S P R E F A C E This chapter is adapted from a paper published by Moyes, L.T.Buck and P.W.Hochachka (Am.J.Physiol. 258:R756-R762, 1990). Some of the mitochondria respiration rates were obtained in collaboration with L.T.Buck. A l l other aspects of collection of data and presentation of results were performed by myself. I N T R O D U C T I O N Long term aerobic muscle activity in vertebrates is usually supported primarily by lipid oxidation, often after an early phase of carbohydrate oxidation. The advantage of this strategy appears related to storage capacities of triglyceride vs. glycogen, and the need to spare carbohydrate for tissues which rely primarily on glucose, such as brain (Hochachka and Somero 1984). With few exceptions, such as flight muscle of Prodenia moths (Stevenson 1968), CPT is required for oxidation of L C F A by extrahepatic (heart, skeletal muscle) mitochondria (Figure 1). Consequently, when Zammit and Newsholme (1979) reported that extrahepatic tissues of elasmobranchs lack detectable CPT, attention was focused on possible metabolic adaptations which may be used to bypass this step in lipid oxidation. As an alternative to direct utilization of fatty acids, Zammit and Newsholme (1979) suggested that the heart and red muscle of elasmobranchs may rely on liver-generated ketone bodies as a "lipid" source. Support for this hypothesis is the observation by Driedzic and Hart (1984) that elasmobranch hearts, unlike teleost hearts, are able to utilize ketone bodies as fuel for work. Although elasmobranch heart and skeletal muscle lack CPT, they do have H O A D , a 8-oxidation enzyme (Moon and Mommsen 1987). These observations present the possibility that elasmobranch muscle may utilize fatty acids by an alternate pathway. Several possibilities are apparent. Fatty acids could be oxidized directly, by a carnitine-independent pathway (Figure 1), such as occurs in mammalian liver and heart (Groot et al. 1976). Alternately, fatty acid oxidation may be directed toward fatty acids of short (SCFA) or medium (MCFA) chain lengths, a capacity which wouldn't 24 neccessarily be reflected by CPT activity. Oxidation of fatty acids by the B-oxidation pathway was long thought the domain of the mitochondrion. Lazarow and deDuve (1976) have described a peroxisomal B-oxidation pathway in rat liver which occurs at maximal activities that approach the mitochondrial rate (Mannaerts et al. 1979). The peroxisomal pathway is unlike the mitochondrial pathway in that CPT is not involved in L C F A oxidation, as fatty acids are apparently transported as CoA esters (Appelvist and Dallner 1980). Also L C F A are not oxidized completely, which results in the production of M C F A (Lazarow and deDuve 1976). At present there is no concensus on the relative importance of the peroxisomal and mitochondrial pathways of 8-oxidation in generation of acetyl CoA (Masters and Crane 1984). The induceability of the peroxisomal pathway in response to metabolic stimuli (nature and quantity of dietary lipid (Osmundsen 1982), food deprivation (Ishii et al. 1980) and cold (Nedegaard et al. 1980)) suggests the relative importance of the two pathways in mammalian tissues is malleable. Fish naturally experience a tremendous range in the factors which are known to affect peroxisomal abundance in mammals. The fatty acids which are particularly effective in causing proliferation of peroxisomal B-oxidation in rats (long chain, unsaturated, Osmundsen 1982) are abundant in the marine food chain (Sargent 1976). A potential role for peroxisomal jj-oxidation in facilitating the CPT-deficient mitochondrial pathway in elasmobranchs is obvious. In the present study I investigate the capacities of dogfish heart and red muscle mitochondria to oxidize various metabolic fuels. The capacity of elasmobranch muscle mitochondria to oxidize fatty acids is determined using fatty acyl carnitines and free fatty acids, each of various chain lengths. I compare the activities of several of the key enzymes involved in peroxisomal and mitochondrial fatty acid oxidation in liver, heart and red muscle of selected species of teleosts (common carp, rainbow trout), chondrichthians (dogfish, raffish) and agnathans (hagfish). M A T E R I A L S A N D M E T H O D S Animals Al l fish were held at seasonal temperatures (10-15°C) in continuously flowing water without 25 feeding (except trout). Dogfish (Squalus acanthias) were caught by hook and line and held for 2-10 days at either Bamfield Marine Station or Dept. of Fisheries and Oceans, West Vancouver. Ratfish (Hydrolagus colliei) were caught by otter trawl and used within 1 to 3 days of capture at Bamfield Marine Station. Hagfish (Eptatretus stouti) were caught by baited trap and held 2-7 days at Bamfield Marine Station or the University of British Columbia. Carp (Cyprinus carpio) were purchased from Latek Enterprises (Ruskin, B.C.) and held 2 to 5 days at the University of British Columbia. Trout (Oncorhyncus myldss) were purchased from Lakeland Trout Farms (Langley, B.C.) and held on a maintenance ration at the University of British Columbia. Preparation of tissues for enzyme assays Heart, red muscle and liver were sampled from animals which were killed by double pith (dogfish) or decapitation. Single (dogfish, carp, trout) or 2-4 pooled (hagfish, ratfish) ventricles were taken for each homogenization. Lateral red muscle was taken from the tail region of dogfish (near the second dorsal fin), near the dorsal fin of carp and trout and in hagfish, at the level of the liver, just below the skin. Red muscle from ratfish was sampled from the muscle pads of the pectoral fins. Enzyme assays were performed on either tissue homogenates or crude peroxisomal fractions. Tissue homogenates were prepared by Polytron disperal of tissues in 6 volumes of buffer A (1 mM E D T A , 1% Triton X-100 and 20 m M HEPES at pH 7.4) followed by two 10 sec bursts of sonication (Kontes Ultrasonic Cell Disrupter). The homogenates were centrifuged at 4°C for 10 min at 12000xg. The aqueous supernatant fraction was pipetted from between the sedimented particulate matter and surface lipid and used for enzyme assays. Tissues used for preparation of crude peroxisomal fractions were dispersed in 10 volumes of buffer using a Potter-Elvejhem homogenizer. Buffer B (150 KC1, 500 mM sucrose, 1 mM E D T A , 20 mM HEPES at pH 7.2) was used for dogfish, ratfish and hagfish tissues. Buffer C (150 KC1, 1 mM E D T A , 20 mM HEPES at pH 7.2) was used for teleost (carp, trout) tissues. Homogenates were centrifuged 5 min at 3000xg to sediment unbroken tissue and heavy mitochondria. Supernatants were collected and 26 centrifuged 20 min at 18000xg to make a peroxisome-enriched pellet. The pellet was resuspended and sonicated in a small volume of buffer A and used for enzyme assays. Enzyme assays A l l assays were performed at 15°C using either a Pye-Unicam SPl800 or SP6-550 at the appropriate wavelength. Preliminary studies were performed to establish that substrate concentrations were saturating. Peroxisomal B-oxidation was assayed according to Lazarow and deDuve (1976) and Neat et al. (1981). Peroxisome-enriched fractions were obtained by differential centrifugation as described above. A 1 ml cuvet contained 0.025 m M F A D , 0.1 m M coenzyme A , 0.2 m M N A D + , 0.1% Triton X-100, 20 mM HEPES (pH 7.2), 100 u M palmitoyl CoA (omitted for control). In this assay peroxisomal B-oxidation is distinguished from the mitochondrial pathway by solubilization with Triton X-100. In mitochondrial B-oxidation by intact mitochondria, electrons accepted at the fatty acyl CoA dehydrogenase step are donated to the electron transport system. In the solubilized system, the electron transport system is disintegrated, and the lack of electron acceptors inhibits the dehydrogenase. In the peroxisomal pathway, electrons accepted by fatty acyl CoA oxidase are transferred directly to molecular oxygen. In this study, solubilization clearly blocked the mitochondrial pathway, as N A D H production was not detected in the mitochondria-rich fractions prepared from heart and muscle. N A D H is produced in the H O A D reaction of the peroxisomal enzyme complex. The rate of appearance of N A D H was monitored at 340 nm. Whole tissue activities were adjusted for peroxisomal yield, using catalase as the peroxisomal marker. The assay for CPT included 0.2 m M D T N B , 20 mM TRIS-HC1 (pH 8.0) and 0.1 m M palmitoyl CoA. CPT activity, initiated by the addition of 2.5 m M L-carnitine, was monitored as the change in absorbance at 412 nm. In tissues with low CPT activity relative to background thioesterase activity (dogfish and hagfish tissues, livers of other species), assays were performed with the reference cell containing the complete assay mix with L-carnitine omitted (thioesterase activity). In tissues with low 27 background thioesterase activity relative to CPT (teleost extrahepatic tissues), the rate prior addition of carnitine was subtracted from that after the addition of carnitine. The assay for catalase activity is described by Aebi (1974). The enzyme fraction (homogenate or peroxisome-enriched pellet) was diluted with 20 m M potassium phosphate buffer (pH 7.0) to give a 0.02-0.06 unit change in absorbance (240 nm) in 30 sec. The reference cuvet contained 2 ml diluted enzyme with 1 ml potassium phosphate buffer (20 m M , pH 7.0). To initiate catalase activity, 1 ml of hydrogen peroxide (30 m M in potassium phosphate buffer) was added to the sample cuvet which also contained 2 ml diluted enzyme. The rate is expressed as k/g wet weight, where k is the rate constant (2.3/30 sec x log (A ( /A 3 0 ) ) . H O A D activity was assayed using tissue homogenates by following the rate of change in absorbance at 340 nm of a cuvet containing acetoacetyl CoA (0.1 m M in heart and muscle preparations, 0.2 m M in liver), 0.15 m M N A D H in 20 m M imidazole at pH 7.0. Mitochondrial studies on dogfish red muscle and heart Dogfish red and cardiac muscle mitochondria were isolated using the following medium: 500 mM sucrose, 150 mM KC1, 10 mM E D T A , 5 m M MgCl 2 , 0.1% bovine serum albumin (BSA) in 20 mM HEPES pH 7.2 at 20°C. Muscle tissue was minced with scissors and homogenized using a Potter-Elvejhem tissue grinder. The homogenate was centrifuged 5 min at 1400 x g. The supernatant was poured through 3 layers of cheesecloth and centrifuged 5 min at 7650 x g. The pellet was resuspended in isolation medium minus BSA and recentrifuged 5 min at 7650 x g. The resultant pellet was resuspended in the same medium to approximately 5 mg mitochondrial protein/ml. Mitochondrial protein was determined by the biuret method, using 10% deoxycholate to solubilize the mitochondrial protein. Mitochondrial oxygen consumption was monitored using a Clark-type electrode in a glass cell at 15°C. One volume of mitochondrial suspension was added to 9 volumes of reaction medium (modified from Moyes et al. 1986) containing 150 m M KC1, 350 m M urea, 175 m M trimethylamine oxide, 300 m M sucrose, 5 m M NajHPO*, 0.1 % BSA , 20 m M HEPES (pH 7.2). Assays were also performed under 28 hypoosmotic conditions using this reaction medium with sucrose omitted. Osmotically-induced swelling increases the rate of oxidation of various metabolites (Ballantyne et al. 1981). This technique was used to try to stimulate oxidation of those metabolites which were not oxidized at detectable rates, thereby increasing the sensitivity of the assay. After 1-2 minutes of incubation with 0.1 m M malate and 0.5-0.8 m M A D P , the rate of oxygen consumption decreased to a slow, linear rate and saturating amounts of substrates were added to the suspension. State 3 is the rate of oxygen consumption in the presence of A D P and substrate. The respiratory control ratio, (RCR) is the ratio of State 3 to State 4, which is the rate of oxygen consumption after all A D P is phosphorylated. R E S U L T S CPT and HO AD The activities of enzymes involved in fatty acid oxidation by mitochondria are summarized in Table 1. In the teleosts, CPT and H O A D activities were similar in heart, red muscle and liver. In the dogfish and hagfish, H O A D and CPT activities were generally higher in liver than other tissues. If one considers the relative activites of these enzymes as indices of the capacity for fat oxidation in extrahepatic tissues of different species, conflicting conclusions would arise from the activities of H O A D vs. CPT. What is most striking in the inter-species comparisons is the general similarity of H O A D activites in heart and muscle of each species, with dramatic differences in CPT activity. The activities of CPT in heart and red muscle of hagfish were about 10% that in the teleosts, and CPT was not detectable in these dogfish tissues. The observations reported in the present study, and previous studies dealing specifically with heart (Sidell et al. 1984) prompted a more detailed investigation into possible carnitine-independent pathways for fatty acid oxidation. Mitochondrial studies The goals of the mitochondrial studies (Tables 2 and 3) were to quantify the capacity of heart and red muscle mitochondria to utilize fatty acids relative to other fuels (pyruvate, ketone bodies) and establish the importance of the carnitine-dependent and carnitine-independent pathways. Mitochondrial 29 Table 1. Maximal activities of the mitochondrial enzymes jS-hydroxyacyl CoA dehydrogenase (HOAD) and carnitine palmitoyl transferase (CPT). H O A D CPT units/g (SEM) N units/g (SEM) N carp liver 1.4 (0.06) 10 0.39 (0.05) 6 heart 2.3 (0.09) 10 0.17 (0.04) 6 red muscle 1.6 (0.04) 10 0.32 (0.05) 6 trout liver 2.9 (0.6) 11 0.30 (0.03) 6 heart 7.2 (0.6) 9 0.28 (0.02) 5 red muscle 8.1 (0.6) 11 0.51 (0.04) 6 dogfish liver 7.9 (0.5) 6 0.09 (0.02) 3 heart 1.6 (0.1) 6 < 0.010 3 red muscle 2.9 (0.2) 6 < 0.010 3 hagfish liver 4.5 (0.3) 6 0.077 (0.01) 12 heart 4.4 (0.09) 6 0.044 (0.005) 12 lat. muscle 2.6 (0.1) 6 0.022 (0.005) 12 Assays were performed on tissue homogenates as described in Materials and Methods. Units for both H O A D and CPT are umoles substrate/min/g wet weight. 30 Table 2. Mitochondrial oxidation of fatty acids, ketone bodies and pyruvate by mitochondria from dogfish red muscle. Isoosmotic Hypoosmotic Substrate mean (SEM) N mean (SEM) N 0.1 m M malate* 6.1 (0.7) 12 6.3 (0.9) 12 a-ketoglutarate (0.1 mM) 14.6 (1.7) 5 15.7 (2.3) 5 pyruvate (5 mM) 80.7 (4.2) 5 101.6 (4.5) 5 j?