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Glucose utilization in fish muscle during sustained swimming and recovery from maximal swimming West, Timothy Gordon 1994

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GLUCOSE UTILIZATION IN FISH MUSCLE DURING SUSTAINED SWIMMING AND RECOVERY FROM MAXIMAL SWIMMING.  By Timothy G. West B.Sc. Biology. Daihousie University, 1982 M.Sc. Biology. Daihousie University, 1986  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 standard  THE UNIVERSITY OF BRITISH COLUMBIA  January 1994 © Timothy G. West, 1994  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.  (Signature)  Department of  o  -  The University of British Columbia Vancouver, Canada  Date  DE-6 (2/88)  /1  tN  11  ABSTRACT The importance of circulatory glucose as a fuel for skeletal muscle and heart during sustained swimming and as a glycogenic substrate for skeletal muscle during C-2-deoxyglucose exercise-recovery was examined. In vivo glucose utilization, using 14 14 in swimming trout (Oncorhynchus mykiss) and carp (Cyprinus carpio) ( C-2-DG), indicated similar rates of glucose utilization in red muscle of both species (20 ). g 1 nmolmin  -  30  Estimates of the energetic importance of glucose, assuming red  muscle accounted for the bulk of the active oxygen consumption, suggest that glucose could be more important to the slower swimming carp potentially accounting for 25 -  -  30 % of total energy provision. Glucose uptake in red muscle of the more active trout was estimated to contribute < 10 % of total substrate demands during exercise. Cardiac energetics was estimated to be largely independent of glucose utilization in ) was g 1 trout. In vivo utilization of glucose in trout heart at rest (about 5 nmolmin ) and measurements g 1 considerably lower than in vitro determinations (120 nmolmin g’ based on hexokinase activities). nmolmin , of glucose utilization capacity (8300 1 Nearly complete inhibition of glucose flux in vivo is likely the result of preference for alternative substrates for oxidation. In carp, cardiac glucose utilization was dependent on plasma glucose concentration, but was also estimated to be of minor energetic importance at plasma concentrations comparable to those observed in swimming trout (2  -  4 mmolL ). Reduced dependence of trout on glucose for swimming energetics t  agrees with the generalization that species with superior aerobic exercise capacities may be dependent on fat based fuels for muscle energetics. Plasma glucose concentration varied from 6  -  1 in trout during 38 mmolL  recovery from intense burst swimming. Hyperglycemia was not related specifically to exercise.  H-glucose tracer, increased 3 Estimated glucose turnover rate, using 6-  proportionally with plasma glucose availability.  Similar concentration-dependent  kinetics were observed in carp at rest and during sustained swimming. Regardless of  11].  the association between glucose turnover and plasma glucose level, glucose utilization C-2-DG 1 ( 4 phosphorylation) in trout white muscle accounted for < 10 % of the glycogen repletion after exercise.  In trout red muscle, glucose utilization was  dependent on plasma glucose concentration up to 10  was estimated to account for 25 muscle type.  -  -  12 mmoPL 1 and glycogenesis  60 % of post-exercise glycogen repletion in this  Moderate post-exercise hyperglycemia sometimes seen in recovering  salmonids seems pertinent to red muscle carbohydrate status, but white muscle recovery occurs independently of plasma glucose availability. These results provide evidence for the predominance of a glyconeogenic fate of white muscle lactate in trout. Investigation of lactate/glycogen recovery profiles in the fast swimming skipjack tuna  (Katsuwonus pelamis) provides evidence that, despite the relatively high oxidative capacity of tuna white muscle and previous determinations of high circulatory fluxes of lactate, this species also seems to convert white muscle lactate to glycogen via intramuscular glyconeogenesis.  A  glyconeogenic  fate  for lactate  conserves  carbohydrate status and ensures relatively rapid replenishment of the principal carbon source in fish white muscle.  iv  TABLE OF CONTENTS  ABSTRACT  ii  TABLE OF CONTENTS  iv  LIST OF FIGURES  viii  LIST OF TABLES  ix  ACKNOWLEDGEMENTS  x  CHAPTER 1. SUBSTRATE UTILIZATION IN FISH MUSCLE DURING EXERCISE AND RECOVERY.  1  Sustained swimming  2  Fuel selection  3  Protein and amino acid utilization  6  Utilization of fat fuels  8  Carbohydrate utilization  10  White muscle  15  White muscle recruitment and exercise metabolism  16  Carbohydrate recovery in fish muscle  18  In vivo measurement of glucose utilization  20  CHAPTER 2. GLUCOSE UTILIZATION DURING SUSTAINED SWIMMING IN  RAINBOW TROUT (Oncorhynchus mykiss) AND CARP (Cyprinus carpio). In vivo utilization of glucose by heart and locomotory muscles of exercising rainbow trout (Oncorhynchus mykiss).  23  Introduction  23  Materials and Methods  24  Animals  24  2-DG and the lumped constant  24  Estimation of LC in muscle slices  25  V  Estimation of LC in isolated trout hearts  27  Glucose utilization in swimming trout  28  Statistics  29 30  Results Tissue LC’s and the effect of glycolytic rate  30  Glucose utilization in swimming trout  30  Discussion  34  The 2-DG lumped constant  34  Glucose utilization in red muscle  37  White muscle glucose uptake  40  Cardiac glucose utilization  41  Muscle glucose utilization during sustained swimming in the carp.  43  Introduction  43  Materials and Methods  44  Animals  44  Estimation of Sustained swimming in cannulated carp  44  Bolus-injections and tissue sampling  45  Metabolite assays  46  Calculations  47  Statistics  48 48  Results  Carp swimming performance  48  Glucose turnover  50  Muscle glycogen and glucose utilizaiton  50  Discussion Oxygen consumption  56 56  vi  Glucose turnover  58  Muscle glucose uptake  59  CHAPTER 3. CARBOHYDRATE RECOVERY IN SKELETAL MUSCLE OF RAINBOW TROUT (ONCORHYNCHUS MYKISS) AND SKIPJACK TUNA  (KATSUWONUS PEL4MIS) AFTER BURST EXERCISE.  62  Implication of hyperglycemia for post-exercise resynthesis of glycogen in trout skeletal muscle.  62  Introduction  62  Materials and Methods  63  Animals  63  Surgery  63  Exercise and recovery protocol  65  Isotope injection and tissue sampling  66  Metabolite assays  67  The 2-DG lumped constant  67  Glucose turnover and muscle uptake  68  Statistics  68  Results  68 Glucose concentration and turnover rate  68  Muscle glucose utilization  71  Glycogen replenishment in skeletal muscle  76  Discussion  76  Glucose turnover  77  White muscle recovery  79  Red muscle recovery  80  vii  Lactate, glycogen and PCr profiles in the white muscle of skipjack tuna during recovery from burst swimming.  82  Introduction  82  Materials and Methods  83  Animals  83  Exercise and biopsy protocol  83  Tissue preparation and analysis  85  Data analysis  86  Results  86 Muscle metabolites after exercise  86  Lactate, glucose and glycogen in recovery.  88  Estimates of recovery time (lactate and PCr)  91  Discussion  95  Muscle lactate and glycogen recovery  95  Plasma laãtate and glucose  99  Lactate and PCr  102  CHAPTER 4. GENERAL DISCUSSION.  104  Glucose as a fuel for swimming  104  Muscle tissue as a site of glucose disposal  104  Maximal glucose flux  107  Regulation of glucose utilization  110  Exercise energetics in trout and carp  111  Post-exercise recovery of muscle glycogen  113  Muscle glyconeogenesis  115  Glucose and lactate in fish white muscle  117  LITERATURE CITED  119  viii LIST OF FIGURES  1.  Glucose utilization in normoxic and NaCN-treated trout hearts.  33  2.  C-2-DG washout in resting and swimming trout. 14  35  3.  Oxygen consumption in relation to swimming speed in carp.  49  4.  Washout of glucose tracers in resting and swimming carp.  51  5.  Effect of glucose availability on glucose turnover and clearance in carp.  53  6.  Effect of muscle glycogen content on glucose utilization in carp red muscle.  55  7.  Effect of glucose availability on glucose utilization index in carp muscles.  57  8.  Post-exercise glucose and lactate levels in trout plasma.  64  9.  Washout of glucose tracers in resting trout.  69  10.  Washout of glucose tracers in recovering trout.  70  11.  Effect of glucose availability on glucose turnover and clearance in trout.  73  12.  Carbohydrate recovery and glucose utilization rate in trout white muscle.  74  13.  Carbohydrate recovery and glucose utilization in trout red muscle.  75  14.  Recovery profiles of glycogen, lactate and PCr in tuna white muscle.  89  15.  Plasma glucose and lactate levels in tuna.  90  16.  Break points in lactate and PCr recovery in representative tuna.  92  17.  Recovery time in relation to tuna white muscle metabolite content.  93  18.  Association between lactate and PCr recovery time in tuna white muscle.  97  19.  Association between lactate and PCr content in tuna white muscle.  98  ix LIST OF TABLES  1.  Fuel stores in a ‘ ypcal salmonid’.  5  2.  Lumped constants in trout muscle slices.  31  3.  Trout heart lumped constant: effect of glycolytic rate.  32  4.  Glucose utilization in muscles of resting and swimming trout.  38  5.  Glucose kinetics and muscle GUI in resting and swimming carp.  52  6.  Glucose turnover and utilization in resting and recovering trout.  72  7.  Metabolites in white muscle of exercised and unexercised tuna.  87  8.  Mean recovery times for lactate and PCr in tuna white muscle.  96  9.  Glucose disposal in carp muscles as a proportion of whole-body turnover.  106  x ACKNOWLEDGEMENTS Thanks first and foremost to Dr. Peter W. Hochachka for your guidance, enthusiasm, and financial support, for forgetting (once, at least) the ‘just-one-conference-ayear’ thing and for handing over the keys to the lab and keeping a sense of humor about my not returning them until recently. The emphasis here has always been on discussion and collaboration and I’m greatly appreciative of, what I believe was, an extraordinary group of lab personnel. Their assistance and constant feedback was (is) immeasurable. I am very grateful to Dr. ‘s R.K. Suarez, P.G. Arthur, and C.D. Moyes for sharing their lab know-how and for many discussions. Thanks also to Peter Arthur for his skill with the isolated trout heart preparation and for measuring PCr in tuna white muscle and to Trish Schulte for the assistance with the trout swimming trials and for helpful discussions. Thanks also to Dr. ‘s Buck, Doll, Land, Staples and Stanley for their interaction in the lab. People outside of Peter’s lab to whom I am grateful are; Dr. Dave Randall for providing the swim-tunnel, Cohn Brauner for his expertise with fish respirometry in the carp study, Dr. R.W. Brill (Southwest Fisheries Science Center Honolulu Laboratory) for help cannulating and spinal-blocking tuna, and the Bamfield Marine Station staff for technical support in the study of sustained swimming in trout. Thanks: to Steve for corroborating those many encounters with landlorditis and for  pointing out the, not so obvious, predictive value of faulty water heating.  Thanks for  introducing the Brownies, Laphroaigs, MacAllans, and Youngers. Sorry about the cider chunks!; to Sohail for camaraderie, from hallway frisbee to toasting the data god from the top of %$*%#@ Mount Harvey.. .always a laugh, man; to the HABS faithful, and the jays faithful  -  they know who they are; to Cwis for inspiration, CD for death-ball frustration,  LB for poker-face exhibition, JS for road-trip transportation, housemates for demonstration of cooperation, all squash partners for enervation, and to Raul for the extra cash. Thanks especially to Mom and Dad for understanding about the infrequent visits home.  1  CHAPTER 1: SUBSTRATE UTILIZATION IN FISH MUSCLE DURING EXERCISE AND RECOVERY.  Comparative studies of fish energetics indicate a striking diversity in exercise performance capability across species. Aerobic capacity of skeletal muscle is about 20-fold higher in fast than in slow swimming species and rates of recovery from short-term maximal exercise extend over more than two orders of magnitude (Moyes et a!. 1992a; 1993). With such diversity in performance characteristics, it might be expected that fish would also display variable patterns of substrate utilization for oxidative energy provision during sustained swimming and for restoration of intramuscular fuel depots during recovery from exercise. For example, indirect observations indicate that fish species with  superior swimming capabilities have elevated muscle oxidative capacities as well as greater capacities for fat utilization compared to less active fish (Moyes et a!. 1992a).  Similar  tendencies are noted among mammals, where aerobically adapted species seem more dependent on lipid based fuels than sedentary species (Weber 1992). However, there is no compeling evidence among fishes that activity metabolism in slower-swimming species is supported mainly by one particular type of substrate (i.e., fat versus carbohydrate based fuels) over another (Moyes et al. 1989; Driedzic 1988; Sidell et a!. 1987). Although generalizations are often made regarding the importance of circulatory glucose in activity related processes in fish, few studies have addressed glucose fluxes during exercise and recovery and there are no estimates of in vivo glucose utilization in specific muscle-types during exercise.  Glucose dependency for swimming energetics  across species may reflect differences in aerobic swimming capabilities.  Similarly,  although the use of circulatory glucose for post-exercise restoration of muscle carbohydrate stores is minimal in most species that have been studied (Wood 1991; Moyes et a!. 1993), glycogenic capacity may nevertheless depend on the aerobic capability of fish.  For  example, tuna possess elevated aerobic capacities and also have the capacity for high post exercise lactate fluxes (Weber et al. 1986). There is the possibility that a major portion of  2  the post-exercise lactate load is oxidized in fast-swimming species, rather than used for muscle glyconeogenesis.  The following studies exploit the differences in aerobic  swimming capacity across species to investigate the in vivo role of circulatory glucose in muscle energetics of rainbow trout (Oncorhynchus mykiss) and carp (Cyprinus carpio) and in post-exercise glycogenesis of rainbow trout and skipjack tuna (Katsuwonus pelamis).  Sustained swimming.  Fish are able to swim in a continuous “steady-state” fashion  over a broad range of speeds. In a swim tunnel, salmonids can be challenged with water speeds that correspond to up to about 80 % of fatigue or critical (Uit) speed before nonsteady state swimming (charcterised by burst-and-glide behaviour) is noticed. Steady-state swimmming, even at very high intensity, can be endured indefinitely. Electromyographical studies show that red muscle is recruited during both slow and fast sustained swimming in species that have been examined (Johnston and Moon 1980a,b; Johnston et al. 1977; Rome et a!. 1984, 1988).  However, it is not so clear that white  muscle recruitment occurs only at high, non-sustainable swim speeds.  In good aerobic  performers (eg., salmonids and tunas) white muscle activity is evident at submaximal swim speeds (Johnston and Moon 1981b; Brill and Dizon 1979). Observations that argue against white fiber involvement at low swim speeds are that only red muscle showed significant hypertrophy after endurance training in salmonids (Johnston and Moon 1980b) and that EMG’s recorded from deep white muscle show activity only at high sustainable swim speeds in coalfish, Pollachius virens (Johnston and Moon 1980a).  The carp is the one  species where sequential recruitment of muscle types has been examined extensively and in this species the division of labour between fiber masses seems fairly sharp. As in other species, red fibers are recruited at low swim speeds, followed by intermediate (similar to fast twitch oxidative glycolytic, or FOG, fibers in mammals) fiber activity at higher sustainable speeds and finally by white muscle recruitment (Johnston et a!. 1977) which, in the most recent analyses, is always accompanied by unstable swimming and rapid fatigue  3  (Rome et al. 1984).  In none of these studies was recruitment examined in relation to  fatigue speed for individual fish, so it is not certain at what percentage of U white fibers became active.  It is not known in either carp or salmonid species whether swimming  specifically at 80 % of Uj is accompanied by white muscle recruitment, although exercise at this intensity is expected to push red muscle metabolic demands close to maximal in both species (Jones 1982; Randall and Daxboeck 1982; Rome et a!. 1988). In species where white muscle recruitment might occur at high sustainable speeds, it would be generally expected that such activity would be supported largely by glycolysis. Endurance training induces increased activity of key fat utilizing enzymes (Moon and Johnston 1980a,b), suggestive of aerobic recruitment of white fibers at sustainable swim speeds. However, the low aerobic capacity of this muscle (Moyes et a!. 1989, 1992b) and  the disthbution of blood flow to mainly active red muscle (Randall and Daxboeck 1982) indicates that aerobic demands of white muscle are minimal compared to red muscle. If in trout, for example, the aerobic demands of high intensity red muscle activity (assumed similar to measurements in the bulirout, Myoxocephalus scorpius, of 5 (Altringham and Johnston 1990) or about 18  -  -  1 8 mWg  29 mol ATP min’g’) were imposed on  the recruited white muscle, its aerobic metabolism could provide < 15% of the ATP (e.g., rainbow trout white muscle mitochondria can produce 3.5 mol ATP min , Moyes et g 1 a!. 1992b).  The situation could be different in the skipjack tuna which have a white  muscle mitochondrial content, and presumably aerobic capacity, that is exceptionally high, 2 5 times that of trout (Moyes et al. 1992b) and carp (Moyes et a!. 1992a). -  Fuel selection.  Fuel use in specific tissues can be implied from in vitro measurement  of maximal enzyme activity (V) and mitoehondrial oxidation rates, both of which provide maximal capacities to utilize particular substrates. It is understood that these in vitro estimates of fuel utilization may not resemble flux capacity in tissues of the whole animal (Moyes et a!. 1990; Wright and Albe 1990), but in comparative analyses such  4  observations have provided clues about which carbon sources may be used in vivo under conditions of maximal aerobic demand.  In endotherms for instance, cardiac enzyme  profiles suggest that maximal aerobic metabolism is supported by fat utilization to a greater extent in animals adapted for aerobic endurance than in more sedentary animals (Driedzic Mammalian models of substrate preference based on in vivo metabolite  et al. 1987).  kinetics also suggest a greater reliance on fat in more aerobic species during exercise (Weber 1992).  Similar enzyme analyses of fish muscle from different species suggest  either that highly aerobic tissues rely more on fat (Moyes et a!. 1992a) or that there is no association between fat utilizing capacity and increasing tissue aerobic capacity (Sidell et a!. 1987). Furthermore, within fish species, the effects of endurance training include small shifts in fuel availability (Hochachka et a!. 1961) and in the activity of some key muscle and heart enzymes (Farrell et a!. 1991; Johnston and Moon 1980), but modified substrate flux through a single pathway is usually not implied. In vivo fuel preference studies attempt to quantify substrate utilization with indirect  calorimetry, tracer infusions or aterial-venous differences in metabolite concentration across muscle groups. Where physiologically significant quantities of fuels exist, rates of substrate supply, competition between pathways and the influence of regulatory factors will determine the pattern of fuel utilization. Unfortunately few studies address these situations directly in fish.  However, it is possible to gain initial insight into the metabolic  partitioning of different substrates during sustained exercise from whole-body sites and levels of the main storage substrates. Representative salmonid tissue masses and metabolite levels, per g organ mass and per kg body mass, are summarized in Table 1.  These  metabolite levels represent the maximum mobilizable fuels in non-feeding salmonids. In the following review the importance of each of the major substrate-types for sustained exercise will be considered in turn. The analyses focus on using maximal capacities for flux and estimates of maximal red muscle energy demand as guides to the potential dependence  on  different  fuels.  As  exercise  becomes  less  intense  or  5  Table 1. Tissue masses and fuel stores of a resting salmonid.  ECF  liver  adipose  white  red  g tissue/kg body mass  250  15  20  600  70  glycogen (as glucosyl) (jLmol/g tissue) (mmol/kg body mass)  0 0  100-200 1.5-3.0  40 24  40 2.8  glucose (jmol/g tissue) (mmol/kg body mass)  2-5 0.5-2.0  2-5 0.03-0.075  2 1.2  2 0.14  lactate (jmol/g tissue) (mmol/kg body mass)  7 1.75  3 0.045  3 1.8  5 0.35  triglyceride (JLmol/g tissue) (mmollkg body mass)  22 5.5  10-30 0. 15-0.45  15 9  40 3  fatty acids (JLmol/g tissue) (mmol/kg body mass)  3 0.75  3 0.045  2 1.2  2 0.14  protein (as amino acyl) (jmol/g tissue) (mmollkg body mass)  2000 500  1000 15  1000 600  500 35  amino acids (jLmol/g tissue) (mmol/kg body mass)  40 10  50 30  9 0.63  800 16  Data were compiled from various sources (French et al. 1983; Idler and Bitners 1959; Leger  eta!. 1981; Mulligan and Wood 1986b; Mommsen et a!. 1987; Parkhouse et al. 1982; Plisetskaya 1980; Sheridan 1988; Weber 1991; Wright et al. 1989). These values are estimates for total fuels and are not neccessarily available metabolically.  6  prolonged, the less abundant fuels and depots may become increasingly important.  In  instances where no direct studies have been performed, indirect studies using enzyme analysis and isolated mitochondria can give further insight into the capacity and preference for fuels during exercise.  Protein and amino acid utilization.  In fish, dietary protein levels of 30-50% (w/w)  seem optimal for growth (Cowey and Sargent 1972).  With protein providing a  considerable portion of the total caloric intake in omnivores, it would not be surprising if daily routine metabolism was covered in large part by amino acid catabolism. On average, it is estimated that amino acid oxidation accounts for 10-20% of maintenance energy costs (Jobling 1980; Kutty, 1978). This value can possibly double after long-term starvation and may be elevated considerably, on a shorter-term basis, immediately after feeding (Kutty 1978). With starvation, increased degradation of muscle-protein is most evident after body reserves of fat have been depleted (Black and Love 1986).  Similarly, amino acid  utilization seems important in migratory salmonids late in the run when other fuel stores have been nearly exhausted (Mommsen et al. 1980). Taken together, these observations suggest that in normal (fed, non-migratory) fish the availability of amino acids for routine oxidative metabolism is influenced by post-feeding absorptive mechanisms more so than by the actual turnover of body-protein. The effects of activity metabolism on protein or amino acid dynamics in fish have not been examined extensively. Continuous low level swimming increases protein turnover and degradation in feeding rainbow trout (Houlihan and Laurent 1987), but the proportion of amino acids utilized for swimming energetics has not been determined directly. Increased aspartate amino-transferase activity in endurance trained red muscle of brook trout (Moon and Johnston 1980b) may be more of a response to enhancement of the level of protein synthesis that is retained as muscle growth (Houlihan and Laurent 1987), rather than catabolic use of amino acid. In salmon starved for 22 days, an indirect assessment of  7  whole-body amino acid utilization (ammonia excretion) indicated that, despite a probable increase in protein degradation, daily increases in routine metabolism were seemingly not supported by amino acids as long as reserves of fat and carbohydrate were available (Brett and Zala 1975; Kutty 1978).  Ammonia excretion largely reflects amino acid  transdeamination in the liver (Van Waarde 1983) and will therefore over-estimate rates of amino acid oxidation by the extent that the resulting carbon skeletons are used in gluconeogenesis, which increases during activity and starvation (Suarez and Mommsen 1987). In any case, it seems likely that the higher aerobic demands of muscle resulting from a rest to exercise transition, in both normal and food-deprived fish, would rely even more on non-protein related energy reserves since the tissue pools of free amino acids are small and are not necessarily available for oxidation (Table 1). In addition, the oxygen cost of ATP production from fat and carbohydrate oxidation is about 30% lower than that of amino acids (Ferrannini 1988). In support of low rates of amino acid oxidation, it has been noted that oxidation of neutral amino acids (serine, glycine, alanine) does not occur at detectable rates in mitochondria isolated from carp red and white muscle (Moyes et al. 1989).  Furthermore, one study of in vivo substrate oxidation rates indicated that  glutamate, alanine and leucine, the latter classed as a branch-chained amino acid which is possibly oxidized at elevated rates in exercising skeletal muscle (Kasperek et al. 1985, 1987), seemed to contribute very little to whole-body oxidative metabolism of rainbow trout swimming at 80% of Ucrit (van den Thillart 1986). Exercising fish may be able to draw on dietary amino acids after feeding as oxidative substrates for swimming.  However, confirmation of such use of amino acids  awaits more detailed examination of post-feeding fluxes and oxidation rates.  In most  studies of swimming metabolism the researcher avoids post-feeding energetics by design in order to minimize inter-individual variability in basal oxygen consumption.  Direct  circulatory infusions of amino acid mixtures (see Tappy et a!. 1992) could be a more useful experimental approach for looking at the effects of amino acid availability on fuel  8  selection in fish. In this way it may be possible to mimic post-feeding levels of plasma amino acids while at the same time eliminate potential influences of gut absorptive rate on whole-body metabolism.  Utilization of Fat Fuels  Currently, the major gap in our understanding of fuel  utilization during steady-state exercise concerns the relative importance of lipid fuels. In total, the body stores of triacylglycerol (TAG) represent the largest energy reserve in fish (Table 1) which could, if delivered exclusively to the red muscle, supply enough free fatty acid (FFA) to fuel about 100 h of maximal aerobic red muscle contraction in a 1 kg animal (assuming maximal power output of 5  -  1 for red muscle (Altringham and 8 mWg  Johnston 1990) which converts to about 0.21  -  , about 23 mol g 1 0.34 mmol 02 min  02 utilized per mol fatty acid oxidized). However, there are few empirical measurements that help to identify which of major depots are mobilized in vivo during exercise. Such studies are complicated by the fact that TAG is deposited in, and mobilized from, muscle, liver and the viscera (Sheridan 1988). Quantitative approaches are needed to increase our awareness of the importance of circulating FFA, extramuscular depot fats and of the dynamics of skeletal muscle intracellular and pericellular TAG stores.  