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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 SUSTAINEDSWIMMING AND RECOVERY FROM MAXIMAL SWIMMING.ByTimothy G. WestB.Sc. Biology. Daihousie University, 1982M.Sc. Biology. Daihousie University, 1986A THESIS SUBMITTED IN PARTIAL FULFILLMENT OFTHE REQUIREMENTS FOR THE DEGREE OFDOCTOR OF PHILOSOPHYinTHE FACULTY OF GRADUATE STUDIES(DEPARTMENT OF ZOOLOGY)THE UNIVERSITY OF BRITISH COLUMBIAJanuary 1994© Timothy G. West, 1994We accept this thesis as conformingstandardIn presenting this thesis in partial fulfilment of the requirements for an advanceddegree at the University of British Columbia, I agree that the Library shall make itfreely available for reference and study. I further agree that permission for extensivecopying of this thesis for scholarly purposes may be granted by the head of mydepartment or by his or her representatives. It is understood that copying orpublication of this thesis for financial gain shall not be allowed without my writtenpermission.(Signature)Department of o -The University of British ColumbiaVancouver, CanadaDate /1 tNDE-6 (2/88)11ABSTRACTThe importance of circulatory glucose as a fuel for skeletal muscle and heartduring sustained swimming and as a glycogenic substrate for skeletal muscle duringexercise-recovery was examined. In vivo glucose utilization, using14C-2-deoxyglucose(14C-2-DG), in swimming trout (Oncorhynchus mykiss) and carp (Cyprinus carpio)indicated similar rates of glucose utilization in red muscle of both species (20 - 30nmolmin1g). Estimates of the energetic importance of glucose, assuming redmuscle accounted for the bulk of the active oxygen consumption, suggest that glucosecould 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 troutwas estimated to contribute < 10 % of total substrate demands during exercise.Cardiac energetics was estimated to be largely independent of glucose utilization introut. In vivo utilization of glucose in trout heart at rest (about 5 nmolmin1)wasconsiderably lower than in vitro determinations (120 nmolming)and measurementsof glucose utilization capacity (8300 nmolmin1g’,based on hexokinase activities).Nearly complete inhibition of glucose flux in vivo is likely the result of preference foralternative substrates for oxidation. In carp, cardiac glucose utilization was dependenton plasma glucose concentration, but was also estimated to be of minor energeticimportance at plasma concentrations comparable to those observed in swimming trout(2 - 4 mmolLt). Reduced dependence of trout on glucose for swimming energeticsagrees with the generalization that species with superior aerobic exercise capacities maybe dependent on fat based fuels for muscle energetics.Plasma glucose concentration varied from 6 - 38 mmolL1 in trout duringrecovery from intense burst swimming. Hyperglycemia was not related specifically toexercise. Estimated glucose turnover rate, using 6-3H-glucose tracer, increasedproportionally with plasma glucose availability. Similar concentration-dependentkinetics were observed in carp at rest and during sustained swimming. Regardless of11].the association between glucose turnover and plasma glucose level, glucose utilization(14C-2-DG phosphorylation) in trout white muscle accounted for < 10 % of theglycogen repletion after exercise. In trout red muscle, glucose utilization wasdependent on plasma glucose concentration up to 10 - 12 mmoPL1 and glycogenesiswas estimated to account for 25 - 60 % of post-exercise glycogen repletion in thismuscle type. Moderate post-exercise hyperglycemia sometimes seen in recoveringsalmonids seems pertinent to red muscle carbohydrate status, but white muscle recoveryoccurs independently of plasma glucose availability. These results provide evidence forthe 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 oxidativecapacity of tuna white muscle and previous determinations of high circulatory fluxes oflactate, this species also seems to convert white muscle lactate to glycogen viaintramuscular glyconeogenesis. A glyconeogenic fate for lactate conservescarbohydrate status and ensures relatively rapid replenishment of the principal carbonsource in fish white muscle.ivTABLE OF CONTENTSABSTRACT iiTABLE OF CONTENTS ivLIST OF FIGURES viiiLIST OF TABLES ixACKNOWLEDGEMENTS xCHAPTER 1. SUBSTRATE UTILIZATION IN FISH MUSCLE DURINGEXERCISE AND RECOVERY. 1Sustained swimming 2Fuel selection 3Protein and amino acid utilization 6Utilization of fat fuels 8Carbohydrate utilization 10White muscle 15White muscle recruitment and exercise metabolism 16Carbohydrate recovery in fish muscle 18In vivo measurement of glucose utilization 20CHAPTER 2. GLUCOSE UTILIZATION DURING SUSTAINED SWIMMING INRAINBOW TROUT (Oncorhynchus mykiss) AND CARP (Cyprinus carpio).In vivo utilization of glucose by heart and locomotory muscles ofexercising rainbow trout (Oncorhynchus mykiss). 23Introduction 23Materials and Methods 24Animals 242-DG and the lumped constant 24Estimation of LC in muscle slices 25VEstimation of LC in isolated trout hearts 27Glucose utilization in swimming trout 28Statistics 29Results 30Tissue LC’s and the effect of glycolytic rate 30Glucose utilization in swimming trout 30Discussion 34The 2-DG lumped constant 34Glucose utilization in red muscle 37White muscle glucose uptake 40Cardiac glucose utilization 41Muscle glucose utilization during sustained swimming in the carp. 43Introduction 43Materials and Methods 44Animals 44Estimation ofSustained swimming in cannulated carp 44Bolus-injections and tissue sampling 45Metabolite assays 46Calculations 47Statistics 48Results 48Carp swimming performance 48Glucose turnover 50Muscle glycogen and glucose utilizaiton 50Discussion 56Oxygen consumption 56viGlucose turnover 58Muscle glucose uptake 59CHAPTER 3. CARBOHYDRATE RECOVERY IN SKELETAL MUSCLE OFRAINBOW TROUT (ONCORHYNCHUS MYKISS) AND SKIPJACK TUNA(KATSUWONUS PEL4MIS) AFTER BURST EXERCISE. 62Implication of hyperglycemia for post-exerciseresynthesis of glycogen in trout skeletal muscle. 62Introduction 62Materials and Methods 63Animals 63Surgery 63Exercise and recovery protocol 65Isotope injection and tissue sampling 66Metabolite assays 67The 2-DG lumped constant 67Glucose turnover and muscle uptake 68Statistics 68Results 68Glucose concentration and turnover rate 68Muscle glucose utilization 71Glycogen replenishment in skeletal muscle 76Discussion 76Glucose turnover 77White muscle recovery 79Red muscle recovery 80viiLactate, glycogen and PCr profiles in the white muscleof skipjack tuna during recovery from burst swimming. 82Introduction 82Materials and Methods 83Animals 83Exercise and biopsy protocol 83Tissue preparation and analysis 85Data analysis 86Results 86Muscle metabolites after exercise 86Lactate, glucose and glycogen in recovery. 88Estimates of recovery time (lactate and PCr) 91Discussion 95Muscle lactate and glycogen recovery 95Plasma laãtate and glucose 99Lactate and PCr 102CHAPTER 4. GENERAL DISCUSSION. 104Glucose as a fuel for swimming 104Muscle tissue as a site of glucose disposal 104Maximal glucose flux 107Regulation of glucose utilization 110Exercise energetics in trout and carp 111Post-exercise recovery of muscle glycogen 113Muscle glyconeogenesis 115Glucose and lactate in fish white muscle 117LITERATURE CITED 119viiiLIST OF FIGURES1. Glucose utilization in normoxic and NaCN-treated trout hearts. 332. 14C-2-DG washout in resting and swimming trout. 353. Oxygen consumption in relation to swimming speed in carp. 494. Washout of glucose tracers in resting and swimming carp. 515. Effect of glucose availability on glucose turnover and clearance in carp. 536. Effect of muscle glycogen content on glucose utilization in carp red muscle. 557. Effect of glucose availability on glucose utilization index in carp muscles. 578. Post-exercise glucose and lactate levels in trout plasma. 649. Washout of glucose tracers in resting trout. 6910. Washout of glucose tracers in recovering trout. 7011. Effect of glucose availability on glucose turnover and clearance in trout. 7312. Carbohydrate recovery and glucose utilization rate in trout white muscle. 7413. Carbohydrate recovery and glucose utilization in trout red muscle. 7514. Recovery profiles of glycogen, lactate and PCr in tuna white muscle. 8915. Plasma glucose and lactate levels in tuna. 9016. Break points in lactate and PCr recovery in representative tuna. 9217. Recovery time in relation to tuna white muscle metabolite content. 9318. Association between lactate and PCr recovery time in tuna white muscle. 9719. Association between lactate and PCr content in tuna white muscle. 98ixLIST OF TABLES1. Fuel stores in a ‘ ypcal salmonid’. 52. Lumped constants in trout muscle slices. 313. Trout heart lumped constant: effect of glycolytic rate. 324. Glucose utilization in muscles of resting and swimming trout. 385. Glucose kinetics and muscle GUI in resting and swimming carp. 526. Glucose turnover and utilization in resting and recovering trout. 727. Metabolites in white muscle of exercised and unexercised tuna. 878. Mean recovery times for lactate and PCr in tuna white muscle. 969. Glucose disposal in carp muscles as a proportion of whole-body turnover. 106xACKNOWLEDGEMENTSThanks first and foremost to Dr. Peter W. Hochachka for your guidance,enthusiasm, and financial support, for forgetting (once, at least) the ‘just-one-conference-a-year’ thing and for handing over the keys to the lab and keeping a sense of humor aboutmy not returning them until recently. The emphasis here has always been on discussionand collaboration and I’m greatly appreciative of, what I believe was, an extraordinarygroup of lab personnel. Their assistance and constant feedback was (is) immeasurable. Iam very grateful to Dr. ‘s R.K. Suarez, P.G. Arthur, and C.D. Moyes for sharing their labknow-how and for many discussions. Thanks also to Peter Arthur for his skill with theisolated trout heart preparation and for measuring PCr in tuna white muscle and to TrishSchulte 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 providingthe 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 helpcannulating and spinal-blocking tuna, and the Bamfield Marine Station staff for technicalsupport in the study of sustained swimming in trout.Thanks: to Steve for corroborating those many encounters with landlorditis and forpointing out the, not so obvious, predictive value of faulty water heating. Thanks forintroducing the Brownies, Laphroaigs, MacAllans, and Youngers. Sorry about the ciderchunks!; to Sohail for camaraderie, from hallway frisbee to toasting the data god from thetop of %$*%#@ Mount Harvey.. .always a laugh, man; to the HABS faithful, and the jaysfaithful- 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 demonstrationof cooperation, all squash partners for enervation, and to Raul for the extra cash.Thanks especially to Mom and Dad for understanding about the infrequent visitshome.1CHAPTER 1: SUBSTRATE UTILIZATION IN FISH MUSCLE DURINGEXERCISE AND RECOVERY.Comparative studies of fish energetics indicate a striking diversity in exerciseperformance capability across species. Aerobic capacity of skeletal muscle is about 20-foldhigher in fast than in slow swimming species and rates of recovery from short-termmaximal 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 fishwould also display variable patterns of substrate utilization for oxidative energy provisionduring sustained swimming and for restoration of intramuscular fuel depots duringrecovery from exercise. For example, indirect observations indicate that fish species withsuperior swimming capabilities have elevated muscle oxidative capacities as well as greatercapacities for fat utilization compared to less active fish (Moyes et a!. 1992a). Similartendencies are noted among mammals, where aerobically adapted species seem moredependent on lipid based fuels than sedentary species (Weber 1992). However, there is nocompeling evidence among fishes that activity metabolism in slower-swimming species issupported mainly by one particular type of substrate (i.e., fat versus carbohydrate basedfuels) over another (Moyes et al. 1989; Driedzic 1988; Sidell et a!. 1987).Although generalizations are often made regarding the importance of circulatoryglucose in activity related processes in fish, few studies have addressed glucose fluxesduring exercise and recovery and there are no estimates of in vivo