-hydroxybutyrate (5 mM) 75.3 (6.1) 12 82.9 (7.0) 12 +a-ketoglutarate (0.1 mM) 93.4 (4.6) 5 117.5 (6.2) 5 acetoacetate (5 mM) 7.9 (0.9) 5 8.6 (1.0) 5 +a-ketoglutarate (0.1 mM) 63.9 (6.1) 10 92.1 (6.1) 10 acetyl-DL-carnitine (3 mM) 21.9 (1.9) 5 26.5 (1.0) 5 octanoyl-DL-carnitine (400 uM) 8.1 (1.5) 5 15.9 (2.4) 5 palmitoyl-DL-carnitine (100 uM) N D 5 9.9 (1.0) 5 hexanoate (100, 500 uM) N D 3 -octanoate (100, 500 uM) N D 4 N D 3 + 5 m M A T P N D 3 N D 3 decanoate (100, 500 uM) N D 3 -palmitate (24, 100 uM) N D 5 N D 3 +5 mM A T P N D 3 N D 3 A l l substrates were assayed in the presence of 0.1 m M malate. Albumin concentration was 0.014 mM (1 mg/ml). Rates are nmoles O/min/mg mitochondrial protein, expressed as mean (SEM). ND- not detected above the rate with malate alone. Mean RCR (8-hydroxybutyrate) <30 (n=5). 31 Table 3. Mitochondrial oxidation of fatty acids, ketone bodies and pyruvate by mitochondria from dogfish heart. Isoosmotic Hypoosmotic Substrate mean (SEM) N mean (SEM) N 0.1 m M malate* 4.8 (0.4) 10 5.0 (0.6) 10 a-ketoglutarate (0.1 mM) 21.4 (1.6) 10 22.4 (1.8) 10 pyruvate (5 mM) 91.6 (4.5) 5 99.8 (4.8) 5 B-hydroxybutyrate (5 mM) 40.2 (3.2) 10 41.0(1.9) 10 +a-ketoglutarate (0.1 mM) 58.1 (7.5) 5 67.7 (5.1) 5 acetoacetate (5 mM) 7.4 (0.6) 5 10.4 (1.3) 5 +a-ketoglutarate (0.1 mM) 48.3 (4.5) 10 50.4 (5.6) 10 acetyl-DL-carnitine (3 mM) 7.0 (0.4) 5 8.3 (0.6) 5 octanoyl-DL-carnitine (400 uM) 6.5 (0.7) 5 15.1 (0.7) 5 palmitoyl-DL-carnitine (100 uM) N D 5 7.3 (1.1) 5 octanoate (100, 500 uM) N D 3 N D 3 + 5 m M A T P N D 3 N D 3 palmitate (24, 100 uM) N D 3 N D 3 +5 m M A T P N D 3 N D 3 See legend for Table 2. Mean RCR (pyruvate) 22.0 (SEM 1.2, n=5) 32 quality of both preparations was high, as indicated by high RCR values with preferred substrates in red muscle (Table 2) and heart (Table 3). Fatty Acids The studies of oxidation of S C F A , M C F A and L C F A by dogfish red muscle and heart mitochondria are summarized in Tables 2 and 3, respectively. Oxidation of free fatty acids was not detectable in either tissue. Hypoosmotic incubation, which stimulated oxidation of other substrates did not elevate free fatty acid oxidation to detectable levels (Tables 2, 3). The possibility that fatty acid oxidation was not observed due to A T P depletion intramitochondrially is unlikely as addition of S mM A T P , giving A T P / A D P > 10, did not stimulate oxidation of free fatty acids under isoosmotic or hypoosmotic conditions. Fatty Acyl Carnitines The results observed with fatty acyl carnitines were similar for red muscle and heart mitochondria. When incubated under isosmotic conditions, palmitoyl carnitine was not oxidized at rates greater than the control rate (malate + ADP) . Octanoyl carnitine was oxidized at several-fold higher rates than palmitoyl carnitine, when adjusted for the control rates. In red muscle the best acyl carnitine substrate tested was acetyl carnitine but in heart acetyl carnitine oxidation was lower than octanoyl carnitine, suggesting the activity of carnitine acetyl transferase was rate-limiting in this tissue. Hypoosmotic incubation stimulated oxidation of all acyl carnitines, and oxidation of palmitoyl carnitine became consistently detectable, although the rate observed remained low. Carbohydrate and Ketone Bodies Elasmobranch extrahepatic tissues are thought to use glucose or ketone bodies to support metabolism (Zammit and Newsholme 1979, Driedzic and Hart 1984). Glucose and lactate oxidation requires mitochondrial pyruvate oxidation; pyruvate was oxidized at high rates in both tissues (Table 2). In dogfish heart mitochondria, pyruvate was oxidized at higher rates than 6-hydroxybutyrate, but in red muscle they were oxidized at similar rates. In the absence of a-ketoglutarate, acetoacetate oxidation 33 occurred at very low rates (data not shown). Sparking mitochondrial respiration with a-ketoglutarate stimulated acetoacetate oxidation by providing a source of succinyl CoA which is needed for activation of acetoacetate into its CoA ester. In both tissues, B-hydroxybutyrate was oxidized at rates similar to acetoacetate. Peroxisomal fi-oxidation Peroxisomal B-oxidation occurred at similar activities in liver of each species examined, except hagfish where it was below detectible limits (Table 4). The rates of peroxisomal B-oxidation in chondrichthians are more impressive when the higher lipid content is taken into consideration. The activities of peroxisomal B-oxidation in extrahepatic tissues of all species were below detectable levels. Catalase activity in each species was highest in liver, followed by heart and red muscle, each 1/7-1/40 that of liver (Table 4). Interestingly hagfish liver catalase activity was similar to dogfish liver, yet these tissues had markedly different capacities for peroxisomal B-oxidation. Also, while carp had 50% higher rates of peroxisomal B-oxidation than trout, catalase activity was 2.5-fold greater in trout. DISCUSSION Oxidation of glucose and lipid demands mitochondrial oxidation of pyruvate and fatty acids. L C F A oxidation in mammalian muscle mitochondria requires CPT activity and carnitine-acyl carnitine translocase (Figure 1). This is not related to free fatty acid permeability but, rather, depends upon the distribution of acyl CoA synthetases capable of activating L C F A . Heart and red muscle mitochondria typically lack matrical long chain acyl CoA synthetases, and are therefore dependent on the CPT-mediated pathway (Groot et al. 1976). The tissue distribution of CPT and H O A D in several fish species (Table 1) demonstrates the unusual situation found in elasmobranchs and hagfish. Although extrahepatic H O A D activities are in the same range as the teleost species, CPT activity is lower in elasmobranch liver and below detectable limits in both heart and red muscle. Hagfish also possess low CPT activities, but H O A D activity in each tissue is similar to the respective teleost tissues. As is the situation with elasmobranchs, isolated hagfish hearts 34 Table 4. Maximal activities of the peroxisomal enzymes catalase and peroxisomal jS-oxidation (PBO). PBO catalase units/g (sem) N units/g (sem) N carp trout dogfish hagfish liver 0.049 (0.002) 8 31.3 (2.4) 14 heart < 0.005 3 2.25 (0.2) 12 red muscle < 0.005 3 0.77 (0.04) 12 liver 0.033 (0.004) 5 79.7 (7.7) 14 heart < 0.005 3 1.45 (0.12) 9 red muscle < 0.005 3 0.87 (0.15) 11 liver 0.020 (0.002) 5 3.53 (0.4) 5 heart < 0.005 3 -red muscle < 0.005 3 0.44 (0.09) 4 liver 0.038 (0.004) 4 11.4 (1.2) 7 heart < 0.005 3 0.16(0.03) 5 red muscle < 0.005 3 0.18 (0.05) 6 liver < 0.005 6 18.2 (2.75) 6 heart < 0.005 3 1.30 (0.09) 9 lat. muscle < 0.005 3 0.30 (0.04) 9 Assays for PBO were performed on peroxisome-enriched fractions and corrected for peroxisomal yield using catalase. Catalase units (k/g wet weight) are as described in Materials and Methods. Peroxisomal 8-oxidation units are umoles N A D H produced/min/g wet weight. Results are expressed as mean (SEM). 35 are unable to utilize palmitate effectively (Sidell et al. 1987). The possibility that L C F A can be oxidized in elasmobranch red muscle by a carnitine-independent pathway, as in vertebrate liver, (Figure 1) appears unlikely. Palmitate was not oxidized at detectable rates by isolated dogfish red muscle (Table 2) or heart (Table 3) mitochondria, even under osmotically-stimulatory conditions, suggesting a lack of matrical CoA synthetase capable of activating L C F A , as is the case in mammalian skeletal muscles (Groot et al. 1976). Dogfish heart and red muscle also apparently lack the ability to utilize M C F A in a carnitine-independent manner. Neither tissue can utilize octanoate even under osmotically-stimulatory conditions (Tables 2 and 3). The acyl carnitine chain length preference of isolated muscle mitochondria suggests that medium chains may be more effective fuels than long chains. Low rates of palmitoyl carnitine oxidation by isolated mitochondria were evident under hypoosmotic conditions. The minimum in situ CPT activity which would be required to account for the observed rate of palmitoyl carnitine oxidation is about 2 nmole/min/g, a fraction of the detectable limit of CPT activity (Table 1, < 10 nmoles/min/g). (Complete oxidation of 1 nmole palmitate to C 0 2 yields 46 nmoles reducing equivalents ((2 reducing equivalents/B-ox idat ion cycle x 7) + (4 T C A cycle reducing equivalents/acetyl CoA x 8 acetyl CoA)). This translates to an oxygen consumption of 46 nmoles O consumed/nmole palmitate. The observed maximal rate of palmitoyl carnitine oxidation, 3.6 nmoles O/min/mg above control (Table 3), demands a minimum CPT rate of 0.08 nmoles/min/mg mitochondrial protein. Assuming dogfish red muscle possesses about 25 mg mitochondrial protein/g tissue, as in carp red muscle (Moyes et al. 1989), the minimum CPT activity required to account for the observed rates of oxygen consumption is 0.002 units/g wet weight.) The higher rates of octanoyl carnitine oxidation relative to palmitoyl carnitine suggest two things. Since it contains half the acetyl CoA content of palmitoyl carnitine yet stimulates twice the rate of oxygen consumption, octanoyl carnitine must be transported across the mitochondrial membrane at approximately 4-fold greater rates than palmitoyl carnitine. Secondly, as the chain length preference of CPT typically favours palmitoyl CoA over octanoyl CoA (e.g., rat liver mitochondrial CPT, Miyazawa et al. 1983), it is likely that the enzyme involved in octanoyl carnitine oxidation is specific for shorter chain fatty acyl 36 carnitines, possibly C A T . Mitochondria typically lack a carnitine acyl transferase which is specific for medium chain fatty acids (Clarke and Bieber 1981). Vertebrate heart and red muscle mitochondria generally oxidize pyruvate and palmitoyl carnitine at similar high rates (see Chapter 1). The inability of elasmobranch muscle mitochondria to oxidize acyl carnitines at high rates is unlike other vertebrate muscles, but similar to certain invertebrate muscles (e.g., heart mitochondria of squid, Ballantyne et al. 1981). Although the oxidation rates for octanoyl carnitine presented in Tables 2 and 3 appear low relative to pyruvate, the potential relevance of medium chain fatty acids as metabolic fuels deserves consideration. Such a pathway would explain the paradoxical H O A D and CPT activities, but the in vivo significance of these fuels would depend on availability. Plasma and tissue storage levels of S C F A and M C F A and triglycerides have not been determined in elasmobranchs as routine analyses either fail to extract or detect these fatty acids. Although elasmobranchs lack the plasma fatty acyl binding proteins required to transport L C F A (Fellows and Hird 1981), the solubility of fatty acids in plasma increases with decreasing chain length, obviating the requirement for a plasma binding protein for transport of M C F A . Furthermore the rate of transport of fatty acids across the cell membrane is dependent on the concentration of free fatty acid, not bound or total (see Stremmel 1987). Thus to achieve a given level of free (unbound) fatty acids, much lower total levels of the more soluble M C F A (mostiy free) are required relative to L C F A (mosdy bound). One obvious source of M C F A in elasmobranchs is peroxisomal B-oxidation, which shortens L C F A (Lazarow and deDuve 1976). Studies attempting to assess the relative quantitative importance of the mitochondrial and peroxisomal pathways in mammalian liver report conflicting results. Maximal activities of the two pathways are comparable in liver (Mannaerts et al. 1979) but studies using inhibitors of the mitochondrial pathway have lead to the conclusion that the peroxisomal pathway is of little quantitative importance in acetyl CoA production from palmitate or oleate (Mannaerts et al. 1979, Christensen et al. 1986, Hagve and Christophersen 1986). Conversely, Kondrup and Lazarow (1984) conclude that peroxisomes initiate about one third of palmitate oxidation in rat hepatocytes, using techinques which did not involve 37 metabolic inhibitors. Further support of a significant role for peroxisomal 8-oxidation in mammalian fatty acid oxidation comes from the marked inducibility of the pathway in vivo (Masters and Crane 1984). Mammalian heart and muscle have considerably lower maximal activities of fatty acyl CoA oxidase and peroxisomal 8-oxidation, 5-10% the rates of liver (Norseth and Thomassen 1983), suggesting that the peroxisomal pathway cannot compete with the mitochondrial pathway for acetyl CoA production, although it may be important in oxidation of fatty acids which are poor substrates for mitochondria (Norseth and Thomassen 1983). The potential contribution of dogfish peroxisomal 8-oxidation to muscle lipid metabolism is difficult to determine. As in mammals, the activity of peroxisomal 8-oxidation was much lower in muscle than liver, below detectable limits in heart or red muscle of dogfish, as well as the other species examined (Table 4). It should be noted that this lower limit of detection (< 5 nmoles N A D H (or acetyl CoA)/min/g) is close to the calculated mitochondrial rate of fatty acid oxidation of approximately 16 nmoles acetyl CoA produced/min/g (minimum red muscle CPT activity of 0.08 nmoles palmitoyl CoA/min/mg mitochondrial protein x 8 moles acetyl CoA/mole palmitate x 25 mg mitochondrial protein/g tissue). Elasmobranch liver peroxisomal 8-oxidation occurs at activities similar to teleost tissues, which are comparable to other studies, when adjusted for assay temperature (e.g., rainbow trout, Henderson et al. 1982). Interestingly, peroxisomal 6-oxidation in hagfish liver is below detectable levels. The capacity of elasmobranch liver for export of peroxisomally produced M C F A (as free fatty acids or medium chain triglyceride) is not known. In fish species which lack adipose tissue, such as elasmobranchs, the liver becomes more important in lipid storage, but the extent to which it replaces the metabolic role of adipose tissue as a hormone-sensitive source of plasma free fatty acids has not been established (Sheridan 1988). In summary, previous studies have shown that glucose is an effective fuel for skate cardiac metabolism (Driedzic and Hart 1984). While the capacity of the mitochondria to utilize pyruvate is clear, the presence of extrahepatic H O A D suggests fatty acids may be utilized to some extent. Low activities of CPT, coupled with an inability of mitochondria to utilize palmitate directly by a carnitine-independent 38 pathway under various metabolic conditions argue against a significant pathway for direct utilization of L C F A by elasmobranch extrahepatic tissues. While long chain fatty acyl carnitine is a poor substrate, medium chain fatty acyl carnitine is oxidized at higher rates. Consequently, it is likely that SCFA or M C F A could also support low rates of muscle activity if they were available to the mitochondria. However, based on relative rates of mitochondrial oxidation, it appears that carbohydrate and ketone bodies are the predominant fuels for extrahepatic tissues of elasmobranchs. Dramatic modifications of the conventional pattern of hepatic fatty acid metabolism are neccessary if elasmobranch steady state exercise is to be fuelled by ketone body oxidation. As discussed in Chapter 1, ketone body production in mammals occurs in a food-deprived state. Constitutive ketogenesis would require demand either fundamentally different hormonal or metabolic controls (i.e. signals that the animals is always "starving", regardless of dietary status) or inherent enzyme insensitivity to the endocrine/metabolite indicators of dietary status (i.e. enzymes that don't recognize signs of being well fed). Although elasmobranch peroxisomal 8-oxidation is similar to other fish species (Table 4), CPT activity is markedly lower (Table 1). It is possible that the enzyme is consituitively more active in vivo. CPT from dogfish liver mitochondria, as in mammals, is highly sensitive to malonyl CoA (Anderson 1990) but it is possible that malonyl CoA levels are maintained at lower levels in vivo. Alternately sustained ketogenesis could be acheived if liver free fatty acids are kept constantly elevated. Hepatic peroxisomal 8-oxidation may have a role in providing M C F A for muscle but it may also have an indirect role in hepatic ketogenesis. Acetyl CoA produced in peroxisomal 8-oxidation could be an important substrate for ketogenesis if given access to the mitochondrial ketogenic enzymes, although this route is not important in mammals. 