Presently, any  conclusions regarding the relative importance of the various lipid stores, and lipid fuels in general, are limited to studies which focus on either changes in lipid content (eg. observations of whole-body and organ fat depletion during starvation/migration; Idler and Bitners 1959; Jezierska et a!. 1982; Black and Love 1986; Jobling 1980) or on in vitro enzyme measurements and mitochondrial capacities for fatty acid utilization. Such studies  support a major role for lipids in steady-state exercise in fish. Although hepatic and skeletal muscle TAG contents are similar (per g tissue), the smaller liver can be viewed as a minor storage organ with respect to steady-state exercise, regardless of the kinetics of hepatic mobilization.  On a quantitative basis, visceral fat  deposits are the richest store of lipid per g tissue, but skeletal muscle, by virtue of its mass  9  possesses similar TAG stores per kg body mass (Table 1). Each of these stores might be drawn upon during aerobic exercise. The degree to which the white muscle and visceral fat may supply FFA to red muscle should be reflected in estimates of circulatory delivery. However, one study of in vivo palmitate oxidation suggests that circulatory fatty acids contribute minimally to the total oxidative fuel required by trout swimming at 85% of Uj (Van den Thillart 1986). Possible delays in the equilibration of labeled circulatory precursor C-palmitate) 14 with the intracellular TAG pool at the site of utilization may ( result in underestimates of circulatory FFA oxidation (Heiling et a!. 1991), but it is not known if this is a problem in fish at different levels of activity.  Among endothermic  species, the likelihood that intramuscular lipid supports aerobic energy demands, at least to some extent, is supported by the observations of increased stores in aerobically trained muscle (Hoppeler and Billeter 1991) and by the proximity and presumed functional association between mitochondria and intracellular lipid droplets in oxidative fibers (Hoppeler and Billeter 1991; Grunyer and George 1969).  The possibility that  intramuscular TAG serves simultaneously as a sink for circulatory fuels and a source of mitochondrial substrate needs to be evaluated more closely in fish. As with most vertebrate oxidative muscles, red muscle from teleost demonstrates a high capacity to utilize FFA, as indicated by mitochondrial oxidation rates (Moyes et at. 1989; 1990; 1992a). While this in itself reveals little about the relative importance of fatty acids, comparisons of mitochondrial oxidation rates and carnitine palmitoyl transferase (CPT) activities between species and tissues are more illuminating. As in aerobic training of mammalian muscles, where an increase in the capacity to utilize fatty acids is apparent, a comparison of fish species and muscle types indicates that CPT activity increases with aerobic capacity of the tissue (Moyes et at. 1992a). This change is due to both an increase in quantity of mitochondria and CPT activity per mg of mitochondrial protein. Thus, fatty acids may become relatively more important in tissues (white muscle vs. red) and species (carp vs. tuna) with greater aerobic capacities (Moyes et a!. 1992a).  In salmonids,  10  endurance training has been shown to increase activities of fat utilizing enzymes (Moon and Johnston 1980b), also suggesting enhancement of FFA oxidation in response to improved aerobic capacity. These observations agree fundamentally with conclusions from comparisons of fuel demands in endotherms and suggest that adaptation for increased aerobic capacity is related to an increased dependence on fat fuels (Driedzic et al. 1987; Weber 1992).  Carbohydrate utilization. There are potentially three ways to supply carbohydrate to active fish red muscle  -  (i) mobilization of intramuscular glycogen reserves, (ii)  translocation of lactate from white muscle to red via the circulation (iii) circulatory glucose derived from hepatic gluconeogenesis/glycogenolysis. Intramuscular glycogen is not expected to be the primary fuel for steady state exercise since total red muscle glycogen is a fairly small depot (Table 1).  If used  exclusively, red muscle glycogen would support sustained maximal aerobic swimming for less than 1 hr (again, estimated assuming red muscle power output of 5 (Altringham and Johnston 1990) which converts to an oxygen demand of 0.21  -  -  1 8 mWg 0.34 mmol  ). However, red muscle glycogen content in carp species appears to be 2 g 1 02 min  -  8  fold higher than that of white muscle (Johnston and Goldspink 1973a; Johnston 1977; Johnston et al. 1977) providing some evidence for a possibly greater reliance on intramuscular carbohydrate for swimming metabolism than is expected in salmonids. Mobilization of red muscle glycogen has been shown to increase in relation to swim speed in carp during a graded swimming protocol (Johnston and Goldspink 1973b). In other species, the importance of glycogen is implied indirectly in endurance training trials which result in about a 5-fold average increase in red muscle glycogen content of trout (Hochachka et al. 1961) and of coalfish (Johnston and Moon 1980a), where similar  changes in phosphofructokinase (PFK) activity are observed. In brook trout, endurance training had no effect on red muscle glycogen content (a slight decrease perhaps) and no  11  effect on PFK activity (Johnston and Moon 1980b). These changes in brook trout occurred concomitantly with increased B-hydroxyacyl dehydrogenase activity, consistent with the possibility of enhanced fat dependence in salmonids with higher aerobic capacities. In all species, glycogen may be a minor oxidative fuel for extended aerobic exercise, although it may augment fuel supply, particularly at higher intensities and in the early phases of a rest to work transition (Spiret et a!. 1990). At lower exercise intensities red muscle glycogen could theoetically increase in importance, if only because its depletion would be extended over a longer period. Glycogen mobilization should be examined more closely at differnt exercise intensities and in connection with relative fluxes of competing substrates.  Another route for the delivery of exogenous carbohydrate fuel to working oxidative muscle may be via the steady-state release of lactate from white glycolytic muscle (Weber 1991). In mammals, the fate of such lactate is predominantly oxidation resulting from the shuttling of lactate to oxidative tissues via the circulation (Brooks 1987). It is intriguing to consider that white muscle in fish, with its large glycogen stores, could supply lactate to red muscle in the same manner. The total white muscle mass represents a glycogen store that is about 20-fold greater than hepatic stores (Table 1) and could support hours of intense steady-state swimming (assuming that the lactate released is delivered exclusively to red muscle and a red muscle lactate demand of 70  -  112 molmin’kg’ body mass  again, a demand based on peak red muscle oxygen consumption of 0.21  -  -  0.34 02  ). If such a pathway operated, lactate would have to be produced by white g 1 molmin muscle at rates which would be reflected in plasma lactate turnover, assuming a conventional circulatory transfer to red muscle. It was pointed out earlier that recruitment of white muscle may occur at high steady state swimming speeds, but there is no evidence that metabolism in white muscle is primarily aerobic under these conditions. Hence, studies in which lactate dynamics have been examined in swimming rainbow trout (Wokoma and Johnston 1981; van den Thillart  12  1986; Weber 1992) have imposed a high level of sustained exercise (80  -  85% of Ucjt)  with the expectation that glycolysis primarily supports white muscle function and lactate released into the circulation serves as a potential oxidative fuel in more aerobic tissues (red muscle and heart). The problem with relying on circulatory lactate as a red muscle fuel seems to be that turnover cannot match estimates of oxidative demand in red muscle (0.21 min 2 0 ) g 1 .  -  0.34 mmol  Lactate turnover would have to increase by about 10-fold to cover the  maximal red muscle metabolic rate. Empirical studies indicate that, on average, the lactate turnover rate during sustained exercise at 80% Ucrit doubles over resting fish (to about 10 kg body mass), similar to that in a salmonid recovering from burst exercise molmin 1 (Milligan and McDonald 1988), but this change does not occur consistently in individual fish (Weber 1992). Thus, it would seem that lactate remains of minor importance as an oxidative fuel during endurance swimming (Weber 1992), although it may be more important at higher levels of Ucrit or in smaller oxidative tissue masses like heart. Although elevated rates of lactate oxidation are predicted in swimming trout (van den Thillart 1986), this does not neccessarily reflect red muscle metabolic events. Rather, this merely points out the fate of the lactate which appears in, and disappears from, the plasma compartment and is not inconsistent with the limited whole-body dependence on lactate as a fuel. As in mammals, oxidation may in fact be the major fate of lactate since hepatic gluconeogenesis from circulatory lactate is minimal in most instances in a variety of fish species (Milhigan and McDonald 1988; van den Thillart 1986; Weber et al. 1986). Although other fuels seem more important for overall metabolism, tissues that conthbute relatively little to the absolute change in metabolic rate with the onset of exercise could still utilize lactate as an oxidizable fuel. Trout myocardium for example, increases its oxygen consumption several-fold at high work intensities, but could rely heavily on circulatory lactate turnover because of its small relative mass and fractional contribution to whole-body  13  oxygen consumption during swimming (Milligan and Farrell 1986; West et al. 1993). This possibility requires further investigation.  The capacity for red muscle to utilize circulatory glucose is indicated indirectly from activities of hexokinase that generally range from 0.5 to 1.5 mol glucose/g tissue/mm or total activity of 35  -  100 molmin 1 in a 1 kg fish (Crabtree and Newsholme  1972; Johnston 1977). It can be calculated (again, assuming 0.21  -  0.34 mol 02 min  kg and assuming about 70 g red muscle per kg body mass) that similar rates of glucose 1 oxidation are required for maximal red muscle function (35 mass).  -  1 body 60 molmin’kg  As an estimate of fuel use, hexokinase measurements suggest that glucose flux  alone could account for the maximal rate of oxygen consumption of fish red muscle. Crabtree and Newsholme (1972) have similarly estimated that the glucose utilizing capacity of trout red muscle could meet the aerobic fuel demands. Can the delivery of circulatory glucose keep up with the fuel demand during exercise? Glucose release from the liver of fish results from net gluconeogenic flux and glycogenolysis.  Both processes are regulated by the action of various hormones and  functional peptides (Mommsen et a!. 1987; Petersen et a!. 1987; Suarez and Mommsen 1987; Mommsen et a!. 1988; Wright et a!. 1989; Reid et a!. 1992), but it is not known if one pathway dominates hepatic glucose production in vivo.  The responsiveness of  glycogen stores to catecholamines (Mommsen et a!. 1988; Wright et al. 1989; Reid et a!. 1992) coupled with the observation that plasma levels of these hormones do not change during submaximal exercise (Ristori and Laurent 1985; Butler et al. 1986), suggests that glucose derived from hepatic glycogen may not be different from resting conditions. In  any case, liver glycogen levels alone (Table 1) could not support extended periods of maximal aerobic swimming. Regardless of the pathway of hepatic glucose production, the rate of production and release to plasma in vivo is probably within the range of 1  -  10  mol/min/kg, estimating from plasma turnover in teleosts under different experimental  14  conditions (Garin et al. 1987). Unfortunately, reliable determinations of salmonid glucose utilization during exercise are not available. Even the upper end of the range of available kg measured in hypoxic trout (Dunn and Hochachka 1987), would 1 molmin data, 10 , represent an imbalance between plasma glucose supply and red muscle glucose demand during exercise.  It is of course expected that glucose in transit though the plasma  compartment is distributed to more tissues than just red muscle, indicating that glucose kg to match the present estimate of 1 delivery would have to exceed 10 molmin oxidative demand for substrate during high intensity aerobic exercise.  There are no  estimates of glucose turnover in exercising fish, although estimates of glucose oxidation do suggest limited dependence on this substrate in exercising rainbow trout. The capacity of red muscle to use glucose (estimated from hexokinase activity) may mean that red muscle uptake can be expanded, perhaps in periods of hypoxia or during exercise in the absence of other fuel-types.  As is the case for lactate, possibly low glucose turnover rates during  exercise could, nevertheless, be significant to smaller tissue masses. Since plasma glucose is poorly regulated in fish (Palmer and Ryman 1972; Hilton and Atkinson 1982; Mommsen and Plisetskaya 1991), in vivo uptake of glucose in fish tissues might be expected to correlate with parameters that affect glucose availability (eg., plasma concentration or changes in regional blood flow). Although an initial estimate of glucose utilization in trout red muscle indicates an imbalance between glucose delivery and oxidative demand, the quantitative importance of glucose specifically in red muscle is unknown.  It also is not known to what extent the estimations made for trout can be  generalized across species which vary in aerobic swimming capabilities.  In addition,  glucose could be particularly important in the relatively small heart tissue which is poised for either glucose or FFA utilization in a number of fishes (Sidell et a!. 1987).  More  investigation of plasma glucose turnover and uptake during exercise, starvation and hypoxia may reveal tissues and conditions in which glucose is quantitatively important in  15  vivo and comparative studies will help identify differences in glucose dependency across species.  White muscle.  In contrast to red muscle, fish white muscle makes up more than 60  % of the body mass (Bone 1978), has a relatively low aerobic capacity (Moyes et a!. 1989; 1992a,b) and relies primarily on intramuscular glycogenolysis and phosphagen hydrolysis for active energy provision across species (Barrett and Connor 1964; Driedzic and Hochachka 1975; Batty and Wardle 1979; Milligan and Wood 1986b; Schulte et a!. 1992). The white muscle mass is recruited during maximal swimming episodes which, if prolonged, lead to exhaustion and dramatic shifts in intramuscular metabolite levels. Although the changes observed in muscle metabolites are fundamentally similar to those that occur in exercised mammals, the slow process of recovery from exercise (and anoxia) in the large homogenous white muscle of fish has proved useful for integrating pathways of metabolite restoration (Mommsen and Hochachka 1988; Schulte et al. 1992) and for examining possible limitations to the rate of recovery (Milligan and McDonald 1988; Moyes et a!. 1992b; Moyes et a!. 1993). Most studies indicate species differences in rates of recovery, yet an entirely general means of restoring carbohydrate status to pre-exercise levels involving the intramuscular reconversion of lactate to glycogen (Batty and Wardle 1979; Milligan and Wood 1986b; Schulte et a!. 1992). It follows that glycogen recovery should occur essentially independently of circulatory glucose disposal, but direct quantification of glucose phosphorylation rate in recovering white muscle, and red muscle, has not been examined. Comparisons of carbohydrate recovery patterns in skeletal muscles of different species is of interest since more aerobically active species tend to display high post-exercise lactate turnover rates, suggesting that lactate is used oxidatively in species with superior aerobic capacities.  16  White muscle recruitment and exercise metabolism.  Recruitment order in swimming  fish is such that white muscle becomes active when the velocity of red muscle exceeds its capacity to produce mechanical power (Rome et a!. 1988). The estimated maximal power output of fish white muscle exceeds that of red muscle by 5 Johnston 1990).  -  7 fold (Altringham and  Unlike the situation in high speed sustained swimming where some  overlap in the recruitment of fiber masses may occur, maximal swimming is dominated by the large homogenous white muscle mass.  Although red muscle continues to shorten  during burst swimming, its contribution to overall power output is negligible (Rome et a!. 1988). In the lab, burst exercise is achieved in a swim tunnel or by manually chasing the fish.  The standard qualitative end-point used in different exercise protocols is fatigue,  characterised by the inability to burst swim continues (Wood 1991).  -  although low-speed cruising capacity  The time-to-fatigue at a given prolonged swim speed is  sometimes reported (Beamish 1978), but the most distinctive quantitative correlates at the time of fatigue relate to the biochemistry of white muscle. White muscle fuel stores are nearly depleted at the time of fatigue, with glycogen, phosphocreatine (PCr) and ATP levels often expended to less than 30 % of pre-exercise concentrations (Milligan and Wood 1986b; Dobson and Hochachka 1987; Mommsen and Hochachka 1988; Schulte et at. 1992). Depletion of these metabolites can be accounted for at exhaustion by intracellular accumulation of lactate, creatine + inorganic phosphate (Pt) and inosine monophosphate (IMP) respectively. These dramatic changes arise at high power output because the high ATP demand cannot be balanced by the low capacity of mitochondria to supply ATP aerobically. The intramuscular retention of certain metabolites also allows estimates of in vivo fluxes in active white muscle. For example, lactate accumulation is estimated at about 2 g in burst swimming trout (Moyes et a!. 1993). At the end of exercise the 1 molnn lactate accumulated is usually twice the molar amount of glycogen that has been depleted,  17  suggesting essentially exclusive dependence on this intramuscular carbon source. Liver has a large glycogen contentlg tissue, but given that total hepatic glycogen represents less than 10% of the intra-muscular stores (Table 1) and it does not change much with exercise (Milligan and Wood 1986b; 1987), it is a relatively unimportant carbon source during burst exercise. The involvement of plasma glucose is likely to be minor, not just because its pool size is small (Table 1), but also because glucose uptake rates presented in chapter 2 (<1 nmolmin’g ) would have to be upregulated by more than three orders of magnitude 1 to account for the lactate accumulation rates expected in trout white muscle. Actual plasma glucose kinetics are difficult to measure in burst swimming fish as a result of the short period needed to induce fatigue (10  -  20 mm) and because of the likelihood that burst  exercise protocols lead to unstable plasma glucose concentrations (Barton and Schreck 1987). With glycogen levels in trout white muscle typically ranging 20  -  40 mo1 glucosyl  units g , white muscle is expected to become quickly substrated-limited during maximal 1 ATP fluxes. Intramuscular ATP levels are buffered for a short period by PCr hydrolysis, a fuel store expected to last less than 30 sec in skeletal muscle during high work rates (Hochachka and Matheson 1992).  Increased P 1 from PCr breakdown is probably  stimulatory to glycogenolysis, while increased proton production from glycolysis can be buffered to some degree by PCr breakdown  -  reasons to expect that these two processes  occur interactively in active muscle (Hoppeler and Billeter 1991), as indicated previously in anoxic fish muscle (van Waarde et a!. 1990). Depletion of ATP content itself occurs rapidly when glycolysis and PCr hydrolysis no longer support energy metabolism.  At  exhaustion, IMP levels mirror ATP depletion (Mommsen and Hochachka 1988; Schulte et a!. 1992), as in mammalian muscle (Meyer and Terjung 1979; Soderlund and Hultman 1991), but the extent of ATP reduction (up to 70 seen in exercised mammalian muscle.  -  90 %) is considerably higher than that  18  It is not known whether these metabolite changes have any direct causal association with fatigue in fish. Exhaustion is sometimes reported with white muscle glycogen levels still in excess of 10 mol glucosyl units g 1 (Schulte et at. 1992), suggesting that, as in mammals, excitation-contraction coupling may be involved with fatigue (Westerbiad et at. 1991). Recovery from such severe perturbations is slow in fish and depends largely on the interaction of intramuscular recovery pathways.  Closer examination of metabolite levels  and fluxes during exercise and recovery is necessary before a quantitative picture of these interactions in fatigue and recovery processes will emerge.  Carbohydrate recovery in fish muscle.  A great deal of research has focused on the  generation and clearance of lactate in fish muscle. Its metabolism is distinguished from that of other vertebrates by the intramuscular retention of lactate produced during exercise and by the relatively slow rate of clearance (8  -  24 h) from muscle.  Some lactate is  released to the circulation during recovery and the peak level varies among species, with higher levels evident in more active fish (Perry et at. 1985; Milligan and Wood 1986b, 1987; Schulte et at. 1992).  In all species, however, the blood compartment is small  relative to the total white muscle mass and peak concentrations of plasma lactate, ranging 25 1),  -  50 % of post-exercise lactate concentrations in white muscle (about 10  -  40 mmolL  represent a small portion of the post-exercise white muscle lactate load. Concentrations  of lactate are only indirect measures of flux and it is evident that post-exercise turnover rates, measured with 14 C-lactate, are increased above rest by 3- and 9-fold (at 10 °C) respectively in flounder (Platichthys stettatus) and salmon (Oncorhynchus kisutch) (Milligan and McDonald 1988) while turnover rates in exercised tuna (25 °C) are on par with rates seen in mammals (Weber 1986).  Lactate turnover rates, like post-exercise  lactate concentrations, are highest in the more active species.  However, turnover rates  may be somewhat misleading in recovery situations in fish since there is evidence for active re-uptake of lactate from the extracellular fluid into the lactate-producing white  19  muscle mass (Turner and Wood 1983).  Turnover will not reflect simple movement of  lactate from the white muscle into the circulation for eventual processing in other tissues. Nevertheless, fluxes of lactate indicate that, at least in salmonids and flatfishes, estimated recovery time would be approximately twice the observed rates if muscle clearance of lactate occured exclusively by efflux into the circulation (Milligan and McDonald 1988). Thus, turnover rates, even if overestimates of net lactate removal from white muscle, cannot completely account for the rate of lactate metabolism in fish white muscle. This is in accord with the repeated contention that fish white muscle lactate is metabolized largely  in situ (Batty and Wardle 1979; Milligan and Wood 1986b; Tang and Boutilier 1991; Schulte et al. 1992) and that Con cycling of lactate carbon is minimal (Weber et al. 1986; Milligan and McDonald 1988). In contrast, other vertebrates metabolize lactate to a large extent by shuttling’ it via the circulation to other tissues for oxidation or gluconeogenesis, as in mammals (Brooks 1986), or perhaps for glycogen synthesis in red muscle fibers, as has been suggested in reptiles (Gleeson and Dalessio 1990). The fate of lactate in fish white muscle has not been traced quantitatively.  A  number of indirect observations point to primarily intramuscular reconversion of lactate to glycogen (glyconeogenesis).  Oxidation is possible (Scarabello et a!. 1991), considering  that a carbon source is needed to fuel glyconeogenesis, but oxidation of only 10  -  20 % of  the post-exercise lactate load would provide sufficient ATP for recovery. The possibility that fat is used for recovery processes, sparing lactate for glyconeogenesis, is suggested from mammalian studies (Bahr et a!. 1991; Bangsbo et a!. 1991) and from the in vitro and  in vivo regulatory features of trout white muscle pyruvate dehydrogenase, which displays only transient activation after exercise and inhibition in the presence of free fatty acid (Moyes et a!. 1992b). Coincident replenishment of glycogen and clearance of lactate, in conjunction with expected low rates of glucose utilization, in fish white muscle strongly suggests in situ conservation of carbohydrate carbon during exercise and recovery. However, quantitative estimates of glycogenic (glycogen formed from glucose) and  20  glyconeogenic fluxes are not available. A better understanding of the relative importance  of glucose to the overall process of glycogen repletion in trout white muscle can help place carbohydrate recovery profiles into a more quantitative framework.  Species like tuna  which display ‘mammal-like’ lactate fluxes, high aerobic capacities and the need to swim continuously may show increased capacity to utilize lactate oxidatively and therefore rely to a greater extent on glycogenesis for recovery of white muscle glycogen stores.  In vivo measurement of glucose utilization.  Most of our knowledge of carbohydrate  utilization for the swimming energetics of fish has come from whole body depletion of fuel stores in migrating species or from in vitro measurements of maximal flux capacity (HK activity or mitochondrial oxidation rates). In vivo observations of glucose kinetics in fish are seldom performed, even though it is often emphasised that such observations would be particularly useful in helping to sort out species differences in the regulation and interaction of inter-organ pathways of metabolite exchange in fishes (see Mommsen and Plisetskaya 1991). Observations of the rate of transit of a metabolite through the circulation are more informative than monitoring shifts in plasma concentration since physiological changes or experimental manipulations which may influence turnover rate are not necessarily accompanied by changes in metabolite concentration.  