glucose utilization inspecific muscle-types during exercise. Glucose dependency for swimming energeticsacross species may reflect differences in aerobic swimming capabilities. Similarly,although the use of circulatory glucose for post-exercise restoration of muscle carbohydratestores 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. Forexample, tuna possess elevated aerobic capacities and also have the capacity for high postexercise lactate fluxes (Weber et al. 1986). There is the possibility that a major portion of2the post-exercise lactate load is oxidized in fast-swimming species, rather than used formuscle glyconeogenesis. The following studies exploit the differences in aerobicswimming capacity across species to investigate the in vivo role of circulatory glucose inmuscle energetics of rainbow trout (Oncorhynchus mykiss) and carp (Cyprinus carpio) andin post-exercise glycogenesis of rainbow trout and skipjack tuna (Katsuwonus pelamis).Sustained swimming. Fish are able to swim in a continuous “steady-state” fashionover a broad range of speeds. In a swim tunnel, salmonids can be challenged with waterspeeds that correspond to up to about 80 % of fatigue or critical (Uit) speed before non-steady state swimming (charcterised by burst-and-glide behaviour) is noticed. Steady-stateswimmming, even at very high intensity, can be endured indefinitely.Electromyographical studies show that red muscle is recruited during both slow andfast 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 whitemuscle recruitment occurs only at high, non-sustainable swim speeds. In good aerobicperformers (eg., salmonids and tunas) white muscle activity is evident at submaximal swimspeeds (Johnston and Moon 1981b; Brill and Dizon 1979). Observations that argue againstwhite fiber involvement at low swim speeds are that only red muscle showed significanthypertrophy after endurance training in salmonids (Johnston and Moon 1980b) and thatEMG’s recorded from deep white muscle show activity only at high sustainable swimspeeds in coalfish, Pollachius virens (Johnston and Moon 1980a). The carp is the onespecies where sequential recruitment of muscle types has been examined extensively and inthis species the division of labour between fiber masses seems fairly sharp. As in otherspecies, red fibers are recruited at low swim speeds, followed by intermediate (similar tofast twitch oxidative glycolytic, or FOG, fibers in mammals) fiber activity at highersustainable speeds and finally by white muscle recruitment (Johnston et a!. 1977) which, inthe most recent analyses, is always accompanied by unstable swimming and rapid fatigue3(Rome et al. 1984). In none of these studies was recruitment examined in relation tofatigue speed for individual fish, so it is not certain at what percentage of U white fibersbecame active. It is not known in either carp or salmonid species whether swimmingspecifically at 80 % of Uj is accompanied by white muscle recruitment, althoughexercise at this intensity is expected to push red muscle metabolic demands close tomaximal 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, itwould be generally expected that such activity would be supported largely by glycolysis.Endurance training induces increased activity of key fat utilizing enzymes (Moon andJohnston 1980a,b), suggestive of aerobic recruitment of white fibers at sustainable swimspeeds. However, the low aerobic capacity of this muscle (Moyes et a!. 1989, 1992b) andthe 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 introut, for example, the aerobic demands of high intensity red muscle activity (assumedsimilar to measurements in the bulirout, Myoxocephalus scorpius, of 5 - 8 mWg1(Altringham and Johnston 1990) or about 18 - 29 mol ATP min’g’) were imposed onthe 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 min1g,Moyes eta!. 1992b). The situation could be different in the skipjack tuna which have a whitemuscle 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 measurementof maximal enzyme activity (V) and mitoehondrial oxidation rates, both of whichprovide maximal capacities to utilize particular substrates. It is understood that these invitro estimates of fuel utilization may not resemble flux capacity in tissues of the wholeanimal (Moyes et a!. 1990; Wright and Albe 1990), but in comparative analyses such4observations have provided clues about which carbon sources may be used in vivo underconditions of maximal aerobic demand. In endotherms for instance, cardiac enzymeprofiles suggest that maximal aerobic metabolism is supported by fat utilization to a greaterextent in animals adapted for aerobic endurance than in more sedentary animals (Driedzicet al. 1987). Mammalian models of substrate preference based on in vivo metabolitekinetics also suggest a greater reliance on fat in more aerobic species during exercise(Weber 1992). Similar enzyme analyses of fish muscle from different species suggesteither that highly aerobic tissues rely more on fat (Moyes et a!. 1992a) or that there is noassociation between fat utilizing capacity and increasing tissue aerobic capacity (Sidell eta!. 1987). Furthermore, within fish species, the effects of endurance training include smallshifts in fuel availability (Hochachka et a!. 1961) and in the activity of some key muscleand heart enzymes (Farrell et a!. 1991; Johnston and Moon 1980), but modified substrateflux through a single pathway is usually not implied.In vivo fuel preference studies attempt to quantify substrate utilization with indirectcalorimetry, tracer infusions or aterial-venous differences in metabolite concentrationacross muscle groups. Where physiologically significant quantities of fuels exist, rates ofsubstrate supply, competition between pathways and the influence of regulatory factors willdetermine the pattern of fuel utilization. Unfortunately few studies address these situationsdirectly in fish. However, it is possible to gain initial insight into the metabolicpartitioning of different substrates during sustained exercise from whole-body sites andlevels of the main storage substrates. Representative salmonid tissue masses and metabolitelevels, per g organ mass and per kg body mass, are summarized in Table 1. Thesemetabolite levels represent the maximum mobilizable fuels in non-feeding salmonids. Inthe following review the importance of each of the major substrate-types for sustainedexercise will be considered in turn. The analyses focus on using maximal capacities forflux and estimates of maximal red muscle energy demand as guides to the potentialdependence on different fuels. As exercise becomes less intense or5Table 1. Tissue masses and fuel stores of a resting salmonid.ECF liver adipose white redg tissue/kg body mass 250 15 20 600 70glycogen (as glucosyl)(jLmol/g tissue) 0 100-200 40 40(mmol/kg body mass) 0 1.5-3.0 24 2.8glucose(jmol/g tissue) 2-5 2-5 2 2(mmol/kg body mass) 0.5-2.0 0.03-0.075 1.2 0.14lactate(jmol/g tissue) 7 3 3 5(mmol/kg body mass) 1.75 0.045 1.8 0.35triglyceride(JLmol/g tissue) 22 10-30 800 15 40(mmollkg body mass) 5.5 0. 15-0.45 16 9 3fatty acids(JLmol/g tissue) 3 3 2 2(mmol/kg body mass) 0.75 0.045 1.2 0.14protein (as amino acyl)(jmol/g tissue) 2000 1000 1000 500(mmollkg body mass) 500 15 600 35amino acids(jLmol/g tissue) 40 50 9(mmol/kg body mass) 10 30 0.63Data were compiled from various sources (French et al. 1983; Idler and Bitners 1959; Legereta!. 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 areestimates for total fuels and are not neccessarily available metabolically.6prolonged, the less abundant fuels and depots may become increasingly important. Ininstances where no direct studies have been performed, indirect studies using enzymeanalysis and isolated mitochondria can give further insight into the capacity and preferencefor 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 aconsiderable portion of the total caloric intake in omnivores, it would not be surprising ifdaily 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 andmay be elevated considerably, on a shorter-term basis, immediately after feeding (Kutty1978). With starvation, increased degradation of muscle-protein is most evident after bodyreserves of fat have been depleted (Black and Love 1986). Similarly, amino acidutilization seems important in migratory salmonids late in the run when other fuel storeshave been nearly exhausted (Mommsen et al. 1980). Taken together, these observationssuggest that in normal (fed, non-migratory) fish the availability of amino acids for routineoxidative metabolism is influenced by post-feeding absorptive mechanisms more so than bythe actual turnover of body-protein.The effects of activity metabolism on protein or amino acid dynamics in fish havenot been examined extensively. Continuous low level swimming increases protein turnoverand degradation in feeding rainbow trout (Houlihan and Laurent 1987), but the proportionof amino acids utilized for swimming energetics has not been determined directly.Increased aspartate amino-transferase activity in endurance trained red muscle of brooktrout (Moon and Johnston 1980b) may be more of a response to enhancement of the levelof protein synthesis that is retained as muscle growth (Houlihan and Laurent 1987), ratherthan catabolic use of amino acid. In salmon starved for 22 days, an indirect assessment of7whole-body amino acid utilization (ammonia excretion) indicated that, despite a probableincrease in protein degradation, daily increases in routine metabolism were seemingly notsupported by amino acids as long as reserves of fat and carbohydrate were available (Brettand Zala 1975; Kutty 1978). Ammonia excretion largely reflects amino acidtransdeamination in the liver (Van Waarde 1983) and will therefore over-estimate rates ofamino acid oxidation by the extent that the resulting carbon skeletons are used ingluconeogenesis, which increases during activity and starvation (Suarez and Mommsen1987). In any case, it seems likely that the higher aerobic demands of muscle resultingfrom a rest to exercise transition, in both normal and food-deprived fish, would rely evenmore on non-protein related energy reserves since the tissue pools of free amino acids aresmall and are not necessarily available for oxidation (Table 1). In addition, the oxygencost of ATP production from fat and carbohydrate oxidation is about 30% lower than thatof amino acids (Ferrannini 1988). In support of low rates of amino acid oxidation, it hasbeen noted that oxidation of neutral amino acids (serine, glycine, alanine) does not occur atdetectable rates in mitochondria isolated from carp red and white muscle (Moyes et al.1989). Furthermore, one study of in vivo substrate oxidation rates indicated thatglutamate, alanine and leucine, the latter classed as a branch-chained amino acid which ispossibly oxidized at elevated rates in exercising skeletal muscle (Kasperek et al. 1985,1987), seemed to contribute very little to whole-body oxidative metabolism of rainbowtrout swimming at 80% of Ucrit (van den Thillart 1986).Exercising fish may be able to draw on dietary amino acids after feeding asoxidative substrates for swimming. However, confirmation of such use of amino acidsawaits more detailed examination of post-feeding fluxes and oxidation rates. In moststudies of swimming metabolism the researcher avoids post-feeding energetics by design inorder to minimize inter-individual variability in basal oxygen consumption. Directcirculatory infusions of amino acid mixtures (see Tappy et a!. 1992) could be a moreuseful experimental approach for looking at the effects of amino acid availability on fuel8selection in fish. In this way it may be possible to mimic post-feeding levels of plasmaamino acids while at the same time eliminate potential influences of gut absorptive rate onwhole-body metabolism.Utilization of Fat Fuels Currently, the major gap in our understanding of fuelutilization during steady-state exercise concerns the relative importance of lipid fuels. Intotal, 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 fattyacid (FFA) to fuel about 100 h of maximal aerobic red muscle contraction in a 1 kg animal(assuming maximal power output of 5 - 8 mWg1 for red muscle (Altringham andJohnston 1990) which converts to about 0.21 - 0.34 mmol 02 min1g,about 23 mol02 utilized per mol fatty acid oxidized). However, there are few empirical measurementsthat