39 C H A P T E R 3: D O F A S T F ISH H A V E F A S T M U S C L E M I T O C H O N D R I A ? P R E F A C E This chapter is currently in press in Can.J.Zool. (Moyes, OA.Mathieu-Costello, R.W.Br i l l , and P.W.Hochachka). Tuna mitochondrial ultrastructure was performed by O.A.Mathieu-Costello as part of the collaborative project. Her data are summarized in the discussion of this chapter (Table 8). R.W.Bri l l and P.W.Hochachka provided funding, laboratory facilities, logistic support and critical review of the manuscript. I N T R O D U C T I O N An important determinant of the maximal aerobic power output of a muscle is the rate at which its mitochondria can produce ATP. Modest increases in aerobic capacity are observed in individual muscles with endurance training. Much greater differences in muscle aerobic capacity are observed when comparisons are made between muscle types (ie. red, white, cardiac) or between species. Such comparisons suggest that there are two general mechanisms by which increased aerobic A T P production capacities are achieved in muscle. 7. More tissue (i.e. increased muscle mass/kg body weight). II. Higher mitochondrial capacity/g tissue. This can be achieved both quantitatively and qualitatively. Higher mitochondrial volume density (Vv(mt,f)) packs more mitochondria into a given tissue volume. Qualitative differences in mitochondria make more effective use of the intracellular volume devoted to mitochondria. One type of qualitative difference is more closely packed inner membrane cristae, measured as m 2 cristae surface area/cm3 mitochondrial volume (Sv(im,m)). Because the protein content of mitochondrial inner membranes is extemely high (Srere 1985), more closely packed cristae allows more mitochondrial membrane protein per unit mitochondrial volume. A second type of qualitative difference involves changes in the nature or efficiency of the mitochondrial enzymes, which would be manifested as an increased ATP production/mg mitochondrial protein. Qualitative differences in mitochondria between high and low performance tissues and species are not well established. It has been suggested as part of the symmorphosis hypothesis that individual 40 mitochondria from mammalian skeletal muscles are fundamentally similar regardless of tissue aerobic capacity (Hoppeler and Lindstedt 1985). Similar oxygen consumption rates are indeed observed with in vitro mitochondrial preparations from red and white muscles of mammals (Pande and Blanchaer 1971, Schwerzmann et al. 1989). There is evidence for relatively minor changes in the nature of the mitochondria in response to endurance training, where not all mitochondrial enzymes increase to the same extent (see Holloszy and Coyle 1984). However when isolated mitochondria from trained and control rats are compared, rates of oxidation of physiological substrates (/mg mitochondrial protein) are not affected by training (Davies et al. 1981). Qualitative differences in mitochondria are more obvious when comparisons are broadened to include non-mammalian vertebrates. A series of studies by Else and Hulbert (Else and Hulbert 1981, 1983; Hulbert and Else 1989) demonstrated that mammalian mitochondria have more densely packed cristae than mitochondria from the same tissues of similarly sized reptiles. Differences in muscle mitochondrial capacities/mg protein have also been observed. Red muscle mitochondria from elasmobranchs (Moyes et al. 1990) demonstrate 2-fold higher rates of oxygen consumption than mitochondria from red or white muscle of teleosts (Moyes et al. 1989). Heart mitochondria tend to show higher rates of oxygen consumption/mg protein than do skeletal muscle mitochondria (e.g. Krieger et al. 1977 vs Palmer et al. 1977). Furthermore, two intracellular populations of mitochondria (subsarcolemmal, interfibrillar) appear to have different oxidative capacities in both skeletal muscle (Kreiger et al. 1977) and heart (Palmer et al. 1977). Apart from questions of oxidative capacity, increased aerobic performance appears to be accompanied by a change in tissue fuel preference. Part of the transition to higher aerobic work capacities, induced by endurance training (Holloszy et al. 1985) or natural selection (Driedzic et al. 1987), appears to be a greater reliance on fatty acids. The extent to which these changes are facilitated by specialization of the individual mitochondria has not been thoroughly assessed. Relative rates of oxidation of pyruvate and palmitoyl carnitine do not change in mitochondria from endurance trained rats 41 (Davies et al. 1981). Training in fish increases tissue activities of H O A D in both red and white muscle, with little change in other mitochondrial enzymes, suggesting a greater reliance on fatty acids (Johnston and Moon 1980a, 1980b). Tne present study compares the oxidative properties of heart (atrium, spongy and compact ventricle) and skeletal muscle (red and white) from the high performance species skipjack tuna (Katsuwonuspelamis) and a low performance species, the common carp (Cyprinus carpio). The goal was to assess the relative importance of mitochondrial differences in determining the aerobic capacity of fish muscle. M A T E R I A L S A N D M E T H O D S Animals Skipjack tuna (Katsuwonuspelamis), were purchased from local commercial fishermen. Fish from 1-3 kg body weight and of undetermined sex were held for up to 4 days in 10 m diameter tanks supplied with continuously flowing sea water (25 +. 1°C) at the Kewalo Research Facility, National Marine Fisheries Service, Honolulu. Carp (Cyprinus carpio) of 1-2 kg body weight and of both sexes, were netted locally and held for up to three months in outdoor tanks supplied with continuosly flowing fresh water (10-15°C) at the Dept. of Zoology, University of British Columbia. Carp were fed trout chow twice weekly. Mitochondrial isolation Fish were netted, quickly stunned by blow to the head, and decapitated. A 2 cm steak was taken from the tuna immediately posterior to the anus. Tuna red and white muscle samples were collected from both sides of the vertebral column. Carp red and white muscle was sampled at the level of the dorsal fin (posterior edge). Pink muscle was carefully sheared from the red muscle bands. Whole ventricles from single tuna or 2-3 carp were used for isolated mitochondrial studies. Mitochondria were prepared as described for elasmobranch mitochondria using a homgenization medium composed of 20 mM HEPES, 140 mM KC1, 10 m M E D T A , 5 m M MgC l 2 , 0.5% BSA at pH 7.1. Initial centrifugations of 5 min at 42 1400 g (4°C) were used to sediment undispersed tissue. Mitochondria were collected and washed by 7 min centrifugations at 9000g (4°C). Aliquots of the resuspended pellet (4-8 mg protein/ml) were frozen immediately in liquid nitrogen for enzyme assays. Mitochondria were used within 1.5 hours for the oxygen consumption experiments. Mitochondrial oxygen consumption Incubations of tuna and carp mitochondria were performed at 25°C and 15°C, respectively. Small volumes of mitochondrial suspensions (25 ul tuna heart, 50 ul tuna red muscle, 100 ul carp heart, red and white muscle) were added to 2 ml incubation medium of the following composition: 140 mM KC1, 20 mM H E P E S , 5 m M NajHPO*, 0.5% bovine serum albumin, pH 7.3 at 20°C. Oxygen consumption rates were monitored by Clark-type electrode. The RCR values were determined on each preparation by addition of carbon substrates followed 4-6 min later by 0.5 m M A D P . To determine oxidative rates, mitochondria were given 0.1 m M malate and 0.5 m M A D P , then incubated 1-3 min until oxygen consumption reached a low, linear rate. At this point, saturating amounts of substrates were added to the cuvettes to elicit the State 3 oxygen consumption rates. Enzyme assays Tissues for enzyme analyses were quickly collected, frozen in liquid nitrogen, and stored on dry ice. Atrium, compact myocardium and spongy myocardium were separated prior to freezing. A l l assays were performed within 7 days of tissue collection. Tissue samples were weighed and added to 6 vol (w/v) ice cold homogenization medium (20 mM HEPES, pH 7.4, 1 m M E D T A , 0.1% Triton X-100). Homogenates were prepared by dicing the tissue, sonicating (1 burst of 10 sec on Kontes sonifier), homogenizing (3 bursts of 10 sec on Ultra-Turrax homogenizer) and sonicating (3 bursts of 10 sec). Particulate matter was sedimented by centrifugation (5 min at 9000g, 4°C). Mitochondrial suspensions were diluted with the same homogenization medium (1/9 v/v) and sonicated (3 bursts of 10 sec). Activities lost in the sedimented pellet were not determined but are assumed to be minimal, as this extraction procedure routinely yields measurements of enzyme activity comparable to, or as much as 3-43 fold higher than, values previously reported for these species (e.g. Guppy et al. 1979, Sidell et al. 1988). Maximal enzyme activities were determined using a Perkin-Elmer Lambda 2 spectrophotometer interfaced with an IBM computer using PECSS software. The cuvet holder was at a temperature of 2S°C for both species. A l l assays were performed in duplicate, as follows: Citrate synthase (CS). 20 mM Tris-HCl (pH 8.0), 0.2 m M D T N B , 0.3 m M acetyl CoA, 0.5 mM oxaloacetate. Oxaloacetate was omitted for the control. Control rates were typically <5% CS rate. Carnitine palmitoyltransferase (CPT). 20 m M Tris-HCl (pH 8.0), 0.2 m M D T N B , 0.1 m M palmitoyl CoA, 5 m M L-carnitine. Carnitine was omitted for the control assays. CPT rates were typically 2- to 3-fold greater than control rates. Hexokinase (HK). 20 m M imidazole (pH 7.4), 1 m M A T P , 0.5 m M N A D P + , 5 m M glucose, 5 m M MgCI 2 ,5 m M dithiothreitol and excess glucose-6-phosphate dehydrogenase. Control assays omitted glucose. H K was typically 2- to 3-fold greater than the control rate. R E S U L T S Mitochondrial oxidation The results of the isolated mitochondrial studies are summarized in Table 5. In both species maximal oxygen consumption rates were several fold higher in mitochondria from heart than skeletal muscle; 4-fold in tuna, 3-fold in carp. In each of the tissues examined, pyruvate was oxidized at the highest rates among all substrates tested. Tuna red muscle mitochondria oxidized palmitoyl carnitine and pyruvate at similar rates, as did carp red muscle mitochondria. Heart mitochondria of both species oxidized palmitoyl carnitine at lower rates than pyruvate (55% lower in tuna, 30% lower in carp). This pattern of pyruvate preference over fatty acids also occured in carp white muscle. Comparisons of mitochondrial respiration rates between species is complicated by the differences in assay temperatures (25°C for tuna, 15°C for carp), which were chosen to mimic acclimation temperatures. Carp red muscle mitochondria demonstrate a Q 1 0 of 1.5, using lauroyl carnitine as substrate (Moyes et al. 1988). It is not known if a similar Q 1 0 is applicable for all substrates or if the rates would 44 Table 5. Oxidation of various substrates by mitochondria from heart and skeletal muscle of carp and tuna. T U N A C A R P heart red heart red white R C R 14.2 (2.8) 6 14.9 (2.2) 6 >20 6 15.1 (1.5) 7 7.8 (0.6) 8 malate 20(2) 6 8(2) 7 24(3) 6 6(1) 8 5(1) 9 pyruvate 403 (41) 6 106 (12) 7 146 (5) 6 55(7) 6 52(7) 7 palmitoyl earn. 178 (29) 6 104 (12) 7 103 (4) 6 54(6) 7 31(4) 7 lauroyl earn. 150 (14) 6 87 (11) 6 89(4) 6 56 (8) 7 36(4) 7 octanoyl earn. 83(4) 6 50(6) 6 49 (4) 6 45(9) 7 18 (2) 6 acetyl earn. 87 (10) 6 36(4) 6 91(3) 6 acetoacetate 58 (5) 5 16(2) 3 9(1) 5 8(1) 5 glutamate 53(5) 4 23(4) 4 33 (6) 6 20(4) 6 Results are expressed as mean (SEM) in nmoles O/min/mg mitochondrial protein with n (number of preparations) below. Tuna data was collected at 25°C; carp at 15°C. 45 change in such a manner if acclimation temperature was 25°C. For the purposes of this study a Q 1 0 of 1.5 is assumed for all substrates with the recognition that this may be an oversimplification. With these reservations in mind, it appears that carp and tuna mitochondria are similar in most respects within tissue types. While red muscle mitochondria respire maximally at similar rates in each species, maximal oxygen consumption (pyruvate) by cardiac mitochondria is almost 2-fold higher in tuna man carp. Acetyl carnitine oxidation by carp heart mitochondria is about 150% that of tuna. Glutamate oxidation by carp red muscle mitochondria is about twice that of tuna. Tissue enzymes A summary of tissue enzyme activities is presented in Table 6. Citrate synthase activity/g wet weight is used as an index of mitochondrial capacity. Of all tissues examined, the highest activities were observed in tuna atrium. Within tuna heart, CS activity is ranked atrium:spongy myocardium:compact myocardium (100:85:72). Within carp heart, CS activity is ranked spongy myocardium:compact myocardium:atrium (100:67:42). Comparing the two species, tuna CS is higher in all tissues examined (compact myocardium (2.1x), spongy myocardium (1.8x) and red muscle (1.6x)) but particularly atrium and white muscle (4.5-5x). CS activities, expressed as units/g wet weight (Table 6) and units/mg mitochondrial protein (Table 7), allow the calculation of mg mitochondrial protein/g tissue (Table 7). Tuna tissues have a higher mitochondrial protein content than those of comparable carp tissue (30% higher in heart, 80% in red muscle). The maximal ventricular oxygen consumption can be estimated from mitochondrial protein content (Table 7) and oxygen consumption/mg protein (Table 5). When the differences in assay temperature are considered, tuna tissues have 130-140% higher maximal oxygen consumption/g than the respective carp tissues. CPT activities were also higher in tuna tissues; atrium, 4.2x; compact, 7.4x; spongy, 3.2x; red muscle, 3.4x. Within tuna, CPT activities were similar in red muscle, compact and spongy ventricle, each being approximately twice that of atrium. CPT in carp atrium and compact ventricle were each about half 46 Table 6. Tissue activities of citrate synthase (CS), carnitine palmitoyl transferase (CPT), and hexokinase (HK) of tuna and carp. CS/g CPT/g mU CPT H K / g U H K U CS U CPT T U N A Atrium (n=7) 87.88 1.26 14 13.55 10.8 (2.45) (0.18) (2) (0.42) (1.4) Compact (7) 63.37 2.52 40 6.66 2.6 (4.15) (0.26) (3) (0.68) (0.20) Spongy (7) 74.58 2.51 35 8.10 3.2 (5.76) (0.25) (6) (0.78) (0.29) Red muscle 79.8 2.40 30 2.03 0.85 (7) (2.15) (0.05) (0.4) (0.16) (0.05) White muscle 16.1 N D 1.06 -(7) (0.95) (0.1) C A R P Atrium (6) 18.12 0.30 17 6.97 23.2 (1.27) (0.022) (1) (1.24) (3.6) Compact (5) 29.60 0.34 12 10.45 30.7 (1.88) (0.012) (1) (1.12) (4.0) Spongy (5) 43.32 0.78 21 11.07 14.2 (6.35) (0.042) (5) (0.54) (1.3) Red muscle 49.00 0.71 14.5 0.55 0.77 (6) (4.4) (0.06) (0.06) -White muscle 3.50 N D _ _ (5) (0.2) Values are mean (SEM) where 1 U/g is 1 umole substrate used/min/g. A l l enzymes were assayed at 25°C. 47 Table 7. Mitochondrial enzymes assayed in isolated mitochondria of carp and tuna. CS/mg CPT/mg mg mito. V0 2MAX protein/g (umoles 0 2 /min x g) T U N A Heart Red muscle C A R P Heart Red muscle White muscle 1.94 (0.16) 1.64 (0.06) 1.34 (0.11) 1.80 (0.16) 1.13 (0.08) 0.030 (0.002) 0.039 (0.003) 0.016 (0.001) 0.015 0.004 35.6 48.7 26.9 27.2 3.09 7.1 2.6 2.0 2 0.75 2 0.0812 Enzyme units are umoles substrate converted/min/mg mitochondrial protein determined for each species at 25°C and expressed as mean (SEM) for 6 animals. 