Tracer determined fluxes offer a  more quantitative view of the capacity for substrate involvement in whole-animal metabolism during exercise, compared to in vitro enzyme measurements and mitochondrial oxidation rates, and provide a means to investigate potential regulatory influences on metabolite availability and disposal in tissues. With determinations of glucose turnover, it is understood that the appearance of glucose in the circulation results from the stimulation of a single process, glucose release  (i.e. glycogenolysis of gluconeogenesis), from essentially a single tissue source, the liver. However, glucose disappearance is divided among the peripheral tissues, making it difficult to assess muscle glucose utilization in specifically the recruited muscle mass  21  during sustained exercise.  In steady state conditions, one way to determine glucose  phosphorylation (utilization) in specific tissues is with bolus injections of 2-deoxyglucose (2-DG), an analogue of glucose that accumulates as 2-DG-phosphate (2-DGP) in non hepatic tissues (Sokoloff et al. 1977).  Irreversible 2-DG phosphorylation can provide  estimates of metabolic rate in tissues that are highly dependent on circulatory glucose for energy metabolism (eg., brian). In tissues with the capacity to utilize multiple substrates, 2-DG phosphorylation can be used to estimate the contribution of glucose to total in vivo tissue energy provision, providing that tissue metabolic rate can be estimated. Furthermore, glucose that is taken up by muscle during a state of net glycogen synthesis is mostly converted to glycogen (Kusunoki et a!. 1993), indicating that 2-DG uptake should provide a reasonable quantitative estimate of in vivo glycogenic flux during recovery from maximal exercise.  When 2-DG uptake measurements are made in concert with  determinations of glucose turnover, it is possible to quantify the relative contribution of specific tissues to total glucose disposal. In chapter 2, radiolabeled tracers of in vivo glucose metabolism were used to examine the importance of circulatory glucose to the energy provision of trout  (Oncorhynchus mykiss) and carp (Cyprinus carpio) during intense aerobic swimming. Previous comparative studies of ‘fast and slow’ fish indicate that fat fuels may be preferred in the better aerobic swimmers (Moyes et a!. 1992a), suggesting indirectly that glucose may be important in poorer aerobic performers.  This pattern of substrate dependency  would be consistent with a more general hypothesis suggested for mammals in which greater use of fat stores is predicted in highly aerobically adapted animals (see Weber 1992). By comparing the results for two species swimming at the same relative speed (80 % U) it is possible to discuss whether glucose might be more important in the poorer aerobic performer. Within this comparative framework, the slower swimming carp was hypothesized to show greater dependence on glucose for energy provision during sustained exercise than the faster swimming trout.  22  Studies of exercise recovery have emphasized white muscle metabolite recovery  profiles and their interactions in the rainbow trout.  Carbohydrate recovery patterns  generally suggest that lactate is a major substrate for glycogen replenishment in white muscle, but the implication that plasma glucose is of minimal importance is largely indirect.  Moderate post-exercise hyperglycemia, sometimes seen in recovering trout  (Milligan and Wood 1986b, Mommsen et al. 1988; Pagnotta and Milligan 1991) may be relevant in this regard.  In chapter 3, glucose utilization in red and white muscle of  recovering trout was examined to test the idea that glycogenesis is a negligible component of white muscle glycogen resynthesis whereas red muscle, because of its small mass and higher perfusion rate, may be more readily able to utilize circulatory glucose for postexercise glycogen restoration.  For comparison, carbohydrate recovery status was also  examined in the white muscle of fast swimming skipjack tuna (Katsuwonus pelamis) to test whether in situ glyconeogenesis was less important in a species that is known to exhibit high post-exercise turnover rates of lactate, the prime glyconeogenic substrate in white muscle.  PUBLICATION STATUS:  Part of the preceding review, the discussion of fuels for  sustained swimming, is my contribution to a book chapter that has been accepted for publication: Moyes, C.D. and West, T.G. (in press). Exercise metabolism of fish. In.  The Biochemistry and Molecular Biology of Fishes vol 4 (eds. P.W. Hochachka and T.P. Mommsen). Amsterdam: Elsevier.  23  CHAPTER 2.  GLUCOSE UTILIZATION DURING SUSTAINED SWIMMING IN  RAINBOW TROUT (Oncorhynchus myldss) AND CARP (Cyprinus carplo)  In vivo utilization of glucose by heart and locomotory muscles of exercising rainbow  trout. INTRODUCTION Skeletal muscle is the principal peripheral site of circulatory glucose disposal in mammals during exercise and, along with fatty acids and lactate, its uptake contributes substantially to energy provision for endurance activity (Weber 1988). In contrast, less is known about the kinetics of glucose and its oxidative role in fish tissues. Plasma turnover and oxidation in most teleosts is slow compared to mammals (van den Thillart 1986; Garin  et a!. 1987; Machado et a!. 1989) and long-term perturbations like starvation/migration suggest limited reliance on plasma glucose as an oxidizable substrate (Mommsen et a!. 1980; Black and Love 1986). In vivo measurements of the circulatory turnover of glucose, while undoubtedly indicative of overall steady-state glucose disposal, are unsatisfactory for estimating utilization by specific fish tissues. Based on fish red muscle and heart enzymes and observations with isolated hearts, there seems to be enough capacity for oxidative muscle to utilize glucose for aerobic contractions in vitro (Crabtree and Newsholme 1972; Lanctin et a!. 1980; Driedzic and Hart 1984). However, verification of the contribution of glucose to tissue oxidation in vivo is more difficult because of the complexities of substrate storage, mobilization, transport and pathway interaction processes in the whole animal (see Weber 1988; 1992).  In  addition, exercise intensity is an important consideration when evaluating muscle fuel use  in vivo since fuel types or stores utilized at high aerobic intensites may be different from that at low intensity. In salmonids, swimming speeds of 80  -  85 % of fatigue or critical  speed (Ujt) can be maintained indefinitely and are expected to place close to maximum  24  steady-state aerobic demands on oxidative muscle.  In the present study the glucose  analogue 2-deoxyglucose (2-DG) was used to assess in vivo glucose uptake in rainbow trout (Onchorhynchus mykiss) red muscle, heart and white muscle during this type of intense aerobic exercise. Direct determinations of glucose utilization in specific tissues give us the oppotunity to test the degree to which the circulatory delivery of glucose supports estimated maximal rates of tissue oxidative metabolism in vivo.  MATERIALS AN]) METHODS Animals.  Rainbow trout (Onchornhynchus mykiss) of both sexes (300  purchased from local suppliers.  -  1200 g) were  Smaller fish (< 600 g) were used for in vitro isolated  heart studies, conducted at the University of British Columbia. In vivo glucose utilization was determined in larger fish at the Bamfield Marine Station, Bamfield B.C. All fish were held in freshwater (15  -  17 °C) and were fed to satiation twice weekly.  2-DG and the lumped constant.  Glucose utilization was calculated from the tissue  content (disintegrations per minute, dpm) of 14 C-2-DG phosphate (2-DGP) and areas under plasma 14 C-2-DG washout curves according to the equation of Ferré et a!. (1985), Tissue dpm 2-DGP UTILIZATION  =  ) 1 (jhmolmin LC  J  (dpm 2-DG/mol glucose)dt  Utilization was corrected for tissue mass and is presented as nmol per gram of muscle per minute. The term LC is a dimensionless ‘lumped’ constant which is necessary to correct for differences between glucose and 2-DG uptake and phosphorylation rates in vivo (see Sokoloff 1983).  Simultaneous determinations of glucose and 2-DG uptake were  determined in vitro from rates of 3 0 (Ashcroft et a!. 1972) and 14 2 H C-2-DGP formed  25  from H 3 -5-glucose and 14 C-2-DG, respectively. The LC was calculated using the equation defined by Ferré et al. (1985),  LC=  tissue 2-DGPI2-DG in medium  glucose utilized/glucose in medium Tissue 2-DGP and 2-DG in the incubation or perfusion medium are presented as dpm 14 C,  while the glucose in the medium and total glucose utilized during the tissue incubation or heart perfusion period are in units of nmol.  Estimation of LC in muscle slices. Fish were netted from the holding tank and killed quickly with a sharp blow on the head. A rectangular incision that encompassed the lateral  line was made in the musculature between the dorsal and adipose fins.  The skin was  peeled back to expose the muscle and the area was flooded with ice cold saline (containing in mM: 127 NaCl, 4.9 KC1, 1.0 CaCl , 3.7 NaHCO 2 , 1.2 MgSO 3 , 2.9 4 4 PO 1.2 2 NaH , PO 11.5 4 2 KH , 4 HPO pH 2 Na ;  =  7.4). A strip of skeletal muscle was dissected free by  gently manouvering a razor blade through the exposed muscle along the long axis of the fish. The strips were trimmed of white muscle and the red muscle slices (0.5 -  -  1.0 cm , 1 2  2 mm thick) were transferred to a vessel of fresh saline (100 ml, bubbled with 99 % 02:  1 % C0 ) containing 5mM glucose. The vessel was placed in a shaking water bath (15 ± 2 1 °C) and the tissues were preincubated for 20 mm.  Ventricular strips were similarly  prepared from hearts that were immersed in saline and cut in half from the base to the apex.  Strips were made by slicing parallel to the freshly exposed surface of spongy  myocardium. Two to four slices of each tissue were prepared from every fish. Following the preincubation, individual slices were transferred to scintillation vials containing 2.5 ml of fresh saline which contained 5H-glucose (0.3 PCi) and 14 3 C-2deoxyglucose (0.15 MCi) (Amersham) and 5 mM unlabelled glucose. The incubation vials were sealed with rubber plugs through which plastic access tubes were affixed for gentle  26  gas infusion. The vials were positioned in the shaker and incubations were run for no longer than 45 mm. Preliminary trials with 3 H-5-glucose indicated that glucose utilization 0 production) remained constant over this period. 2 H 3 (  Two to three blanks containing  only saline and radioisotope were performed with every run to assess background levels of 0. 2 H 3  5H -glucose and dpm 14 Specific activity (SA) of 3 C-2-DG in the medium were  determined by counting 200 l aliquots mixed with 10 ml ACS II (Amersham) on a LKB Rackbeta scintillation counter. Samples of saline (200 JLl) retrieved from the vials following incubation were placed on columns of Dowex-1-borate (1 ml bed volume) to separate glucose from 3 0 2 H produced (Hammerstedt 1973). The samples were allowed to percolate into the column bed by gravity flow before washing the resin with water (5 x lml washes). The washes were collected separately into 20 ml glass scintillation vials assayed for dpm 3 H. Total dpm 3 H for each sample was corrected for background dpm and adjusted to the total incubation volume to calculate 3 0 accumulated. Levels of 14 2 H C-2-DGP were determined in tissues that were homogenized (3 x 30 sec bursts with an Ultra-turrax tissue homogenizer) immediately post-incubation in 1.5 ml of ice cold 7 % perchioric acid (PCA). Homogenates were centrifuged (10000 rpm for 10 mm) and the supernatants were neutralized with 3 M 3 C0 in 0.5 M triethanolamine. 2 K  An aliquot of neutralized  supematant was counted to determine total 14 C (2-DG + 2-DGP) present. Phosphorylated DG was removed from an equal volume by either precipitation with 0.3 N Ba(OH) 2 and 0.3 N Zn(S0 ) (sample:reagents, 1:1.5:1.5) or by anion exchange on a column of DEAE 4 Sephadex AL125 (1 ml bed volume) equilibrated with 20 mM imidazole-Cl, pH 7.2. Extraction on columns involved first counting aliquots (500 d) of neutralized tissue extracts to determine total tissue 1 ’C (2-DG + 2-DGP). Equal volumes of extract were then placed on columns of DEAE-Sephadex A-125 (2 ml bed volume) that had been pre equilibrated with 20 mM imidazole chloride, pH 7.2.  The sample was allowed to  percholate into the resin, then the column was washed with 10 ml of water.  Fractions  27  collected were mixed with 10 ml of aqueous scintillant, counted, corrected for background C-2-DG retrieved from the column. and summed together to give the 14  This value  C-2-DGP. Preliminary subtracted from total 14 C radioactivity gave an estimate of tissue 14 C-2-DG and 14 C-2-DGP, the latter made from reacting 14 Ctesting with known levels of 14 2-DG/2-DG to completion with hexokinase, prepared in fish muscle extracts indicated that C-2-DGP, with similar levels of 14 C-2the anion-exchange column retained > 95 % of 14 DG recovery.  Lumped constants were calculated from the amount of glucose utilized  (calculated from 3 0 production) and 2-DGP produced with the equation presented 2 H earlier.  Estimation of LC in isolated trout hearts.  The effect of glucose utilization rate on  the LC was investigated using isolated perfused trout hearts.  Isolation and cannulation  procedures were as outlined by Farrell et a!. (1989). Isolated hearts were suspended in a saline bath and air-equilibrated perfusate was introduced from water jacketed reservoirs (15 °C).  Output flow from the ventral aorta was monitored continuously as outlined by  Graham and Farrell (1989) and oxygen consumption was calculated from the difference in  oxygen content, measured with a Radiometer electrode and meter, of perfusate entering and leaving the heart. Additionally, an inlet branch was placed at the level of the preload pressure head for infusing sodium cyanide (NaCN). Electrodes attached to the input and output cannula allowed for the electrical control of heart rate. Changes in circulatory glucose utilization were invoked by manipulation of heart  rate and input and output pressure heads to reduce or increase cardiac power output, and by infusion of NaCN (1 mM) at low power output.  Power output, in milliwatts/gram  ventricle mass (mWIg), was calculated according to Graham and Farrell (1989). Isotope perfusions were initiated only after output flow recordings became steady under the desired conditions with ‘unlabelled perfusate’.  Low power perfusions with and without NaCN  28  were run for 40  -  60 mm, while the higher power output perfusions were stopped after 15  -  20mm. Glucose utilization and 2-DGP formation were determined in hearts with single pass perfusions of 5H-glucose (0.1 3  ) 1 ,iCimr  and 14 C-2-DG (0.02 1 Ciml ) .  Following  perfusion, hearts were removed from the saline bath, blotted, and immersed in pre weighed vials of 7 % PCA (2 ml).  Tissues were homogenized, then neutralized and  assayed for 14 C-2-DGP. Perfusate collected at the afterload outlet was assayed for lactate according to Bergmeyer (1985) and 3 0 by vacuum distillation. For the latter, a flask 2 H containing outflow perfusate (3 ml) was attached to a micro condenser with an 8 cm still head, then immersed in a water bath (60  -  65 °C) where the perfusate was brought to  boiling under constant vacuum (-600 mm Hg). aliquots (1 ml) were counted for 3 H and 14 C.  All of the distillate was collected and The lack of detectable 14 C above  background level indicated no contamination of raw perfusate in the distillate. Perfusion times and flow rates from the afterload outlet were monitored to calculate cardiac output and to determine total 3 0 produced and total amounts of isotope and substrate to pass 2 H each heart to calculate LC’s.  Glucose utilization in swimming trout.  Rainbow trout were anesthetized (0.5 g MS  2221k buffered with 1 3 g.NaHCO 1 ’) and placed in a standard fish operating sling. Oxygenated water containing a light level of buffered MS-222 (0.1 gt ) was circulated 1 over the gills, and dorsal aortic cannulae (PE-50, Clay Adams) were implanted. recovered from anesthesia (12  -  Fish  18 h) in dark plastic tubes (15 cm diameter) which were  submerged in a flow through water (15  -  17 °C) resevoir at the downstream end of a Brett  type swim tunnel. Once recovered, each fish was handled separately. A fish was guided to the entrance of the tunnel and each swam at its own accord from the recovery tube into the swim space.  Water speed was adjusted to about 0.5 body-lengths(bl) sec 4 (15-20  ) and the fish was left for approximately 10 h before the experiment was started. 1 cmsec  29  Fish used to determine resting rates of glucose utilization recovered from anesthesia in black perspex boxes, supplied with fresh flow-through water (15 °C). A high sustainable swimming speed was found by increasing water velocity every t increments until the fish began to demonstrate ‘burst and glide’ 15 mm by 20 cmsec swimming. Water velocity was then reduced until the fish held its position against the flow of water. This velocity was maintained for 1 h prior to isotope injection. Bolus injections consisted of 14 C-2-DG (5 MCi) that was pre-dried under nitrogen and reconstituted in 250 1 .d of Cortland saline (Wolf 1963). An aliquot (10 ILl) was taken to determine the dpm of 14 C injected and the remaining volume was injected into the dorsal aorta via the cannula. The cannula was flushed with two volumes of saline. Blood (150 d) was drawn through the same cannula at 1, 2, 3, 4, 5, 10, 20, 40, 60, 90 and 120 mm 1 post-injection. Neutralized PCA (1:1, 7 % PCA: sample) extracts were assayed for dpm C in glucose and plasma glucose concentration (Sigma glucose assay kit). 14  Plasma  washout kinetics were determined using a curve-stripping program (JANA; Statistical Consultants, Lexington, KY) and areas under curves were determined (MATHCAD; Mathsoft Inc., Cambridge Ma.). When blood sampling was completed in each fish the spinal cord was severed just posterior to the head and tissues were removed and freezeclamped between Wollenberger tongs that were pre-cooled on dry ice. A double bladed cleaver was used to rapidly cut a steak from the musculature immediately posterior to the dorsal fm.  The heart was removed and the ventricle was quickly cleared of blood and  freeze-clamped. The delay between death of the fish and freeze clamping of tissues was < 20 sec. Tissues were stored at -80 °C until assayed for the content of 2-DGP.  Statistics.  The significance (p < 0.05) of differences between mean (± SE) LC’s and  glucose utilization rates was determined using unpaired t-tests or ANOVA with Tukey’s multiple comparison test.  30  RESULTS Tissue LC’s and the effect of glycolytic rate.  Table 2 lists LC’s for red muscle and  heart ventricle from tissue incubations. The LC’s calculated were quite variable, although  most values were between 0.2  -  0.6. Box plots in Table 2 provide an indication of the  range and clustering of data points around the median LC for each tissue. LC’s calculated for red muscle and heart were not significantly different. The isolated trout heart preparation provided a means for assessing whether changes in perfusate glucose uptake affected the 2-DG LC (Table 3). The use of perfusate glucose ranged from trace levels in the substrate free perfusions to 250 1 min in the NaCN nmolg  perfusions. However, there was no effect of utilization rate on the LC’s calculated from the various perfusions. Interestingly, power output of the NaCN hearts spanned a narrow subphysiological range (0.04  -  0.17 mWg ), yet potent enhancement of perfusate glucose utilization 1  occurred within this group in response to small increases in power output (Figure 1). The  variability is likely the reason for the glucose utilization not being significantly different from the high work rate hearts (Table 3). We did not attempt experiments to confirm that power output was in fact the principal factor affecting glucose utilization during NaCN perfusions. As presented in Figure 1, the regression of glucose utilization against cardiac power output is y  =  2003x + 49, r  =  0.70. Individual LC’s for these hearts are also  depicted, but there is no indication that the formation of 3 0 glucose and the 2 H accumulation of 14 C-2-DGP changed disproportionately with increased perfusate glucose utilization. All in vivo red muscle and heart glucose utilization rates were calculated using the average LC determined from these in vitro experiments (LC = 0.40).  Glucose utilization in swimming trout.  Plasma  profiles  of  dpm  C-2-DGmo1 14  1 and of glucose concentration in trout are shown in Figure 2. Small fluctuations glucose  Lumped constants (LC) calculated from in vitro incubations of trout red muscle slices and heart  0.2  0  LC  0.4  I  0.6  I  0.43 ± 0.06  LC  I  48.1 ± 9.7  GLUCOSE UTILIZED (nmol)  I  760 ± 160  TISSUE 2-DGP (dpm)  RED MUSCLE (n=15)  0.8  I  .  1.0  I  red muscle and heart LC’s with data outside 5th and 95th percentiles shown  0  I  (.).  0.2  I  0.4  I  LC  0.6  I  0.38 ± 0.05  76.2 ± 13.8  1130 ± 240  HEART (n=18)  0.8  I  1.0  I  dpm and 421900 ± 48300 dpm for the red muscle and heart incubations, respectively. Box plots are presented for  ventricular strips with 14 C-2-DG and 3 5H -glucose. Total 14 C-2-DG in 2.5 ml of saline averaged 459800 ± 57800  TABLE 2.  w  24.8 ± 2.0  6.6 ± 1.3  High Work (N=4)  Cyanide (N=6)  5.8 ± 0.7  Cyanide (N=3) 0.10 ± 0.02  1.22 ± 0.11  0.10 ± 0.02  1.69 ± 0.09  0.07 ± 0.02  ) 1 (mWg  POWER OUTPUT  1296 ± 178’  1468 ± 139  212 ± 34  LACTATh PRODUCTION  791 ± 72’  182 ± 59’  1012 ± 129a,d  118 ± 28°  49 ± 13  1 min (nmolg ) 1  OXYGEN CONSUMPTION  a. Significantly different from low work load (p < 0.05) b. Not significantly different from glucose perfused high work load (p = 0.17). c. Significantly different from glucose free, high work load (p < 0.001). d. Significantly different from glucose perfused high work load (p <0.01). * Not different from oxygen and lactate in the corresponding glucose perfused hearts.  22.7 ± 2.3  High Work (N=3)  Glucose Free Prefusion  4.5 ± 0.8  Low Work (N=4)  t min (mlkg ) 1  Perfusate Glucose, 5 mM  CARDIAC OUTPUT  consumption and lactate production per gram of ventricle mass were also determined.  43500 ± 2400w’  15400 ± 500  ) t (dpmg’ min  57a,b  0.41 ± 0.03  0.39 ± 0.04  0.40 ± 0.03  0.41 ± 0.06  121 ± 22a 253 ±  0.41 ± 0.03  L.C.  29 ± 6  GLUCOSE UTILIZATION  (LC.). Cardiac output was normalized per kilogram body mass, while power output was normalized per gram ventricle mass. Oxygen  TABLE 3. The effects of cardiac power output and NaCN (1mM) perfusion on glucose utilization rate and the 2-DG lumped constant  33  V  0.45 .  400  Z  o  300  200  100  133  I”  0.3  O.32J0.45 V..,  I  0.48  • .  •$  0.0  FIGURE 1.  1.5 POWER OUTPUT (mW.j’) 0.5  1.0  2.0  Glucose utilization in relation to power output in  normoxic (circles) and NaCN treated (triangles) hearts.  LC’s  calculated for individual NaCN perfusions are also depicted.  34  in plasma glucose were typical of individual resting and exercising fish. Dynamic steadystate was assumed if during the sampling period the coefficient of variation (CV) for the mean plasma glucose level was  15 %. The 14 C activity washout curves were all fit to  equations described by two exponential tenns. Table 4 lists data for individual resting and swimming trout. Relative swimming speed was very reproducible with the exercise protocol used. The average relative speed, in bi/sec, corresponded to an absolute velocity of 71.5 ± 0.78 1 .cmsec P1asma glucose ranged from 2  -  5 mM among both control and exercised fish. The in vivo rate of muscle  glucose utilization in unexercised trout was highest in heart ventricle, being about 6-fold higher than in red muscle.  However, during sustained steady-state exercise, cardiac  glucose utilization did not change significantly compared to resting fish. Conversly, red muscle utilization increased from less than 1 nmolg min at rest to about 21 nmolg 1 min representing a 24-fold change in the use of circulatory glucose. White muscle , 1 utilization, estimated assuming that the LC =0.4 for this tissue, averaged less than 0.5 1 at rest and during exercise. nmolg’min  DISCUSSION The 2-DG lumped constant.  The LC as described by Sokoloff (1983) for normal  brain tissue of mammals is different depending on the species studied (ranging from 0.3  -  0.6), but seems invariable within a species when determined under a variety of physiological conditions. Similarly, the LC calculated for rat skeletal muscle is influenced minimally by treatment induced changes in glucose utilization (Mészáros et al. 1987; Furhler et cii. 1991).  Despite the apparent stability of the LC with respect to tissue  treatment it is still difficult to generalize, even within a species.  The LC chosen as  characteristic for rat skeletal muscle in different studies ranges from 0.4 to 1.0 (Ferré et cii. 1985; Furhler et cii. 1991; Mészáros et al. 1987), which perhaps indicates that  35  100000 Cd)  o o  C)  10000  1000  E 100 0  20  40  60  100  80  120  Time 100000 ‘—4  Cl)  0 0  ‘  10000  C  II  o  U  •  I  _I_  I  0 20406080100120 Time (mm)  1000  E 100  I  20  40  I  I  60  80  I  100  120  Time  14 in resting (upper panel) and FIGURE 2. Plasma washout of C-2-DG swimming (lower panel) rainbow trout.. Inset axes show profiles of plasma glucose concentration during the blood sampling period.  36  attributes of individual muscle groups, such as fiber type distribution and glucose transporter type or density, can affect LC determination. In addition, mammalian tissues other than brain and skeletal muscle may be more sensitive to changes in experimental conditions. Indeed, the LC for rabbit myocardium is on average 0.6, but extremes of flow rate and contraction frequency, albeit beyond the range of physiological relevance, were shown to have significant effects on the LC in perfusions of isolated interventricular septa (Krivokapich et a!. 1987).  The use of 2-DG methodology will clearly produce fewer  ambiguities when extrapolating LC’s to the whole animal if in vitro determinations are made in the tissues of interest in concert with the in vivo experimental design. One point to make about the in vitro analysis of trout tissues is that the LC determined with isolated heart preparation was generally more reproducible than with the slices. Despite the fact that slices were preincubated for 20 mm, the broad range of LC’s may in part reflect a varying degree of cellular damage since disruption of the plasma membrane could have altered the importance of the transport component of the LC. Determinations of the LC with the more intact isolated heart preparation were always between 0.3  -  0.6 (see Figure 1, for example). Nevertheless, the LC produced from the  two protocols is similar and the indication is that an LC of 0.4 is a reasonable application to both heart and red muscle calculations of in vivo glucose utilization. The additional utility of the isolated trout heart was in the determination of LC’s during variable rates of perfusate glucose uptake. The high power output hearts used in the present study matched basal power output levels set normally in isolated and in situ trout hearts (1.0  -  2.0 mWg’ , Farrell et a!. 1989; Milligan and Farrell 1991). Maximal work 1  loads in our isolated preparations were not stable long enough to reliably coordinate steadystate cardiac parameters with isotope perfusions. For the same reason, the NaCN perfused hearts had to be set at subphysiological work levels.  However, the advantage of this  protocol was that a broad range, of glucose utilization could be induced.  