help to identify which of major depots are mobilized in vivo during exercise. Suchstudies 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 ourawareness of the importance of circulating FFA, extramuscular depot fats and of thedynamics of skeletal muscle intracellular and pericellular TAG stores. Presently, anyconclusions regarding the relative importance of the various lipid stores, and lipid fuels ingeneral, 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 andBitners 1959; Jezierska et a!. 1982; Black and Love 1986; Jobling 1980) or on in vitroenzyme measurements and mitochondrial capacities for fatty acid utilization. Such studiessupport a major role for lipids in steady-state exercise in fish.Although hepatic and skeletal muscle TAG contents are similar (per g tissue), thesmaller 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 fatdeposits are the richest store of lipid per g tissue, but skeletal muscle, by virtue of its mass9possesses similar TAG stores per kg body mass (Table 1). Each of these stores might bedrawn upon during aerobic exercise. The degree to which the white muscle and visceralfat 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 acidscontribute minimally to the total oxidative fuel required by trout swimming at 85% ofUj (Van den Thillart 1986). Possible delays in the equilibration of labeled circulatoryprecursor 14C-palmitate) with the intracellular TAG pool at the site of utilization mayresult in underestimates of circulatory FFA oxidation (Heiling et a!. 1991), but it is notknown if this is a problem in fish at different levels of activity. Among endothermicspecies, the likelihood that intramuscular lipid supports aerobic energy demands, at least tosome extent, is supported by the observations of increased stores in aerobically trainedmuscle (Hoppeler and Billeter 1991) and by the proximity and presumed functionalassociation between mitochondria and intracellular lipid droplets in oxidative fibers(Hoppeler and Billeter 1991; Grunyer and George 1969). The possibility thatintramuscular TAG serves simultaneously as a sink for circulatory fuels and a source ofmitochondrial substrate needs to be evaluated more closely in fish.As with most vertebrate oxidative muscles, red muscle from teleost demonstrates ahigh 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 fattyacids, comparisons of mitochondrial oxidation rates and carnitine palmitoyl transferase(CPT) activities between species and tissues are more illuminating. As in aerobic trainingof 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 withaerobic capacity of the tissue (Moyes et at. 1992a). This change is due to both an increasein quantity of mitochondria and CPT activity per mg of mitochondrial protein. Thus, fattyacids 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,10endurance training has been shown to increase activities of fat utilizing enzymes (Moonand Johnston 1980b), also suggesting enhancement of FFA oxidation in response toimproved aerobic capacity. These observations agree fundamentally with conclusions fromcomparisons of fuel demands in endotherms and suggest that adaptation for increasedaerobic 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 toactive fish red muscle - (i) mobilization of intramuscular glycogen reserves, (ii)translocation of lactate from white muscle to red via the circulation (iii) circulatory glucosederived from hepatic gluconeogenesis/glycogenolysis.Intramuscular glycogen is not expected to be the primary fuel for steady stateexercise since total red muscle glycogen is a fairly small depot (Table 1). If usedexclusively, red muscle glycogen would support sustained maximal aerobic swimming forless than 1 hr (again, estimated assuming red muscle power output of 5 - 8 mWg1(Altringham and Johnston 1990) which converts to an oxygen demand of 0.21 - 0.34 mmol02 min1g). However, red muscle glycogen content in carp species appears to be 2 - 8fold higher than that of white muscle (Johnston and Goldspink 1973a; Johnston 1977;Johnston et al. 1977) providing some evidence for a possibly greater reliance onintramuscular carbohydrate for swimming metabolism than is expected in salmonids.Mobilization of red muscle glycogen has been shown to increase in relation to swim speedin carp during a graded swimming protocol (Johnston and Goldspink 1973b). In otherspecies, the importance of glycogen is implied indirectly in endurance training trials whichresult 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 similarchanges in phosphofructokinase (PFK) activity are observed. In brook trout, endurancetraining had no effect on red muscle glycogen content (a slight decrease perhaps) and no11effect on PFK activity (Johnston and Moon 1980b). These changes in brook trout occurredconcomitantly with increased B-hydroxyacyl dehydrogenase activity, consistent with thepossibility of enhanced fat dependence in salmonids with higher aerobic capacities. In allspecies, glycogen may be a minor oxidative fuel for extended aerobic exercise, although itmay augment fuel supply, particularly at higher intensities and in the early phases of a restto work transition (Spiret et a!. 1990). At lower exercise intensities red muscle glycogencould theoetically increase in importance, if only because its depletion would be extendedover a longer period. Glycogen mobilization should be examined more closely at differntexercise intensities and in connection with relative fluxes of competing substrates.Another route for the delivery of exogenous carbohydrate fuel to working oxidativemuscle may be via the steady-state release of lactate from white glycolytic muscle (Weber1991). In mammals, the fate of such lactate is predominantly oxidation resulting from theshuttling of lactate to oxidative tissues via the circulation (Brooks 1987). It is intriguing toconsider that white muscle in fish, with its large glycogen stores, could supply lactate tored muscle in the same manner. The total white muscle mass represents a glycogen storethat is about 20-fold greater than hepatic stores (Table 1) and could support hours ofintense steady-state swimming (assuming that the lactate released is delivered exclusively tored 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 02molmin1). If such a pathway operated, lactate would have to be produced by whitemuscle at rates which would be reflected in plasma lactate turnover, assuming aconventional circulatory transfer to red muscle.It was pointed out earlier that recruitment of white muscle may occur at high steadystate swimming speeds, but there is no evidence that metabolism in white muscle isprimarily aerobic under these conditions. Hence, studies in which lactate dynamics havebeen examined in swimming rainbow trout (Wokoma and Johnston 1981; van den Thillart121986; 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 lactatereleased into the circulation serves as a potential oxidative fuel in more aerobic tissues (redmuscle and heart).The problem with relying on circulatory lactate as a red muscle fuel seems to bethat turnover cannot match estimates of oxidative demand in red muscle (0.21 - 0.34 mmol02min1g). Lactate turnover would have to increase by about 10-fold to cover themaximal red muscle metabolic rate. Empirical studies indicate that, on average, the lactateturnover rate during sustained exercise at 80% Ucrit doubles over resting fish (to about 10molmin1kgbody mass), similar to that in a salmonid recovering from burst exercise(Milligan and McDonald 1988), but this change does not occur consistently in individualfish (Weber 1992). Thus, it would seem that lactate remains of minor importance as anoxidative fuel during endurance swimming (Weber 1992), although it may be moreimportant at higher levels of Ucrit or in smaller oxidative tissue masses like heart.Although elevated rates of lactate oxidation are predicted in swimming trout (vanden 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, theplasma compartment and is not inconsistent with the limited whole-body dependence onlactate as a fuel. As in mammals, oxidation may in fact be the major fate of lactate sincehepatic gluconeogenesis from circulatory lactate is minimal in most instances in a varietyof 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 conthbuterelatively little to the absolute change in metabolic rate with the onset of exercise could stillutilize lactate as an oxidizable fuel. Trout myocardium for example, increases its oxygenconsumption several-fold at high work intensities, but could rely heavily on circulatorylactate turnover because of its small relative mass and fractional contribution to whole-body13oxygen 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 indirectlyfrom activities of hexokinase that generally range from 0.5 to 1.5 mol glucose/gtissue/mm or total activity of 35 - 100 molmin1in a 1 kg fish (Crabtree and Newsholme1972; Johnston 1977). It can be calculated (again, assuming 0.21 - 0.34 mol 02 min1kg and assuming about 70 g red muscle per kg body mass) that similar rates of glucoseoxidation are required for maximal red muscle function (35- 60 molmin’kg1bodymass). As an estimate of fuel use, hexokinase measurements suggest that glucose fluxalone could account for the maximal rate of oxygen consumption of fish red muscle.Crabtree and Newsholme (1972) have similarly estimated that the glucose utilizing capacityof trout red muscle could meet the aerobic fuel demands. Can the delivery of circulatoryglucose keep up with the fuel demand during exercise?Glucose release from the liver of fish results from net gluconeogenic flux andglycogenolysis. Both processes are regulated by the action of various hormones andfunctional peptides (Mommsen et a!. 1987; Petersen et a!. 1987; Suarez and Mommsen1987; Mommsen et a!. 1988; Wright et a!. 1989; Reid et a!. 1992), but it is not known ifone pathway dominates hepatic glucose production in vivo. The responsiveness ofglycogen 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 changeduring submaximal exercise (Ristori and Laurent 1985; Butler et al. 1986), suggests thatglucose derived from hepatic glycogen may not be different from resting conditions. Inany case, liver glycogen levels alone (Table 1) could not support extended periods ofmaximal aerobic swimming. Regardless of the pathway of hepatic glucose production, therate of production and release to plasma in vivo is probably within the range of 1 - 10mol/min/kg, estimating from plasma turnover in teleosts under different experimental14conditions (Garin et al. 1987). Unfortunately, reliable determinations of salmonid glucoseutilization during exercise are not available. Even the upper end of the range of availabledata, 10 molminkg measured in hypoxic trout (Dunn and Hochachka 1987), wouldrepresent an imbalance between plasma glucose supply and red muscle glucose demandduring exercise. It is of course expected that glucose in transit though the plasmacompartment is distributed to more tissues than just red muscle, indicating that glucosedelivery would have to exceed 10 molminkg to match the present estimate ofoxidative demand for substrate during high intensity aerobic exercise. There are noestimates of glucose turnover in exercising fish, although estimates of glucose oxidation dosuggest limited dependence on this substrate in exercising rainbow trout. The capacity ofred muscle to use glucose (estimated from hexokinase activity) may mean that red muscleuptake can be expanded, perhaps in periods of hypoxia or during exercise in the absence ofother fuel-types. As is the case for lactate, possibly low glucose turnover rates duringexercise could, nevertheless, be significant to smaller tissue masses.Since plasma glucose is poorly regulated in fish (Palmer and Ryman 1972; Hiltonand Atkinson 1982; Mommsen and Plisetskaya 1991), in vivo uptake of glucose in fishtissues might be expected to correlate with parameters that affect glucose availability (eg.,plasma concentration or changes in regional blood flow). Although an initial estimate ofglucose utilization in trout red muscle indicates an imbalance between glucose delivery andoxidative demand, the quantitative importance of glucose specifically in red muscle isunknown. It also is not known to what extent the estimations made for trout can begeneralized across species which vary in aerobic swimming capabilities. In addition,glucose could be particularly important in the relatively small heart tissue which is poisedfor either glucose or FFA utilization in a number of fishes (Sidell et a!. 1987). Moreinvestigation of plasma glucose turnover and uptake during exercise, starvation andhypoxia may reveal tissues and conditions in which glucose is quantitatively important in15vivo and comparative studies will help identify differences in glucose dependency acrossspecies.