1 CS/g (mean of spongy and compact)/ CS/mg 2 estimates of oxygen consumption rates for carp are at 15°C 48 that of spongy ventricle and red muscle. In oxidative muscles, H K / C P T may reflect the fuel preferences of the tissue. Red muscle H K activities are several fold lower than the heart tissues. In tuna and carp red muscle, H K is slighdy lower than CPT resulting in H K / C P T ratios less than unity. HK/g is similar in the heart tissues of each species but, unlike red muscle, it is several fold higher than CPT. H K / C P T in tuna ventricle is approximately 3 but in carp this ratio is between 14 (spongy) and 30 (compact). Mitochondrial enzymes Direct comparisons of mitochondrial enzymes (activity/mg mitochondrial protein) between species (Table 7) is possible because all enzymes were assayed at 25°C. No clear pattern emerges in CS activity/mg protein between tissues or species. Activities are marginally higher in heart than red muscle in tuna but the reverse is seen in carp. Activities in tuna heart are slighdy higher than carp heart. Activities are similar in red muscles of each species. CPT is often used as an index of the capacity for fatty acid utilization by a tissue. This study demonstrates that differences in CPT activity are not neccessarily accompanied by differences in the maximal mitochondrial capacity to utilize fatty acids. CPT/mg protein is 2-2.5-fold higher in tuna relative to the same tissues in carp, but carp and tuna have similar tissue-specific mitochondrial palmitoyl carnitine oxidation rates, when differences in assay temperature are considered. Although CPT activities (U/mg protein) were 30% higher in tuna red muscle than tuna ventricle, mitochondria from heart oxidized palmitoyl carnitine at 70% greater rates than red muscle. A similar situation occurs in carp, where 2-fold greater palmitoyl carnitine oxidation rates in heart occurred despite similar activities of CPT/mg protein. The reverse situation occurs in rat heart mitochondria from two intracellular populations. Interfibrillar mitochondria oxidize palmitoyl carnitine and palmitoyl CoA+carnitine at 55-65% greater rates than subsarcolemmal mitochondria despite similar activities of CPT (Palmer et al. 1977). The maximal mitochondrial capacities for palmitoyl carnitine oxidation rates utilize only a fraction of the measured CPT activity. For instance, in situ activity of 0.01 units CPT/mg mitochondrial protein can provide 49 enough fatty acid substrate to result in an oxygen consumption of 460 nmoles O/min/mg mitochondrial protein, well beyond the maximal rates observed in vitro. Even this may be an underestimate, as the activity of CPT in the direction required for palmitoyl carnitine oxidation is approximately 9-fold greater than that in the direction which is assayed by the DTNB method (Palmer et al. 1977). The metabolic advantages of higher CPT/mg mitochondrial protein in the different tissues and species may be kinetic in nature and be related more to regulation or substrate selection than the actual in situ capacity. DISCUSSION Aerobic capacity There are two potential strategies to increase tissue aerobic capacity (i) more aerobic tissue/ kg body mass or (ii) higher mitochondrial capacity/g tissue (increased V v(mt,f), increased Sv(im,m) or increased oxidation/mg protein). When tuna and carp are compared, each of these strategies occur to some extent and the relative importance of each depends on the tissue. In general, it appears that highly aerobic tissues (heart, red muscle) differ between species primarily in functional mass and to a lesser extent in mitochondrial capacity/g. In contrast, muscles which are mitochondria-poor in carp (white muscle, atrium) have several fold higher mitochondrial capacities/g in tuna. The interspecies differences in red muscle mitochondrial properties are modest. Substrate oxidation rates/mg mitochondrial protein are similar between species, when differences in assay temperature are considered. Tuna red muscle, however, has about 80% more mitochondrial protein/g tissue than carp. As the Vv(mt,f) found in tuna red muscle (32%) is within the range found in other teleosts (25-38%, see Johnston 1981), the higher mitochondrial protein/g must be manifested as more densely packed cristae. The intracellular volume devoted to mitochondria in fish red muscle may approach the spatial limit allowed by cells that must also perform mechanical work (Weibel 1985, Hochachka 1988). Higher mitochondrial volume densities are observed in tissues such as heater organ of billfish (Block 1986), which demonstrate high metabolic rates but do not perform mechanical work. If such a constraint occurs in fish muscle, adaptations toward increased mitochondrial capacity/ g tissue may 50 neccessarily involve increased Sv(im,m). Cristae packing in tuna red muscle is exceptionally high for a vertebrate skeletal muscle. This avenue for increasing tissue aerobic capacity may also be limited in tuna red muscle. The value of 63-70 nrVcm3 (Table 8) reported for tuna red muscle S^im.m) in tuna red muscle is greater than the range reported from skeletal muscles of Antarctic fish (25-37 nrVcm3, Archer and Johnston 1991), a wide variety of mammals (20-40 m 2 /cm 3 , Hoppeller and Lindstedt 1985), hummingbird flight muscle (58 nrVcm 3, Suarez et al. in press), mammalian and reptilian hearts (38-66 nrVcm3, Else and Hulbert 1983). Srere (1985) calculated that a Sv(im,m) of 83 m 2/cm 3 is the maximum degree of cristae packing that will still allow enough matrix space for 2 average sized Krebs cycle enzymes, each in contact with an opposing membrane. Thus it appears that the only structural strategy available to tuna to increase the total aerobic capacity of red muscle by several fold is to increase recruitable tissue mass. Active, pelagic fish generally possess more red muscle than benthic or sluggish fish (Greer-Walker and Pull 1975). Higher aerobic capacity in skeletal muscle could also be achieved by utilizing white muscle during steady state swimming. The limits to the use of white muscle at low swim speeds are based on innervation pattern and muscle ultrastructure (Johnston 1981). The fiber orientation neccessary in white muscle to power burst exercise compromises its ability to function at lower swimming speeds (Rome et al. 1988). Electro-physiological studies suggest tuna white muscle is indeed recruited at sustainable swim speeds (Brill and Dizon 1979) as in several other fish species (Johnston 1981) but not carp (Johnston et al. 1977). Previous metabolic and ultrastructural studies support an aerobic role for tuna white muscle (Guppy et al. 1979, Hulbert et al. 1979). It is highly vascularized and contains significant stores of intracellular l ipid, a fuel which can only be used aerobically. In the present study, tuna white muscle mitochondrial content, based on citrate synthase/g, was found to be 5-fold higher than carp, supporting the suggestion that tuna white muscle is used aerobically. The major differences between species in myocardial oxidative capacity/kg body weight are attributable to muscle mass. Mass alone affords tuna a 5-fold advantage in oxidative capacity/kg body 51 Table 8. Tuna muscle mitochondrial ultrastructure (from Moyes et al. in press). V v(m,f) SvOm.m) S^ im . f ) (%) (m2/cm3 mitochondria) (m 2/cm 3 tissue) V v(ms,f) Vv(mt,f) S¥(im,ms) S v(im,mi) Heart Compact 1.7 25.4 - 57.88 14.70 (0.1) (1.0) (1.57) Spongy 1.7 25.3 - 56.63 14.33 (0.5) (0.7) (1.36) Red muscle 12.3 32.4 63.20 70.61 21.89 (2.4) (1.9) (1.60) (1.67) White muscle 0.3 2.5 - 31.32 0.78 (0.2) (0.3) (1.66) Vv(m,f)-volume of mitochondria/volume fiber, espressed as % Vv(ms,f)-as above, for subsarcolemmal mitochondria Vv(mt,f)-as above, for total mitochondria Sv(im,m)-surface area of inner mitochondrial membrane/volume mitochondria SvOm .ms^as above, for subsarcolemmal mitochondrondria Sv(im,mi)-as above, for interfibrillar mitochondria. Sv(im,f)-surface area of inner mitochondrial membrane/volume tissue. 52 mass relative to carp (4.0 vs. 0.76 g ventricle/kg). Skipjack tuna ventricular mass is higher than in most fish species (Poupa et al. 1981, Sidell et al. 1987), close to the the range found in mammals (Poupa and Ostadal 1969, Driedzic et al. 1987). Higher mitochondrial capacity/g ventricle is also obvious in tuna, but this is of lesser quantitative importance than the increased ventricle mass. Tuna have more mg mitochondrial protein/g ventricle (30% greater) and a 2-fold higher oxidative capacity /mg mitochondrial protein than carp. These three factors combine to give tuna a 13-fold greater myocardial oxidative capacity /kg body mass, but much of this difference is due to ventricular mass. Compared to tuna red muscle, ventricle has lower Vv(mt,f) and lower Svfim .m). Although teleological, it is instructive to ask why tuna ventricle mitochondria are less packed than they could be, based on what is observed in red muscle. Large hearts are needed to obtain the high ventral aortic pressures observed in vivo (Brill and Bushnell in press). However, power output/g ventricle in tuna is similar to other fish species (Brill and Bushnell in press, Farrell et al. submitted) so selection for higher aerobic capacities/g tissue may have been minimal. As hemodynamic considerations probably determine atrial size, it is not surprising that the ratio of atrial mass to ventricular mass is similar in each species (15% in tuna, 18% in carp). As was the case with ventricle, the greater mass of tuna atrium (0.60 vs. 0.14 g atrium/kg body mass in carp) provides a 4-fold higher aerobic capacity. However tuna atrium also has an approximately 5-fold greater mitochondrial capacity/g than carp, based on CS/g tissue. The combined effect of higher mitochondrial capacity/g and increased tissue mass provides tuna atrium with a 20-fold greater aerobic capacity/kg body mass than carp. Studies using combined physiological, biochemical and morphometric techniques suggest that mammalian skeletal muscle mitochondria in vivo may operate close to the maximal rates obtained with isolated mitochondria in vitro (Schwertzman et al. 1989). In the case of tuna heart mitochondria, the VOJMAX we predict from in vitro studies is close to that estimated to be required by the heart at whole animal V0 2MAX. Myocardial oxygen consumption of trout swam at maximal sustained speed has been estimated to be 80 ul oxygen/min/kg body mass, corresponding to 95 ul (or 4.0 umoles) oxygen/min/g 53 ventricle (based on 15°C trout with 0.87 g ventricle/kg working at 7.2 mW/g, Graham and Farrell 1990). The maximal cardiac power output observed in skipjack tuna in vivo is 25 mW/kg body mass determined on spinally-blocked fish (Brill and Bushnell in press). Using trout data relating oxygen consumption and power output (Graham and Farrell 1990), this work rate would require a myocardial oxygen consumption of 3.3 umoles/min/g ventricle (based on 25°C tuna with 4 g ventricle/kg). Maximal mitochondrial capacity found in this study (7.1 umoles oxygen/min/g) is more than 2-fold greater than the maximal myocardial oxygen consumption rate observed in vivo. It has been suggested that the maximal cardiac output may be 1.5-2 fold greater than that observed with spinally-blocked tuna, close to 12.5 mW/g or 6.6 umoles 0 2 /min/g (Brill and Bushnell in press). If this were the case, the maximum mitochondrial capacity (7.1 umoles 0 2/min/g) is similar to that expected to be required under these conditions. This is unlike the situation in cat skeletal muscle where in vitro rates of pyruvate oxidation determined in isolated mitochondria could not account for the rate of 0 2 consumption known to occur in the tissue in vivo (Schwerzmann et al. 1989). If the assumptions regarding myocardial power output are sound, then oxygen or substrate delivery, N A D H / N A D + ratio or availability of A D P may prevent ventricle mitochondria from reaching their maximal capacity in vivo. Given the high rates of lactate release by isolated tuna hearts (Farrell et al. submitted), it is likely that its impressive glycolytic capacity would augment aerobic A T P production at such high power outputs. Fuel preference One of the consequences of training in mammals is greater reliance on fatty acids as fuels. Although a considerable part of this shift is related to fatty acid supply to the mitochondria, clear metabolic changes also occur at the muscle. Mitochondrial density in mammals increases with endurance training and fatty acid catabolic enzymes increase in parallel, resulting in greater enzyme activity/g tissue (Holloszy 1985). Endurance training of coalfish and brook trout causes increases in H O A D activities in red and particularly white muscle, suggesting an increased capacity for fat oxidation also occurs in fish (Johnston and Moon 1980a, 1980b). Analagous to the intra-tissue changes that occur with endurance 54 training is increasing dependence on fatty acids as fuel in hearts of species with higher cardiac power outputs. When resting power output was plotted against enzyme activity, Driedzic et al. (1987) found that glycolytic enzymes (e.g. hexokinase) plateaued at low power demands, whereas fatty acid catabolizing enzyme activity (e.g. CPT) continued to increase. Along this line, the HKVCPT ratios may indicate the relative importance of exogenous glucose and fatty acids as fuels. The data from Driedzic et al. 1987 show that heart HK7CPT ratios of birds (pigeon 1.7) and mammals (rat 1.1) are considerably lower than reptiles (fence lizard 7.3, turtle 33), amphibians (grass frog 7.8, mudpuppy 21) and fish (trout 10.9, ocean pout 12, sea raven 22, hagfish 28). In this study, H K / C P T of carp ventricular tissue is as high as in other poikilotherms (14-30) but in tuna, the ratios are closer to those found in homeotherms (approx. 