The results  indicate that with stimulation of glycolytic rate by anoxia or changes in power output the  37  rate of 2-DG uptake and phosphorylation should remain a constant proportion of glucose utilization. Furthermore, both the aerobic and NaCN perfusions lacking unlabelled glucose shows that even if cardiac function is dominated by endogenous fuel utilization, the LC calculated from tracer uptake is unaffected.  In vivo glucose utilization in red muscle. For rainbow trout of the size used in this study, an average steady-state swimming speed of 1.7 body lengthssec 1 corresponds to approximately 80 % of U (Kiceniuk and Jones 1977). From rest to this high level of sustained swimming, rates of 2-DGP accumulation indicate that glucose utilization increased by 28-fold in the lateral red muscle.  What proportion of the in vivo energy  production in red muscle is accounted for by glucose? The red muscle of trout constitutes about 5  -  10 % of the total muscle mass (Webb  1971). A 1 kg trout, possessing up to 70 g red muscle, consumes about 25 mol 0 kg 2 min at rest and increases this by about 7 fold when swimming aerobically at close to 80 1 % of U (Kiceniuk and Jones 1977). Swimming at this intensity results mainly from the power output of the active red muscles (reviewed by Jones 1982) and given the concomitant redistribution of blood flow to these fibers (Randall and Daxboeck 1982), it is likely that the cost of swimming, above routine maintenance costs, becomes associated largely with the increased aerobic demand of this muscle mass. Assuming that glucose is fully oxidized in red muscle (6 nmol 02 per nmol glucose), it is then apparent that, given an uptake rate of 21 nmol 1 min circulatory glucose is maximally 5 % of the glucoseg , total red muscle substrate oxidation in vivo. Other exercise-induced changes in whole-body metabolic costs are not accounted for in the preceding calculation. It is an oversimplification to attribute 100 % of the cost of swimming to red muscle and this first approximation of in vivo red muscle glucose oxidation is probably an underestimate. However, osmoregulatory costs do not appear to change  with  increased  aerobic  exercise,  although  this  is  probably  species  1.68 ± 0.02  803 ± 62 1066 898 1079 1096 798 805 740  926 ± 57  Mean ± SE  EXERCISED 1 2 3 4 5 6 7  Mean ± SE  2.11 2.32 2.43 2.35 2.71 3.52 2.78  2.32 2.77 2.83 2.51 3.69 4.61 2.49 3.93 3.29  ± ± ± ± ± ± ±  0.17 0.08 0.07 0.10 0.09 0.14 0.11  ± 0.10 ± 0.09 ± 0.10 ± 0.07 ± 0.14 ± 0.20 ± 0.10 ± 0.15 ±0.11  PLASMA [GLUCOSE] (mM)  significantly different from resting control, p < 0.001.  1.68 1.61 1.62 1.67 1.76 1.76 1.66  983 650 875 601 944 1120 607 675 776  SWIMMING SPEED (bl/sec)  RESTING 1 2 3 4 5 6 7 8 9  BODY MASS (g)  V  V  5.31 ± 1.04 0.87 ± 0.15  20.85 ± 5.52  18.91 4.93 30.93 11.17 48.44 12.96 18.63  0.36 ± 0.10  8.91 8.85 2.50 1.78 1.94 5.22  0.60 0.40 0.96 0.12 0.83 0.20 0.22 0.48 ± 0.12  1.81 1.85 6.31 4.46 13.05 0.90 4.58 4.62 ± 1.60  ----  0.20 0.40 1.04 0.20 0.29 0.20 0.23 0.31  6.63 6.65  min 1 (nmolg )  HEART  1.39 0.83 1.34 0.37 1.11 0.20 0.75 0.99  ----  RED MUSCLE  WHITE MUSCLE  An in vivo 2-  GLUCOSE UTILIZATION  In vivo glucose utilization in red muscle and heart of resting and swimming rainbow trout.  deoxyglucose lumped constant of 0.40 was used for calculations of tissue glucose uptake.  TABLE 4.  cz  39  dependent (Pérez-Pinzón and Lutz 1991). In addition, combined oxidation in other muscle types, heart because of its small mass and white muscle because its peak aerobic demand is expected at speeds < 80 % of Uj or in recovery from intense burst exercise (Moyes et al. 1992b; Scarabello et al. 1991), is probably minor compared to red muscle oxidative demand during exercise. Nevertheless, EMG’s have been recorded from the white muscle  of salmonids during sustained swimming (Johnston and Moon 1980) and given that the oxidative capacity of trout white muscle is relatively large compared to slower swimming fish like carp (9.3 moPmin g citrate cynthase and about 8 mg mitochondrial protein 1 per gram white muscle in trout versus 3.5 mo1min’g’ and 3.1 mg protein in carp; Moyes et al. 1989, 1992b) and given the large mass of this tissue, aerobic white muscle recruitment could be a significant metabolic cost in swimming trout.  Activity related  changes in costs of white muscle recruitment, ion regulation and ventilation are difficult to quantify in vivo, but it is apparent that if such costs diverted one-half of the whole-animal oxygen consumption at 80 % U then an estimate of glucose oxidation in red muscle would change only slightly, to 10 % of total substrate oxidation. It is interesting that a large increase in glucose utilization in exercising red muscle (28-fold above resting) is seemingly unimportant to red muscle energetics. It may be that a major determinant of red muscle uptake of circulatory glucose is simply the redistribution of blood flow during exercise. Most of the cardiac output is delivered to the active red fibers in swimming trout (Randall and Daxboeck 1982) and it is evident that muscle glucose uptake correlates postively with bloodflow (Chaliss et a!. 1986).  Although the  metabolic demand of red muscle increases, the use of glucose might be limited by the availability of plasma glucose or by potent glycolytic inhibition brought on by the use of fatty acids in highly oxidative muscle (Kobayashi and Neely 1979).  As a result, the  potential contribution of glucose to overall red muscle substrate oxidation would be small, as calculated, despite a several fold increase in uptake over resting rates.  40  It may be that lipid based substrates are utilized for sustained aerobic swimming in trout. This possiblity is strengthened by the likelihood that carbohydrates from all in vivo sources are a small component of red muscle oxidation. Blood-borne lactate is a minor fuel in trout during an endurance-type swim (Weber 1991) and red muscle glycogen, although possibly important during submaximal swimming in some species (Johnston and Goldspink 1973), is available in limited supply in trout (20 molg 1 Parkhouse et al. 1987) and cannot solely furnish fuel for intense aerobic swimming. The use of fat fuels has not been quantified with in vivo methodologies, but the potential for reliance on fatty acids in teleosts is suggested at least indirectly from intramuscular storage and mobilization capacities, from triglyceride depletion studies in migrating/starving species and from in  vitro measurements of flux capacity (Black and Love 1986; Mommsen et al. 1980; Sheridan 1988; Moyes et a!. 1992a).  White Muscle Glucose Uptake.  Recruitment of white fibers in rainbow trout is not  expected at swimming speeds up to about 80 % of Uj (Webb 1971). As was discussed earlier, however, recruitment of this fiber mass cannot be ruled out, particularly since just one velocity increment above 80 % Uj induced burst and glide behavior in these trout. However, low glucose utilization in white muscle is not unexpected since circulatory substrates are probably of minimal importance in this tissue during exercise. A reduction in blood flow to the white fibers at 80 % U (Randall and Daxboeck 1982) is consistent with the view that any recruitment of this muscle mass is fueled mostly by mobilization of intracellular fuels.  About 95 % of the blood flow to the total mosaic musculature of  rainbow trout at 80 % U could be recieved by the small proportion of ‘red’ fibers dispersed throughout the white muscle mass (Randall and Daxboeck 1982), suggesting that white fiber recruitment may be fueled by intramuscular glycogenolysis. Glucose kinetics in the deeper red fibers might well behave as in the lateral red muscle during exercise, but  41  changes would most likely always be masked by glucose uptake, or a lack of glucose uptake, in the larger white fiber population.  Cardiac glucose utifization. In vivo glucose utilization in trout heart was not affected by increased aerobic swimming.  Before estimating the contribution of glucose to cardiac  energy production, it should be noted that cardiac power output in isolated trout hearts varies from 1  -  2 mWg’ ventricle mass to maximally 6  -  8 mWg (see Milligan and  Farrell 1991). Interpreting this as an in vivo scope for cardiac power output means that gmin 2 realistic limits of oxygen consumption range from 1 (rest) to 4 (exercise) mol 0 1 (assumes a cardiac efficiency of 20 %, Graham and Farrell 1989; and a caloric  equivalent of 02 of about 4.8 kcalml). Oxygen consumption by isolated hearts in this  study agree with the lower end of this estimated in vivo range (see Table 3). An in vivo rate of glucose utilization of approximately 5 nmolgmin indicates that circulatory glucose accounted for about 6 % of the expected cardiac oxygen uptake at rest and less than 1 % during exercise. As with red muscle metabolism, there are still some inadequacies in our understanding of the importance of various circulatory and endogenous fuels in fish hearts. Evidence that fat is utilized in the absence of any glycolytic flux comes from in vitro observations that iodoacetate treated hearts remain functional when provided palmitate in the perfusate (Driedzic and Hart 1984) and possibly via endogenous triglyceride mobilization (see Milligan and Farrell 1991). However, the effects of fatty acid utilization on glycolytic flux in fish hearts is not known. It may be that the trout heart is similar to mammalian heart models in which fatty acids restrict glucose oxidation, possibly through  combined effects on transport and glycolytic enzymes, and enable sustained cardiac function over a broad work range (Kobayashi and Neely 1979; Saddik and Lopasehuk 1991).  This agrees with calculations of low glucose oxidation in vivo and with  observations that glucose in the absence of other substrates accounts for up to 50 % of the  42  oxygen consumption of isolated trout hearts operating at 1  -  1 (calculated from 2 mWg  Table 2 values). Endogenous fuel stores presumably account for a substantial portion of the oxygen consumed in trout hearts, but quantification of the specific role of myocardial glycogen and triglyceride in relation to work intensity requires more study. Myocardial fuel utilization in vivo could also switch from fat preference during low cardiac work to lactate during elevated aerobic activity, as speculated for skipjack tuna Katsuwonus pelamis (Moyes et a!. 1992a). High concentrations (10 mM) of circulatory  lactate can serve as the sole oxidative substrate in isolated trout hearts over the range of power output (Milligan and Farrell 1991).  Indeed, even at concentrations observed in  plasma of trout performing endurance-type exercise (2  -  3 mM; Weber 1991), lactate will  inhibit cardiac glucose oxidation in vitro (Lanctin et a!. 1980) and can partially alleviate trends of diminishing performance seen in glycolytically inhibited hearts (Driedzic and Hart 1984).  While it is doubtful that lactate is the preferred fuel for whole-body  metabolism in trout (Weber 1991), the doubling of in vivo lactate turnover from rest to 85 % of U (Weber 1991) may be pertinent to the oxidative needs of the relatively small cardiac tissue mass.  Either case of a fat-to-lactate transition in relation to increased  myocardial energy demand or of an overall dependence on fatty acids is compatible with low glucose oxidation in vivo and prompts the generalization that cardiac glucose metabolism in teleosts is directed mainly toward biosynthesis of storage substrates (glycogen and triglyceride), with a potential oxidative role only during periods of fatty acid limitation.  PUBLICATION STATUS:  The preceding study has been published,  see West et a!.  1993 in reference list. I am grateful to my main collaborator on this project, Dr. P.G. Arthur, for setting up the isolated heart preparation and for determining myocardial oxygen consumption.  43  Muscle glucose utilization during sustained swimming in carp. INTRODUCTION The carp (Cyprinus carpio) is typical of teleosts in the sense that estimated in vivo glucose kinetics at rest are relatively slow (Garin et al. 1987). However, comparison of the fat utilizing capacities of carp and the fast-swimming skipjack tuna (Katsuwonus pelamis), suggests that the general hypothesis that better aerobic performers depend to a greater extent on fat-based fuels (eg., see Weber 1992) may also hold for teleosts with different aerobic swimming capacities (Moyes et al. 1992). The small relative mass of heart and red muscle in conjunction with the relatively sluggish swimming characteristics of carp may mean that glucose disposal is significant in oxidative muscle, despite comparatively low in vivo glucose kinetics. In the present study, we investigate the importance of circulatory glucose in the swimming energetics of carp by examining in vivo glucose kinetics and tissue specific glucose uptake, determined with bolus-injections of 3 6H -glucose and 14 C-2-deoxyglucose 14 Simultaneous determination of flux and uptake is useful for evaluating the ( C-2-DG). importance of skeletal muscle in whole-body glucose disposal. Furthermore, measurement of tissue 2-DG uptake provides a means of detecting exercise-stimulation of glucose disposal in small tissue masses (eg., heart and red muscle) which may be mediated by redistribution of blood flow or changes in muscle transporters in the absence of direct effects on whole-body glucose flux.  As in the previous study of trout, the starting  assumption for estimating oxidation of glucose was that red muscle accounted for the bulk of the whole body metabolic costs at 80 %  Since the fate of glucose in working  muscle need not be oxidation (L.aughlin et a!. 1992), 2-DG uptake is therefore used as an indicator of the maximum potential for oxidative glucose disposal in muscle during  exercise.  44  MATERIALS AND METHODS Animals.  Carp (1  -  2 kg) of both sexes were purchased from a local supplier and  maintained in dechlorinated fresh water (10  -  13 °C) in the aquarium facility at the  Department of Zoology, U.B.C. Fish were held for at least one month before being used in experiments and were fed twice weekly with trout pellets (Moore-Clark, Vancouver).  Estimation of U.j . U, representing the maximum speed sustainable for 30 mm using 1 the present exercise protcol, was determined for individual carp (986  -  1790 g, 35  -  45 cm  total length) using a Brett-type swim-tunnel respirometer (Gehrke et a!. 1990). A fish was placed in the swim-tunnel and water velocity was adjusted to between 0.6 . 1 lengthssec  -  0.7 body  Exercise intensity was maintained at this low level for 2 h.  The  respirometer was then closed to inflowing water for 20 mm and oxygen consumption (mg kg was determined by circulating water from the respirometer at a constant rate h ) 02 1 over a P°2 electrode (Radiometer, Copenhagen). P 2 data was collected and stored using 0 a computerized, data aquisition system described previously (Gehrke et a!. 1990). Thereafter, swimming speed was increased every 30 mm  in 0.3 body lengthssec 1  increments and oxygen consumption was determined at each new speed unless the fish fatigued within the first 10  -  15 mm.  Between estimates of oxygen uptake, the  respirometer was flushed with fresh water and oxygen level was readjusted to 156 mm Hg. The time to fatigue, designated as the time when a mild electric shock (5 volts) no longer induced the fish to swim, was recorded and Uj was calculated using equations reviewed by Beamish (1978). After exercise, fish were allowed to recover in black Perspex fish boxes that were supplied with areated, flow-through fresh water.  Sustained swimming in cannulated carp. Following exercise recovery (48 h), individual carp were anesthetized in buffered MS-222 (tricaine methane sulfonate, 0.2 gL 1 with 0.4 1 3 gL NaHCO ) . Each fish was placed in an operating sling and the gills were irrigated  45  with oxygenated water containing a light level of buffered anesthetic (0.08 g MS-222 L ). 1 A 50 cm cannula (PE-90; Clay Adams, New Jersey) was implanted in the dorsal aorta, sutured to the roof of the mouth and passed through a piece of PE-200 that was secured through the membrane of the distendable upper lip. The cannula was filled with Cortland saline (Wolf 1963) that contained 5 i.u. ml 1 of heparin (Glaxo, Montreal) and the fish was placed in a black perspex fish-box to recover from anesthesia (24 36 h). -  The protocol used to bring carp to a high sustainable swim speed was similar to that used in the preceding study of trout glucose kinetics (West et al. 1993). A fish was placed in the swim tunnel and forced to swim at a low intensity (about 1/3 of a body lengtfrsec ) 1 for 2 h.  Swimming speed was then increased every 5 mm  (in 0.3 body 1engthsec 1  increments) until the fish displayed obvious burst and glide swimming behavior. At this point, water velocity was reduced until the fish could maintain its position in the swim tunnel. Fish were then exercised at this high intensity for a total of 120 mm. Radiolabeled metabolites were administered after the first 60 mm. During the interval between 30  -  60  mm post-injection of radioisotopes (described in the next section), the oxygen consumption of swimming carp was determined as described previously.  Bolus-injections and tissue sampling.  Appropriate volumes of 6H-glucose (15 MCi) 3  and 14 C-2-DG (5 PCi) were dried under a constant stream of N 2 gas. The radiolabeled metabolites were reconstituted together in saline (140  -  180 1 .d). An aliquot (10 l) was  removed immediately, mixed with aqueous scintillant (10 ml of ACS II, Amersham) and counted using a dual-label assay on an LKB Rackbeta scintillation counter. The remaining solution was drawn into a glass Hamilton syringe to determine the volume of bolus to be delivered. A bolus injection was delivered to individual fish through one port of a 3-way stopcock that was attached to the free end of the implanted cannula.  Another syringe  connected to the second port of the stopcock served as a saline rinse (2 volumes) for flushing solution from the glass syringe and the cannula into the fish.  46  Blood (125  -  150 ul) was sampled from both resting and swimming fish at 1, 2, 3,  4, 5, 10, 20, 30 and 60 mm post-injection of radioisotopes. Blood removed from the fish was replaced with an equal volume of saline. After retrieving the final blood sample (60 ) was injected through the cannula in order 1 mm), 2 ml of sodium pentobarbital (65 mgml to kill the fish with minimal struggling. When opercular movements had ceased, a single cross-sectional slice  (-  1 cm thick) of skeletal muscle was removed from immediately  posterior to the dorsal fin and freeze-clamped between aluminum tongs that were pre cooled in liquid nitrogen. The heart was removed and also freeze-clamped. Less than 30 sec elapsed from the time the fish was killed to the moment the last tissue was immersed in liquid nitrogen. Tissue and plasma samples were stored at -80 °C.  Metabolite assays.  Plasma was  separated immediately from blood  samples by  centrifugation and a precise volume was deproteinized with one volume of 0.6 N perchloric acid (PCA).  Protein was precipitated by centrifucation and the plasma was neutralized  d) of the final with 3 M 3 C0 in 0.5 M triethanolamine hydrochloride. Aliquots (50 1 2 K supernatant were placed in 20 ml scintillation vials and evaporated to driness under streams 0. Residues were reconstituted in 1 ml of water and assayed 2 H of N 2 gas to remove any 3 using a dual-label protocol on a LKB Rackbeta scintilation counter to estimate radioactivity C-2-DG. in plasma 6H-glucose and 14 3 The frozen cross-sections of skelatal muscle were immersed in a shallow bath of liquid nitrogen and samples (0.5  -  1.0 g) of red and white muscle were dissected free. The  muscles were chopped into fragments and transferred to pre-weighed tubes that contained approximately 4 volumes of ice-cold 0.6 N PCA. The tubes were re-weighed and samples were homogenized with 2 x 20 sec passes of an Ultra-turrax tissue homogenizer. Two aliquots (100 1 .d) of this homogenate were removed for glycogen analysis.  After  centrifugation (10 mm, 7500 g), the tissue extracts were neutralized, spun again and the  47  resulting supernatant frozen at -80 °C. Cardiac muscle was treated in the same manner, but homogenate samples were not removed for glycogen analysis. Glycogen was measured in aliquots of muscle homogenate and is presented as mol  glucosyl unitsg tissue. Muscle glycogen was digested by incubating the homogenates for 3 h at 40°C with amyloglucosidase (Boehringer Mannheim, 2 mgmr , 1 ml total 1 volume) in acetate buffer, pH 4.8 (Bergmeyer 1985). Perchioric acid (25 d, 70 %) was used to halt the incubation and glucose was determined in the extract after neutralization. Red and white muscle glucosyl units, as well as muscle lactate and plasma glucose, were determined with standard enzymatic assays Bergmeyer (1985) that were modified for use with microtitration plates (0.3 ml) and a Titertek Multiskan plate reader. Muscle 14 C-2-DGP was separated from total tissue 14 C radioactivity using the anion-exchange chromatography method described in the preceding trout study (Chapter 1.2, West et al. 1993).  Calculations. Radioactivity in dried plasma samples was used to estimate ratios of 6H3 glucose/glucose and 14 C-2-DG/glucose (in dpmmoP ). The change in each ratio for an 1 individual fish was plotted against time (mm post-injection) from 1 to 60 mm and fit to a double exponential equation.  To estimate glucose turnover the appropriate curve was  extrapolated to time zero and infinity and the area under the curve was calculated (Katz 1992). The ratio of the injected dose of 6H-glucose (dpmkg 3 1 body mass) to area under the curve (dpmminmol ) was formulated to estimate glucose turnover rate (JLmolmin 1 kg ) 1 . The 14 C-2-DG washout curve was integrated from time 0 to 60 mm and an index of muscle glucose utilization (GUI) was calculated as tissue 14 C-2-DGP (dpmg 1 tissue) divided by the area under the plasma 14 C-2-DG washout curve (dpmmin’nmol ). 1 Calculation of glucose utilization rate (GUR) in specific tissues requires that GUI is corrected with a lumped constant (LC) to account for slower 2-DG phosphorylation compared to glucose (Sokoloff 1983).  For discussion of the energetic importance of  48  glucose in carp muscle, GUR’s were in carp tissues were estimated by dividing GUI by LC’s determined previously for rainbow trout (West et a!. 1993; West et a!. accepted J. Fxp. Biol).  Statistics.  The signicance (p <0.05) of differences between group means was assessed  with ANOVA or Mann-Whitney U-test for non-normal data. Regression analysis was used to examine relationships between glucose kinetics and plasma glucose concentration.  RESULTS Carp swimming performance.  Considerable variability existed in oxygen uptake  determinations between individual carp, but all showed similar rates of change in log oxygen consumption with increasing exercise intensity (Figure 3).  Maximal oxygen  consumption of the 4 fish that had reached a swimming speed of 1.5 body lengthssec’ averaged 175 ± 36 mg 1 kg Extrapolation of the general regression to a swimming h 2 0 .  speed of 0 body lengthssec 1 indicates that the oxygen consumption of resting carp at 15 °C is expected to average about 43 mg 0 kg (95 % confidence limits indicate a 1 h 2 range of between 25 to 65 mg 02 1 kg h ) . for the 6 exercised carp ranged from 1.00 to 1.68 body lengthssec , 1 averaging 1.38 ± 0.11 body lengthssec 1 (61 cmsec 1 for the average body length of 44 cm). Two fish had U’s of about 1 body 1 1engthssec and these were eliminated from further swimming tests.  The 4 remaining carp, with one new fish added to the group,  were cannulated and re-exercised 24  -  36 h later at an average sustained speed of 1.26 ±  0.1 body lengthssec 1 (54 cmsec’). This speed corresponded to about 80 % of the Ucrit determined for the 4 re-exercised fish (1.56 ± 0.06 body lengthssec ). 1 1 or 69 cmsec As indicated (Figure 3), the measured rate of oxygen consumption for the cannulated fish (130 ± 10 mg ) kg was in agreement with the rate expected for uncannulated carp. 1 h 2 0 Given that cannulated carp were able to sustain swimming at the test speed for 120 mm and  49  ‘‘  2.5  ——  y  =  1.64 + 0.35(x), r=0.68 —  0  E  D  1.0  C  C C •-  0.0 0.0  0.5  1.0  1.5  2.0  Swimming speed (body lengths.sec’) FIGURE 3. Oxygen consumption of 6 carp during a stepwise increase in swimming speed. Fatigue speed (Ucrit) was deter mined and 4 of these same fish, plus one additional fish, were  cannulated and re-exercised at 80 % U. Arrows denote ave rage speed and oxygen uptake of the swimming, cannulated carp.  50  that average oxygen consumption could be predicted from the relationship between oxygen uptake and swimming speed determined for uncannulated carp, it would seem that the implanted cannulae had minimal effect on swimming performance. Since burst and glide activity was observed in carp at one velocity increment (0.30 body lengthssec ) above the 1 fmal test speed 1.26 ± 0.10, it would seem likely that Ujt was similar before and after placement of the dorsal aortic cannulae.  Glucose turnover. ranged 5  -  Plasma glucose ranged 3  10 mM among swimming fish.  -  17 mM among individual carp at rest and  Nevertheless, plasma glucose remained in  steady state in individual fish from both groups (see Figure 4)  -  the coefficient of variation  of plasma glucose was < 15 % for individual fish throughout the blood sampling period. Although glucose turnover rate was elevated in swimming carp, the effect of exercise on average turnover rate was not significant (Table 5). However, turnover was clearly dependent on plasma glucose concentration and the slopes of these retation ships were different (p  =  0.05) for resting and exercising fish (Figure 5). Intercepts of the two  lines were not significantly different.  Interestingly, despite the difference in regression  slopes, turnover in both swimming and resting carp changed proportionally with concentration. This meant that at any given concentration, at least within the range of 5 10 mM, exercise caused a roughly constant relative stimulation of turnover  -  -  about 2-fold.  The concentration-independent effect of exercise was more apparent from the plasma clearance of glucose which ranged from 0.3 to 1.3 mlmin kg in carp (Figure 5) and 1 was 2-fold higher in swimming fish than in those at rest (p  =  0.032, Mann-Whitney U-  test).  Muscle glycogen and glucose utilization. Average glycogen content in red muscle of exercised carp (22.1 ± 10.0 mol glucosyl unitsg’) was lower than that of resting fish (41.1 ± 8.1 mol glucosyl 1 unitsg ) , but the difference was not significant as a result of  51  ‘—5 a) C’,  0  C) -  C35 C’,  -  I  I  I  I  _I  -  03  6  0 ‘I  E  I  I  0  102030405060 Time (minutes)  I  I  I  I  I  I  I  0  10  20  30  40  50  60  Time (minutes) FIGURE 4.  Plasma glucose concentration and washout of  -glucose in resting (hollow symbols) and 6H -2-DG and 3 ‘ C 4 exercising (filled symbols) carp following bolus-injection of tracers.  1.7 ± 0.6  1.0 ± 0.1#  6.9 ± 1.3  Swimming  =  7•9*  0.03).  18.4 ±  3.3 ± 1.5  24.5 ± 4.8*  8.8 ± 2.9  Red Heart Muscle 1g (nmo1min ) 1  signinicantly different from restin control (Mann-Whitney rank sum, p significantly different from resting control (ANOVA, p < 0.05).  