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 hydrolysisfor active energy provision across species (Barrett and Connor 1964; Driedzic andHochachka 1975; Batty and Wardle 1979; Milligan and Wood 1986b; Schulte et a!. 1992).The white muscle mass is recruited during maximal swimming episodes which, ifprolonged, lead to exhaustion and dramatic shifts in intramuscular metabolite levels.Although the changes observed in muscle metabolites are fundamentally similar to thosethat 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 ofmetabolite restoration (Mommsen and Hochachka 1988; Schulte et al. 1992) and forexamining 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 ratesof recovery, yet an entirely general means of restoring carbohydrate status to pre-exerciselevels involving the intramuscular reconversion of lactate to glycogen (Batty and Wardle1979; Milligan and Wood 1986b; Schulte et a!. 1992). It follows that glycogen recoveryshould occur essentially independently of circulatory glucose disposal, but directquantification of glucose phosphorylation rate in recovering white muscle, and red muscle,has not been examined. Comparisons of carbohydrate recovery patterns in skeletal musclesof different species is of interest since more aerobically active species tend to display highpost-exercise lactate turnover rates, suggesting that lactate is used oxidatively in specieswith superior aerobic capacities.16White muscle recruitment and exercise metabolism. Recruitment order in swimmingfish is such that white muscle becomes active when the velocity of red muscle exceeds itscapacity to produce mechanical power (Rome et a!. 1988). The estimated maximal poweroutput of fish white muscle exceeds that of red muscle by 5 - 7 fold (Altringham andJohnston 1990). Unlike the situation in high speed sustained swimming where someoverlap in the recruitment of fiber masses may occur, maximal swimming is dominated bythe large homogenous white muscle mass. Although red muscle continues to shortenduring 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 thefish. The standard qualitative end-point used in different exercise protocols is fatigue,characterised by the inability to burst swim - although low-speed cruising capacitycontinues (Wood 1991). The time-to-fatigue at a given prolonged swim speed issometimes reported (Beamish 1978), but the most distinctive quantitative correlates at thetime of fatigue relate to the biochemistry of white muscle. White muscle fuel stores arenearly depleted at the time of fatigue, with glycogen, phosphocreatine (PCr) and ATPlevels often expended to less than 30 % of pre-exercise concentrations (Milligan and Wood1986b; Dobson and Hochachka 1987; Mommsen and Hochachka 1988; Schulte et at.1992). Depletion of these metabolites can be accounted for at exhaustion by intracellularaccumulation of lactate, creatine + inorganic phosphate (Pt) and inosine monophosphate(IMP) respectively. These dramatic changes arise at high power output because the highATP demand cannot be balanced by the low capacity of mitochondria to supply ATPaerobically.The intramuscular retention of certain metabolites also allows estimates of in vivofluxes in active white muscle. For example, lactate accumulation is estimated at about 2molnn1gin burst swimming trout (Moyes et a!. 1993). At the end of exercise thelactate accumulated is usually twice the molar amount of glycogen that has been depleted,17suggesting essentially exclusive dependence on this intramuscular carbon source. Liver hasa large glycogen contentlg tissue, but given that total hepatic glycogen represents less than10% 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 duringburst exercise. The involvement of plasma glucose is likely to be minor, not just becauseits 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 magnitudeto account for the lactate accumulation rates expected in trout white muscle. Actual plasmaglucose kinetics are difficult to measure in burst swimming fish as a result of the shortperiod needed to induce fatigue (10 - 20 mm) and because of the likelihood that burstexercise protocols lead to unstable plasma glucose concentrations (Barton and Schreck1987).With glycogen levels in trout white muscle typically ranging 20 - 40 mo1 glucosylunits g1, white muscle is expected to become quickly substrated-limited during maximalATP 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 P1 from PCr breakdown is probablystimulatory to glycogenolysis, while increased proton production from glycolysis can bebuffered to some degree by PCr breakdown - reasons to expect that these two processesoccur interactively in active muscle (Hoppeler and Billeter 1991), as indicated previouslyin anoxic fish muscle (van Waarde et a!. 1990). Depletion of ATP content itself occursrapidly when glycolysis and PCr hydrolysis no longer support energy metabolism. Atexhaustion, IMP levels mirror ATP depletion (Mommsen and Hochachka 1988; Schulte eta!. 1992), as in mammalian muscle (Meyer and Terjung 1979; Soderlund and Hultman1991), but the extent of ATP reduction (up to 70 - 90 %) is considerably higher than thatseen in exercised mammalian muscle.18It is not known whether these metabolite changes have any direct causal associationwith fatigue in fish. Exhaustion is sometimes reported with white muscle glycogen levelsstill in excess of 10 mol glucosyl units g1 (Schulte et at. 1992), suggesting that, as inmammals, 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 theinteraction of intramuscular recovery pathways. Closer examination of metabolite levelsand fluxes during exercise and recovery is necessary before a quantitative picture of theseinteractions in fatigue and recovery processes will emerge.Carbohydrate recovery in fish muscle. A great deal of research has focused on thegeneration and clearance of lactate in fish muscle. Its metabolism is distinguished fromthat of other vertebrates by the intramuscular retention of lactate produced during exerciseand by the relatively slow rate of clearance (8 - 24 h) from muscle. Some lactate isreleased to the circulation during recovery and the peak level varies among species, withhigher 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 smallrelative to the total white muscle mass and peak concentrations of plasma lactate, ranging25- 50 % of post-exercise lactate concentrations in white muscle (about 10 - 40 mmolL1), represent a small portion of the post-exercise white muscle lactate load. Concentrationsof lactate are only indirect measures of flux and it is evident that post-exercise turnoverrates, measured with 14C-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 parwith rates seen in mammals (Weber 1986). Lactate turnover rates, like post-exerciselactate concentrations, are highest in the more active species. However, turnover ratesmay be somewhat misleading in recovery situations in fish since there is evidence foractive re-uptake of lactate from the extracellular fluid into the lactate-producing white19muscle mass (Turner and Wood 1983). Turnover will not reflect simple movement oflactate 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, estimatedrecovery time would be approximately twice the observed rates if muscle clearance oflactate 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 isin accord with the repeated contention that fish white muscle lactate is metabolized largelyin 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 largeextent 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, ashas been suggested in reptiles (Gleeson and Dalessio 1990).The fate of lactate in fish white muscle has not been traced quantitatively. Anumber of indirect observations point to primarily intramuscular reconversion of lactate toglycogen (glyconeogenesis). Oxidation is possible (Scarabello et a!. 1991), consideringthat a carbon source is needed to fuel glyconeogenesis, but oxidation of only 10 - 20 % ofthe post-exercise lactate load would provide sufficient ATP for recovery. The possibilitythat fat is used for recovery processes, sparing lactate for glyconeogenesis, is suggestedfrom mammalian studies (Bahr et a!. 1991; Bangsbo et a!. 1991) and from the in vitro andin vivo regulatory features of trout white muscle pyruvate dehydrogenase, which displaysonly 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, inconjunction with expected low rates of glucose utilization, in fish white muscle stronglysuggests in situ conservation of carbohydrate carbon during exercise and recovery.However, quantitative estimates of glycogenic (glycogen formed from glucose) and20glyconeogenic fluxes are not available. A better understanding of the relative importanceof glucose to the overall process of glycogen repletion in trout white muscle can help placecarbohydrate recovery profiles into a more quantitative framework. Species like tunawhich display ‘mammal-like’ lactate fluxes, high aerobic capacities and the need to swimcontinuously may show increased capacity to utilize lactate oxidatively and therefore relyto a greater extent on glycogenesis for recovery of white muscle glycogen stores.In vivo measurement of glucose utilization. Most of our knowledge of carbohydrateutilization for the swimming energetics of fish has come from whole body depletion of fuelstores in migrating species or from in vitro measurements of maximal flux capacity (HKactivity or mitochondrial oxidation rates). In vivo observations of glucose kinetics in fishare seldom performed, even though it is often emphasised that such observations would beparticularly useful in helping to sort out species differences in the regulation and interactionof inter-organ pathways of metabolite exchange in fishes (see Mommsen and Plisetskaya1991). Observations of the rate of transit of a metabolite through the circulation are moreinformative than monitoring shifts in plasma concentration since physiological changes orexperimental manipulations which may influence turnover rate are not necessarilyaccompanied by changes in metabolite concentration. Tracer determined fluxes offer amore quantitative view of the capacity for substrate involvement in whole-animalmetabolism during exercise, compared to in vitro enzyme measurements and mitochondrialoxidation rates, and provide a means to investigate potential regulatory influences onmetabolite availability and disposal in tissues.With determinations of glucose turnover, it is understood that the appearance ofglucose 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 itdifficult to assess muscle glucose utilization in specifically the recruited muscle mass21during sustained exercise. In steady state conditions, one way to determine glucosephosphorylation (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 nonhepatic tissues (Sokoloff et al. 1977). Irreversible 2-DG phosphorylation can provideestimates of metabolic rate in tissues that are highly dependent on circulatory glucose forenergy 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 vivotissue 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 ismostly converted to glycogen (Kusunoki et a!. 1993), indicating that 2-DG uptake shouldprovide a reasonable quantitative estimate of in vivo glycogenic flux during recovery frommaximal exercise. When 2-DG uptake measurements are made in concert withdeterminations of glucose turnover, it is possible to quantify the relative contribution ofspecific tissues to total glucose disposal.In chapter 2, radiolabeled tracers of in vivo glucose metabolism were used toexamine 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 preferredin the better aerobic swimmers (Moyes et a!. 1992a), suggesting indirectly that glucosemay be important in poorer aerobic performers. This pattern of substrate dependencywould be consistent with a more general hypothesis suggested for mammals in whichgreater use of fat stores is predicted in highly aerobically adapted animals (see Weber1992). 