3), due primarily to higher CPT rates. Part of the differences in CPT/g tissue observed are due to mitochondrial content, but there is also a difference in CPT per unit mitochondria. Comparison of tissues with widely different aerobic capacitites, from carp white muscle through to tuna ventricle, reveals a 10-fold difference in CPT/mg mitochondrial protein that correlates with mitochondrial content of the tissue (Figure 2). It is likely that the higher CPT activity per unit mitochondria confers kinetic advantages to favour fatty acid utilization. Taken together these data support the general hypothesis that more aerobic tissues rely more on fatty acids than glucose as fuel. Previous calculations suggest that tuna cardiac mitochondrial capacity is similar to that which would be required at whole animal V0 2MAX. If this mitochondrial capacity were realized in vivo, ventricular aerobic power output could reach 53 mW/kg body mass (14 mW/g ventricle) with pyruvate (from lactate, glucose) as substrate. However a maximum of only 24 mW/kg (6 mW/g ventricle) could be generated with fatty acids as primary fuel. It should be noted that this fat oxidizing capacity is sufficient to support the highest cardiac outputs measured in vivo (Brill and Bushnell in press). This difference is due to the 2-fold mitochondrial fuel preference of pyruvate over fatty acids. As the enzyme analyses suggest fatty acids are relatively more important in tuna ventricle, the fuel preference of these mitochondria for pyruvate over fatty acids may reflect the capacity to use lactate as a fuel for heart 55 metabolism. Lactate produced in tuna white muscle during exercise can reach 80-100 umoles/g (Guppy and Hochachka 1978) and appears to be partially released from the tissue post-exercise (Arthur et al. in prep.). Significant lactate release following exercise may occur in channel catfish (Cameron and Cech 1990) but not in salmon and flounder (Milligan and McDonald 1988). Arthur et al. (in prep.) found tuna blood lactate concentration did not fall below 10 m M even when the white muscle lactate returns to a low value. Given the high rate of mitochondrial pyruvate oxidation relative to fat, it is tempting to speculate that fat may be used to fuel myocardial metabolism at low cruising speeds whereas lactate may be used during or following high intensity exercise, when metabolic rate is high, blood lactate is elevated and increased cardiac outputs are required. 56 0 I 1 1 1 1 1 ' 1 i i 0 10 2 0 3 0 4 0 5 0 60 7 0 8 0 9 0 U C S / g t i s s u e Figure 2. Relationship between mitochondrial content (CS/g) and mitochondrial specific activity of CPT. A unit (U) represents the activity pf enzyme required to convert 1 umole of substrate to product in 1 min. Abbreviations:C-, carp tissue;T-, tuna tissue;-RM, red muscle; - W M , white muscle;-CM, compact myocardium;-SM, spongy myocardium. 57 C H A P T E R 4 : R E C O V E R Y M E T A B O L I S M O F T R O U T W H I T E M U S C L E : T H E R O L E O F T H E M I T O C H O N D R I A P R E F A C E This study was done in parallel with work done by P.M.Schulte. Some data are repeated in this chapter to facilitate comparisons. Her study characterized the changes in tissue metabolites with burst exercise and is currendy submitted to the Journal of Applied Physiology (P.M.Schulte, CD.Moyes and P.W.Hochachka). This chapter examines the influence of the changes in tissue metabolites on mitochondrial metabolism, specifically as it relates to recovery. I N T R O D U C T I O N High intensity exercise in mammalian white muscle is fuelled by breakdown of glycogen, with formation of lactate and accompanied by a tissue acidosis. Recovery metabolism involves reducing [lactate] to resting levels and repletion of glycogen stores. In mammals, if blood [lactate] is low, lactate is released and primarily oxidized in aerobic tissues, with a smaller proportion used as a glyconeogenic substrate (Brooks and Gesser 1980, Brooks 1987). If perfusion [lactate] is high, conversion of lactate to glycogen in situ (glyconeogenesis) becomes increasingly important, especially in the more glycolytic fiber types (Bonen et al. 1990, Pagliassotti and Donovan 1990). Recovery from such exercise bouts typically takes several minutes to an hour (e.g. Hermansen and Vaarge 1977). Fish burst exercise and recovery differs from the mammalian pattern in a number of important aspects. The changes in metabolites are greater and more prolonged than in mammalian white muscle. For instance, lactate can reach 145 umoles/g (skipjack tuna, Arthur et al. in prep). Salmon and flounder recovery requires 8 to 12 hr or more (Milligan and McDonald 1988). Also, a number of indirect studies suggest that white muscle lactate is metabolized primarily in situ. Injections of 14C-lactate into the blood remain primarily as lactate (skipjack tuna, Weber et al. 1986; flounder and salmon, Milligan and McDonald 1988). Lactate turnover cannot account for the changes observed in white muscle lactate (salmon and flounder, Milligan and McDonald 1988), suggesting the blood and muscle compartments do 58 not equilibrate. Studies using the glucose analogue deoxyglucose suggest that less than 10% of the glycogen resynthesized in white muscle following burst exercise in rainbow trout can be attributed to exogenous glucose uptake (T.G. West et al. unpublished). Furthermore, hepatocyte studies suggest that fish liver has insufficient capacity for gluconeogenesis from lactate to account for the rate of lactate disappearance (Walsh 1989, gulf toadfish; Buck et cd. submitted, skipjack tuna). Since there is a net conservation of carbohydrate (glucosyl units + 2xlactate) during exercise and recovery (Schulte et cd. submitted), it is clear that oxidation is not an important route of lactate metabolism post-exercise. The route for in situ glycogen resynthesis from lactate is unclear. White muscle lacks adequate activities of pyruvate carboxylase, one of the enzymes required for gluconeogenesis in liver (Opie and Newsholme 1967). Alternate routes could involve malic enzyme + P E P C K (Connett 1979) or reversal of pyruvate kinase (Dyson et cd. 1975). White muscle from most fish species lacks both pyruvate carboxylase and P E P C K (Cowey et cd. 1977, Hulbert and Moon 1978, Moon and Johnston 1980, Mosse 1980), suggesting pyruvate kinase reversal may be the more important route of glyconeogenesis in fish. Whatever the route of glycogen resynthesis, the ATP demands imposed by recovery metabolism (net glyconeogenesis) must be met by mitochondrial (aerobic) pathways. Comparison between the rates of recovery reported for various species (tuna > salmon > plaice, Arthur et al. submitted, Milligan and McDonald 1988) reveals an apparent correlation with the mitochondrial density of the white muscle. The present study investigates if the A T P producing capacity of the white muscle mitochondria limits the rate of glyconeogenesis in situ. Surprisingly, when the maximal ATP-producing capacity of isolated mitochondria is expressed per g tissue, it is in vast excess to the A T P demands of recovery metabolism. The influence of physiological changes in A T P / A D P , pH , phosphate and carbon substrates suggest that mitochondrial metabolism of fish white muscle is stimulated in recovery by the changes in [phosphate], not adenylates, and is influenced by pH. If fatty acids are available post-exercise, it is likely that fatty acid oxidation would fuel recovery metabolism, sparing pyruvate and lactate for glyconeogenesis. 59 M A T E R I A L S A N D M E T H O D S Animals Rainbow trout (Oncorhyncus mykiss) of both sexes 400-600 g were held in continuously flowing fresh water at seasonal temperatures. Animals were fed ad libitum 4 times a week. Exercise protocol The methodology involving exercise of fish is described in detail by Schulte et al. (submitted). Briefly, fish were exercised to exhaustion in a swim tunnel and either sacrificed immediately or allowed to recover for 2, 4, 8, or 24 nr. A anaesthetic (Somnotol) overdose technique was developed to quickly kill fish with little struggling in order to minimize changes in labile metabolites. Enzyme activities Tissue homogenates were prepared in 9 vol of the following medium: 100 m M potassium phosphate, 5 m M E D T A , 1 m M dichloroacetate, 0.1% Triton X-100 at pH 7.0. K F , often used in mammalian studies to inhibit PDH-kinase was not used as it inhibited P D H directly. Tissue was homogenized by 3 bursts of an UltraTurrax grinder followed by 3 bursts of a Kontes sonicator. Homogenates were centrifuged 10 min at lOOOOg, with the resulting supernatant used for assays of both P D H and citrate synthase. Activity of P D H was determined by measuring acetyl CoA-dependent acetylation of /?-nitroaniline in the presence of pigeon liver arylamine acetyltransferase. The coupling enzyme was purified from pigeon liver based on Tabor et al. (1953). Pigeon liver was frozen in liquid nitrogen and kept up to 2 months before extraction. Tissue (100 g) was homogenized in 200 ml of 5 mM H E P E S , 1 m M E D T A at pH 7.0. Homogenates were centrifuged 20 min at 12000g. The pellets were rehomogenized in 200 ml of buffer and recentrifuged. The combined supernatants were brought to 500 ml and added to 400ml ice-cold acetone. After centrifugation (10 min at 10000 g), the combined supernatants were added to IL acetone and centrifuged (5000 g for 10 min). The pellet was dissolved in 20 ml water for 60 min at 4°C and added to 5 g alumina C r (pelleted from 50 ml suspension-Sigma Chem Co.). After 1 hr of swirling 60 (2Hz) the suspension was centrifuged 7 min at lOOOg. The pellet was washed twice with 100 ml water before elution with 250 ml potassium phosphate buffer (100 m M , pH 7.7) in 4 aliquots. The enzyme was concentrated by Amicon Centriprep units and frozen (-80°C) until needed. Each purification was checked for contaminating P D H activity and none was detected. P D H activity in tissue extracts was assayed using a Perkin-Elmer Lambda 2 spectrophotometer, interfaced with an IBM computer using PECSS software to calculate enzyme activities. The 1 ml assay mixture contained 1 mM M g C l 2 , 0.5 m M E D T A , 0.5 m M cocarboxylase, 0.15 m M coenzyme A , 2 m M N A D * , 5 mM 6-mercaptoethanol, 0.1 m M /7-nitroaniline, 200 mU arylamine acetyltransferase, 40 ul homogenate, 100 mM Tris (pH 7.7). Reaction was initiated by addition of 1 m M pyruvate and was linear for at least 5 min. Activity was measured within 30 min of homogenization. Attempts to fully activate fish muscle P D H by purified pig heart PDH-phosphatase were unsuccessful. To standardize for inter-individual differences in mitochondrial density, P D H activity is expressed against CS activity determined in the same homogenateas described in Chapter 3. Al eijme assays on tissues were performed at 25°C to obtain maximal sensitivity of the P D H assay and minimize coupling enzyme requirements. Q 1 0 determinations were performed to allow conversion between tissue enzyme analyses and mitochondrial studies performed at 15°C. Isolated mitochondrial studies Trout were killed by blow to the head. Approximately 40 g white muscle were collected from an area extending from the dorsal fin to 3 cm anterior to the caudal peduncle, including all tissue dorsal to the lateral line. After skinning the tissue, great care was taken to shear off any superficial red muscle. Tissue was placed on an ice-cold petri dish, chopped with razor blades, divided into 3 portions and added to 3 Potter-Elvejhem homogenizers, each containing 30 ml isolation buffer. The composition of isolation buffer is as described previously (Chapter 3). Tissue was dispersed by 2 passes of a loosely fitting pestle followed by 3 passes of a more tighdy fitting pestle. Homogenates were combined and diluted to 500 ml with more isolation buffer and centrifuged 3 min at 2600g. The supernatants were poured through 4 61 layers of cheesecloth and recentrifuged 5 min at 2600g. The supernatant was poured through 8 layers of cheesecloth and centrifuged 10 min at 9000g. Mitochondrial pellets were resuspended in 10 ml isolation medium (minus BSA) and centrifuged 5 min at 8000g. The mitochondrial pellet was resuspended in 1-2 ml BSA-free isolation medium and used within 60 min. Mitochondrial oxygen consumption Determinations of the rate of oxidation of physiological substrates was as described previously (Chapter 3) at pH 7.3 and 15°C. The oxygen consumption rate was also determined as a function of A T P / A D P ratio. Assays were performed with 0.3-0.5 mg protein/ml assay medium. Injections of 7.5 m M A T P , 10 m M glucose, then increasing hexokinase levels were used to obtain A T P / A D P ratios close to those observed in recovering fish (Schulte et al. submitted). Aliquots (500ul) were removed from parallel incubations and added to 500 ul ethanol for deproteinization. The supernatants (10 min at 15000g) were frozen for A T P and A D P determinations by H P L C as described (Schulte et al. submitted). The influence of M g C l 2 was determined at subsaturating [ADP]. Mitochondria were added to a cuvet containing assay medium (minus BSA) , 1 m M pyruvate, 0.1 mM malate, 150 U hexokinase with and without 5 mM MgC l 2 . The high activity of hexokinase was used to preclude an indirect effect of M g 2 + due to M g - + -dependent stimulation of hexokinase. Reactions were started by addition of 5 or 2uM A D P . The A D P levels were chosen to elicit submaximal rates which were dependent on A D P transport rate into the mitochondria, analagous to high A T P / A D P ratios. The actual A T P / A D P ratio was not determined. In experiments where a hexokinase ATP-regenerating system was used, A T P production rates were determined by assaying glucose-6-phosphate + fructose-6-phosphate