1.1 ± 0.4  White Muscle  0.5 ± 0.1  Glucose Clearance (mlmin’ kg ) 1  3.8 ± 1.0  Glucose Turnover (mo1 min 1 kg ) 1  GUI  Whole-body glucose turnover and muscle glucose utilization index (GUI) in resting and swimming carp.  Resting  TABLE 5.  t’.)  53  15  • swimming y o resting y  = =  0.6 + 0.4(x) (r =0.92, p 0.03) -2.3 + 1.3(x) (r =0.89, p =0.04)  •  O1O  •  .—  o>I  I  •  o :‘ .—  • •.  0  EO.5  0.0  •  0 0  0  0  0 I  I  I  I  5  10  15  20  Plasma glucose (mM) FIGURE 5. Glucose turnover (upper panel) and plasma clearance of glucose (lower panel) in carp. Turnover was dependent on glucose concentration in both resting and swim ming fish. Plasma clearance was concentration-independent.  54  the large degree of variability in the data.  Similarly, the difference between glycogen  ) and after 1 levels in white muscle of carp at rest (21.3 ± 4.3 mol glucosyl unitsg  ) was not significant. Effects of exercise on 1 exercise (12.4 ± 6.8 mol glucosyl unitwg 1 the terminal level of lactate in red (3.2 ± 1.3 molg’ at rest and 8.0 ± 1.6 molg after exercise) and white (5.1 ± 1.2 molg 1 at rest and 13.2 ± 4.9 molg 1 after exercise) muscle of carp were not significant. Sustained exercise resulted in an approximately 6-fold increase in GUI in red muscle, while GUI in heart increased by 3-fold (Table 5). GUI in white muscle increased by 54 % during exercise, but the average rate was not significantly different from that of resting controls.  Glucose utilization would appear to be associated with the terminal glycogen content of muscle. Figure 6 illustrates that at very low glycogen levels, red muscle GUI was elevated by more than 4-fold above the average rate for fish in which red muscle glycogen levels were in excess of 10 mol glucosyl unitsg . Low glycogen content was 1 observed in the red and white muscle of only one exercised carp in the present study. Glycogen level in the heart of the same fish was not determined, but an elevated rate of cardiac glucose uptake (see Figure 7) suggests that glycogen level was possibly reduced in all muscles of this one animal. Muscle GUI in individual fish was examined (Figure 7) for evidence that plasma glucose level may have influenced uptake rate in the different muscle-types. GUI in white muscle varied over a relatively narrow range (0.1  -  ) in both swimming g 1 3.5 nmolmin  and resting fish. Although trends indicate positive slopes in plots of white muscle glucose utilization, regression slopes were not significant.  In red muscle, GUI appeared  independent of plasma glucose level in both resting and swimming carp. The one high estimate of glucose uptake (50 nmolmin ) occured in a fish which displayed low g 1 terminal red muscle glycogen (< 1 mol glucosyl unitsg ). 1  Excluding this one  determination, GUI in the 4 other exercised carp averaged 10.7 ± 2.6 nmolmin , g 1  55  ,60 0  50  •  .  resting swimming  E 040  E  30 0 N •—  Cl)  •  ow  0  C.)  0  .  o  i  0  10  20  01  30  10  40  50  60  70  Glycogen content (mo1 glucosyl units g) .  FIGURE 6. The relationship between glucose utilization rate and terminal glycogen content in red muscle of carp that had been at rest or swimming at 80 % Ucrjt•  56  displaying no tendency to change with increasing plasma glucose concentration. In heart, significant regressions were calculated for data from resting fish (utilization 1.2[glucose]; r =  0.91, p  different (p  =  =  =  0.83, p  =  0.05) carp.  0.04) and swimming (utilization  =  =  -1.3 +  -14.5 + 4. 7[glucose]; r  Slopes of these concentration-dependent relationships were  0.05) and showed trends similar to the pattern observed for glucose turnover  rate.  DISCUSSION The present study examined the role of circulatory glucose in the swimming energetics of carp. Glucose turnover increased with exercise, but the absolute difference in flux between resting and swimming fish was dependent on plasma glucose concentration. Despite this concentration effect on turnover, glucose availability was apparently of less importance to the active locomotory muscle than to other tissues, like myocardium, since GUI in specifically the active lateral red muscle was concentration-independent. Nevertheless, GUI in red muscle increased 3-fold above resting rates and we argue in the following discussion that glucose disposal could potentially have accounted for 25  -  30 %  of the fuel demands of this muscle mass during sustained swimming. It is evident that carp were more reliant on circulatory glucose than were better aerobic performers like rainbow trout, with the latter using glucose for an estimated < 10 % of total fuel utilization during exercise at 80 % U.  Oxygen consumption. previous measurements.  Carp oxygen uptake in the present study compares well with The extrapolation of active oxygen uptake to zero activity  concurred with other such procedures and with determinations of oxygen consumption in inactive carp at 15 °C (about 50 mg O kg in 0.8 2.0 kg animals: Christiansen et a!. 1 h 2 -  1982; Hughes et a!. 1983; Ultsch et a!. 1980). Maximum active oxygen consumption in the present study was about 4-fold higher than the estimated resting rate, resulting in a  57  4 white muscle  3  0  2  -  • 0  11  —  o  E  0  I• 0  0  I  I  I  50red muscle  -  30 .—  0  0  .  ‘,  100 40-  0 (1•pp  I  fl  I  I  .  — .— .._.e  -n  c,  20  heart  -  0  o C.)  .  100  0  0  Q  0  0 I  5  0  I  I  I  10  15  20  Plasma glucose (mM)  FIGURE 7. Muscle glucose utilization index (GUI) in resting (hollow symbols) and swimming (filled symbols) carp in relation to plasma glucose concentration.  58  metabolic scope of 132 mg 1 h’kg This expansion of oxygen consumption from rest 2 0 . to near maximal exercise is less than one-half of the metabolic scope in salmonids of similar mass (Kiceniuk and Jones 1977). Although resting rates of oxygen consumption are comparable in -1 kg carp and trout, maximal oxygen consumption is higher in salmonids as a result of their superior swimming capacity (Ut of 2 body lengthssec ; 1 Kiceniuk and Jones 1977). Oxygen consumption in carp swimming at 80 % of U (130 mg ) kg was about 3-fold higher than the predicted resting rate. This metabolic 1 h 2 0 scope was similar to the aerobic scope observed previously in spontaneously active carp (Ultsch et at. 1980).  Glucose turnover.  Glucose turnover in resting carp was somewhat higher than had been  measured previously for other teleosts (Garin et a!. 1978).  This resulted largely from  higher glucose levels in carp in the present study. The regression describing proportional increases in resting glucose turnover with glucose concentration allows us to predict that at 2  -  4 mM glucose, whole-body glucose utilization would be within the range estimated in  most species of teleosts that have been studied (1.4  -  2.2 molmin kg (Garin et at. 1  1978)). The effect of exercise on glucose kinetics in carp was revealed upon inspection of the concentration-dependent glucose fluxes during rest and exercise (Figure 5). Over the range of 5  -  10 mM plasma glucose, exercise was clearly accompanied by a constant 2-fold  stimulation of turnover.  The absolute difference in turnover between resting and  swimming fish was dependent on glucose concentration, as a result of the difference in slopes of the data regressions. However, proportionality in these relationships meant that glucose clearance (turnover divided by glucose concentration) was independent of plasma glucose and the doubling of clearance rate (Figure 5) confirmed the concentration independent effect of exercise on glucose kinetics, at least over the range of glucose concentrations measured.  59  The regression for swimming carp would be predicted to overlap that for the resting fish at 2  -  4 mM plasma glucose, suggesting that an exercise effect would not be detected  at low plasma glucose concentrations. The implication (discussed further in chapter 4) is that glucose is used increasingly for swimming energetics depending on its plasma  concentration. However, this would appear to not be the case since GUI in red muscle, the main locomotory muscle at 80 %  was concentration-independent. The possibility  that other activity dependent costs (ventilation, cardiac function, pink muscle recruitment, etc.) use glucose to a greater extent is given some support by the fact that changes in cardiac GUI parallel concentration-dependent turnover rates in the whole-animal (Figure 7).  Muscle glucose uptake.  The effect of exercise on red muscle glucose utilization would  appear to be pertinent to the oxidative demands of this fiber mass.  If, as a starting  assumption, the metabolic scope for sustained swimming (87 mg 1 kg in carp is h 2 O ) attributed solely to the increased aerobic functioning of the lateral red muscle, then oxidative demand for glucose would have been equivalent to about 7400 nmol’min kg. 1 In turn, assuming about 70 g of red muscle per kg body mass (Bone 1978), the oxidative glucose demand in active red muscle per gram of tissue mass is estimated at just over 100 . This compares to an estimated red muscle GUR of about 28 nmolmin’g g 1 nmolmin 1  (excluding the one value for the glycogen-depleted fish and using a red muscle lumped  constant of 0.40; West et a!. 1993). Therefore, a significant portion (25  -  30 %) of the  oxidative energy demand in swimming carp could potentially have been derived from circulatory glucose.  This estimate would increase in the likely event that other aerobic  metabolic costs also contributed to the change in whole-body oxygen consumption. For instance, electromyograms from previous work on swimming carp (40 cm length and acclimated to 15 °C) indicate that intermediate pink muscle fibers are recruited at 1.3 body lengthssec (Johnston et cii. 1977), a speed comparable to 80 % Uj in the present study.  60  Given that sustained swimming was established for individual carp by visual inspection of swimming behaviour (i.e., water speed was slowed until burst-and-glide swimming ceased), we cannot rule out the possibility of aerobic involvement of intermediate fibers in the present study. White muscle recruitment, however, was probably minimal since such recruitment in carp is generally accompanied by rapid fatigue (Rome et al. 1984). Intramuscular  glycogen  could  have  augmented,  or  even  dominated,  the  carbohydrate demands of red muscle in swimming carp, as suggested in a previous study (Johnston and Goldspink 1973). Post-exercise glycogen and lactate levels in the present study were inconclusive because of the variability in resting glycogen levels and because of the potential intermittent use of muscle glycogen during step changes in swimming speed. Nevertheless, it is noteworthy that GUI increased markedly when red muscle glycogen had been depleted to below 10 mol glucosyl unitsg 1 in swimming fish (Figure 6). At the moment, it is not known whether glucose transporters in fish muscle are sensitive to either the effects of hormones or increased contractions, or both as in mammals (Ploug et al. 1992), but the apparent capacity to expand uptake does suggest that transport was not saturated in the carp in this study.  An interpretation that favors carbohydrate as the  preferred substrate in active carp red muscle is that this situation reflected reversal of the inhibitory effects of glycogenolysis on hexokinase activity (Williamson 1965) as glycogen depletion progressed.  In humans with McArdle’s disease, the absence of muscle  phosphorylase and thus of glycogenolysis, is in a sense analogous since glucose utilization is enhanced in patients exercising at intensities that normally induce muscle glycogen mobilization (Vissing et al. 1992). As has been argued for swimming trout, however, a similar inhibition of red muscle glucose utilization could also occur if lipid-based fuels were preferred for oxidation (West et al. 1993).  In any case, it would seem that red  muscle glucose utilization was dependent less on the availability of glucose than on the availability of other competing substrates. Given that the endurance of carp at high relative swim speeds (at least 25 h can be sustained; Van Dijk et al. 1993) would over-burden  61  typical red muscle glycogen stores, the possibility of shifts in red muscle fuel sources  (intramuscular to extramuscular) and types (carbohydrate versus lipid) throughout an endurance swim should be examined more closely. Unlike skeletal muscle, GUI in cardiac muscle showed defmite concentrationdependent uptake and an effect of exercise that paralled the responses seen in whole-body turnover.  Complete oxidation of glucose at the utilization rates estimated in this study  would have accounted for 10  -  40 % of total myocardial fuel consumption in resting and  swimming carp (assumes resting power output of about 1 mWg 1 for slow swimming species (Farrell 1984), exercising power output 4 x resting (A.P. Farrell, personal communication), 20 % cardiac efficiency (Graham and Farrell 1990), caloric equivalent of 20.1 Joulemr’ and a myocardial lumped constant of 0.4 (West et a!. 1993)). The range (10  -  40 %) in this calculation reflects the concentration-dependence of GUI in heart,  indicating that glucose availability could have strongly influenced the pattern of myocardial substrate use for energy provision. Fuels other than glucose would appear to dominate cardiac energetics at low glucose levels. Possible shifts toward greater reliance on glucose oxidation at higher plasma glucose concentrations would be consistent with previous arguements (Sephton et a!. 1990, Sidell et a!. 1987) that the teleost myocardium seems poised to utilize carbohydrate- and lipid-based substrates equally well.  PUBLICATION STATUS:  The preceding study has been submitted for publication.  West, T.G., Brauner, C.J. and P.W. Hochachka.  Muscle glucose utilization during  sustained swimming in the carp (Cyprinus carpio). Am. J. Physiol. (submitted).  I am  grateful to C.J. Brauner for helping with the swimming trials and for measuring oxygen consumption in active carp.  62  CHAPTER 3.  CARBOHYDRATE RECOVERY IN SKELETAL MUSCLE OF  RAINBOW TROUT (Oncorhynchus myldss) AND SKIPJACK TUNA (Katsuwonus pelamis) AFTER BURST EXERCISE.  Evaluating the role of hyperglycemia in post-exercise resynthesis of glycogen in trout skeletal muscle.  INTRODUCTION  Burst swimming in teleosts causes dramatic changes in metabolite levels which can leave the white muscle depleted of fuel reserves at the end of an exercise bout (Milligan and Wood 1986b; Mommsen and Hochachka 1988; Schulte et a!. 1992).  The resulting  accumulation of lactate occurs in stiochiometries that quantitatively reflect depletion of intracellular glycogen.  Subsequent post-exercise clearance/recovery of white muscle  metabolites is generally a slow process with little metabolite exchange occuring between muscle and plasma (Tang and Boutilier 1991).  With respect to glycogen recovery,  evidence suggests that the quantitative significance of glycogenesis from glucose is minimal because of the predominance of an, as yet undefined, intramuscular glyconeogenic pathway  from lactate (Arthur et a!. 1992; Moyes et a!. 1992; Schulte et a!. 1992; Pagnotta and Milligan 1991; Tang and Boutilier 1991). However, the importance of white muscle as a site of circulatory glucose disposal cannot be overlooked, if only because of the large relative mass of the tissue. In contrast to the situation in white muscle, the stoichiometry of lactate accumulation in red muscle does not appear to be matched with glycogen depletion at the end of exercise (Parkhouse et a!. 1987), which is perhaps indicative of greater transfer of metabolites between plasma and the well perfused red muscle mass.  Post-exercise  glycogen repletion in red muscle of other species is very much dependent on the uptake of substrate from the circulation, primarily glucose in mammals (Bonen et a!. 1990; Johnson  63  and Bagby 1988; Pagliassotti and Donovan 1990) and lactate in some terrestrial ectotherms (Gleeson and Dalessio 1990). While it is clear that glycogen stores of salmonid red muscle are greatly depleted by high-intensity exercise (Parkhouse et a!. 1987), the relative utility of lactate vs. glucose as a precursor for glycogen synthesis and the rate of fuel repletion is not known for this tissue. Post-exercise hyperglycemia is sometimes seen in recovering salmonids (Mulligan and Wood 1986b, Mommsen et a!. 1988; Pagnotta and Milligan 1991) and was evident in a preliminary examination of blood metabolites in the present study (Figure 8).  Whether or not post-exercise glucose availability can influence glucose  utilization in different muscle types can be addressed with 2-DG methodolgy. 2-DG uptake provides in vivo rates of tissue-specific glucose phosphorylation and, in turn, this measure of glucose utilization can serve as a maximal estimate of in vivo skeletal muscle glycogenesis, particularly during a period of net glycogen synthesis (Kusunoki et a!. 1993). In addition to determinations of muscle glucose uptake, glucose turnover was estimated simultaneously in recovering rainbow trout (Oncorhynchus mykiss) using bolus-injections of 6H-glucose. The results are discussed in terms of the pertinence 3 of possible changes in post-exercise glucose availability to glycogenesis in different skeletal muscle types.  MATERIALS AND METHODS Animals.  Rainbow trout (Oncorhynchus mykiss) of both sexes and weighing 300  -  600  g were obtained from a local supplier and maintained in the aquarium facility at the Department of Zoology, U.B.C. Fish were held in flow-through freshwater (10  -  15 °C)  and fed to satiation three times weekly. Fish were maintained in these conditions for at least 1 month prior to experimentation.  Surgery.  Procedures followed for cannulating rainbow trout were precisely as  outlined in chapter 1.2. A single 60 cm cannula (PE-50, Clay-Adams) was placed in the  64  15 o  lactate  • glucose -I  0  E  0  0 P r0 exercise  2  4  6  8  10  12  Time (mm post-exercise)  FIGURE 8. Preliminary observations of plasma glucose and lactate in trout during recovery from enforced burst swimming.  65  dorsal aorta of each fish.  The cannula was filled with Cortland saline (Wolf 1963)  1 heparin (Glaxo Canada, Ont.) and was flushed twice daily. Each containing 5 i.u. mL experimental fish was allowed to recover for 48 h in a black perspex box supplied with a continuous flow of aerated freshwater (10  Exercise and recovery protocol.  -  15 °C).  Fish were netted from the recovery boxes and  transferred to a Brett-type swim tunnel. A protocol similar to the one used by Schulte et a!. (1992) was followed to ensure recruitment of white muscle and reduction of glycogen stores.  Briefly, water speed was increased slowly over a period of 2  -  3 mm  to a  maximum level, defined as the point where the fish first displayed burst and glide swimming behavior. This speed was maintained until the fish started to rest against the rear screen of the swim space. Water velocity was immediately reduced to a point where the fish could be induced to swim again and then once more increased gradually to the maximum. The cycle was repeated for about 15 mm until burst swimming could no longer be induced. At this point the fish was quickly removed to a holding box for recovery and eventual administration of radiolabels. Studies of white muscle metabolite status in fish following maximal exercise show that complete recovery is a long process, requiring 12  -  24 h for some metabolites.  However, most studies indicate that up to 50 % of glycogen replenishment can be expected between 2  -  8 h post-exercise (Milligan and Wood 1986b; Pagnotta and Milligan 1991;  Pearson et a!.  1990; Scarabello et a!. 199 la,b; Schulte et a!. 1992).  Isotope  administrations were designed to bracket a portion of this interval in recovery so that glucose utilization could be related to changes in muscle metabolite levels.  Glucose  turnover and uptake in muscle were determined in a total of 9 exercised fish with bolus  injections of isotope given at 2.5, 4 or 5 h (3 fish per group) post-exercise. Glucose fluxes and tissue glucose utilization rates were also determined in 6 unexercised animals.  66  Isotope injection and tissue sampling.  Bolus injections of 6H-glucose (15 ILCi) and 3  C-2-deoxyglucose (2-DG, 5 MCi), purchased from Amersham Canada, were used to 14  determine in vivo circulatory glucose turnover and muscle glucose utilization, respectively. For each preparation, the appropriate volume of each isotope was dried under a continuous stream of N 2 gas and these were reconstituted together in a single bolus of Cortland saline  (150 ILL).  An aliquot (10 ILL) was removed to determine the dose of each radiolabel  injected while the remainder was taken up in a glass Hamilton syringe to verify the bolus volume. The syringe was connected to a 3-way valve positioned on the free end of the dorsal arotic cannula and the bolus was administered to the fish. Fresh saline (250 ILL) was drawn into the glass syringe and then delivered to the fish in order to flush the  cannula. Blood samples (125 ILL) were collected through the same cannula at 1, 2, 3, 4, 5, 10, 20, 30 and 60 mm post-injection. After sampling, the cannula was flushed with saline (125 ILL) to replace the blood volume. Red blood cells were separated immediately from each sample by centrifugation and the plasma retrieved was deproteinized with an equal 1 volume of 0.6 N perchloric acid (PCA). The samples were neutralized with 3 molL C0 in 0.5 molL 2 K 3 1 triethanolamine and kept frozen (-70 °C) until analysed for metabolites and radioactivity. At the end of the sampling period, a mixture of sodium pentobarbital (1 ml of 65 ; MTC Pharmaceuticals, Ont) and d-tubocurarine chloride (0.1 mgmU 1 mgmL 1 dissloved in 1 mL of saline; Sigma Chemicals, Mo) was delivered through the cannula to immobilize the fish. When opercular movements ceased (30  -  60 see) the spinal cord was severed  immediately posterior to the head. Red and white muscle was taken with a single pass of a double-bladed cleaver through the musculature of the tail just posterior to the dorsal fm. The steak was clamped between Wollenberger tongs, pre-chilled in liquid N , and 2 immersed in liquid N . Only 10 2  -  15 sec passed from the time the fish was killed to the  moment that the tissues were frozen. The samples were kept frozen at -70 °C.  67  Metabolite assays.  Muscle was prepared for homogenization by dissecting away the skin  and separating red and white fiber masses with the steak in a liquid N 2 bath. Pieces of tissue (0.5  -  1 g) were placed in pre-weighed tubes and quickly reweighed. Ice cold 0.6 N  PCA (4 vol) was added while the tissues were still frozen. The tissues were immediately homogenized with three 20 sec passes of an Ultra-turrax homogenizer. Aliquots (2x100 L) of homogenate were removed for glycogen determination as described previously (Chapter 2). The remainder of the homogenate was centrifuged (12000g for 10 mm at 4 °C) and the acidic supernatant was neutralized and stored at -70 °C. Assays for tissue lactate and glucosyl units and plasma glucose were adapted for use with microtitration plates (0.3 mL) and a Titertek Multiskan plate reader. The  tissue  content  of  C-2-deoxyglucose-6-phosphate 14  C-2-DGP) 1 ( 4  was  determined in neutralized tissue extracts either by separation of 14 C-2-DGP from 14 C-2DG on columns of DEAE Sephadex A-125 (Sigma Chemicals, Mo) or by precipitation with 0.3 N Ba(OH) 2 and 0.3 N Zn(S0 ), as described previously (Chapter 2). 4  The 2-DG lumped constant.  The lumped constant (LC) is a correction factor  needed in the calculation of tissue specific glucose utilization to account for different rates of 2-DG and glucose phosphorylation in tissues (Sokoloff 1983). In the study of trout red muscle and heart the LC was estimated to be 0.40 (West et al. 1993), a value that was unaffected by changes tissue glycolytic activity. In the present study, estimation of the LC for trout white muscle was made by determining simultaneous in vitro rates of glucose utilization and 2-DGP formation in slices of white muscle as described previously for red muscle (West et a!. 1993). Briefly, slices of white muscle were prepared and preincubated in saline as described previously. Slices were then incubated individually for 40 mm with 0.3 Ci 5H-glucose (Amersham Canada) and 0.15 Ci 14 3 C-2-DG in fresh saline (2.5 mL). At the end of the incubation the slice was removed, weighed and analyzed for 14 C2-DGP content.  0 was separated from 52 H 3 H-glucose by distillation of the PCA 3  68  deproteinized/neutralized incubation medium as described previously (West et a!. 1993). The LC was formulated as the rate of 14 C-2-DGP formation divided by the rate of glucose utilized (calculated from 3 0 production). 2 H  Glucose turnover and muscle uptake.  Aliquots (20 ,LL) of deproteinized/neutralized  plasma were dried under a continuous stream of nitrogen gas for 5 h to remove 3 0. 2 H c.2.DG was determined and 3 14 H-glucose was estimated after reconstituting the dried plasma with 1 mL H 0 and adding 10 mL aqueous scintillant (ACS II; Amersham 2 Canada). Radioactivity was assayed using a dual counting protocol prepared on an LKB Rackbeta scintillation counter.  Analysis of tracer washout for estimation of glucose  turnover and muscle glucose utilization was conducted as described previously (Chapter 2).  Statistics.  Data is presented as means ± se. The significance of differences between  groups was assessed using ANOVA and Tukey’s HSD.  RESULTS Glucose concentration and turnover rate, Average plasma glucose concentrations and washout curves for the injected radiolabels are depicted in Figure 9 (resting fish) and Figure 10 (recovering fish).  Plasma glucose concentration varied considerably among  experimental animals (ranging 6  -  38 mmolL’), but individual fish remained in steady  state for the period following bolus-injection of radiolabels. No differences in plasma glucose concentration were evident between experimental groups (Table 6). High variability and small sample sizes during recovery likely accounted for the lack of differences between groups, even though plasma glucose at 4  -  5 h of  recovery was comparatively low. Average glucose turnover rate was also depressed during the 4  -  5 h interval, with no statistical differences between groups (Table 6). Turnover rate  was linearly associated with steady state glucose concentration for individual fish at rest  69  s  1500  -  E ‘e  1200  200 -  0  0 0  150  0  900-  F  100  U F  600  •0  0  0  10  20  30  40  50  60  TIME (mm)  0  10  20  30  40  50  60  TIME (mm) FIGURE 9. Plasma glucose concentration (upper panel) and H-glucose 3 C-2-DG and 64 washout curves (lower panel) for ‘ in resting trout.  70  -  -  E I)  25 20  15 10  0 C)  —  -‘  I  I  I  I  I  I  I  1800  C,,  1500  300  0  bf.  0  C)  1200  —  0  c)  Cd)  0 0  E  250 200  150 100  50  o  600  0 0  :1  10  20  30  40  50  60  TIME (mm)  300  E 0’  I  I  0  10  I  I  I  20 30 40 TIME (mm)  50  63  FIGURE 10. Plasma glucose concentration (upper panel) and washout curves (lower panel) for ‘ C-2-DG and 64 H-glucose 3 in trout during recovery from burst exercise.  