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 pooreraerobic performer. Within this comparative framework, the slower swimming carp washypothesized to show greater dependence on glucose for energy provision during sustainedexercise than the faster swimming trout.22Studies of exercise recovery have emphasized white muscle metabolite recoveryprofiles and their interactions in the rainbow trout. Carbohydrate recovery patternsgenerally suggest that lactate is a major substrate for glycogen replenishment in whitemuscle, but the implication that plasma glucose is of minimal importance is largelyindirect. Moderate post-exercise hyperglycemia, sometimes seen in recovering trout(Milligan and Wood 1986b, Mommsen et al. 1988; Pagnotta and Milligan 1991) may berelevant in this regard. In chapter 3, glucose utilization in red and white muscle ofrecovering trout was examined to test the idea that glycogenesis is a negligible componentof white muscle glycogen resynthesis whereas red muscle, because of its small mass andhigher perfusion rate, may be more readily able to utilize circulatory glucose for post-exercise glycogen restoration. For comparison, carbohydrate recovery status was alsoexamined in the white muscle of fast swimming skipjack tuna (Katsuwonus pelamis) to testwhether in situ glyconeogenesis was less important in a species that is known to exhibithigh post-exercise turnover rates of lactate, the prime glyconeogenic substrate in whitemuscle.PUBLICATION STATUS: Part of the preceding review, the discussion of fuels forsustained swimming, is my contribution to a book chapter that has been accepted forpublication: 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.23CHAPTER 2. GLUCOSE UTILIZATION DURING SUSTAINED SWIMMING INRAINBOW TROUT (Oncorhynchus myldss) AND CARP (Cyprinus carplo)In vivo utilization of glucose by heart and locomotory muscles of exercising rainbowtrout.INTRODUCTIONSkeletal muscle is the principal peripheral site of circulatory glucose disposal inmammals during exercise and, along with fatty acids and lactate, its uptake contributessubstantially to energy provision for endurance activity (Weber 1988). In contrast, less isknown about the kinetics of glucose and its oxidative role in fish tissues. Plasma turnoverand oxidation in most teleosts is slow compared to mammals (van den Thillart 1986; Garinet a!. 1987; Machado et a!. 1989) and long-term perturbations like starvation/migrationsuggest 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 forestimating 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 aerobiccontractions in vitro (Crabtree and Newsholme 1972; Lanctin et a!. 1980; Driedzic andHart 1984). However, verification of the contribution of glucose to tissue oxidation in vivois more difficult because of the complexities of substrate storage, mobilization, transportand pathway interaction processes in the whole animal (see Weber 1988; 1992). Inaddition, exercise intensity is an important consideration when evaluating muscle fuel usein vivo since fuel types or stores utilized at high aerobic intensites may be different fromthat at low intensity. In salmonids, swimming speeds of 80 - 85 % of fatigue or criticalspeed (Ujt) can be maintained indefinitely and are expected to place close to maximum24steady-state aerobic demands on oxidative muscle. In the present study the glucoseanalogue 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 intenseaerobic exercise. Direct determinations of glucose utilization in specific tissues give us theoppotunity to test the degree to which the circulatory delivery of glucose supportsestimated maximal rates of tissue oxidative metabolism in vivo.MATERIALS AN]) METHODSAnimals. Rainbow trout (Onchornhynchus mykiss) of both sexes (300 - 1200 g) werepurchased from local suppliers. Smaller fish (< 600 g) were used for in vitro isolatedheart studies, conducted at the University of British Columbia. In vivo glucose utilizationwas determined in larger fish at the Bamfield Marine Station, Bamfield B.C. All fish wereheld in freshwater (15 - 17 °C) and were fed to satiation twice weekly.2-DG and the lumped constant. Glucose utilization was calculated from the tissuecontent (disintegrations per minute, dpm) of 14C-2-DG phosphate (2-DGP) and areas underplasma 14C-2-DG washout curves according to the equation of Ferré et a!. (1985),Tissue dpm 2-DGPUTILIZATION =____________________________(jhmolmin1)LC J (dpm 2-DG/mol glucose)dtUtilization was corrected for tissue mass and is presented as nmol per gram of muscle perminute. The term LC is a dimensionless ‘lumped’ constant which is necessary to correctfor differences between glucose and 2-DG uptake and phosphorylation rates in vivo (seeSokoloff 1983). Simultaneous determinations of glucose and 2-DG uptake weredetermined in vitro from rates of 3H20 (Ashcroft et a!. 1972) and 14C-2-DGP formed25from3H-5-glucose and 14C-2-DG, respectively. The LC was calculated using the equationdefined by Ferré et al. (1985),tissue 2-DGPI2-DG in mediumLC=glucose utilized/glucose in mediumTissue 2-DGP and 2-DG in the incubation or perfusion medium are presented as dpm 14C,while the glucose in the medium and total glucose utilized during the tissue incubation orheart perfusion period are in units of nmol.Estimation of LC in muscle slices. Fish were netted from the holding tank and killedquickly with a sharp blow on the head. A rectangular incision that encompassed the lateralline was made in the musculature between the dorsal and adipose fins. The skin waspeeled back to expose the muscle and the area was flooded with ice cold saline (containingin mM: 127 NaCl, 4.9 KC1, 1.0 CaCl2, 3.7 NaHCO3, 1.2 MgSO4, 2.9 NaH2PO4,1.2KH2PO4, 11.5 Na2HPO4;pH = 7.4). A strip of skeletal muscle was dissected free bygently manouvering a razor blade through the exposed muscle along the long axis of thefish. The strips were trimmed of white muscle and the red muscle slices (0.5 - 1.0 cm2, 1- 2 mm thick) were transferred to a vessel of fresh saline (100 ml, bubbled with 99 % 02:1 % C02) containing 5mM glucose. The vessel was placed in a shaking water bath (15 ±1 °C) and the tissues were preincubated for 20 mm. Ventricular strips were similarlyprepared from hearts that were immersed in saline and cut in half from the base to theapex. Strips were made by slicing parallel to the freshly exposed surface of spongymyocardium. Two to four slices of each tissue were prepared from every fish.Following the preincubation, individual slices were transferred to scintillation vialscontaining 2.5 ml of fresh saline which contained 5-3H-glucose (0.3 PCi) and 14C-2-deoxyglucose (0.15 MCi) (Amersham) and 5 mM unlabelled glucose. The incubation vialswere sealed with rubber plugs through which plastic access tubes were affixed for gentle26gas infusion. The vials were positioned in the shaker and incubations were run for nolonger than 45 mm. Preliminary trials with3H-5-glucose indicated that glucose utilization(3H20 production) remained constant over this period. Two to three blanks containingonly saline and radioisotope were performed with every run to assess background levels of3H20. Specific activity (SA) of5-3H-glucose and dpm 14C-2-DG in the medium weredetermined by counting 200 l aliquots mixed with 10 ml ACS II (Amersham) on a LKBRackbeta scintillation counter.Samples of saline (200 JLl) retrieved from the vials following incubation were placedon columns of Dowex-1-borate (1 ml bed volume) to separate glucose from 3H20produced (Hammerstedt 1973). The samples were allowed to percolate into the columnbed by gravity flow before washing the resin with water (5 x lml washes). The washeswere collected separately into 20 ml glass scintillation vials assayed for dpm 3H. Totaldpm 3H for each sample was corrected for background dpm and adjusted to the totalincubation volume to calculate 3H20accumulated. Levels of14C-2-DGP were determinedin tissues that were homogenized (3 x 30 sec bursts with an Ultra-turrax tissuehomogenizer) 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 wereneutralized with 3 M K2C03 in 0.5 M triethanolamine. An aliquot of neutralizedsupematant was counted to determine total 14C (2-DG + 2-DGP) present. PhosphorylatedDG was removed from an equal volume by either precipitation with 0.3 N Ba(OH)2 and0.3 N Zn(S04)(sample:reagents, 1:1.5:1.5) or by anion exchange on a column of DEAESephadex 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 tissueextracts to determine total tissue 1’C (2-DG + 2-DGP). Equal volumes of extract werethen placed on columns of DEAE-Sephadex A-125 (2 ml bed volume) that had been preequilibrated with 20 mM imidazole chloride, pH 7.2. The sample was allowed topercholate into the resin, then the column was washed with 10 ml of water. Fractions27collected were mixed with 10 ml of aqueous scintillant, counted, corrected for backgroundand summed together to give the 14C-2-DG retrieved from the column. This valuesubtracted from total 14C radioactivity gave an estimate of tissue 14C-2-DGP. Preliminarytesting with known levels of14C-2-DG and 14C-2-DGP, the latter made from reacting 14C-2-DG/2-DG to completion with hexokinase, prepared in fish muscle extracts indicated thatthe anion-exchange column retained > 95 % of14C-2-DGP, with similar levels of 14C-2-DG recovery. Lumped constants were calculated from the amount of glucose utilized(calculated from 3H20 production) and 2-DGP produced with the equation presentedearlier.Estimation of LC in isolated trout hearts. The effect of glucose utilization rate onthe LC was investigated using isolated perfused trout hearts. Isolation and cannulationprocedures were as outlined by Farrell et a!. (1989). Isolated hearts were suspended in asaline bath and air-equilibrated perfusate was introduced from water jacketed reservoirs (15°C). Output flow from the ventral aorta was monitored continuously as outlined byGraham and Farrell (1989) and oxygen consumption was calculated from the difference inoxygen content, measured with a Radiometer electrode and meter, of perfusate enteringand leaving the heart. Additionally, an inlet branch was placed at the level of the preloadpressure head for infusing sodium cyanide (NaCN). Electrodes attached to the input andoutput cannula allowed for the electrical control of heart rate.Changes in circulatory glucose utilization were invoked by manipulation of heartrate and input and output pressure heads to reduce or increase cardiac power output, and byinfusion of NaCN (1 mM) at low power output. Power output, in milliwatts/gramventricle mass (mWIg), was calculated according to Graham and Farrell (1989). Isotopeperfusions were initiated only after output flow recordings became steady under the desiredconditions with ‘unlabelled perfusate’. Low power perfusions with and without NaCN28were 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 passperfusions of 5-3H-glucose (0.1 ,iCimr) and 14C-2-DG (0.02 Ciml1). Followingperfusion, hearts were removed from the saline bath, blotted, and immersed in preweighed vials of 7 % PCA (2 ml). Tissues were homogenized, then neutralized andassayed for 14C-2-DGP. Perfusate collected at the afterload outlet was assayed for lactateaccording to Bergmeyer (1985) and 3H20 by vacuum distillation. For the latter, a flaskcontaining outflow perfusate (3 ml) was attached to a micro condenser with an 8 cm stillhead, then immersed in a water bath (60- 65 °C) where the perfusate was brought toboiling under constant vacuum (-600 mm Hg). All of the distillate was collected andaliquots (1 ml) were counted for 3H and 14C. The lack of detectable 14C abovebackground level indicated no contamination of raw perfusate in the distillate. Perfusiontimes and flow rates from the afterload outlet were monitored to calculate cardiac outputand to determine total 3H20 produced and total amounts of isotope and substrate to passeach heart to calculate LC’s.Glucose utilization in swimming trout. Rainbow trout were anesthetized (0.5 g MS2221k buffered with 1 g.NaHCO31’) and placed in a standard fish operating sling.Oxygenated water containing a light level of buffered MS-222 (0.1 gt1) was circulatedover the gills, and dorsal aortic cannulae (PE-50, Clay Adams) were implanted. Fishrecovered from anesthesia (12 - 18 h) in dark plastic tubes (15 cm diameter) which weresubmerged in a flow through water (15 - 17 °C) resevoir at the downstream end of a Bretttype