production. Because such high activities of hexokinase were used, the relatively low contamination by phosphoglucoisomerase became significant. Approximately 10% of the glucose-6-phosphate was converted to fructose-6-phosphate. Mitochondrial incubations (0.9ml) were added to 0.1 ml 70% perchloric acid and centrifuged 5 min at 15000g. Aliqouts of the supernatant were neutralized by addition of 1 M Tris (40 m M final) and 10 M 62 K O H . Glucose-6 phosphate + fructose-6-phosphate were assayed in 50 m M Tris (pH 8.0) and 1 mM N A D P + . The reaction was started by addition of 1 U glucose-6-phosphate dehydrogenase + 1 U phosphoglucoisomerase and was complete within 2 min. Mitochondrial pyruvate oxidation Pyruvate oxidation was assayed using 1 4C(l)pyruvate. As the radiolabelled carbon is removed in the decarboxylation reaction of the P D H complex, 1 4 C 0 2 production represents the flux through P D H in situ. This technique was used to examine the influence of a number of potential effectors of mitochondrial metabolism. Also, some comparisons were performed between mitochondria from trout white muscle and carp red muscle. Carp mitochondria were prepared as described in Chapter 3 and assayed as described for the trout preparation. Small volumes of mitochondria were added to the assay medium used for oxygen consumption experiments to a total volume of 2 ml. Incubations were performed in 20ml scintillation vials. A 5 min preincubation period was sufficient time to allow the vials to be capped with rubber stoppers fitted with plastic center wells. Reaction was started by injection of lOOul of pyruvate ( 1 2 C+ , 4 C; 0.25 uCi/umole) through the rubber cap and vials were shaken (1 Hz) at 15°C for 10 min, unless noted otherwise. Incubations were terminated by a 200ul injection of 70% perchloric acid. Hyamine hydroxide (150ul) was added to the center well and vials were shaken 90 min at room temperature to collect 1 4 C 0 2 . Filter papers were added to 10ml of ACS-2 (Amersham), with 0.1% acetic acid to prevent background colour formation and eliminate chemiluminescence. Samples were counted for 10 min in a L K B 1214 RackBeta scintillation counter with programmed quench correction. A l l assays were done in at least duplicate. A similar protocol was used to determine P D H activities with solubilized mitochondria. The assay components were as descibed for tissue P D H analysis except /--nitroaniline and arylamine acetyltransferase were omitted and coenzyme A was elevated to 0.6 m M to prevent significant depletion in the longer incubations. Decarboxylation of 14C(l)-pyruvate was measured as before. The reaction was shown to be linear for at least 10 min using P D H that was inactivated by 20 min incubation of 63 mitochondria with SO u M palmitoyl carnitine. Routinely assay incubations were run for 6 min in at least duplicate. Studies examining the kinetics of pyruvate oxidation by intact mitochondria were performed under the conditions described above. Oxygen consumption rates provided estimates of pyruvate oxidation which were used to establish the highest mitochondrial concentration which would not deplete pyruvate by more than 10% at the lowest [pyruvate] used in the radiolabel studies. Preliminary studies showed the rate of pyruvate oxidation was linear with respect to [protein] over the ranges used in this study and linear for at least IS min. The influence of A D P availability was examined under two conditions. Experiments in State 4 (no added ADP) used high [protein] to obtain adequate ' *C0 2 collection. State 3 experiments (+ADP) were performed with 0.3 mM A D P , which was not substantially depleted because mitochondria were diluted and incubations were limited to 10 min. In some experiments lactate (40 mM) was added 5 min prior to addition of pyruvate. Statistics Where required, data were compared by A N O V A with Tukeys test post hoc (a=0.05). RESULTS Mitochondrial pyruvate oxidation i: kinetics During recovery from burst exercise tissue [pyruvate] increases several fold (Schulte et al. submitted). The effects of physiological changes in [pyruvate] on the oxidation of pyruvate, and consequendy lactate are shown in Figure 3. At low respiration rates (State 4) the oxidative pathway was saturated with pyruvate well below physiological levels, with an apparent K m estimated to be below 5 u M . The maximal State 4 P D H flux rates were approximately 1 nmole/min/mg protein. When pyruvate oxidation rates were examined in State 3, the maximal flux rates increased more than 20-fold, and the kinetics showed marked changes. A much higher [pyruvate] was needed to saturate the oxidative pathway. The apparent Km increased to 37.4 (2.5 SEM) uM. Comparisons with carp red muscle mitochondria (Fig. 4) were used to determine if the pyruvate 64 120 100 80 60 40 20 o 0 E E 120 •»- 100 o 80 c o 60 o 40 *x w n 20 w > E 0 a! 120 0.2 0.4 0.6 0.8 1.0 [pyruvate] (mM) Figure 3. Kinetics of pyruvate oxidation by trout white muscle mitochondria. The values are mean (SEM) determined in 10 min incubations, expressed as a per cent of the rate determined with 1 m M pyruvate. The absolute rates for each condition are as follows (mean (SEM,n), nmoles , 4C(l)-pyruvate decarboxylated/min/mg protein); State 4 (upper), 0.82 (0.08,5): State 3 (middle), 23.4 (3.0,4): State 3 with 40 mM lactate (lower), 19.5 (1.4,4). 65 O L_ 1 4 0 13 1 2 0 O o CN 1 0 0 o V ' 8 0 c o • 6 0 D "x 4 0 O <D D 2 0 > —* i i_ >s Q. 0 carp red musc le State 3 0 0.1 0 .2 0.3 [pyruvate] (mM) 0 .4 0 .5 Figure 4. Kinetics of carp red muscle mitochondrial pyruvate oxidation. N=3 animals. The apparent Km was not determined, but is estimated to be below 5 uM pyruvate, the lowest concentration tested. 66 kinetics were a property of fish muscle or particular fiber type. Carp red muscle in State 3 demonstrated kinetics similar to those observed in trout white muscle in State 4 i.e. not dependent on [pyruvate] over the physiological range. It has been reported that lactate competes for the rat heart mitochondrial pyruvate transporter, but with relatively low affinity (Km 12 m M , Halestrap 1978). Since fish develope extremely high post exercise lactate levels that remain for long periods (Milligan and Wood 1987) it was of interest to examine whether physiological lactate levels could inhibit pyruvate oxidation through transport competition. Pyruvate transport is potentially limiting only at high flux rates and low [pyruvate]. We examined the influence of 40 mM lactate under these conditions. No effects of 40 m M lactate on pyruvate oxidation were observed (Fig. 3). The apparent Km for pyruvate was 35.8 (3.9 SEM) u M , compared to 37.4 (2.5) u M in the absence of lactate. Mitochondrial pyruvate oxidation ii: effects of fatty acid oxidation The effects of fatty acid availability were examined at high (200 uM) and low (20 uM) [pyruvate], each at high (State 3) and low (State 4) respiratory rates (Fig. 5). The control data also illustrate that the rate of pyruvate decarboxylation by isolated mitochondria was linear for the 10 min period used in the kinetics studies. P D H flux within mitochondria is dependent on covalent modification of P D H activation state (phosphorylation-dephosphorylation) and allosteric control of the active enzyme (Hansford 1980). In mammals, fatty acid oxidation reduces pyruvate oxidation by both covalent and allosteric mechanisms, sparing carbohydrate when fatty acids are available. In the present study, P D H flux in isolated mitochondria was markedly reduced by fatty acid availability under most conditions. However, at 20 uM pyruvate and high respiratory rates (State 3), fatty acid availability had no effect on pyruvate oxidation, which is consistent with pyruvate transport limiting pyruvate oxidation. The effects of fatty acid availability on P D H activation state are summarized in Table 9 for different respiratory states. Incubation in different states had litde influence on P D H activation state (i.e. 67 incubation time (min) Figure 5. Effects of 50 u M palmitoyl carnitine on pyruvate dehydrogenase flux in trout white muscle mitochondria. Circles are 20 u M pyruvate. Triangles are 200 u M pyruvate. Open symbols are control incubations (- palmitoyl carnitine). Closed symbols are incubations where 50 u M palmitoyl carnitine was added simultaneously. Rates obtained for 200 u M pyruvate controls, in nmoles , 4C(l)-pyruvate decarboxylated/min/mg, were as follows (mean (SEM.n)): State 4 (upper) 1.24 (0.22,3): State 3 (lower), 24.03 (2.58,5). 68 Table 9. Effects of palmitoyl carnitine on P D H activation state in isolated trout white muscle mitochondria. [palmitoyl carnitine] 0 50 u M State 3 (300 u M ADP) mU PDH/mg 21.3 (1.8) 25.6 (2.5) m U P D H / U C S 1 28.0 33.7 State 4 (no ADP) mU PDH/mg 15.5 (1.7) 19.4 (3.5) mU P D H / U CS 20.4 25.5 A l l incubations contained 200 u M pyruvate. Results are expressed as mean (SEM) for 5 animals, 'mean mU PDH/mg / 0.76 U citrate synthase/mg. The assay represents a Vmax determination at this particular (clamped) activation state, not P D H T 0 T A L , where all P D H is dephosphorylated and maximally activated. There was no significant effect of palmitoyl carnitine addition in State 3 or 4. There was no significant effect of respiratory state, in the presence or absence of palmitoyl carnitine. 69 little covalent modification). Although there was no significant difference in P D H activation state in State 3 vs. 4 (Table 9), there was a 95% reduction of P D H flux in intact mitochondria (Fig. 5). Thus, most of the reduction of P D H flux observed in intact mitochondria in State 4 is due to allosteric inhibition of P D H , rather than covalent modification. Similarly, incubation with 50 u M palmitoyl carnitine (in the presence of pyruvate) had no significant effect on P D H activation state, but P D H flux in isolated mitochondria decreased markedly (Fig. 5). Incubations with palmitoyl carnitine in the absence of pyruvate markedly inactivated P D H in situ (Table 9), confirming that the enzyme can also be covalentiy regulated. PDH activity during recovery PDHa/CS ratios (Fig. 6) showed relatively little change with the exercise regime. A minor increase in activation state was observed immediately post-exercise. During recovery, PDHa activity was relatively constant, near resting activities. PDH in vitro vs in vivo The in vitro activities of P D H in isolated mitochondria (Table 9) was close to that observed in resting fish in vivo (Fig. 6). Conversion between mitochondria rates and tissue activities must take into consideration both temperature differences and CS/mg mitochondrial protein. A mitochondrial P D H flux (15°C) of 10 nmole pyruvate/mg protein is equal to 13.16 nmoles/U CS (0.76 U CS/mg at 15°C). The Q, 0 values for P D H and CS are similar (1.48, 1.52 respectively) so the effects of temperature on PDHa/CS is negligible (Table 11). Mitochondrial oxygen consumption Estimates of the maximal rates of substrate oxidation obtained with trout white muscle mitochondria are presented in Table 10. Pyruvate was oxidized at higher rates than fatty acids but both substrates together elicited higher oxygen consumption rates than pyruvate alone. This may be related to the observation that palmitoyl carnitine availability in the presence of pyruvate did not covalendy inactivate of P D H . Tissue VOJMAX can be calculated from mitochondrial protein/g tissue (7.77 mg/g, Table 11) and 70 OT C o OT c o X Q CL R Ex 2 4 8 recovery time (hr) 24 Figure 6. Pyruvate dehydrogenase (PDH) activation state in white muscle of burst exercised rainbow trout. Activation state is expressed as mean ratio (+ SEM) of P D H (munits)/citrate synthase (units), measured at 25°C. Number of animals at each time is reported with each data point. The asterisk denotes a significant difference from rest. 71 Table 10. Oxidation of physiological fuels by trout white muscle mitochondria (15°C). substrate nmoles O/min/mg malate, 0.1 m M 7 (1.2) pyruvate, 1 m M 133 (9.8) palmitoyl carnitine, 50 u M 78 (7.1) pyruvate + palmitoyl carnitine 150 (12.7) lauroyl carnitine, 100 u M 75 (6.1) octanoyl carnitine, 300 u M 70 (8.3) acetyl carnitine, 1 m M 74 (5.2) R C R determinations (pyruvate + malate) State 3 144 (11.2) State 4 7 (0.5) RCR 22 (1.9) Rates are expressed as mean (SEM) for 5 animals determined at 15°C. A l l incubations contained 0.1 mM malate. The State 4 rate is obtained after all the A D P is phosphorylated. 72 Table 11. Activities of citrate synthase and pyruvate dehydrogenase in trout white muscle and isolated mitochondria at 15°C and 25°C. enzyme source °C activity (SEM) N citrate synthase tissue mitochondria P D H Q 1 0 (15-25°C) citrate synthase P D H PDHa/CS mitochondria mitochondria 25 9.32 (0.67) U/g 25 1.20 (0.05) U/mg 15 0.76 (0.032) U/mg 34 11 6 25 27.9 (2.7) mU/mg 15 18.9 (1.7) mU/mg 5 5 1.52 (0.05) 1.48 (0.06) 0.97 Mitochondrial protein/g tissue, from (CS/g)/(CS/mg mito. protein), is 7.77 mg protein/g. 73 the highest mitochondrial rate observed (150 nmoles O/min/mg, Table 10) and is estimated at 1166 nmoles O/min/g wet weight. This represents an ATP producing capacity of 3.5 umoles ATP/min/g, assuming an A D P / O ratio of 3. Effect of ATP/ADP ratio The results of the parallel study (Schulte et al. submitted) on the effects of exercise on A T P / A D P f are presented in Figure 7. Several important features are obvious. At exhaustion there is a drop in A T P / A D P f ratio by approximately 50%. By 2 hr, at which point phosphocreatine has recovered, the A T P / A D P f ratio has climbed to 2000. By 8 hr it is still elevated compared to rest, but it has recovered by 24 hr. In the present study we examined the respiratory rates of mitochondria presented with physiological A T P / A D P ratios (Fig. 7). We assume that the total A T P / A D P measured in isolated mitochondrial studies corresponds to A T P / A D P f in vivo because the proteins which bind A D P in vivo are fractionated out of the preparation. At the high ratios which occur during recovery, it is apparent that very low rates of oxygen consumption would be elicited if this ratio were the sole controlling factor for mitochondrial respiration. At rest and exhaustion, A T P / A D P f is in the range where changes in the ratio would be expected to affect mitochondrial respiration (i.e. <300). In the previous experiment M g 2 + was omitted to allow establishment of high A T P / A D P ratios. M g 2 + addition so dramatically stimulated contaminating ATPases (possibly mitochondrial-bound hexokinase, myofibrillar or mitochondrial ATPases) that it was impossible to obtain high A T P / A D P ratios in vitro. As much of the ADP, present in the cell is bound to M g 2 + , the effects of M g 2 + on this mitochondrial preparation were examined using a different protocol. If Mg-ADP was the required substrate for the mitochondrial adenylate translocase, then a Mg2+-dependency would be most pronounced under ADP-limiting conditions. As there was no stimulation of ADP-limited respiration in the presence of 5 mM M g 2 + (Table 12), either Mg-ADP is not the substrate for the mitochondrial adenylate translocase or the transporter does not discriminate between chelated and non-chelated A D P . In any case the effects 74 2500 . J — i — i — i — i — i — i — i — i i i i • • • • • » 0 100 200 300 400 500 600 7 0 0 800 ATP/ADP Figure 7. Influence of A T P / A D P ratios on trout white muscle mitochondrial respiration. Upper figure is taken from mean values of A T P and calculated free A D P during recovery (Schulte et al. submitted). Lower figure is oxygen consumption of mitochondria isolated from trout white muscle, exposed to variable A T P / A D P ratios. Each symbol types represents a different mitochondrial preparation. 