71  and throughout the 2.5  -  6 h of recovery (Figure 11). Separate regressions for resting and  recovering fish were not different, suggesting that changes in plasma glucose status had a direct influence on glucose kinetics while effects of exercise-state of the fish were less important. The plasma clearance of glucose, calculated as turnover rate divided by glucose concentration, was shown to be independent of plasma glucose level (Figure 11). This means that the volume of plasma cleared of glucose was relatively constant at 0.5  -  1.5  kg regardless of plasma glucose concentration. mLmin , 1  Muscle glucose utilization. Pre- and post-exercise rates of glucose utilization in white muscle were calculated using the lumped constant determined from muscle slice incubations (0.51 ± 0.06, n= 12). There were no significant differences (ANOVA p >  0.05, glucose concentration was a significant covariate p  =  0.02) in utilization rate  between control and recovering fish (Table 6). Like turnover rate, glucose utilization in white muscle between 4  -  5 h of recovery tended to be 1/3 to 1/2 of the rates estimated at  other recovery times and at rest. Glucose uptake in white muscle varied within relatively  narrow limits (Figure 12) and plasma glucose concentration had a less noticable influence on utilization rate compared to its effect on turnover rate. As indicated, the significance of a regression through these data was strongly influenced by a single observation which, emphasizing that utilization was only marginally dependent on glucose availability (Figure 12). As is suggested in Figure 13, glucose uptake in red muscle of recovering trout changed with plasma glucose concentration only between 6 1 utilization seemed independent of concentration. mmoiU  -  12 mmol1J . 1  Above 12  Since all of the glucose  utilization values for resting fish had relatively high plasma glucose levels (> 12 mmolU 1), the average resting utilization rate from 9 fish with lower plasma glucose levels (<  , from West et al. 1993) is included in Figure 13 for comparison. 1 mmolL  5  The data  suggest that the trends in glucose uptake were similar for both resting and recovering trout.  Whole-body glucose turnover and glucose utilization in skeletal muscle during recovery from enforced maximal  18.2 ± 5.9  20.7 ± 6.8  8.1 ± 1.2  7.1 ± 0.6a  20.2 ± 8.9  19.7 ± 8.8  19.3 ± 2.6  19.1 ± 2.3  PLASMA GLUCOSE ) 1 (mmoFL  GLUCOSE TURNOVER ) 1 (molkg’ min  2.0 ± 0.3 0.8 ± 0.6 2.1 ± 0.9  1.5 ± 0.3  WHITE MUSCLE  b. p = 0.008 for comparison to pre-exercise red muscle glucose utilization.  a. p = 0.07 for comparison to pre-exercise glucose turnover.  10.9 ± 3.6 5.2 ± 1.1”  12.2 ± 3.2  18.5 ± 1.7  RED MUSCLE  GLUCOSE UTILIZATION ) 1 (nmolg’ min  5-6h  4-5h  2.5-3.5h  RECOVERY INTERVAL PRE-EXERCISE  was used.  exercise. Utilization in red muscle was calculated using a lumped constant of 0.4 (West et at. 1993), while for white muscle 0.51  TABLE 6.  73  40  I  Lv  o  PRE-EXERCISE 3.5h, • 5h, • 6h  30  0  0  E2o  0  0  81 0 2.0 1.5 .  0 V  1.0  0 0 0  .  0.5  I  0.0 0  I  I  10 20 i 30 PLASMA GLUCOSE (mmol.L)  40  FIGURE 11. Glucose turnover in resting and recovering trout as a function of plasma glucose concentration (upper panel), turnover  =  0.97[glucose] +0.57, r=0.93 (solid line). Separate  lines for resting (long dash) and exercised (short dash) fish were not different. Plasma clearance (lower panel) was concentrationindependent.  74  ‘5’3O  5O3’  ..  4O  2O 2O10I  Io PRE  EXERCISE  I  I  4.0  5.0  I•  6.0  RECOVERY (hrs.)  PRE-EXERCISE 3.5h, • 5h, • 6h 0  ,  4  v  0  8i  + p  0  . I  I  10 20 30 PLASMA GLUCOSE 1 (mmol.L )  I  40  FIGURE 12. White muscle glycogen and lactate in resting and recovering trout (upper panel). *different from pre-exercise (p < 0.05); **different from pre-exercise and 6 h (p <0.001). Glucose level had only a minor affect on utilization rate (lower panel), with utilization = 0.07 [glucose] + 0.31, r = 0.68). *Denotes an observa tion with large influence on any regression (slope is not significant without this datapoint, r=0.48). Data from 9 resting trout with low glucose levels (from West et al. 1993) is also shown (cross).  75  15  15 •  C.)  10  L  l0  *  oI  .c-) 0  ‘—0 L  0 ‘J  //  PREEXERCISE  _l  p.  4_I  4.0  6.0  5.0  RECOVERY (hrs.)  —  C.)  PRE-EXERCISE 3.5h, • 5h, • 6h  o  -  0  20  0  -  0  0  V  •  0  V  .10 CE  0  C..2  •  •  V  I.  0 —  I  0  I  I  10 20 30 PLASMA GLUCOSE (mmol.U)  I  40  FIGURE 13. Red muscle glycogen and lactate in resting and recovering trout (upper panel). *different from pre-exercise value (p <0.05); **different from pre-exercise and 6 h (p <0.01). Glucose utilization in red muscle (lower panel) appeared to have been bi-phasic with respect to plasma glucose availability, be coming concentration-independent only at higher (>10mM) glucose levels. Uptake in 9 resting trout with lower plasma glucose levels (see West et al. 1993) are also shown (cross).  76  There were differences (ANOVA p  =  0.04, glucose concentration was an insignificant  covariate) among groups and the lower utilization at the end of 5 h of recovery (Table 6) appears to have resulted from the data for this one group falling completely within the range where utilization is sensitive to plasma glucose level (Figure 13).  Glycogen replenishment in skeletal muscle.  Post-exercise recovery of white and red  muscle glycogen is illustrated in Figures 12 and 13, respectively. Red muscle glycogen content was still reduced 3.5 h into the recovery period and had recovered to 60 % of the pre-exercise value by the end of 6 h. The data indicate a rate of replenishment of about 20 nmol glucosyl unitsmin g red muscle over the interval from 3.5 to 6 h post-exercise. 1 Glucose utilization in red muscle during recovery was 5  -  12 nmolmin g (Table 6), 1  indicating that plasma glucose possibly supported a large portion, 25 replenishment in this tissue.  -  60 %, of glycogen  In comparison, although white muscle glycogen content  remained significantly reduced 6 h post-exercise, the rate of glycogen replenishment between 2 and 6 h (about 40 nmolmin ) was 20 g 1 glucose phosphorylation (1  -  -  40 times higher than the rate of  21 nmolmin g ’, Table 6).  DISCUSSION An unexpected result in this study was the range of plasma glucose concentrations among the experimental animals. This might suggest variable levels of stress (Barton and Schreck 1987), but identifying the nature of any stress is difficult. In previous studies that  used the same pre-experimental care, anesthetization, surgical and handling procedures there was no hyperglycemia evident (West et a!. 1993). In the present study, glucose was higher in resting fish than in those that were exercised, indicating that hyperglycemia was not a direct result of the exercise protocol.  In turn, trends in glucose turnover were  explained primarily by plasma glucose status rather than exercise state (rest or recovery). However, while hyperglycemia and concentration-dependent glucose turnover are  77  remarkable from a glucoregulatory perspective, these kinetics seemed dissociated from the carbohydrate recovery pattern in trout white muscle. It is important to note that turnover increased proportionally with plasma glucose concentration, suggesting that whole-body utilization of glucose may have occurred predominantly via substrate-mediated disposal. Despite this, glucose utilization rate in white muscle, the largest homogenous tissue mass in trout, was low and was only slightly sensitive to plasma glucose concentration. This situation emphasizes that glucose availability was pertinent to tissues other than white muscle and,  further, that white muscle glycogen was resynthesized essentially  independently of glucose availability.  The interpretation in red muscle is less  straightforward since the pattern of glucose utilization in relation to glucose availability was bi-phasic, with an initial concentration-dependent phase followed by concentrationindependence at higher plasma glucose levels. It would seem that glycogen reformation in this muscle type is dependent on both glucose availability and glyconeogenic flux. Postexercise blood glucose status, which is moderately hyperglycemic in some studies (discussed later), could therefore have consi&rable influence on the relative importance of glucose incorporation into red muscle glycogen.  Glucose turnover.  Average glucose turnover rates presented for rainbow trout (Table 7)  were unusually high for salmonids (see Garin et a!. 1987). Turnover was on par with rates determined for warm-bodied skipjack tuna (Weber et a!. 1986) and the American eel (Cornish and Moon 1985), both of which may rely on glucose to a large extent for energy provision. In neither of these species was glucose turnover necessarily accompanied by hyperglycemia. In trout, blood glucose status was closely connected to glucose turnover rate in both resting and recovering fish (Figure 11). Turnover rates in resting salmonids with similar plasma glucose levels are not available for comparison. However, the overall regression does suggest that at plasma glucose levels below 10 mmolL 1 turnover rate is likely to fall within 2  -  10 . kg This estimate is in agreement with previous 1 molmin  78  measurements in resting rainbow trout in which plasma glucose was < 10 mmolL 1 (Dunn and Hochachka 1987; Washburn et a!. 1992) and demonstrates the apparent utility of plasma glucose concentration as a predictor of whole-body glucose flux. Beyond its predictive value, the relationship between plasma glucose concentration and turnover rate implies that whole-body glucose disappearance in trout responds to glucose availability over a wide range.  Proportional changes in turnover with  concentration accounted for the observation that a relatively constant blood volume was cleared of glucose, independent of glucose concentration (Figure 11, lower panel). Concentration-dependent turnover further suggests that glucose concentration itself may have been an important regulator of whole-body glucose disposal.  This would be  consistent with the generalization that glucoregulatory hormones are slow to respond to elevated glucose levels in teleosts (Harmon et a!. 1991), but it is not known what regulatory characteristics are specifically involved with enhanced glucose production in vivo. With respect to glucose disposal, an important point is that, even at relatively low turnover rates (eg. from 4  -  5 h of recovery; Table 6), disposal of glucose in muscle tissue  (calculated from glucose utilization and assuming body mass is 10 % red muscle and 60 %  white muscle) is estimated to have been circulation.  <  15 % of the total glucose removed from the  This also holds for resting salmonids with lower plasma glucose  concentrations (2  -  4 mM), based on total muscle glucose disposal of about 0.5 molmin  kg body mass (West et a!. 1993) and turnover rates of 2 1  -  kg (Lin et a!. molmin 51  1978; Washburn et a!. 1992 or predicted from Figure 12). Therefore, the largest tissue mass in the body utilized only a small portion of the glucose released into the circulation. The suggestion that glucose metabolism is involved largely with mucous production in fish (Bever et a!. 1981), rather than energy provision (West et a!. 1993), would be particularly interesting  to  hyperglycemia.  investigate  in  relation  to  glucoregulation  during  stress-induced  79  White Muscle Recovery.  The present study provides direct evidence that extramuscular  pathways of glucose formation are of minor importance to white muscle glycogen  recovery.  Two observations point to the relative unimportance of changes in glucose  availability to the process of glycogen resynthesis in this muscle mass.  Firstly, post-  exercise changes in white muscle glycogen and lactate appeared unaffected by hyperglycemia and elevated glucose kinetics since recovery rates were comparable to previous findings (25  -  50 nmol glucosyl unitsmin g and 40 1  -  g 1 100 nmol lactatemin  1, Milligan and Wood 1986b; Pagnotta and Milligan 1991; Tang and Boutilier 1991;  Schulte et a!. 1992).  Secondly, the rate of glucose utilization in white muscle spanned  quite narrow limits despite the highly variable plasma glucose levels and utilization was not linked in a discernible way to recovery interval. It can be calculated that, regardless of plasma glucose availability, glucose phosphorylation in white muscle could have potentially accounted for 5  -  10 % of the glycogen formed during recovery (assuming white muscle is  60 % of body mass and that glucose utilization at 1 throughout the tissue).  -  g is uniform 1 2 nmolmin  This estimate of white muscle glycogenesis is somewhat higher  than one other estimate made previously with different methodology (see Pagnotta and Mulligan 1991).  Accounting for the extent to which competing pathways utilize G-6-P  might be expected to bring these different calculations in closer agreement. However, the likelihood that muscle glucose is directed primarily toward glycogen storage during exercise-recovery conditions (> 80 % in rats, Kusunoki et a!. 1993) suggests that the present 2-DG measurements of glucose utilization rate are reasonable estimates of in vivo glycogenesis.  Red Muscle Recovery.  Unlike glucose utilization in white muscle, utilization rate in  red muscle was bi-phasic and could account for a relatively large proportion of glycogen repleted (25  -  60 %). The difference in the rate and concentration-dependence of glucose  utilization between the muscle-types may be related to a higher amount of glucose  80  transporters in red muscle, as in mammalian skeletal muscle (Marette et a!. 1992). In any case, the significance of an increased capacity to phosphorylate glucose in red muscle is the greater potential use of glucose for glycogen synthesis compared to white muscle. Furthermore, the concentration-dependence of glucose uptake suggests that post-exercise blood glucose status could have influenced the rate of glucose incorporation into red muscle glycogen.  The present data indicate that glucose incorporation into red muscle  glycogen is potentially adjustable over a limited range of plasma glucose concentration, up to 10  -  12 mmolL . 1  While hyperglycemia in the present study was not specifically  related to exercise state, it is known from previous studies that moderate, yet sustained, hyperglycemia is sometimes seen in recovering trout (Milligan and Wood 1986a; Mommsen and Hochachka 1988; Pagnotta and Milligan 1991; Scarabello et a!. 1991; Figure 8).  Typically post-exercise plasma glucose level is within the range where  concentration-dependent utilization in red muscle was observed in the present study. We can conclude that glucose availability is likely to influence the rate of glucose incorporation into red muscle glycogen, but given that glycogen synthesis occured at a faster rate than could be accounted for by glucose utilization alone, it is probable that both of the processes of glycogenesis and glyconeogenesis contributed to carbohydrate recovery after exercise. Trout are similar to rats in the sense that both rely on glucose to a greater extent for glycogen synthesis in red muscle than in white muscle (Bonen et a!. 1990; Kusunoki et a!. 1993). In mammals, however, both red and white muscles use relatively more glucose for glycogen synthesis, > 90 % and 30  -  50 % respectively (Bonen et a!. 1990; Johnson and  Bagby 1988; Pagliassotti and Donovan 1990).  Lactate is expected to be the major  glyconeogenic precursor in vertebrate muscle and greater oxidative losses of lactate in mammals (Brooks 1986) than in fish may partly explain greater dependence on glycogenesis in rat muscle-types.  It is evident, however, that rats use glucose almost  exclusively in red muscle, even when lactate is made available for glyconeogenesis (Bonen  et a!. 1990). In trout, the process of glycogen resynthesis in red muscle resembles the  81  situation in  mammalian white muscle since glucose accounts  substantially,  yet  incompletely, for glycogen synthesis. Glyconeogenesis is probably involved in the overall  process, as it is in mammalian white muscle (Bonen et at. 1990), but the specific role for lactate in trout red muscle is unknown.  Changes in red muscle lactate content after  exercise (Figure 13) provide few insights. measurably in salmonids between 2  -  Nevertheless, plasma lactate increases  8 h post-exercise (Milligan and Wood 1986a,b;  Mommsen and Hochachka 1988; Pagnotta and Milligan 1991; Schulte et at. 1992; Figure 8), as does lactate turnover (Milligan and McDonald 1988). The relative importance of glyconeogenesis in trout red muscle might be expected to peak when plasma lactate availability and flux is maximal.  A better understanding of the relative importance of  lactate and glucose in this muscle-type might emerge from studies of lactate incorporation into glycogen in relation to glycogen recovery rate in individual fish  -  combining  techniques of tracer lactate incorporation into glycogen with 2-DG uptake and sequential biopsy analysis of muscle.  PUBLICATION STATUS:  The preceding study has been accepted for publication.  West, T.G., Schulte, P.M. and P.W. Hochachka (in press).  Implications of  hyperglycemia in the post-exercise resynthesis of glycogen in trout skeletal muscle. &p. Blot. Thanks to P.M. Schulte for assistance with the swimming trials.  J.  82  Lactate, glycogen and PCr profiles in the white muscle of skipjack tuna during recovery from burst swimming.  INTRODUCTION 1 indefinitely, yet Tuna can sustain relatively high speeds of 3-5 body lengthss  they are also capable of short term speeds of up to 20 body lengthsc’. This intriguing combination of sprint and endurance abilities is supported by a metabolism with several unusual features.  For example, skipjack tuna (Katsuwonus pelamis) are able to fuel  burst swimming through an exceptional glycolytic capacity in white muscle, which can 1 (Guppy et al. 1979; Hulbert et produce lactate concentrations of about 100 Lmolg a!. 1979).  Removal of elevated plasma lactate and metabolically-produced protons  following exhaustive exercise occurs very rapidly in tuna (Barrett and Connor 1964; Perry et a!. 1985).  In fact, lactate turnover in post-exercise tuna resembles rates  observed in mammals (Weber et a!. 1986), suggesting that the circulatory translocation of lactate accounts for some portion of the white muscle lactate clearance in recovery. In contrast, other teleosts are known to retain most of the lactate formed during exhaustive exercise within the white muscle mass and require relatively long periods for lactate clearance (Batty and Wardle 1979; Milligan and McDonald 1988). In studies of recovery from exercise in other teleosts, such as trout, the fish is exercised and terminal samples are collected by freeze clamping at selected times after exercise.  Muscular contractions during sampling can cause artefacts resulting from  changes in the concentrations of intracellular metabolites such phosphocreatine and ATP. Nevertheless, with care and rapid sampling it has been possible to follow the changes in intracellular metabolites following exercise in trout (Pearson et a!. 1990; Schulte et a!. 1992).  It is not feasible to use this approach for tuna since ‘resting’  skipjack typically swim at 1-2 lengthss 1 to keep from sinking and to ‘ram ventilate’ the gills (Guppy et a!. 1979). The capture of free swimming tuna is not compatible  83  with the requirement to reliably collect tissue samples from tuna that have not struggled. Recently Bushnell et a!. (1990) described a protocol for spinally-blocking tuna which permitted continuous measurement of cardiovascular parameters under controlled conditions. By using this procedure, and sampling muscle with a high speed  biopsy technique, it was possible to separately collect and rapidly freeze muscle samples from the same fish at different times after exercise.  The advantage of this  approach was that it provided a means to analyse detailed muscle metabolite changes from individual fish and consequently to use fewer fish overall to elucidate general patterns of recovery metabolism than would be necessary if a terminal sampling procedure were employed.  MATERIALS AND METHODS Experimental animals.  Live skipjack tuna, Katsuwonus pelamis, (1  -  2 kg) were  purchased from a local fisherman and maintained in outdoor holding tanks at the Kewalo Research Facility (Southwest Fisheries Center Honolulu Laboratory, National Marine Fisheries Service, National Oceanic and Atmospheric Administration). tanks were supplied continuously with aerated sea water at 25 ± 2  DC.  The  Fish were not  fed and were used for experiments within three days of capture.  Exercise and biopsy protocol.  In preliminary studies, terminal muscle samples  were taken to determine resting and post-exercise levels of white muscle metabolites. Eleven unexercised fish were netted from the holding tank and killed with a sharp blow on the head. A sample of epaxial white muscle was dissected from behind the dorsal fm of each fish, freeze-clamped between Wollenberger tongs and immersed in liquid nitrogen as in earlier studies (Guppy et al. 1979).  Very little struggling occurred  during this procedure, and less than 20 sec elapsed from the time of capture to the point of tissue immersion in liquid nitrogen.  Three other fish were sampled in the same  84  manner after first being isolated individually in another holding tank and chased with capture nets for 15 mm. All dissected muscles were stored frozen over dry ice prior to homogenization and assay procedures. The two groups were designated unexercised freezed-clamped and exercised freeze-clamped, respectively. A serial biopsy technique was developed to assess in vivo metabolite changes in tuna white muscle during recovery from exercise.  Each fish was first exercised as  described above, then captured by net and guided into a plastic bag containing buffered MS-222 (tricaine methane sulfonate, 1.0 gL’ with 1.0 gL 4 NaHCO ) dissolved in 3 oxygen saturated sea water.  This group is referred to as exercised biopsied.  Tuna  taken from the holding tank and anesthetized without being chased served as unexercised controls and are referred to as unexercised biopsied.  Immediately  following anesthesia (about 2 mm), each fish was placed on an operating table and the gills were irrigated with recirculated water containing a low level of anesthetic (MS 222, 0.1 gL ). At this time initial biopsy samples were obtained from the epaxial 4 muscle using a 3 mm bore, at 10  -  15 mm depth (ALKO Diagnostic Corporation). The  tissue sample was drawn from the fish by suction into a chamber cooled with liquid nitrogen and freezing occurred almost instantly.  Bleeding was minimal and was  stopped easily by inserting a cotton applicator tip into the puncture. Skin was separated from the white muscle biopsy under liquid nitrogen and the sample stored over dry ice. After the first biopsy was taken, the fish was turned ventral side up on the operating table and the ventral aorta was cannulated following procedures described previously (Jones et a!. 1986). The fish was then righted and a 20-gauge needle, 6 cm long, was inserted through the dorsal musculature (just lateral to the second spine of the dorsal fin) to the level of the spinal cord. Lidocaine hydrochloride (0.3 ml, 2 % w/v) was then administered to establish a spinal block and arrest contractions of the swimming muscles. The needle was left in place for periodic re-injections throughout the experiment. Once secured in a foam-lined brace that was supported in a perspex  85  holding box (see Bushnell et al. 1990), the fish was presented with continuously flowing sea water (35 Lmin) and allowed to recover from anesthesia. All subsequent biopsies were taken from the tuna while in this restrained, spinally-blocked position.  To do this, the water level in the holding chamber was  lowered to expose the back of the fish. The gills remained submerged during this time. Biopsies were taken from an area directly lateral to the dorsal midline, 10  -  20 cm  posterior to the spinal needle insertion site. Samples of white muscle and blood (0.25 ml) were taken at 20, 40, 60, 100, 120, 140 and 180 mm. into the recovery period. Plasma was extracted immediately from red cells by centrifugation, then deproteinized with one volume of 0.6 N perchloric acid and stored on dry ice. A terminal muscle sample was also taken after killing the fish with a blow on the head at the end of the. experiment.  Tissure preparation and analysis. Biopsies were homogenized with an Ultra-turrax tissue homogenizer in 2 ml of 8 % perchloric acid in 40 % ethanol which had been precooled over dry ice. Homogenization temperature was maintained below -20°C. A  50 d aliquot of homogenate was removed for measurement of muscle glycogen and the remainder was centrifuged for 10 mm (7500 g, 2°C) in a Jouan table-top centrifuge. Potassium perchiorate was precipitated from the supernatant with 1 N KOH and the neutralized sample was stored on dry ice. Total creatine, creatine (Cr) + phosphocreatine (PCr), content remains constant for a given tissue (Connett 1988) and was used as initial reference for all muscle metabolites measured. This is a reliable normalization procedure (Sabina et a!. 1983) which avoids possible inaccuracies associated with weighing small, deep-frozen tissue samples. Conversion to moFg , for comparison with other studies, was made after 4 determining total creatine content in tuna white muscle samples of known weight. A factor of 39 (Jhmol total creatineg 4 white muscle) was used for these conversions.  86  Muscle lactate, glucose, Cr and PCr were measured according to Bergmeyer  (1985), modified for use with microtitration plates (0.3 ml) and a Titertek Multiskan plate spectrophotometer.  Glycogen was measured (see Chapter 1.2) in aliquots (100  j.il) of white muscle homogenate and is presented as mol glucosyl unitsg tissue. Assays modified for the plate spectrophotometer were also used to measure glucose (Sigma Diagnostics assay) and lactate (Bergmeyer 1985) in the neutalized plasma samples.  Data analysis.  Linear regression was used to determine break points in the  curves describing the recovery of white muscle lactate and PCr.  Break points were  used as estimates of recovery time for these metabolites and means from exercised and non-exercised tuna were compared using the two-sample t-test.  Comparisons of all  other metabolite concentrations in the white muscle of the various groups of tuna were assessed with Tukey’s multiple comparison. Dependent t-tests were used to compare metabolite levels in biopsies taken at different times throughout the recovery period.  RESULTS Muscle metabolites after exercise. Terminal muscle samples from the exercised freeze-clamped tuna show significantly elevated lactate levels and depressed glycogen and PCr levels (Table 7) consistent with the utilization of anaerobic glycolysis in fueling intense white muscle activity. Similarly, the initial biopsy from the exercised biopsied fish had significantly elevated lactate and depleted PCr relative to unexercised  freeze-clamped tuna (Table 7).  Glycogen in the first biopsy was reduced, but not  significantly compared to unexercised freeze-clamped fish (p reflects the large variation associated with these values.  =  0.09), which likely  Freeze-clamped, unexercised fish were taken directly from the  0.05 ± 0.01  PYRUVATE (JLmol/g)  ±  ±  0.79 ± 0.30c1  ±  32.4c  56.9  88.6  ±  36.7a,b  ±  107.5  10.4a  8.7a,b  20.1 ±  EXERCISED (N=4)  9.6’  0.39 ± 0.14a  127.3  38.5  54.5  UNEXERCISED (N=5)  INflAL  ±  ±  ±  40.1  8.1  3.2  0.05 ± 0.01  145.6  11.9  75.7  UNEXERCISED (N=5)  ±  ±  ±  53.6  15.7  8.5  0.19 ± 0.19  139.3  14.9  76.5  EXERCISED (N=4)  FINAL (180 MIN)  BIOPSIES FROM SPINALLY-BLOCKED TUNA  ±  25.1”  ± 4•9i,b  ± 4•73,i  0.14 ± 0.04e  74.6  75.8  11.5  (N=3)  FREEZE CLAMPED, EXERCISED  exercised.  different from freeze-clamped controls and final biopsy, unexercised; (e) significantly different from all other groups except final biopsy,  unexercised; (c) p = 0.09 for comparison to freeze-clamped controls, and p < 0.05 for comparison to final biopsy, exercised; (d) significantly -J  (a) significantly different from freeze-clamped controls and final biopsy (unexercised & exercised); (b) significantly different from initial biopsy,  34.4  145.0  GLYCOGEN (jLmol/g) ±  3.8  7.3  LACTATE (mol/g)  ±  ±  74.6  PCr (%TOTAL Cr)  3.4  (N=11)  FREEZECLAMPED, UNEXERCISED  exercised and unexercised, spinally-blocked tuna.  holding tank, while the freeze-clamped, exercised fish were chased for 15 mm. and then killed immediately. Biopsies were taken from both  TABLE 7. Metabolites in white muscle of exercised and unexercised tuna.  88  An intermediate metabolite profile was observed in the initial biopsies of the unexercised, biopsied tuna (Table 7) with lactate and PCr levels significantly different from both unexercised and exercised freeze-clamped fish.  These changes were  probably the consequence of the fish struggling prior to complete anesthesia, perhaps combined with a direct effect of the MS-222 anesthetic on muscle metabolism (see Van den Tillart et a!. 1989). It should be noted that the relative amount of creatine in the phosphorylated state is very high in tuna white muscle. In mammals and other teleosts PCr levels of 30 50 % of total creatine are typically measured. Van Waarde et al. (1990) suggested -  that, based on measurements with NMR, in vivo Cr was 80 % phosphorylated in teleost white muscle and assert that lower levels arise because of handling stress and/or less than instantaneous freezing of tissue samples. The comparable values of 75 % in white muscle of resting and recovered tuna presented in this study (Table 7) attest to the reliability of the freezing methods used.  Lactate, glucose and glycogen in recovery.  Changes in white muscle lactate and  glycogen during 180 mm. of recovery from exercise are illustrated in Figure 14. In both groups of biopsied fish the lactate recovery curves declined initially in a linear fashion and after 80  -  100 mm., remained constant at the pre-exercise level. Lactate  concentrations in the final biopsy taken from exercised and unexercised biopsied fish were not significantly different from the resting level (Table 7).  Biopsy glycogen  concentrations were more variable than lactate in recovery and significant differences  between initial and final tissue levels were detected in only the exercised group (Table 7).  However, most of the variation in recovering fish appeared to result from  differences between individual fish. Consequently, t-tests on paired data indicated that glycogen concentrations in the white muscle of exercised biopsied fish was significantly higher  in  the  final  biopsy  than  at  the  start  of  the  recovery  period.  89  200  200  • 150  150  Oci 00  ct  100 50  50  150  0 150  125  125  -10o  100  75  75  E  -  E  1)  I  I  I  I  I  I  I  I  I  I  0  50  50  25  25  0 40  0 40  10•  10  0  E Q  I  0  I  I  I  150 Time (mm)  50  100  I  0  200  150 Time (mm) 100  200  FIGURE 14. Glycogen, lactate and PCr in biopsies of tuna white muscle,  °  unexercised,  analysis is provided in table 7.  •  exercised. Statistical  90  50  50 exercised 40  40  30  30  20  20,  0  .  10  I,  0 50  I  I  I  unexercised 40  40  E 30  30  20  20  10  10  0 0  I  I  50  100  150  —0 200  Time (mm) FIGURE 15. Lactate and glucose levels in plasma of exercised and unexercised skipjack tuna.  c  91  Glycogen concentration in the final biopsy was not different from unexercised freezeclamped fish at rest (Table 7) and, in fact, complete replenishment was evident at 80 100 mm  (glycogen concentration at 100 mm  -  was 154 ± 55.5 JLmolg , not 4  significantly different from the fmal biopsy glycogen content). Plasma lactate concentration in the exercised biopsied tuna was essentially steady, between 35  -  40 mM, during the first half of the recovery period (Figure 15)  and only began to decline after muscle lactate had reached the resting level. In the unexercised biopsied fish, plasma lactate dropped steadily between 20 and 100 mm recovery and remained constant thereafter at about 10 mM. Plasma was not obtained from unexercised freeze-clamped fish, but the apparent resting steady state achieved in recovery by the unexercised biopsied fish is comparable to previous measurements of plasma lactate in resting skipjack tuna (Perry et a!. 1986). The plasma concentration of glucose remained in a constant steady-state throughout the recovery period in biopsied fish (Figure 15).  Glucose concentration  averaged 12.9 ± 2.9 mM and 13.6 ± 3.5 mM, mean ± sd, for the control and exercised biopsied groups, respectively.  Estimates of recovery time (lactate and PCr).  Figure 16 depicts lactate and PCr  concentrations in tuna white muscle from representative exercised and unexercised biopsied fish during recovery. Every fish showed the same general recovery pattern consisting of an initial phase of rapid lactate and PCr change followed by a period in which these metabolites changed much more slowly. Both stages were linear, which permitted the estimation of recovery time by solving a set of regession equations for each of the fish biopsied. This analysis assumes a homogeneous distribution of muscle metabolites, which appears to be valid on the basis of the consistent changes in lactate/PCr observed during recovery for each fish despite random selection of  92  150  100  E  c) ci  150 50 100 Time (mm)  () ci  200  0 0  50  100  150  200  Time (mm) 50 100  40 50  30  20  0 0  10  100 150 50 Time (mm)  200  0  0  50  100  Time  (mm)  150  200  FIGURE 16. Lactate and PCr in white muscle of representative exercised (upper panel) and unexercised (lower panel) tuna, showing regression lines for calcu lating break points in metabolite recovery.  93  ) 4 Initial lactate level (mol.g 0 120  I  I  I  50  25  I  I  I  100  75  125  °  I  I  I  200  175  150  .  90  +15.8, r=O.88  60  .  .  r=O.93  30 0  0 0  I  I  I  20  40  60  80  Initial PCr level (as % of total creatine)  100  .  FIGURE 17. Estimated recovery time in relation to lactate  and PCr concentration in the initial biopsy of individual exercised and unexercised tuna. Points on the lower x-axis are mean lactate and PCr levels determined in 11 resting tuna (see Table 7).  94  biopsy sites. The variability in muscle glycogen during recovery prevented the use of the break-point determination procedure on these data. It became apparent that the experimental protocol produced a wide range of recovery times, rather than two discrete groupings, which reflected the degree of muscle lactate accumulation or PCr depletion. This essentially agrees with the earlier observation that the unexercised biopsied fish exhibited a muscle metabolite profile that was intermediate to the resting and exercised freeze-clamped tuna, probably resulting from the uncontrolled activity during anesthetization. The recovery times overlapped somewhat between exercised and unexercised biopsied fish and therefore it seemed more appropriate to consider recovery time as a continuum with respect to exercise intensity. This is emphasized in Figure 17 where recovery time is displayed as a linear function of the concentration of both lactate and PCr in the first biopsy obtained from individual fish. As a result of this overlap, the average lactate (and PCr) recovery time for the exercised biopsied fish was only marginally different from the lactate (and PCr) recovery time for the unexercised biopsied fish (Table 8). Another interesting outcome of the comparisons in Table 8 was that the time necessary for lactate recovery, for either the exercised or non-exercised biopsied fish, was not significantly different from the respective PCr recovery time. Furthermore, if individual biopsied fish were once again considered, then there was a striking synchrony between the rates of lactate disappearance and PCr repletion (Figure 18). There was also a close relationship between the concentrations of lactate and PCr in the individual sample biopsies (Figure 19). This suggests that, unlike other teleosts which have been studied, some regulatory mechanism seemingly operates to tightly couple major adjustments of these metabolites in tuna white muscle.  95  DISCUSSION Muscle lactate and glycogen recovery.  The disappearance of white muscle lactate  levels at a rate of 1.3 moFg’miif 1 is up to 20 times faster than has been measured for trout (0.055-0.12 1 min jmolg ) .  This unusually rapid clearance of lactate is  intermediate between the rate of clearance for humans (0.66 molg’min’) and rats min following maximal exercise (Hermansen and Vagge 1977; molg ) (about 2.5 1 Meyer and Terjung 1979). Weber et a!. (1986) noted that despite high rates of whole-body lactate turnover in tuna, lactate recovery apparently occurs too quickly in white muscle to result from turnover alone, suggesting that some portion of the lactate remained in the white muscle as a substrate for post-exercise, in situ metabolism. In some teleosts, where rates of lactate turnover are greatly mismatched with actual clearance rates from white muscle during recovery from exercise (Milligan and McDonald 1988), retention and utilization of lactate within the white muscle mass is likely the dominant means of lactate disposal. Such a proposal of in situ metabolism of muscle lactate is prevalent throughout the area of exercise-recovery metabolism of both mammals and ectotherms. Some evidence comes from Hermansen and Vaage (1977) who biopsied human quadriceps muscle following maximal exercise and point out that the closely matched rates of lactate and glycogen change in muscle, combined with a small arterio-venous difference for lactate, suggest direct reconversion of lactate to glycogen. The enzymic steps involved in the process are not understood well (see Bonen et a!. 1989 for a partial review and Chapter 4), but numerous studies on vertebrate muscle propose varying degrees of in situ conversion of lactate to glycogen and demonstrate dependence of the pathway on muscle fiber type, pH, arterial lactate concentration and an extramitochondrial pathway of glycogen resynthesis (Bonen et a!. 1990; Johnson and Bagby 1988; Pagliassotti and Donovan 1990).  96  TABLE 8. Estimated recovery times for PCr and lactate in tuna white muscle. Within  each test group, PCr recovery time closely matches that of lactate (see also Figure 15), while differences between exercised and unexercised fish were only marginally different.  RECOVERY TIME (mm) EXERCISED  m=4  UNEXERCISED (N=5’i  PCr  79.9 ± 2O.9°  42.4 ± 16.4”  LACTATE  76.4 ± 20.3”  42.3 ± 15.2  (a) 0.07 <p < 0.08 for comparison to non-exercised fish.  (b) p >> 0.05 for comparison to lactate recovery time.  97  120-  I  60. 30-  y  =  3.43 + 0.92x =0.99 2 r  0 0  I  I  I  I  30  60  90  120  PCr recovery time (mm) FIGURE 18. The relationship between estimated recovery times for white muscle lactate and PCr in exercised and unexercised tuna.  98  150  x 0  x irr  x  fJlVU  0  0  o  o  E  0 +  C C..)  X  -iv  +  0 0  C  X +  •Q. ++  +.‘o •% 1’i  0 0  5  10  15  20  25  30  35  PCr (imo1.g-i)  FIGURE 19. White muscle lactate in relation to PCr in biopsies from exercised (large symbols; each symbol-type represents a different fish) and unexercised (small crosses; individuals not depicted) tuna. Also shown are measurements from muscle taken from 3 fish following 15 mm of exercise (small filled circles).  99  Results of the present study support and extend the suggestion that tuna white muscle clears lactate in situ, since the nearly stoichiometric changes in the lactate and glycogen recovery profiles (i.e., 2 lactate removed: 1 glucosyl unit reformed) indicate mainly a glyconeogenic fate for most of the lactate produced during burst activity. The extent to which this result approaches a quantitative representation of lactate incorporation into white muscle glycogen is reinforced by estimates of minimal glycogenesis from plasma glucose and low capacities for other tuna tissues to utilize lactate (discussed below).  Plasma lactate and glucose.Plasma lactate concentration in the exercised tuna remained essentially constant for the period during which muscle lactate declined sharply. In terms of the kinetics of lactate, the restrictive condition of a plasma steadystate means that lactate had to enter the plasma pool at the same rate which it was removed. Furthermore, 35  -  40 mM plasma lactate suggests, indirectly, a high rate of  flux through the plasma pool in tuna (see Weber et al. 1986).  In the post-exercise  state, white muscle is the probable source of plasma lactate which could be utilized either oxidatively or for gluconeogenesis in other tissues. In mammals, the primarily oxidative disposal of lactate observed at rest and during sustained, aerobic activity has lead to the formation of the lactate-shuttle hypothesis (Brooks 1986) which proposes the translocation of lactate from producing to consuming muscle fibres. It is not likely that oxidative metabolism is reponsible for the removal of white muscle lactate in tuna.  The oxygen consumption of skipjack tuna swimming at 2-5  1 is calculated to be 0.68 molg lengthss min’ (Gooding et a!. 1981) which is 4 1 times the rate of a recovering trout (Mulligan and McDonald 1988).  This rate of  oxygen consumption is still about 30 times too slow to account for the rate of lactate disappearance by oxidation. It is possible that tuna have higher oxygen consumptions during recovery. However, it is unlikely that red muscle or the heart of tuna even have  100  the capacity to oxidize lactate at the rate required.  Based on maximal estimates of  mitochondrial oxidation of pyruvate, tuna red muscle and heart ventricle could min respectively (Moyes et a!. lactateg , maximally oxidise 0.9 and 2.5 jmol 1  ) could oxidize about 5 1 1992). Thus, in a 1 kg animal, red muscle (assume 70 gkg , Farrell et a!. 1992) could oxidize about 0.8 1 mmol of lactate and the heart (4 gkg mmol of lactate in 80 mm.  Combined oxidation in these tissues would therefore  account for the removal of less than 10% of an initial white muscle lactate load of 72 , table 7). If the temperature of tuna red muscle was 1 mmoles (based on 108 molg 10 °C higher than ambient (Hochachka et a!. 1978) then maximal oxidation would still account for less than 15 % of the lactate clearance. The low capacity of tuna red muscle to oxidize lactate may mean that red muscle is glyconeogenic to some degree, using lactate derived from white muscle in a manner similar to the version of a “lactate shuttle” proposed for reptiles (Gleeson and Dalessio 1990).  The glycogen content of tuna red muscle is reduced following  exhaustive swimming activity (Hulbert et a!. 1979), but the quantitative importance of lactate incorporation into red muscle glycogen is expected to be low because of the relative masses of red and white muscle. Integration of a shuttling mechanism with the lactate and glycogen changes observed in tuna must accommodate the apparent conservation of lactate carbon within white muscle mass. Another way for simultaneous glycogen recovery/lactate disappearance to occur is through the delivery of plasma glucose to muscle for glycogenesis. However, the capacity for skipjack tuna liver to utilize lactate as either a gluconeogenic or an oxidizable substrate is low (Buck et a!. 1992) and in the general context of postexercise lactate removal, contributes negligibly. Weber et a!. (1986) came to a similar conclusion by noting that Cori cycle activity is probably minimal in tuna recovering from exercise. While liver glycogenolysis and gluconeogenesis from amino acids are also potential sources of plasma glucose in teleost fishes (Suarez and Mommsen 1987),  101  it remains unlikely that turnover from all of these sources is high enough to supply glucose at a rate that matches the rapid glycogen recovery evident in tuna white muscle. It is generally difficult to predict glucose flux rates because turnover is often independent of plasma glucose concentration (see Bonen et al. 1989; Weber et al. 1986). However, turnover rate in the present study would have to have been more than an order of magnitude higher than rates measured in tuna by Weber et a!. (1986) to reflect just 50 % repletion of the white muscle glycogen by plasma glucose. The implication remains that much of the lactate formed during exercise stays in the white muscle, or is retrieved from the plasma, and is the principle substrate for glycogen resynthesis. It is interesting that plasma lactate in the exercised tuna declined markedly only after lactate and glycogen recovery in white muscle approached completion. High plasma lactate likely produces a concentration gradient favorable for lactate uptake and utilization by the tissues mentioned above, but may also help to attenuate diffusive efflux of lactate from white muscle. In this regard, tuna resemble the mammals in which prolonged elevation of plasma lactate seems necessary to sustain in situ muscle glyconeogenesis (Johnson and Bagby 1988; Stevenson et a!. 1987). This situation is different from other teleosts where plasma lactate is elevated only transiently relative to the prolonged period required for white muscle lactate and glycogen recovery. Tuna need to swim continuously and this may mean that muscular contractions in free swimming fish influence changes in lactate, and other muscle metabolites, compared to spinally-blocked fish. There are no studies of lactate removal from white muscle of free swimming tuna for comparison of methodologies, but the rate of glycogen replenishment in skipjack tuna measured by terminal sampling in recovery from exercise (Barrett and Connor 1964) is comparable to changes in lactate and glycogen measured in the present study.  In addition, while swimming after intense  exercise might change both the rate of lactate clearance from white muscle and whole-  102  body turnover, it is still likely that the principal fate of lactate in tuna is as we have proposed since the potential to oxidize lactate is low and the proportion of red muscle relative to total muscle mass is small.  Lactate and PCr.  The extraordinary inverse correlation between the concentration  of PCr and the concentration of lactate has not, to our knowledge, been previously reported in muscle. Meyer and Terjung (1979) appeared to find a similar relationship between between concentrations of PCr and lactate in the recovering gastrocnemius of rats.  However, they suggested that the recovery of phosphocreatine was biphasic,  whereas the recovery of lactate was fitted best by a single exponential. A comparable relationship between PCr and lactate has also been noted in hypoxic turtle brains (Lutz  et a!. 1984). We suggest that recovery of pHi, dependent on the clearance of lactate, causes the apparent linkage of phosphocreatine to lactate during recovery from exercise in tuna.  If, as proposed, lactate is a glyconeogenic substrate then there would be  simultaneous consumption of protons within the white muscle.  This suggestion is  supported by the finding of Tang and Boutilier (1991) who found that protons produced in rainbow trout during intense exercise were most likely cleared by metabolic processes within the white muscle compartment. As discussed earlier, net efflux seems to account minimally for clearance of lactate from tuna white muscle. Thus an imbalance, if any, between the clearance of lactate and protons would have only a minimal effect on the relationship between pHi and lactate. Nevertheless, the efflux of lactate from tuna white muscle may also be linked with proton transport. Lactate and proton movements in muscle appear to be coupled in a carrier-mediated transport process (Mason and Thomas 1988; Juel and Wibrand 1989). Furthermore, lactate transport across the mammalian sarcolemma is highly sensitive to a pH-gradient (Roth and Brooks 1990). It is not clear to what extent  103  these findings can be generalized as Wiseman et al. (1989) found that the efflux of proton and lactate equivalents was not tightly coupled in molluscan muscle tissue. In tuna there is an excess of blood lactate over blood metabolic protons after exercise, but Perry et al. (1985) noted that more efficient removal of protons from plasma, rather than differential lactate/proton release from white muscle, probably accounted for this difference.  Directly measuring pHi and lactate in white muscle would resolve the  relationship between the concentrations of protons and lactate.  Nevertheless, the  evidence is consistent with the suggestion that changes in pHi are linked with the concentration of lactate. Phosphocreatine is linked to pHi through the near equilibrium reaction catalysed by creatine kinase where: PCr+ADP+H<---->ATP+Cr In addition to pHi, the ratio of ATP/ADP will also affect the final concentration of PCr. Work on turtle brain and fish white muscle indicates a clear relationship between acidosis, or glycolysis, and PCr depletion induced by hypoxia (Lutz et a!. 1984; Van Waarde et a!. 1990). During recovery from anoxia the rephosphorylation of PCr in fish muscle becomes dissociated from recovery of pHi, and presumably lactate clearance. However, during recovery from anoxia there were substantial increases in  the ratio of ATP/ADP which would dissociate pHi from PCr.  Similarly, adenylate  ratios seem to influence PCr in rainbow trout white muscle where, in recovery from exhaustive exercise, lactate is cleared at a much slower rate than PCr repletion (Schulte et a!. 1992).  The close relationship between lactate and PCr in tuna suggests that  changes in pHi determine the extent of phosphorylation of creatine during recovery.  PUBLICATION STATUS: The preceding study has been published. See Arthur et a!. 1992 in the reference list. Thanks to Dr. P.G. Arthur for measuring PCr and for conceptual and practical collaboration in this study.  104  CHAPTER 4. GENERAL DISCUSSION. Glucose as a fuel for swimming.  A comparison of glucose utilization in trout and carp  is interesting in two respects. Firstly, as hypothesized from observations of superior fat utilization capacity in faster swimming fish (Moyes et a!. 1992b), it is evident that the slow swimming carp utilizes circulatory glucose for muscle energetics to a greater extent than do rainbow trout.  As will be discussed later this interpretation assumes that red muscle  accounts for most of the increased metabolic demands at 80 % U .j, 1 1 which are lower in carp than in trout. Glucose dependency may be related largely to differences in absolute aerobic performance capacity rather than differences in muscle glucose utilization rate. Secondly, these data, particularly the concentration-dependence of glucose turnover in carp and discrepancies in glucose utilization in trout heart at different levels of organization (isolated heart vs in vivo heart), raise some questions about how glucose disposal might be regulated in the whole-animal and in various muscle-types. In addition to the energetic importance of glucose, the following discussion focuses on the significance of muscle  tissue for in vivo disposal of glucose and on some of the glucoregulatory features, consistent with the general glucose intolerance of teleosts, that emerge from a comparison of trout and carp.  Muscle tissue as a site of glucose disposal.  The contribution of various muscle-  types to the disappearance of circulatory glucose depends on not only the rate of glucose  utilization, but also on the relative masses of the muscles involved.  Muscle-type  differences in utilization rate are in accord with observations in trout and carp indicating higher hexokinase (HK) activity in heart and red muscle compared white muscle (at 15 °C, approximately 5 0.1  -  -  g in heart and 1 1 10 molmin  -  21 molmin in red muscle versus g  g in white muscle, Johnston 1977; Dnedzic 1987). It is likely that 1 0.5 molmin  105  muscle-type differences in glucose utilization capacity parallel differences in transport capacity, suggesting that fish may be similar to mammals, in which the relative capacity for glucose transport been shown to be superior in the red muscle fibers (Marette et al. 1992; Megeney et a!. 1993). However, despite relatively low rates of glucose utilization in fish white muscle, this muscle is a potentially significant site for circulatory glucose disposal because of its large mass relative to red muscle and heart. The relative masses of white, red and heart muscle are similar for trout and carp, with the major difference being the presence of a large intermediate (pink) fiber mass in carp that is about 10 % of the total muscle (Johnston 1977; Johnston et al. 1977). Simply because of its large mass, white muscle at rest is estimated to account for a large portion of the glucose disposal in the carp (see table 9). In total, heart, red and white skeletal muscle are estimated to account for 50  -  70 % of glucose diappearance in resting animals. The  relative importance of each skeletal muscle type is similar because of the differences in relative glucose uptake and relative mass of the two tissues. However, glucose disposal in red muscle was somewhat higher when turnover was low and increased in both low and high turnover conditions during exercise.  Note that the one glycogen depleted fish  (discussed earlier) was eliminated from this analysis because of the apparent dependency of glucose uptake on muscle glycogen below levels of 10 mol glucosyl unitsg 1 (Figure 6). In the more general situation, red muscle GUI was stimulated to a relatively constant level in individual swimming carp (Figure 7), despite the concentration-dependence of wholebody turnover.  Concentration effects on glucose flux in swimming carp therefore had  more relevance to tissues other than the active red muscle.  White muscle, given its  preference for carbohydrate fuels (Driedzic and Hochachka 1975; Moyes et a!. 1989), might be expected to be one such tissue. However, exercise- and concentration-effects on GUI in white muscle were not significant, even though trends (Figure 7) looked similar to  those observed for whole-body turnover (Figure 5). On the other hand, while GUI in heart muscle was concentration-dependent (Figure 7), its small relative mass makes it an  106  TABLE 9.  