swim tunnel. Once recovered, each fish was handled separately. A fish was guidedto the entrance of the tunnel and each swam at its own accord from the recovery tube intothe swim space. Water speed was adjusted to about 0.5 body-lengths(bl) sec4 (15-20cmsec1)and the fish was left for approximately 10 h before the experiment was started.29Fish used to determine resting rates of glucose utilization recovered from anesthesia inblack perspex boxes, supplied with fresh flow-through water (15 °C).A high sustainable swimming speed was found by increasing water velocity every15 mm by 20 cmsect increments until the fish began to demonstrate ‘burst and glide’swimming. Water velocity was then reduced until the fish held its position against the flowof water. This velocity was maintained for 1 h prior to isotope injection.Bolus injections consisted of 14C-2-DG (5 MCi) that was pre-dried under nitrogenand reconstituted in 250 1.d of Cortland saline (Wolf 1963). An aliquot (10 ILl) was takento determine the dpm of 14C injected and the remaining volume was injected into the dorsalaorta via the cannula. The cannula was flushed with two volumes of saline. Blood (1501d) was drawn through the same cannula at 1, 2, 3, 4, 5, 10, 20, 40, 60, 90 and 120 mmpost-injection. Neutralized PCA (1:1, 7 % PCA: sample) extracts were assayed for dpm14C in glucose and plasma glucose concentration (Sigma glucose assay kit). Plasmawashout kinetics were determined using a curve-stripping program (JANA; StatisticalConsultants, Lexington, KY) and areas under curves were determined (MATHCAD;Mathsoft Inc., Cambridge Ma.). When blood sampling was completed in each fish thespinal cord was severed just posterior to the head and tissues were removed and freeze-clamped between Wollenberger tongs that were pre-cooled on dry ice. A double bladedcleaver was used to rapidly cut a steak from the musculature immediately posterior to thedorsal fm. The heart was removed and the ventricle was quickly cleared of blood andfreeze-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 andglucose utilization rates was determined using unpaired t-tests or ANOVA with Tukey’smultiple comparison test.30RESULTSTissue LC’s and the effect of glycolytic rate. Table 2 lists LC’s for red muscle andheart ventricle from tissue incubations. The LC’s calculated were quite variable, althoughmost values were between 0.2 - 0.6. Box plots in Table 2 provide an indication of therange and clustering of data points around the median LC for each tissue. LC’s calculatedfor red muscle and heart were not significantly different.The isolated trout heart preparation provided a means for assessing whether changesin perfusate glucose uptake affected the 2-DG LC (Table 3). The use of perfusate glucoseranged from trace levels in the substrate free perfusions to 250 nmolg1min in the NaCNperfusions. However, there was no effect of utilization rate on the LC’s calculated fromthe various perfusions.Interestingly, power output of the NaCN hearts spanned a narrow subphysiologicalrange (0.04 - 0.17 mWg1), yet potent enhancement of perfusate glucose utilizationoccurred within this group in response to small increases in power output (Figure 1). Thevariability is likely the reason for the glucose utilization not being significantly differentfrom the high work rate hearts (Table 3). We did not attempt experiments to confirm thatpower output was in fact the principal factor affecting glucose utilization during NaCNperfusions. As presented in Figure 1, the regression of glucose utilization against cardiacpower output is y = 2003x + 49, r = 0.70. Individual LC’s for these hearts are alsodepicted, but there is no indication that the formation of 3H20 glucose and theaccumulation of 14C-2-DGP changed disproportionately with increased perfusate glucoseutilization.All in vivo red muscle and heart glucose utilization rates were calculated using theaverage LC determined from these in vitro experiments (LC = 0.40).Glucose utilization in swimming trout. Plasma profiles of dpm 14C-2-DGmo1glucose1 and of glucose concentration in trout are shown in Figure 2. Small fluctuationsTABLE2.Lumpedconstants(LC)calculatedfrominvitroincubationsoftroutredmuscleslicesandheartventricularstripswith14C-2-DGand5-3H-glucose.Total14C-2-DGin2.5ml ofsalineaveraged459800±57800dpmand421900±48300dpmfor theredmuscleandheart incubations, respectively.BoxplotsarepresentedforredmuscleandheartLC’swithdataoutside5thand95thpercentilesshown(.).REDMUSCLEHEART(n=15)(n=18)TISSUE2-DGP760±1601130±240(dpm)GLUCOSEUTILIZED48.1±9.776.2±13.8(nmol)LC0.43±0.060.38±0.05.IIIIIIIIIIII00. kilogrambodymass,whilepoweroutputwasnormalizedpergramventriclemass.Oxygenconsumptionandlactateproductionper gramofventriclemasswerealsodetermined.CARDIACPOWEROXYGENLACTAThGLUCOSEOUTPUTOUTPUTCONSUMPTIONPRODUCTIONUTILIZATIONL.C.PerfusateGlucose, 5mM(mlkgtmin1)(mWg1)(nmolg1min1)LowWork4.5±0.80.07±0.02212±3449±1329±60.41±0.03(N=4)HighWork24.8±2.01.69±0.091468±139118±28°121±22a0.41±0.06(N=4)Cyanide6.6±1.30.10±0.021012±129a,d253±57a,b0.40±0.03(N=6) GlucoseFreePrefusion(dpmg’mint)HighWork22.7±2.31.22±0.111296±178’182±59’15400±5000.39±0.04(N=3)Cyanide5.8±0.70.10±0.02791±72’43500±2400w’0.41±0.03(N=3)a.Significantlydifferentfromlowworkload(p<0.05)b.Notsignificantlydifferentfromglucoseperfusedhighworkload(p=0.17).c.Significantlydifferentfromglucosefree,highworkload(p<0.001).d.Significantlydifferentfromglucoseperfusedhighworkload(p<0.01).*Notdifferentfromoxygenandlactateinthecorrespondingglucoseperfusedhearts.33V0.45.400133Z 300 I”o 0.3200 O.32J0.45V..,I100 0.48•.•$0.0 0.5 1.0 1.5 2.0POWER OUTPUT (mW.j’)FIGURE 1. Glucose utilization in relation to power output innormoxic (circles) and NaCN treated (triangles) hearts. LC’scalculated for individual NaCN perfusions are also depicted.34in plasma glucose were typical of individual resting and exercising fish. Dynamic steady-state was assumed if during the sampling period the coefficient of variation (CV) for themean plasma glucose level was 15 %. The 14C activity washout curves were all fit toequations described by two exponential tenns.Table 4 lists data for individual resting and swimming trout. Relative swimmingspeed 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 cmsec1.P1a ma glucoseranged from 2 - 5 mM among both control and exercised fish. The in vivo rate of muscleglucose utilization in unexercised trout was highest in heart ventricle, being about 6-foldhigher than in red muscle. However, during sustained steady-state exercise, cardiacglucose utilization did not change significantly compared to resting fish. Conversly, redmuscle utilization increased from less than 1 nmolgmin at rest to about 21 nmolg1min, representing a 24-fold change in the use of circulatory glucose. White muscleutilization, estimated assuming that the LC =0.4 for this tissue, averaged less than 0.5nmolg’min1at rest and during exercise.DISCUSSIONThe 2-DG lumped constant. The LC as described by Sokoloff (1983) for normalbrain 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 ofphysiological conditions. Similarly, the LC calculated for rat skeletal muscle is influencedminimally 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 tissuetreatment it is still difficult to generalize, even within a species. The LC chosen ascharacteristic 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 that35100000Cd)o C)o100100000Cl)00‘10000CU1000FIGURE 2. Plasma washout of 14C-2-DG in resting (upper panel) andswimming (lower panel) rainbow trout.. Inset axes show profiles of plasmaglucose concentration during the blood sampling period.100001000E0 20 40 60 80 100 120Time‘—4I____ ____I o • _I_ I I0 20406080100120Time (mm)I I I I20 40 60 80 100 120TimeE10036attributes of individual muscle groups, such as fiber type distribution and glucosetransporter type or density, can affect LC determination. In addition, mammalian tissuesother than brain and skeletal muscle may be more sensitive to changes in experimentalconditions. Indeed, the LC for rabbit myocardium is on average 0.6, but extremes of flowrate and contraction frequency, albeit beyond the range of physiological relevance, wereshown 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 fewerambiguities when extrapolating LC’s to the whole animal if in vitro determinations aremade 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 LCdetermined with isolated heart preparation was generally more reproducible than with theslices. Despite the fact that slices were preincubated for 20 mm, the broad range of LC’smay in part reflect a varying degree of cellular damage since disruption of the plasmamembrane could have altered the importance of the transport component of the LC.Determinations of the LC with the more intact isolated heart preparation were alwaysbetween 0.3 - 0.6 (see Figure 1, for example). Nevertheless, the LC produced from thetwo protocols is similar and the indication is that an LC of 0.4 is a reasonable applicationto 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’sduring variable rates of perfusate glucose uptake. The high power output hearts used in thepresent study matched basal power output levels set normally in isolated and in situ trouthearts (1.0- 2.0 mWg’1,Farrell et a!. 1989; Milligan and Farrell 1991). Maximal workloads in our isolated preparations were not stable long enough to reliably coordinate steady-state cardiac parameters with isotope perfusions. For the same reason, the NaCN perfusedhearts had to be set at subphysiological work levels. However, the advantage of thisprotocol was that a broad range, of glucose utilization could be induced. The resultsindicate that with stimulation of glycolytic rate by anoxia or changes in power output the37rate of 2-DG uptake and phosphorylation should remain a constant proportion of glucoseutilization. Furthermore, both the aerobic and NaCN perfusions lacking unlabelled glucoseshows that even if cardiac function is dominated by endogenous fuel utilization, the LCcalculated from tracer uptake is unaffected.In vivo glucose utilization in red muscle. For rainbow trout of the size used in thisstudy, an average steady-state swimming speed of 1.7 body lengthssec1 corresponds toapproximately 80 % of U (Kiceniuk and Jones 1977). From rest to this high level ofsustained swimming, rates of 2-DGP accumulation indicate that glucose utilizationincreased by 28-fold in the lateral red muscle. What proportion of the in vivo energyproduction in red muscle is accounted for by glucose?The red muscle of trout constitutes about 5 - 10 % of the total muscle mass (Webb1971). A 1 kg trout, possessing up to 70 g red muscle, consumes about 25 mol 02kg1min at rest and increases this by about 7 fold when swimming aerobically at close to 80% of U (Kiceniuk and Jones 1977). Swimming at this intensity results mainly from thepower output of the active red muscles (reviewed by Jones 1982) and given theconcomitant redistribution of blood flow to these fibers (Randall and Daxboeck 1982), it islikely that the cost of swimming, above routine maintenance costs, becomes associatedlargely with the increased aerobic demand of this muscle mass. Assuming that glucose isfully oxidized in red muscle (6 nmol 02 per nmol glucose), it is then apparent that, givenan uptake rate of 21 nmol glucoseg1min,circulatory glucose is maximally 5 % of thetotal red muscle substrate oxidation in vivo.Other exercise-induced changes in whole-body metabolic costs are not accountedfor in the preceding calculation. It is an oversimplification to attribute 100 % of the costof swimming to red muscle and this first approximation of in vivo red muscle glucoseoxidation is probably an underestimate. However, osmoregulatory costs do not appear tochange with increased aerobic exercise, although this is probably speciesTABLE4.Invivoglucoseutilizationinredmuscleandheartofrestingandswimmingrainbowtrout.Aninvivo2-deoxyglucoselumpedconstantof 0.40wasusedfor calculationsoftissueglucoseuptake.VGLUCOSEUTILIZATIONBODYSWIMMINGPLASMAREDHEARTWHITEMASSSPEED[GLUCOSE]MUSCLEMUSCLE(g)(bl/sec)(mM)(nmolg1min)RESTING 19832.32±0.10----6.6326502.77±0.091.396.650.2038752.83±0.100.83----0.4046012.51±0.071.348.911.0459443.69±0.140.378.850.20611204.61±±±0.150.751.940.2397763.29±0.110.995.220.31Mean±SE803±620.87±0.155.31±1.040.36±0.10EXERCISED110661.682.11±0.1718.911.810.6028981.612.32±0.084.931.850.40310791.622.43±0.0730.936.310.96410961.672.35±0.1011.174.460.1257981.762.71±0.0948.4413.050.8368051.763.52±0.1412.96V0.900.2077401.662.78±0.1118.634.580.22Mean±SE926±571.68±0.0220.85±5.524.62±1.600.48±0.12significantlydifferentfromrestingcontrol, p<0.001.cz39dependent (Pérez-Pinzón and Lutz 1991). In addition, combined oxidation in other muscletypes, heart because of its small mass and white muscle because its peak aerobic demand isexpected at speeds < 80 % of Uj or in recovery from intense burst exercise (Moyes etal. 1992b; Scarabello et al. 1991), is probably minor compared to red muscle oxidativedemand during exercise. Nevertheless, EMG’s have been recorded from the white muscleof salmonids during sustained swimming (Johnston and Moon 1980) and given that theoxidative capacity of trout white muscle is relatively large compared to slower swimmingfish like carp (9.3 moPmin1gcitrate cynthase and about 8 mg mitochondrial proteinper 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 musclerecruitment could be a significant metabolic cost in swimming trout. Activity relatedchanges in costs of white muscle recruitment, ion regulation and ventilation are difficult toquantify in vivo, but it is apparent that if such costs diverted one-half of the whole-animaloxygen consumption at 80 % U then an estimate of glucose oxidation in red musclewould 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 amajor determinant of red muscle uptake of circulatory glucose is simply the redistributionof blood flow during exercise. Most of the cardiac output is delivered to the active redfibers in swimming trout (Randall and Daxboeck 1982) and it is evident that muscleglucose uptake correlates postively with bloodflow (Chaliss et a!. 1986). Although themetabolic demand of red muscle increases, the use of glucose might be limited by theavailability of plasma glucose or by potent glycolytic inhibition brought on by the use offatty acids in highly oxidative muscle (Kobayashi and Neely 1979). As a result, thepotential contribution of glucose to overall red muscle substrate oxidation would be small,as calculated, despite a several fold increase in uptake over resting rates.40It may be that lipid based substrates are utilized for sustained aerobic swimming introut. This possiblity is strengthened by the likelihood that carbohydrates from all in vivosources are a small component of red muscle oxidation. Blood-borne lactate is a minorfuel in trout during an endurance-type swim (Weber 1991) and red muscle glycogen,although possibly important during submaximal swimming in some species (Johnston andGoldspink 1973), is available in limited supply in trout (20 molg1 Parkhouse et al.1987) and cannot solely furnish fuel for intense aerobic swimming. The use of fat fuelshas not been quantified with in vivo methodologies, but the potential for reliance on fattyacids in teleosts is suggested at least indirectly from intramuscular storage and mobilizationcapacities, from triglyceride depletion studies in migrating/starving species and from invitro 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 notexpected at swimming speeds up to about 80 % of Uj (Webb 1971). As was discussedearlier, however, recruitment of this fiber mass cannot be ruled out, particularly since justone velocity increment above 80 % Uj induced burst and glide behavior in these trout.However, low glucose utilization in white muscle is not unexpected since circulatorysubstrates are probably of minimal importance in this tissue during exercise. A reductionin blood flow to the white fibers at 80 % U (Randall and Daxboeck 1982) is consistentwith the view that any recruitment of this muscle mass is fueled mostly by mobilization ofintracellular fuels. About 95 % of the blood flow to the total mosaic musculature ofrainbow trout at 80 % U could be recieved by the small proportion of ‘red’ fibersdispersed throughout the white muscle mass (Randall and Daxboeck 1982), suggesting thatwhite fiber recruitment may be fueled by intramuscular glycogenolysis. Glucose kineticsin the deeper red fibers might well behave as in the lateral red muscle during exercise, but41changes would most likely always be masked by glucose uptake, or a lack of glucoseuptake, in the larger white fiber population.Cardiac glucose utifization. In vivo glucose utilization in trout heart was not affected byincreased aerobic swimming. Before estimating the contribution of glucose to cardiacenergy production, it should be noted that cardiac power output in isolated trout heartsvaries from 1 - 2 mWg’ ventricle mass to maximally 6 - 8 mWg (see Milligan andFarrell 1991). Interpreting this as an in vivo scope for cardiac power output means thatrealistic limits of oxygen consumption range from 1 (rest) to 4 (exercise) mol02gmin1 (assumes a cardiac efficiency of 20 %, Graham and Farrell 1989; and a caloricequivalent of 02 of about 4.8 kcalml). Oxygen consumption by isolated hearts in thisstudy agree with the lower end of this estimated in vivo range (see Table 3). An in vivorate of glucose utilization of approximately 5 nmolgmin indicates that circulatoryglucose accounted for about 6 % of the expected cardiac oxygen uptake at rest and lessthan 1 % during exercise.As with red muscle metabolism, there are still some inadequacies in ourunderstanding 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 vitroobservations that iodoacetate treated hearts remain functional when provided palmitate inthe perfusate (Driedzic and Hart 1984) and possibly via endogenous triglyceridemobilization (see Milligan and Farrell 1991). However, the effects of fatty acid utilizationon glycolytic flux in fish hearts is not known. It may be that the trout heart is similar tomammalian heart models in which fatty acids restrict glucose oxidation, possibly throughcombined effects on transport and glycolytic enzymes, and enable sustained cardiacfunction over a broad work range (Kobayashi and Neely 1979; Saddik and Lopasehuk1991). This agrees with calculations of low glucose oxidation in vivo and withobservations that glucose in the absence of other substrates accounts for up to 50 % of the42oxygen consumption of isolated trout hearts operating at 1 - 2 mWg1 (calculated fromTable 2 values). Endogenous fuel stores presumably account for a substantial portion ofthe oxygen consumed in trout hearts, but quantification of the specific role of myocardialglycogen and triglyceride in relation to work intensity requires more study.Myocardial fuel utilization in vivo could also switch from fat preference during lowcardiac work to lactate during elevated aerobic activity, as speculated for skipjack tunaKatsuwonus pelamis (Moyes et a!. 1992a). High concentrations (10 mM) of circulatorylactate can serve as the sole oxidative substrate in isolated trout hearts over the range ofpower output (Milligan and Farrell 1991). Indeed, even at concentrations observed inplasma of trout performing endurance-type exercise (2- 3 mM; Weber 1991), lactate willinhibit cardiac glucose oxidation in vitro (Lanctin et a!. 1980) and can partially alleviatetrends of diminishing performance seen in glycolytically inhibited hearts (Driedzic andHart 1984). While it is doubtful that lactate is the preferred fuel for whole-bodymetabolism 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 smallcardiac tissue mass. Either case of a fat-to-lactate transition in relation to increasedmyocardial energy demand or of an overall dependence on fatty acids is compatible withlow glucose oxidation in vivo and prompts the generalization that cardiac glucosemetabolism in teleosts is directed mainly toward biosynthesis of storage substrates(glycogen and triglyceride), with a potential oxidative role only during periods of fatty acidlimitation.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 oxygenconsumption.43Muscle glucose utilization during sustained swimming in carp.INTRODUCTIONThe carp (Cyprinus carpio) is typical of teleosts in the sense that estimated in vivoglucose kinetics at rest are relatively slow (Garin et al. 1987). However, comparison ofthe fat utilizing capacities of carp and the fast-swimming skipjack tuna (Katsuwonuspelamis), suggests that the general hypothesis that better aerobic performers depend to agreater extent on fat-based fuels (eg., see Weber 1992) may also hold for teleosts withdifferent aerobic swimming capacities (Moyes et al. 1992). The small relative mass ofheart and red muscle in conjunction with the relatively sluggish swimming characteristicsof carp may mean that glucose disposal is significant in oxidative muscle, despitecomparatively low in vivo glucose kinetics.In the present study, we investigate the importance of circulatory glucose in theswimming energetics of carp by examining in vivo glucose kinetics and tissue specificglucose uptake, determined with bolus-injections of6-3H-glucose and14C-2-deoxyglucose(14C-2-DG). Simultaneous determination of flux and uptake is useful for evaluating theimportance of skeletal muscle in whole-body glucose disposal. Furthermore, measurementof tissue 2-DG uptake provides a means of detecting exercise-stimulation of glucosedisposal in small tissue masses (eg., heart and red muscle) which may be mediated byredistribution of blood flow or changes in muscle transporters in the absence of directeffects on whole-body glucose flux. As in the previous study of trout, the startingassumption for estimating oxidation of glucose was that red muscle accounted for the bulkof the whole body metabolic costs at 80 % Since the fate of glucose in workingmuscle need not be oxidation (L.aughlin et a!. 1992), 2-DG uptake is therefore used as anindicator of the maximum potential for oxidative glucose disposal in muscle duringexercise.44MATERIALS AND METHODSAnimals. Carp (1 - 2 kg) of both sexes were purchased from a local supplier andmaintained in dechlorinated fresh water (10 - 13 °C) in the aquarium facility at theDepartment of Zoology, U.B.C. Fish were held for at least one month before being usedin experiments and were fed twice weekly with trout pellets (Moore-Clark, Vancouver).Estimation of U.j1. U, representing the maximum speed sustainable for 30 mm usingthe present exercise protcol, was determined for individual carp (986 - 1790 g, 35 - 45 cmtotal length) using a Brett-type swim-tunnel respirometer (Gehrke et a!. 1990). A fish wasplaced in the swim-tunnel and water velocity was adjusted to between 0.6 - 0.7 bodylengthssec1. Exercise intensity was maintained at this low level for 2 h. Therespirometer was then closed to inflowing water for 20 mm and oxygen consumption (mg02h1kg)was determined by circulating water from the respirometer at a constant rateover a P°2 electrode (Radiometer, Copenhagen). P02 data was collected and stored usinga computerized, data aquisition system described previously (Gehrke et a!. 1990).Thereafter, swimming speed was increased every 30 mm in 0.3 body lengthssec1increments and oxygen consumption was determined at each new speed unless the fishfatigued within the first 10 - 15 mm. Between estimates of oxygen uptake, therespirometer 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 longerinduced the fish to swim, was recorded and Uj was calculated using equations reviewedby Beamish (1978). After exercise, fish were allowed to recover in black Perspex fishboxes that were supplied with areated, flow-through fresh water.Sustained swimming in cannulated carp. Following exercise recovery (48 h), individualcarp were anesthetized in buffered MS-222 (tricaine methane sulfonate, 0.2 gL1 with 0.4gL1 NaHCO3). Each fish was placed in an operating sling and the gills were irrigated45with oxygenated water containing a light level of buffered anesthetic (0.08 g MS-222 L1).