75 Table 12. M g 2 * effects on respiratory properties of isolated trout white muscle mitochondria. A D P (uM) control +5 m M M g C l 2 300 u M (state 3) 126 (6.4) 5 u M 82 (3.7) 80 (4.4) 2 u M 56 (3.3) 50 (2.6) Results are as mean (SEM) in nmoles O/min/mg mitochondrial protein, for 5 preparations. A l l incubations contained 1 m M pyruvate and 0.1 m M malate. 76 of A T P / A D P on mitochondrial respiration (Fig. 7) were not influenced by omission of physiological levels of M g 2 * . The pattern of A T P / A D P , ratios observed in exercised fish, and the response of mitochondrial respiration to these ratios presents a paradox. At a time when fish white muscle has an added metabolic cost (lactate metabolism), the A T P / A D P { ratio is more inhibitory to mitochondrial respiration than at rest. Further experiments indicated a potential role for phosphate and pH in stimulating respiration when the mitochondria are inhibited by the A T P / A D P f ratio. Effects of phosphate It was clear that under in vivo conditions mitochondria would be relatively insensitive to changes in ATP/ADP, . Furthermore, if A T P / A D P , ratio was the primary regulator of mitochondrial respiration, the elevated ratios observed post-exercise would imply that there was no metabolic scope to meet the increased demands of recovery metabolism. The influence of [phosphate] and pH-phosphate interactions were examined to explain how increased A T P demands during recovery could be met in spite of the elevated A T P / A D P ratio. During the exercise regime, [phosphate] would be expected to increase from basal concentrations to as high as SO mM due to both phosphocreatine hydrolysis and adenylate catabolism via the purine nucleotide cycle. After 2 hr recovery, phosphocreatine is regenerated, but only 50% of the total adenylates remain as IMP. From 2 to 8 hr, [phosphate] is expected to be 8 m M above rest (Schulte et al. submitted). Our phosphate experiments examined how physiological concentrations affect respiratory rate and efficiency of oxidative phosphorylation (as ADP/O). These data are presented in Figure 8. The highest respiratory rates were observed over a range of 5-10 m M phosphate. At the highest concentration (40 mM), phosphate inhibited both the State 3 rate and the 66%-State 3 rate (5 u M ADP) by 30%. Lower respiratory rates (70% of the 10 mM phosphate rate) were evident at 1 m M phosphate at both [ADP]. The efficiency of oxidative phosphorylation (ADP/O) was not affected by [phosphate] at either [ADP]. The influence of pH on phosphate dependency is summarized in Figure 9 and Table 13. The pH 77 100 r phosphate (mM) phosphate (mM) Figure 8. Effects of phosphate concentration on mitochondrial respiration rate and A D P / O ratio. The State 3 (closed circles) incubations contained 1 mM pyruvate, 0.1 mM malate and phosphate. The 66%-State 3 incubations (5 u M A D P , open circles) were also given 100 units hexokinase, 5 m M M g C l 2 and 10 m M glucose. After a 1-2 min pre-incubation the reactions were started with addition of A D P . In each case, the 5 u M rates of oxygen consumption are corrected for ADP-independent respiration. Results are mean (SEM) for 7 determinations. Left panel-ADP/O ratio for the State 3 incubations were calculated using known A D P addition and oxygen consumed between State 4 transitions. The A D P / O for the 5 uM A D P incubations were determined by measurement of glucose-6-phosphate produced in 5 min as described in Materials and Methods. Right panel-Respiration rates obtained as described above. 78 Figure 9. Influence of pH on phosphate dependency of mitochondrial respiration. Mitochondria were incubated as described above except phosphate was excluded and A D P added at time=0. Left panel (open symbols)-2 u M A D P added. Right panel (closed symbols)-0.5 u M A D P added. At 1-2 min intervals small volumes (totalling <5% of final volume) of concentrated, pH-specific potassium phosphate were added to the cell, resulting in a linear rate of oxygen consumption. 79 Table 13. Influence of pH on the respiratory rates of isolated trout white muscle mitochondria. nmoles 0 consumed/min/mg protein p H 6 . 5 pH6 .9 p H 7 . 3 State 3 (0.3 mM ADP) 99 (11) 105 (11) 111(11) 2 u M A D P added 44.4 (5.9) 48 (8.6) 48.8 (10.3) 0.5 u M A D P added 23.8 (3.4) 25 (2.9) 23.8 (2.2) no A D P 4.9 (0.7) 5.3 (0.7) 6.5 (0.5) Rates are mean (SEM) for 5 determinations. The [ADP] reported refer to added levels: No attempt was made to measure total A D P or identify whether adenylates were as A D P or A T P . The oxygen consumption for each pH and A D P condition correspond to the 100% value in Figure 8. There was no significant effect of pH at any [ADP]. 80 values were chosen to reflect rest (7.3), early (6.5) and late (6.9) recovery. At each [ADP] tested (2,0.5 u M ADP) , acidosis decreased the sensitivity to [phosphate]. At each [ADP], the maximal respiration rates were achieved at lower [phosphate] in response to acidosis. There was a greater phosphate dependency at lower [ADP]. DISCUSSION The dramatic and prolonged changes in lactate, pyruvate, pH and mitochondrial controlling factors (ATP/ADP f , phosphate) accompanying recovery metabolism in fish white muscle have been characterized in a parallel study (Schulte et al. submitted). Since recovery metabolism in white muscle must be fuelled by mitochondrial ATP it is important to determine how these physiological changes in tissue metabolites may affect mitochondrial ATP production and fuel preference. Carbon metabolism The first part of this study attempts to discover which metabolite is oxidized by the mitochondria to fuel glyconeogenesis. A potentially important consideration is the influence of the dramatic changes in [lactate], [pyruvate] and pH which are typical of recovery metabolism in fish white muscle. Comparisons of different muscle types in fish (Moyes et al. 1989) and mammals (Pande and Blanchaer 1971) suggest that white muscle mitochondria are better suited to utilize pyruvate-based fuels, such as lactate, than fatty acids. Milligan and McDonald (1988) measured respiratory exchange ratios in recovering fish and found a value close to 1, suggesting carbohydrate, presumably lactate, was the predominant fuel. Thus, changes in [pyruvate] and [lactate] post-exercise might have important implications to recovery metabolism. Fish muscle [lactate] and [pyruvate] increase several fold post-exercise. Resting tissue pyruvate levels (30-50 uM) are below the Km for the mitochondria pyruvate transporter of rat heart (200 u M , Paradies and Ruggiero 1988) and liver (150 u M , Halestrap 1975). Lactate can compete with the pyruvate transporter with lower affinity (Km= 12 m M , Halestrap 1978). Since these kinetic constants are in the range found in trout white muscle, it was important to establish how changes in metabolite concentration affect the kinetics of mitochondrial pyruvate oxidation. Another 81 complicating factor is the influence of pH on pyruvate transport. Increases in the mitochondrial pH gradient increase the pyruvate gradient (Paradies and Papa 1975), and studies with isolated carp red muscle mitochondria suggest that cytosolic acidosis is accompanied by an increase in the mitochondrial pH gradient (Moyes et al. 1988). Thus, tissue acidosis in vivo could enhance pyruvate transport even if [pyruvate] does not change. A regulatory role for pyruvate transport has been proposed in mammalian tissues (Zweibel et al. 1982). A change in pyruvate transport, mediated via changes in the lipid membrane, is part of the response of mitochondria to hyperthyroidism (Paradies and Ruggiero 1988). The effects of [pyruvate] on pyruvate oxidation were highly dependent on the respiratory rate. At low respiratory rates, [pyruvate] over the physiological range had little effect on its oxidation. At higher respiration rates a strong dependency was observed. The relationship is not ubiquitous to fish muscle, as the kinetics of carp red muscle, also in State 3, are insensitive to [pyruvate] over the physiological range presumably due to different activities of the monocarboxylate exchanger (Fig. 4). When [pyruvate] is low and respiratory rate is high, the rate of transport limits pyruvate oxidation in trout white muscle mitochondria. If lactate inhibits pyruvate transport, its effects would be most evident under these conditions. High [lactate] (40 mM) did not inhibit pyruvate oxidation, suggesting that it does not compete with the pyruvate transporter from this tissue. Brain mitochondrial respiration suffers when isolated mitochondria are incubated with lactate, but the effect is due primarily to the decrease in pH from 7.3 to 6.5 (Hillered et al. 1984). Similar changes in pH , which reflect the changes in tissue pH post-exercise, did not affect trout white muscle mitochondrial respiration (Table 13). Prolonged exposure to high [lactate] inhibits mitochondrial respiration, but again the effects are primarily due to pH (Mukherjee etal. 1979). Later in this discussion I provide evidence that trout white muscle mitochondria in situ are much closer to State 4 rates than State 3 rates at rest and during recovery. In light of this, the kinetic relationships demonstrate that physiological changes in [pyruvate] and [lactate] do not affect pyruvate (lactate) oxidation by the mitochondria. Consequendy P D H , and not pyruvate transport, must be the step 82 governing substrate preference at the level of the mitochondria. Covalent regulation of P D H is not important during recovery. A significant increase in activation state immediately post-exercise (Fig. 6) coincides with a drop in A T P / A D P , ratio typical of trout white muscle post-exercise (Parkhouse et al. 1988 and Fig. 7). As there was no significant difference in P D H activation state observed in situ between mitochondria in State 3 and State 4, it is unlikely that the stimulation observed in vivo is due exclusively to the increase in ATP/ADP, . Isolated mitochondria have a similar PDHa/CS ratio to that found in tissue extracts (Table 9, Figure 5) so they provide a realistic system for studying fuel selection. In vitro, fatty acid availability strongly inhibits pyruvate oxidation. This inhibition occurs at high (200 uM) and low (20um) [pyruvate] in both State 3 and State 4, except where transport is rate limiting (State 3, 20 u M pyruvate). The more physiologically realistic condition for recovering white muscle is State 4, and under these conditions P D H flux is strongly inhibited by the presence of fatty acids. Thus if fatty acids were available in vivo, we expect that fatty acid oxidation would inhibit lactate oxidation, sparing it for glyconeogensis. As this inhibition occurs without major changes in PDHa/CS, the PDHa/CS pattern observed in vivo does not provide evidence for the utilization of either fuel. Although fish white muscle is a lip id-poor tissue, the required mitochondrial flux rates/g are so low that even a modest supply, relative to red muscle, may meet the demands of the tissue. Metabolic rate The rate of recovery (i.e. lactate disappearance) in most fish is very slow relative to mammals. Flounder metabolizes 0.01 umoles lactate/min/g (Milligan and Wood 1987), salmonids 0.05 umoles/min/g (Schulte et al. submitted), and skipjack tuna 1.4 umoles/min/g (Arthur et al. in prep.). With the exception of tuna, these rates are considerably lower than observed in mammals (3.0 umoles/min/g, Meyer and Terjung 1979; 0.66 umoles/min/g, Hermansen and Vaage 1977). In each fish species glyconeogenesis occurs primarily i/i situ during recovery. Since the rate of recovery in fish species apparendy correlates with the mitochondrial density of the white muscle, we undertook a series of experiments to determine 83 if the rate of mitochondrial A T P production for glyconeogenesis may limit the rate of recovery in trout white muscle. We were somewhat surprised to find that trout white muscle possesses many fold excess mitochondrial ATP-producing capacity than would be needed to meet the added demands of lactate metabolism. The cost of glycogen resynthesis in situ via pyruvate kinase reversal is 2.5 ATP/lactate. Lactate disappears at a rate of 0.05 umoles/min/g (Schulte et al. submitted), therefore the metabolic cost of glycogen resynthesis is 0.125 umoles ATP/min/g tissue. With a mitochondrial A T P producing capacity of 3.5 umoles ATP/min/g tissue, there is a 28-fold excess capacity over the added demand of lactate conversion to glycogen post-exercise. Conversely, the cost of lactate metabolism can be met by an added (above basal) mitochondrial rate of 5.4 nmoles O/min/mg protein. At present there is no estimate of resting white muscle oxygen consumption in fish, but it is likely that this is very low. If trout white muscle mitochondria respired at State 3 rates in vivo, whole animal oxygen consumption, without the influence of the other aerobic tissues, would be 23 mmoles Cykg/hr (assuming 660 g white muscle/kg). This rate is well above standard metabolic rate for salmonids (approximately 2.5 mmoles 0 2/kg/hr) (Bushnell et al. 1984, Mill igan and McDonald 1988). It is also at least 2 fold higher than the rate measured for fish recovering from burst exercise (Milligan and McDonald 1988) or swimming at high steady state velocities (Bushnell et al. 1984). It has been suggested that mammalian red muscle mitochondria respire at State 3 rates when the tissue is at V0 2MAX (Schwertzmann et al. 1989). However, the required in vitro rates could only be achieved with succinate as substrate; mitochondrial pyruvate oxidation accounted for less than 50% the rate of oxygen consumption known to occur in cat skeletal muscle in vivo. This inability of isolated muscle mitochondrial preparations to match the required in vivo rates also has been reported for hummingbird