Total glucose disposal in the cardiac and skeletal muscle of carp. It is  assumed that carp body mass is comprised of 66 % white muscle, 7 % red muscle and 1 (Bone 1978; Sidell et a!. 1987). The contribution of that ventricle mass is 0.76 gkg specific muscles to the whole-body disappearance of glucose assumes a 2-DG lumped constant (see text) of 0.40 for red muscle and heart (West et a!. 1993) and 0.51 for white muscle (West et a!., in press) when calculating glucose utilization rate (GUR). To produce the data ranges, resting and swimming carp were subdivided on the basis of turnover rate  -  n was chosen arbitrarily to designate carp as having g’ 1 molmi 5 k  low turnover (< 5) or high turnover (5 and above). The one glycogen depleted fish was excluded.  Wet weight of Carp (g) 1360  Resting  -  1830  Precentage of GUR Glucose Turnover Turnover (nmolrnin Accounted for ) ) 1 1 (nmolmin 3000  -  11400  white red heart Swimming white red heart  800 4250 800 -1250 12-56  26- 37 26-11 <1  1600-5200 2340-2700 34-80  28-34 40-17 <1  -  1175  -  1525  5750  -  15300  107  insignificant disposal site for circulatory glucose (Table 9). Pink skeletal muscle was not sampled in the present study, but given its prominent mass, its glycolytic disposition and its probable recruitment at 1.3 body lengthssec 1 (Johnston et a!. 1977) pink fibers could well have accounted for a large portion of glucose disposal in carp. Turnover rate was not measured in resting rainbow trout. However, glucose uptake in the total red muscle and heart masses (about 70 g and 1 g respecitvely) of a resting 1 kg trout requires a rate of circulatory glucose delivery of about 80 nmolmin 1 in fish at rest. The total white muscle mass (about 660 gkg 1 body mass) is estimated to demand considerably more circulatory glucose at rest (about 300 nmolmin , based on utilization 1 rates in Table 4). However, we cannot be certain what proportion of resting whole-body glucose turnover is accounted for by combined utilization in these tissues. Estimates of glucose turnover range from 1 2  -  -  10 1 kg in trout with resting glucose levels of molmin  5 mM (Dunn and Hochachka 1987; Garin et a!. 1987; Washburn et a!. 1992; or  estimated from Figure 12, chapter 2.1). Uptake in muscle tissue at rest would therefore account for 3  -  30 % of circulatory glucose turnover, suggesting that the bulk of glucose  disposal occurs in other tissues in trout.  During exercise the significant increase in red  muscle glucose utilization would raise this estimate of disposal, but it is not known whether turnover rate changes in swimming trout.  The uncertainty in designating trout glucose  disposal at rest as low compared to carp arises from the possibility that glucose utilization in trout skeletal muscle may display limited sensitivity to plasma glucose concentration which would bring estimates of muscle glucose disposal in closer agreement with the estimations made for carp.  Maximal glucose flux.  As was discussed earlier, HK activity provides a guide to the  relative capacity for glucose flux in different muscle-types. The general pattern of higher glucose utilization rates in more oxidative muscles is consistent with the trends seen in HK activity in carp and trout muscle.  Interestingly, an apparent excess in flux capacity,  108  relative to in vivo utilization rates, exists in all muscle-types examined. The reasons for excess capacity to phosphorylate glucose are not clear, although such observations have been made before. In isolated rat hearts, }{K has been measured at 5-fold in excess of peak glucose oxidation rates (England and Randle 1967). It may be that these capacities are required for situations in which alternative fuels flux  -  -  those that may be inhibiting glucose  are limiting, but it would appear that flux capacity is in excess of even the normal  aerobic metabolic requirements of some tissues. The trout heart is one such system and provides good example for discussion since glucose flux data is available from in vitro and  in vivo experiments. Measurements of trout heart hexokinase in vitro indicate a potential glucose flux of ’min (see Driedzic et a!. 1987), more than 1000 times greater than nmolg up to 8300 1 observed in vivo fluxes. It is very likely however, that actual maximal in vivo fluxes are lower since cellular substrate concentrations are usually less than saturating and possible structural roles for enzymes may influence availability for maximal flux compared to in vitro estimates (Wright and Albe 1990). It may be that the peak glucose utilization seen in the NaCN heart perfusions (i.e., -500 1 min more closely resembles flux t nmolg ) capacity in vivo. The surprising observation is that in vivo rates of glucose utilization are so low (-5 nmolg min’) 1 normoxic isolated hearts (30  considerably lower, in fact, than even rates of uptake in  -  -  120 ) min 1 nmolg .  Seemingly, both endogenous (eg.,  preference for intracellular fat) and circulatory (hormone effects and circulatory fuel supply) factors contribute to nearly complete suppression of glucose utilization in normal post-absorptive trout. From another perspective however, the in vitro results suggest at least the potential to expand myocardial reliance on glucose in vivo. It is relevant to ask under what conditions the apparent capacity for glucose flux might be approached in the whole animal. Hypoxic-stress is one situation in which myocardial glucose utilization is expected to be increased in vivo, with greater energy provision from glycolysis allowing the heart to  109  sustain performance in the absence of adequate oxidative ATP production.  Increased  glucose utilization in the NaCN perfusions of isolated hearts is analogous to the glycolytic response expected, although during complete inhibition of oxidative metabolism in trout hearts can sustain only very low, subphysiological, work rates. progressive hypoxia has been examined in previous studies.  The response to  In sea raven hearts, for  example, power output remains steady through the early stages of progressive hypoxia even though oxygen consumption declines steadily (Farrell et a!. 1985). Cardiac performance may be protected to some extent by glycolytic stimulation until the oxygen level falls below a critical point.  The responses of trout heart to hypoxia are less clear, although  there is an indication that performance drops off gradually with progressive hypoxia (Farrell et a!. 1989) and this roughly coincides with a drop in whole-body oxygen consumption (see Boutilier et a!. 1988). A decline in heart metabolism in this case may mean that the heart is able to function aerobically and shows enhanced glycolytic activity only at very low arterial oxygen levels. Support for this alternative possibility comes from whole animal responses to hypoxia in which metabolic depression occurs earlier in a hypoxic challenge than stimulation of glycolysis (Boutilier et a!. 1988). In either situation, that shown by the sea raven heart or that of the trout heart, eventual stimulation of glycolysis could greatly increase phosphorylation of glucose in an effort to support in vivo cardiac function during periods of low oxygen availability. One way to increase normoxic glucose flux in vivo is to increase the availability of glucose in the circulation. This is evident in both resting and swimming carp (Figure 7) and seems to be consistent with the concentration-dependent changes in glucose turnover rate (discussed next section).  Interestingly, glucose uptake in carp red muscle, while  responsive to exercise, was concentration-independent. However, it would seem unlikely that the glucose phosphorylation process was somehow saturated in active red muscle since flux was able to increase considearbly when muscle glycogen content was low. Glucose uptake and phosphorylation would seem to be regulated differently in the two muscle-  110  types, but the results indicate that glucose flux is indeed flexible in both muscles under different in vivo conditions. Normoxic glucose fluxes probably never match in vitro flux capacity, perhaps because of inhibition of flux in vivo (England and Randle 1967) or intracellular compartmentation of the enzyme or reduced in vivo catalytic potential because of possible structural roles for enzymes (Wright and Albe 1990). HK activities that are seemingly in excess might be viewed more appropriately as reserve capacities (See Diamond and Hammond 1992) that are exploited for shorter term protection of cellular energetics when oxidative metabolism is compromised.  Regulation of glucose utilization. Among resting carp, blood glucose level varied considerably (3  -  17 mM) and glucose turnover increased with plasma glucose  concentration. These observations suggest, firstly, that carp are typical of teleosts in not regulating resting blood glucose concentration to within rigid in vivo limits (Harmon et a!. 1991; Mommsen and Plisetskaya 1991; Nagai and Shizunori 1973; Palmer and Ryman 1972) and, secondly, that glucose availability itself may have been important in mediating uptake in peripheral tissues. Substrate-mediated glucose disposal is evident in mammalian models that display ineffective (diabetics) or underdeveloped (neonate dogs) hormonal glucoregulation (Doberne et al. 1982; Hulman et a!. 1988; Verdonk et a!. 1981). This mode of plasma glucose removal is consistent with a recent suggestion that among salmonids insulin secretion itself may be influenced by interactions with other circulating hormones. While it is evident that insulin serves an important anabolic function in fish (Mommsen and Plisetskaya 1991), potential interactions with other glucose stimulated hormones delay the insulin response, thereby, contributing to the apparent glucose intolerance of salmonids (Harmon et a!. 1991). It is not known to what extent hormonal interactions can be generalized among teleosts, nor is it clear that all species demonstrate the inability to regulate glucose within narrow limits, as evident in carp and in trout. However, proportional changes in glucose utilization with increasing glucose concentration  111  in both species tends to support a possibly dominant role for glucose availability in the regulation of tissue glucose disposal rate, regardless of activity state. The concentration-dependence of glucose turnover rate in carp has further implications for glucoregulation in swimming fish. The difference in regression slopes for resting and swimming carp suggests that it would be effective for individual fish to become hyperglycemic during a rest-to-work transition in order to enhance delivery of glucose to those tissues which have an increased demand for oxidative substrate. We have no direct evidence for this kinetic pattern in carp. However, another study (Van Dijk et a!. 1993) has shown that plasma glucose in carp swimming at a high relative speed for 25 h was unchanged from the pre-exercise level throughout exercise. This implies that stimulation of glucose flux in the present study most likely occurred in the absence of a direct effect of exercise on plasma glucose concentration, supposedly via parallel up-regulation of the rates of glucose appearance and disappearance (Wasserman et a!. 1989).  Thus, potential  expansion of glucose use in swimming carp would seem to be largely fortuitous, depending very much on the prevailing blood glucose status of individual fish prior to exercise. This situation is also consistent with having poor control over plasma glucose concentration and with possible substrate-mediation of glucose uptake.  However, the regulatory features  which would allow up-regulation of hepatic glucose release concomitant with increased glucose disposal require further investigation.  Exercise energetics in carp and trout.  Some differences in the kinetics and utilization  of glucose between carp and rainbow trout imply that glucose is somewhat more important for energy provision in carp. Lower glucose disposal in trout skeletal muscle than in that of carp suggests, indirectly, that glucose may be utilized to a greater extent in the carp for resting metabolism. Skeletal muscle accounted for about 50 % of glucose disappearance in resting carp (plasma glucose, 3  -  17 mM), compared to estimates of less than 20  in trout  112  (plasma glucose, 5 among  -  40 mM). Glucose in resting skeletal muscle is expected to divided  biosynthetic  production) pathways.  (lipid/glycogen  storage)  and  biodegradative  (oxidation/lactate  While higher disposal in carp musculature suggests, at least  qualitatively, increased availability of intracellular glucose-carbon for resting muscle energetics, the uncertainty in assigning any quantitative significance to species differences in resting glucose utilization arises from the lack of satisfactory estimates of in vivo maintenance costs in red and white muscle. During exercise, cardiac energetics are difficult to compare directly between the two species because of the clear concentration-dependence of GUI in carp myocardium. Trout myocardial glucose utilization was determined at plasma glucose levels of 2  -  4 mM  and, unlike carp, an exercise-effect was not detected. Glucose accounted for < 1 % of cardiac energetics in trout swimming at 80 % Uj (West et al. 1993). However, if the concentration-dependent GUI for carp was extrapolated to lower plasma glucose levels (Figure 7), then it is apparent that an exercise effect would have been more difficult to detect.  Moreover, the maximal proportion of glucose that could account for estimated  heart oxygen consumption of active carp would approach estimates for the trout heart.  In contrast to heart, carp red muscle glucose uptake was independent of plasma glucose concentration and estimated glucose utilization was stimulated to similar levels in carp and trout (20  -  ). 1 30 nmolmin’g  However, the smaller metabolic scope for  sustained swimming in the less active carp accounted for a higher estimate of glucose use for energy provision in carp red muscle (25  -  30 % in carp, < 10 % in trout) while  swimming at 80 % U. Obviously, comparison of glucose utilization in these species is most useful if the assumption holds that red muscle accounts for the bulk of the active oxygen consumption. If, in particular, aerobic recruitment of white muscle is an important metabolic cost during exercise, then this would have to be accounted for in calculations of red muscle dependence on glucose.  Simply stated, the nearly 2.5-fold higher aerobic  capacity of trout white muscle compared to carp (based on citrate synthase activity and  113  mitochondrial protein content per gram red muscle, Moyes et a!. 1989; 1992b) may contribute significantly to the difference in aerobic swimming capacity in these species. Greater aerobic involvement of white muscle in trout than in carp at 80 % Uj would tend to bring estimates of glucose oxidation in closer agreement in these species.  While the  other aerobic costs were not accounted for in these studies, it was noted that even if 50 % of whole body oxygen consumption in salmonids is accounted for by tissues other than red muscle, this would have minimal effect on the estimate of the use of glucose for swimming energetics of trout. Greater use of glucose in the less active carp would be consistent with the generalization that aerobic exercise capacity influences the type of substrate used in support of maximal oxidative energy provision.  Post-exercise recovery of muscle glycogen.  Comparison of the rate of lactate  clearance, or glycogen repletion, in trout and tuna indicates a greater rate of recovery, by 10  -  30 times, in tuna white muscle. However, the balance between lactate clearance and  glycogen restoration in tuna white muscle provides evidence that, like other fish species, lactate is metabolized in situ via glyconeogenesis. It is likely that temperature accounts for part of the difference in recovery rate between trout and tuna, but clearance rate across temperate water species correlates with a number of enzymatic indicators of tissue metabolic capacities, including glycolytic capacity (eg., suggested by pyruvate kinase (PK) activities), marker enzymes for muscle glyconeogenesis (eg., malic enzyme (ME) activities) and tissue aerobic capacity (eg., citrate synthase activities), with the highest lactate fluxes evident in species with greater metabolic capacities (Moyes et al. 1993). The tunas demonstrate superior capacities, even when temperature is factored out, while trout display a more intermediate position. However, it is not certain what these correlations mean specifically to the rate of lactate clearance. The relationship with aerobic capacity is unlikely to be indicatative of enhanced oxidative removal of lactate, since conservation of lactate carbon for glyconeogenesis  114  seems consistent across species, including the highly aerobic tuna white muscle. Interestingly, the mitochondrial capacity to produce ATP, although inhibited in vivo by high ATP:ADPfr ratios, is far in excess of the energetic demand for conversion of white muscle lactate to glycogen (Moyes et a!. 1992b). Thus, aerobic capacity would appear not to be rate limiting to carbohydrate dynamics in recovering fish (Moyes et a!. 1992b). Differences in the rate of lactate clearance between trout and tuna are more likely related to the capacity to convert lactate to glycogen. PK and ME activities may have implications for the route, and rate, of glyconeogenic flux in white muscle (next section), but it has yet to be demonstrated experimentally that the capacity for flux through these pathways limits recovery in vivo. Despite differences in recovery rates, teleost white muscle across species can be regarded as almost a self-contained metabolic system with respect to carbohydrate status during burst exercise and recovery. The dependence of white muscle on glycolysis and on intramuscular reconversion of lactate to glycogen allows gross level lactate/glycogen changes in white muscle to be examined in a reasonably quantitative manner (Arthur et a!. 1992; Milligan and Wood 1986a,b; Moyes et al. 1992b; Pagnotta and Milligan 1991; Schulte et a!. 1992; Tang and Boutilier 1991).  However, the fine details of lactate  clearance and the role of glucose are somewhat more difficult to place into a quantitative framework. Apparent deficits in white muscle glycogen in trout range 0  -  50 %, trout  relative to resting control glycogen levels (Mulligan and Wood 1986b; Mommsen and Hochachka 1988; Pagnotta and Milligan 1991; Pearson et al. 1990; Schulte et a!. 1992 and this study).  Oxidative losses of lactate within the white muscle are probably minimal,  since preference for fatty acids for recovery energetics (Bahr et a!. 1991; Bangsbo et al. 1991), even in the presence of high lactate levels (Moyes et at. 1992b), would tend to spare lactate that is retained in white muscle for primarily glyconeogenesis. Most analyses suggest that only 10  -  20 % of the lactate is rerouted away from muscle glyconeogenesis  (Tang and Boutilier 1991; Schulte et at. 1992), possibly via extra(white)muscular oxidation  115  (Scarabello et a!. 1991; Moyes et a!. 1992) or hepatic gluconeogenesis.  Similarly,  maximal losses of lactate in tuna white muscle, sampled with sequential biopsy, are expected to be < 10 %, based on the capacities for oxidation and gluconeogenesis. In vivo rates of lactate oxidation in heart and red muscle and of possible red muscle glyconeogenesis are unknown and the hepatic gluconeogenic rate from lactate is low (Milligan and McDonald 1988; Weber et a!. 1986; Buck et a!. 1992), but together these processes might explain minor mismatches between white muscle glycogen replenishment and lactate clearance rates that are sometimes observed. In turn, low glucose uptake, as determined in trout, would be most useful for post-exercise glycogen restoration over the long-term.  A better understanding of the role of glucose uptake in white muscle may  emerge from studies that include specific examinations of mismatches in lactate/glycogen stoichiometries and of apparent deficits in glycogen recovery. It is expected, however, that circulatory glucose serves mainly as a supplement to the normal turnover of intramuscular glycogen reserves and perhaps in an energetic role for resting muscle metabolism.  Muscle glyconeogenesis.  These studies, like most examinations of white muscle  glyconeogenesis in vertebrates, provide evidence for an intramuscular pathway of lactate reconversion to glycogen. The major gap in our understanding of this process is that the pathway of lactate flux into the phospho(enol)pyruvate (PEP) pool remains elusive. Of the analyses performed over the past 25 years, evidence would seem to eliminate mitochondrial conversion of pyruvate to oxaloacetate (OAA) and subsequent shuttling of OAA to the cytoplasm for decarboxylation to PEP since pyruvate carboxylase has not been detected in muscle extracts (Opie and Newsholm 1967). Furthermore, there is no apparent randomization of 14 C in labeled lactate that was metabolised by perfused rabbit muscles, suggestive of minimal interconversion of lactate carbon with Krebs cycle intermediates (Pagliassotti and Donovan 1990). There are two proposed extramitochondrial pathways; (1) conversion of pyruvate to PEP via malate and OAA, requiring C0 , GTP and the 2  116  presence of malic enzyme (ME) and PEP carboxykinase (PEPCK), and (2) reversal of PK, requiring ratios of pyruvate/PEP and ATP/ADP that are favorable to reversal of flux. PEPCK is detected at low activities in vertebrate muscles (Opie and Newsholme 1967; Crabtree et al. 1972) and Connett (1979) indicates that its inhibition in frog muscle reduces lactate incorporation into glycogen. Conversly, the data of Pagliasotti and Donovan (1990) points to glycogen repletion independent of PEPCK activity in rabbit muscles.  The  situation in fish seems less ambiguous since PEPCK activity has not been detected in white muscle of species for which lactate clearance rates are known (Moyes et al. 1993), including tuna muscle where rapid carbohydrate recovery might be expected to be  associated with relatively high levels of PEPCK.  The only known extramitochondrial  pathway that is independent of PEPCK activity is reversal of PK. It is not known whether mass action ratios in recovering fish white muscle favor flux from pyruvate to PEP through PK.  However, adenylate ratios are elevated during recovery in trout (ATP:ADPfr  =  2000, Schulte et al. 1992) and, in conjunction with elevated pyruvate levels (6-fold in trout, Schulte et a!. 1992), would favor PK reversal. Conditions may well be similar in tuna (pyruvate increases 20-fold over resting levels, for example) and it is evident that reverse flux rates in tuna white muscle would have to equal about 0.1 % of maximal PK activities (about 1300 molmin , Guppy et a!. 1979) to completely account for rates g 1 of lactate clearance a level of reverse flux that is well below maximum reversal rates of 2 -  % of forward flux determined by manipulation of metabolite levels in in vitro assays (Dyson et a!. 1975). In trout, a similar level reverse PK flux (0.04 % of 140 molmin , Moon and Johnston 1980b) would account for lactate clearance in white muscle g 1 (Moyes et a!. 1992b).  Generally, species with lower lactate clearance rates also display  reduced PK activities, thereby maintaining the need for PK reversal at 0.03  -  0.1 % of the  maximal reaction velocity (Moyes and West, in press). It is not known by what mechanism trout red muscle may convert lactate carbon to PEP, although observations in rabbit muscles (Pagliassotti and Donovan 1990) indicate that  117  glyconeogenesis occurs by a general pathway in all muscle types. However, the capacity for glyconeogenesis is related to oxidative capacity, with red fibers displaying the lowest levels of lactate incorporation. In trout, the same tendency is suggested indirectly from the increased potential of red muscle to use glucose for glycogen resynthesis after exercise. Red muscle, should be examined in comparative manner, similar to previous studies on white muscle, to identify regualtory features that help to distinguish lactate dynamics in relation to aerobic capacity, glycolytic potential and fiber-type across species.  Glucose and lactate in fish white muscle. Regardless of species differences in the rate of white muscle lactate clearance, post-exercise lactate turnover and white muscle aerobic capacity, the fmal fate of lactate seems restricted to glycogen restoration in fishes. Low rates of glucose utilization in trout, demonstrate that flux is insufficient to support glycogen synthesis rates observed. In tuna, previous estimates of recovery glucose turnover, even in the unrealistic situation of exclusive delivery to white muscle, are too low (about 15 molmin’kg’, plasma glucose 2  -  ; Weber 1986) for glycogenesis to account 1 6 mmolL  for glycogen repletion in the present study (requiring > 350 1 kg assuming molmin , 660 g white muscle per kg body weight). The stoichiometry between white muscle lactate and glycogen argues against significant rerouting of lactate away from a glyconeogenic fate.  The apparent  inconsistency is that tuna display relatively high lactate turnover rates, as measured with radiolabelled lactate (Weber et al. 1986). In trout, although plasma lactate kinetics appear unrelated to white muscle glycogen replenishment, there is evidence that an initial outward flux of lactate from white muscle is followed by inward transport to favor metabolism in situ (Turner and Wood 1983). A similar kinetic pattern in tuna would mean that wholebody turnover reflects lactate uptake from plasma by the lactate-producing as well as consuming tissues.  Simultaneous lactate extraction and removal has been observed in  exercising human leg muscle, in which the extraction of tracer lactate from plasma occurs  118  despite net lactate release by muscle (Stanley et al. 1986). High plasma concentrations of lactate and mammal-like turnover rates would, therefore, seem to be indicative of the plasma compartment acting as a temporary storage space for lactate that is eventually taken up by mainly the white muscle mass.  This kinetic pattern cannot be confirmed in the  present tuna study, but low capacities for lactate use in other tissues and the observation that plasma lactate remained elevated until the late stages of white muscle clearance provides some indirect evidence. In both trout and tuna the significance of an exercise-recovery cycle of glycogen lactate-glycogen is to conserve carbohydrate status of the white muscle. Glycogen reserves represent a short term fuel reserved for burst swimming and that by conserving lactate, it is possible to conserve glycogen.  The advantage, given that dependence on glycogenesis  alone would greatly lengthen recovery time, is that carbohydrate recovery can occur independently of processes that are incompatible with the rapid replenishment of glycogen (i.e., glucose delivery from food sources or gluconeogenesis from amino acid precursors). An analogy can be drawn with the PCr stores, which are available as a short term energy store that can also be rapidly replenished after exercise in both trout and tuna.  Rapid  metabolite recovery ensures that fish are ready to engage in another burst of high speed swimming as quickly as possible. More comprehensive comparative examinations of post exercise metabolite changes (Moyes et a!. 1992b; Schulte et a!. 1992) should elucidate limitations to the rate of carbohydrate recovery across species.  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