flight muscle (Suarez et al. in press) and locust thoracic muscle (Suarez and Moyes submitted). Kidney cell mitochondria are near State 3 when the tissue is maximally stimulated by addition of nystatin (Harris et al. 1981). Tuna heart muscle mitochondria do not exhibit this shortfall (Chapter 3). In contrast, the trout white muscle mitochondrial preparation is the first to demonstrate many fold excess 84 capacity in vitro. Clearly this tissue does not demonstrate the concept of symmorphosis in relation to oxygen delivery and utilization. This is to be expected in fish, where a large proportion of the body mass works at maximal rates which are independent of oxygen availability. When presented with physiologically realistic A T P / A D P ratios, trout white muscle mitochondria respire at much lower rates (Fig. 7). Skeletal muscle mitochondria from mammals differ from heart mitochondria in that resting [ADP] and [phosphate] are below their S ^ levels (Balaban 1990). Changes in A D P availability to the mitochondria can account for changes in the metabolic rate of skeletal muscle in vivo (Chance et al. 1985, Balaban 1990). The white muscle mitochondrial respiration rate in relation to A T P / A D P (Fig. 7) is similar to that shown for rat heart (Bishop and Atkinson 1984) and liver (Wanders and Westerhoff 1988) mitochondria. The A T P / A D P ratio in trout white muscle is elevated to the point that mitochondrial respiration would be relatively insensitive to changes in this parameter. Mammalian skeletal muscle respiration becomes less sensitive to A T P / A D P with decreasing mitochondrial density (Dudley et al. 1987). Trout muscle has a lower mitochondrial content (10 U citrate synthase/g) than even hypothyroid rats (30 U citrate synthase/g, Dudley et al. 1987). The increase in oxygen consumption required to meet the A T P demands for glyconeogenesis in trout white muscle is 5 nmoles O/min/mg mitochondrial protein. If A T P / A D P f ratio is the primary signal for mitochondrial respiration, the elevated ratios post-exercise leave no scope for recovery metabolism, if basal demands are unchanged. The expected change in phosphate is probably the predominant signal for increased A T P production during recovery. Fish white muscle [phosphate] has not been reliably measured at rest. 3 1 P - N M R studies by van den Thillart and coworkers (1989) demonstrate a phosphate peak that is barely detectible and estimated to be approximately 0.5 mM (van den Thillart pers. comm.). With burst exercise several metabolites change in a manner which suggests marked elevation of white muscle [phosphate]. Phosphocreatine has been estimated to be 95% phosphorylated in vivo (van den Thillart et al. 1989), and it is mosdy hydrolyzed following burst exercise (Schulte et at. submitted). Total creatine is approximately 40 85 umoles/g (Schulte etal. submitted), consequently phosphocreatine hydrolysis would release approximately 35 umoles phosphate/g. Adenylate depletion via A M P deaminase, also accompanies burst exercise with the net effect of conversion of approximately 5.5 umoles ATP to IMP (Schulte et al. submitted). A M P deaminase would elevate [phosphate] by 11 umoles/g. Thus, in early recovery the mitochondria are probably exposed to 40-50 u M phosphate. By 2 hr post-exercise, phosphocreatine has recovered, but the total adenylates remain low until at least 8 hrs. In this phase of recovery, 5-10 u M phosphate is expected. The effects of these physiological changes in [phosphate] on isolated mitochondria can explain many of the observations associated with burst exercise recovery. Phosphate concentrations of 5-10 umoles/g are optimal for mitochondria under a wide variety of [ADP] and pH values (Fig. 8,9). At the A D P levels used in this study, mitochondrial rate was sensitive to [phosphate] over the range 0.2 to 10 m M (Fig. 8). The sensitivity to [phosphate] increased with decreased [ADP] (Fig. 9). Decreases in pH similar in magnitude to the acidosis which accompanies recovery (Schulte et al. submitted) increases the respiratory rate at each sub-optimal [phosphate]. Thus increased A T P production with severe exercise involving adenylate depletion is obtained from the response to an elevated [phosphate]. With moderate exercise, where less adenylate depletion occurs and consequently less phosphate accumulates, acidosis would potentiate the phosphate effect on respiration. High [phosphate] is inhibitory to mitochondria (Fig. 8) and may be expected to affect the aerobic scope in the early phase of recovery. However, this early phase where phosphate inhibition might be expected is the only time when the A T P / A D P f ratio actually declines relative to rest, possibly countering the inhibition. The phosphate inhibition of mitochondrial respiration may explain why the rate of lactate disappearance in the early recovery phase is often slower than in later recovery (e.g. Pearson et al. 1990). The pH-induced stimulation of respiration at sub-optimal [phosphate] may be due to an increase in the relative abundance of ortho-phosphate, the species which is transported by the mitochondrial phosphate/hydroxyl exchanger. Alternately acidosis-induced increases in the mitochondrial pH gradient (e.g. Moyes et al. 1988) may increase the mitochondrial [phosphate] at any particular cytoplasmic 86 [phosphate]. There is also a potential role for pH in altering other aspects of mitochondrial control not examined in this study such as adenylate distribution across the mitochondrial membrane. The adenylate translocase is electrogenic, and alterations in the pH gradient may affect mitochondrial A T P / A D P or [ADP] at a given cytosolic A T P / A D P f . Although acidosis enhances the rate of recovery in mammalian skeletal muscle (Bonen et al. 1980), the site of action has not been established and could be related to the effects described in the present study. The initial goal of this study, to assess the possibility that mitochondrial capacity may limit the rate of glyconeogenesis in vivo, is difficult to assess direcdy. The in vivo increase in mitochondrial ATP production in recovery is estimated from the difference in oxygen consumption between the resting mitochondrial rate (low A T P / A D P but limiting [phosphate]) and the recovery rate (higher A T P / A D P but optimal [phosphate]). The resting rate is best approximated by the State 4 rate, which is about 5% of the State 3 (RCR=20). The recovery rate is best approximated by the values determined in Figure 7, 15% of the State 3 rate. The difference (10%) is 3 fold greater than the recovery requirements for lactate metabolism (3.5% of State 3). If the phosphate and pH signals are adequate to stimulate mitochondrial metabolism enough to fuel glyconeogenesis, what then limits the rate of recovery? Oxygen availability is unlikely to be important in this tissue. Blood flow to trout red muscle is only 5 fold greater than white muscle (Neumann et al. 1983). The difference in maximal respiration rates/g muscle are expected to be much greater than the differences in oxygen delivery to the tissues. White muscle has a lower mitochondrial content and is expected to use a much smaller fraction of its oxidative capacity (i.e. proportion of State 3). The importance of redox in control of mitochondrial respiration has not been examined. Redox control is difficult to study in fish white muscle because the low mitochondrial content may invalidate many of the assumptions required to measure mitochondrial redox. The mitochondrial fraction of any of the relevant metabolites (e.g. N A D H / N A D + or glutamate dehydrogenase reactants) would be expected to represent a much smaller proportion of the total pool in mitochondria-poor white muscle of fish. Instead of an ATP supply short fall, factors affecting the anabolic capacity may be most 87 important. Although not unequivocally established, the pathway for glyconeogenesis in fish white muscle may demand reversal of pyruvate kinase. The pathway for glyconeogenesis favoured by Connett (1979) requires P E P C K , which is absent from trout (Cowey et al. 1977) and other fish white muscles (Hulbert and Moon 1978, Moon and Johnston 1980, Mosse 1980) with the noteable exception of marlin (Suarez et al. 1986). Rabbit pyruvate kinase can be reversed to 2% of its Vmax under the appropriate (non-physiological) conditions (Dyson et al. 1977). If rainbow trout pyruvate kinase activity is similar to brook trout white muscle (140 u/g, Johnston and Moon 1980) it would require only 0.04% of the forward rate to account for the rate of lactate disappearance. It is possible that the high A T P / A D P is primarily a signal to drive pyruvate kinase in the reverse direction, aided by elevated [pyruvate] in the early phases of recovery (Schulte et al. submitted). In summary, several of the differences in skeletal muscle metabolism of mammals and fish have important implications to metabolic control. The equilibrium model of respiratory control (Ericinska and Wilson 1982) does not appear to be applicable at high rates of respiration in mammalian muscle because the reactants of the mitochondrial ATP synthase are not near equilibrium (LaNoue et al. 1986). As trout white muscle may perpetually respire at low rates relative to the maximal capacity, the equilibrium model may be more appropriate in this tissue. This remains to be tested. Fish muscle mitochondria are relatively insensitive to A T P / A D P ratio over the physiological range. Instead, changes in [phosphate], generated as a result of phosphocreatine hydrolysis and adenylate depletion, may be the more important factor regulating mitochondrial respiration. A potential role for pH in enhancing the mitochondrial response to phosphate changes is also proposed. Although the kinetics of pyruvate oxidation suggest that [pyruvate] could affect mitochondrial metabolism at high respiratory rates, this effect is not expected to be important in vivo because of the low respiratory rates. We found no evidence for lactate inhibition of pyruvate transport into the mitochondria. If fatty acids are available to mitochondria post-exercise, fatty acid oxidation would spare pyruvate (lactate), primarily through allosteric inhibition of P D H . 88 G E N E R A L DISCUSSION A comparison between rates of oxygen consumption of isolated cat skeletal muscle mitochondria and in vivo mitochondrial fluxes prompted Schwertzmann et al. (1989) to suggest that mitochondria in situ respire at rates approaching their in vitro V0 2MAX (State 3). Closer examination of their data reveals that in vivo requirements could only be met with succinate as substrate: Rates of oxidation of the more physiological fuel pyruvate (fatty acids were not examined) could not account for respiration which obviously occurred in vivo. When similar comparisons are performed with tissues notable for extremely high mitochondrial fluxes in vivo, a similar deficiency between the in vivo and in vitro occurs (hummingbird flight muscle, Suarez etal. 1991; locust flight muscle, Suarez and Moyes submitted). The calculations for tuna ventricle (Chapter 3) are the first where the in vitro rates actually account for what happens in the muscle in vivo, and in fact match quite closely. Such an observation is important for a number of reasons, not the least of which is providing a frame of reference for isolated mitochondrial studies. The observation that in vivo fluxes are similar to isolated mitochondrial State 3 rates allows confident extrapolation from the in vitro to the in vivo situation when questions of regulation, flux etc. are addressed. Trout white muscle is the most unusual muscle preparation studied to date (Chapter 4). Its in vitro capacity is many fold greater than may ever be utilized in vivo. This is not because white muscle mitochondria have an unusually high oxidative capacity, but rather due to the cytosolic regulatory environment (ATP/ADP, phosphate) suppressing mitochondrial respiration below State 3 rates. It is paradoxical that a tissue considered to be mitochondria-poor has far more mitochondrial capacity than it needs. Muscle metabolic regulation differs between red and white muscle because of the physiological role of each fiber type, and the pathways of ATP synthesis utilized when the muscle is recruited. Priority in white muscle performance is maximizing glycolytic flux: Natural selection that increased mitochondrial capacity would not substantially improve white muscle burst performance because of the high rates of A T P synthesis required. (The high mitochondrial density observed in tuna white muscle is thought to be related to recruitment of this tissue at high steady state velocities rather than aerobic support of burst 89 activity.) Burst exercise recovery in white muscle is fuelled by mitochondria but it is a a secondary priority, occurring whenever it can, as fast as it can, within the regulatory environment associated with being a glycolytic tissue. If high A T P / A D P f is required to direct this glycolytic tissue toward glyconeogenesis, then there must be adequate mitochondrial capacity under those regulatory conditions to meet the ATP demands of recovery. A completely different regulatory environment exists for red muscle function because the capacity of mitochondrial metabolism is critical when muscle is working. Despite a 25-fold difference in mitochondrial density in muscles examined in this thesis, the maximal capacities of mitochondria isolated from skeletal muscles are remarkably similar, varying less than 2-fold between species. It is likely that there are many mitochondrial differences correlating with mitochondrial density which are not revealed by comparisons of State 3 rates. Such subtle differences include those identified in this thesis which are related to regulation of fuel preference. In comparing mitochondria prepared from a spectrum of teleost muscles, a 10-fold difference in CPT/unit mitochondria occurs, with higher specific activities in mitochondria from more oxidative muscles. Comparisons between CPT activities and mitochondrial fatty acid fluxes suggest that, even in carp white muscle, with the lowest CPT specific activity, there is far more measureable CPT than would be required to saturate the mitochondrial oxidative pathway (Chapter 4). The lack of correlation between CPT activities and fatty acyl carnitine oxidation suggests there is more to the story than Vmax measurements of CPT. Complexities associated with CPT activity in vivo may include differential effects of the lipid memebrane, sub-optimal substrate concentrations, intracellular metabolite diffusion gradients and mitochondrial subcellular localization in relation to such gradients. 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