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 securedthrough the membrane of the distendable upper lip. The cannula was filled with Cortlandsaline (Wolf 1963) that contained 5 i.u. ml1 of heparin (Glaxo, Montreal) and the fish wasplaced 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 thatused in the preceding study of trout glucose kinetics (West et al. 1993). A fish was placedin the swim tunnel and forced to swim at a low intensity (about 1/3 of a body lengtfrsec1)for 2 h. Swimming speed was then increased every 5 mm (in 0.3 body 1engthsecincrements) until the fish displayed obvious burst and glide swimming behavior. At thispoint, water velocity was reduced until the fish could maintain its position in the swimtunnel. Fish were then exercised at this high intensity for a total of 120 mm. Radiolabeledmetabolites were administered after the first 60 mm. During the interval between 30 - 60mm post-injection of radioisotopes (described in the next section), the oxygen consumptionof swimming carp was determined as described previously.Bolus-injections and tissue sampling. Appropriate volumes of6-3H-glucose (15 MCi)and 14C-2-DG (5 PCi) were dried under a constant stream of N2 gas. The radiolabeledmetabolites were reconstituted together in saline (140 - 180 1.d). An aliquot (10 l) wasremoved immediately, mixed with aqueous scintillant (10 ml of ACS II, Amersham) andcounted using a dual-label assay on an LKB Rackbeta scintillation counter. The remainingsolution was drawn into a glass Hamilton syringe to determine the volume of bolus to bedelivered. A bolus injection was delivered to individual fish through one port of a 3-waystopcock that was attached to the free end of the implanted cannula. Another syringeconnected to the second port of the stopcock served as a saline rinse (2 volumes) forflushing solution from the glass syringe and the cannula into the fish.46Blood (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 fishwas replaced with an equal volume of saline. After retrieving the final blood sample (60mm), 2 ml of sodium pentobarbital (65 mgml1)was injected through the cannula in orderto kill the fish with minimal struggling. When opercular movements had ceased, a singlecross-sectional slice (- 1 cm thick) of skeletal muscle was removed from immediatelyposterior to the dorsal fin and freeze-clamped between aluminum tongs that were precooled in liquid nitrogen. The heart was removed and also freeze-clamped. Less than 30sec elapsed from the time the fish was killed to the moment the last tissue was immersed inliquid nitrogen. Tissue and plasma samples were stored at -80 °C.Metabolite assays. Plasma was separated immediately from blood samples bycentrifugation and a precise volume was deproteinized with one volume of 0.6 N perchloricacid (PCA). Protein was precipitated by centrifucation and the plasma was neutralizedwith 3 M K2C03 in 0.5 M triethanolamine hydrochloride. Aliquots (50 1d) of the finalsupernatant were placed in 20 ml scintillation vials and evaporated to driness under streamsof N2 gas to remove any 3H20. Residues were reconstituted in 1 ml of water and assayedusing a dual-label protocol on a LKB Rackbeta scintilation counter to estimate radioactivityin plasma6-3H-glucose and 14C-2-DG.The frozen cross-sections of skelatal muscle were immersed in a shallow bath ofliquid nitrogen and samples (0.5 - 1.0 g) of red and white muscle were dissected free. Themuscles were chopped into fragments and transferred to pre-weighed tubes that containedapproximately 4 volumes of ice-cold 0.6 N PCA. The tubes were re-weighed and sampleswere homogenized with 2 x 20 sec passes of an Ultra-turrax tissue homogenizer. Twoaliquots (100 1.d) of this homogenate were removed for glycogen analysis. Aftercentrifugation (10 mm, 7500 g), the tissue extracts were neutralized, spun again and the47resulting 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 molglucosyl unitsg tissue. Muscle glycogen was digested by incubating the homogenatesfor 3 h at 40°C with amyloglucosidase (Boehringer Mannheim, 2 mgmr1, 1 ml totalvolume) in acetate buffer, pH 4.8 (Bergmeyer 1985). Perchioric acid (25 d, 70 %) wasused 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, weredetermined with standard enzymatic assays Bergmeyer (1985) that were modified for usewith microtitration plates (0.3 ml) and a Titertek Multiskan plate reader.Muscle 14C-2-DGP was separated from total tissue 14C radioactivity using theanion-exchange chromatography method described in the preceding trout study (Chapter1.2, West et al. 1993).Calculations. Radioactivity in dried plasma samples was used to estimate ratios of 6-3Hglucose/glucose and14C-2-DG/glucose (in dpmmoP1). The change in each ratio for anindividual fish was plotted against time (mm post-injection) from 1 to 60 mm and fit to adouble exponential equation. To estimate glucose turnover the appropriate curve wasextrapolated to time zero and infinity and the area under the curve was calculated (Katz1992). The ratio of the injected dose of6-3H-glucose (dpmkg1body mass) to area underthe curve (dpmminmol1was formulated to estimate glucose turnover rate (JLmolmin1kg). The 14C-2-DG washout curve was integrated from time 0 to 60 mm and an indexof muscle glucose utilization (GUI) was calculated as tissue 14C-2-DGP (dpmg1 tissue)divided by the area under the plasma 14C-2-DG washout curve (dpmmin’nmol).Calculation of glucose utilization rate (GUR) in specific tissues requires that GUI iscorrected with a lumped constant (LC) to account for slower 2-DG phosphorylationcompared to glucose (Sokoloff 1983). For discussion of the energetic importance of48glucose in carp muscle, GUR’s were in carp tissues were estimated by dividing GUI byLC’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 assessedwith ANOVA or Mann-Whitney U-test for non-normal data. Regression analysis was usedto examine relationships between glucose kinetics and plasma glucose concentration.RESULTSCarp swimming performance. Considerable variability existed in oxygen uptakedeterminations between individual carp, but all showed similar rates of change in logoxygen consumption with increasing exercise intensity (Figure 3). Maximal oxygenconsumption of the 4 fish that had reached a swimming speed of 1.5 body lengthssec’averaged 175 ± 36 mg02h1kg. Extrapolation of the general regression to a swimmingspeed of 0 body lengthssec1indicates that the oxygen consumption of resting carp at 15°C is expected to average about 43 mg02h1kg (95 % confidence limits indicate arange of between 25 to 65 mg 02h1kg).for the 6 exercised carp ranged from 1.00 to 1.68 body lengthssec1,averaging 1.38 ± 0.11 body lengthssec1 (61 cmsec for the average body length of 44cm). Two fish had U’s of about 1 body 1engthssec and these were eliminated fromfurther 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 lengthssec1(54 cmsec’). This speed corresponded to about 80 % of the Ucritdetermined for the 4 re-exercised fish (1.56 ± 0.06 body lengthssec1or 69 cmsec1).As indicated (Figure 3), the measured rate of oxygen consumption for the cannulated fish(130 ± 10 mg02h1kg)was in agreement with the rate expected for uncannulated carp.Given that cannulated carp were able to sustain swimming at the test speed for 120 mm and49‘‘2.5E 1.0DCCC•- 0.00.0 0.5 1.0 1.5 2.0Swimming speed (body lengths.sec’)FIGURE 3. Oxygen consumption of 6 carp during a stepwiseincrease in swimming speed. Fatigue speed (Ucrit) was determined and 4 of these same fish, plus one additional fish, werecannulated and re-exercised at 80 % U. Arrows denote average speed and oxygen uptake of the swimming, cannulatedcarp.y = 1.64 + 0.35(x), r=0.68———050that average oxygen consumption could be predicted from the relationship between oxygenuptake and swimming speed determined for uncannulated carp, it would seem that theimplanted cannulae had minimal effect on swimming performance. Since burst and glideactivity was observed in carp at one velocity increment (0.30 body lengthssec1)above thefmal test speed 1.26 ± 0.10, it would seem likely that Ujt was similar before and afterplacement of the dorsal aortic cannulae.Glucose turnover. Plasma glucose ranged 3 - 17 mM among individual carp at rest andranged 5 - 10 mM among swimming fish. Nevertheless, plasma glucose remained insteady state in individual fish from both groups (see Figure 4) - the coefficient of variationof plasma glucose was < 15 % for individual fish throughout the blood sampling period.Although glucose turnover rate was elevated in swimming carp, the effect ofexercise on average turnover rate was not significant (Table 5). However, turnover wasclearly dependent on plasma glucose concentration and the slopes of these retationshipswere different (p = 0.05) for resting and exercising fish (Figure 5). Intercepts of the twolines were not significantly different. Interestingly, despite the difference in regressionslopes, turnover in both swimming and resting carp changed proportionally withconcentration. 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 plasmaclearance of glucose which ranged from 0.3 to 1.3 mlminkg in carp (Figure 5) andwas 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 ofexercised carp (22.1 ± 10.0 mol glucosyl unitsg’) was lower than that of resting fish(41.1 ± 8.1 mol glucosyl unitsg1), but the difference was not significant as a result of51I - _I‘—5a)C’,0C)-C35C’,- 60‘II I I03EI I0 102030405060Time (minutes)I I I I I I_____I0 10 20 30 40 50 60Time (minutes)FIGURE 4. Plasma glucose concentration and washout of‘4C-2-DG and6-3H-glucose in resting (hollow symbols) andexercising (filled symbols) carp following bolus-injection oftracers.TABLE5.Whole-bodyglucoseturnoverandmuscleglucoseutilizationindex(GUI)inrestingandswimmingcarp.GUIGlucoseGlucoseWhiteRedHeartTurnoverClearanceMuscleMuscle(mo1min1kg1)(mlmin’ kg1)(nmo1min1g1)Resting3.8±1.00.5±0.11.1±0.43.3±1.58.8±2.9Swimming6.9±1.31.0±0.1#1.7±0.618.4±7•9*24.5±4.8*signinicantlydifferentfromrestincontrol(Mann-Whitney ranksum,p=0.03).significantlydifferentfromrestingcontrol(ANOVA, p<0.05).t’.)53FIGURE 5. Glucose turnover (upper panel) and plasmaclearance of glucose (lower panel) in carp. Turnover wasdependent on glucose concentration in both resting and swimming fish. Plasma clearance was concentration-independent.• swimming y = 0.6 + 0.4(x) (r =0.92, p 0.03)o resting y = -2.3 + 1.3(x) (r =0.89, p =0.04)••o>I15O1O.—I :‘o.—EO.50.0••0 ••.0 0000I I I I5 10 15Plasma glucose (mM)2054the large degree of variability in the data. Similarly, the difference between glycogenlevels in white muscle of carp at rest (21.3 ± 4.3 mol glucosyl unitsg1) and afterexercise (12.4 ± 6.8 mol glucosyl unitwg1)was not significant. Effects of exercise onthe terminal level of lactate in red (3.2 ± 1.3 molg’ at rest and 8.0 ± 1.6 molg1after exercise) and white (5.1 ± 1.2 molg1 at rest and 13.2 ± 4.9 molg1 afterexercise) muscle of carp were not significant.Sustained exercise resulted in an approximately 6-fold increase in GUI in redmuscle, while GUI in heart increased by 3-fold (Table 5). GUI in white muscle increasedby 54 % during exercise, but the average rate was not significantly different from that ofresting controls.Glucose utilization would appear to be associated with the terminal glycogencontent of muscle. Figure 6 illustrates that at very low glycogen levels, red muscle GUIwas elevated by more than 4-fold above the average rate for fish in which red muscleglycogen levels were in excess of 10 mol glucosyl unitsg1. Low glycogen content wasobserved 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 ofcardiac glucose uptake (see Figure 7) suggests that glycogen level was possibly reduced inall muscles of this one animal.Muscle GUI in individual fish was examined (Figure 7) for evidence that plasmaglucose level may have influenced uptake rate in the different muscle-types. GUI in whitemuscle varied over a relatively narrow range (0.1 - 3.5 nmolmin1g)in both swimmingand resting fish. Although trends indicate positive slopes in plots of white muscle glucoseutilization, regression slopes were not significant. In red muscle, GUI appearedindependent of plasma glucose level in both resting and swimming carp. The one highestimate of glucose uptake (50 nmolmin1g)occured in a fish which displayed lowterminal red muscle glycogen (< 1 mol glucosyl unitsg1). Excluding this onedetermination, GUI in the 4 other exercised carp averaged 10.7 ± 2.6 nmolmin1,55,6050 0 resting. • swimmingE040E300N• —Cl) •0 owC.) 0.o i 01 100 10 20 30 40 50 60 70Glycogen content (mo1 glucosyl units. g)FIGURE 6. The relationship between glucose utilizationrate and terminal glycogen content in red muscle of carpthat had been at rest or swimming at 80 % Ucrjt•56displaying no tendency to change with increasing plasma glucose concentration. In heart,significant regressions were calculated for data from resting fish (utilization = -1.3 +1.2[glucose]; r = 0.83, p = 0.04) and swimming (utilization = -14.5 + 4. 7[glucose]; r= 0.91, p = 0.05) carp. Slopes of these concentration-dependent relationships weredifferent (p = 0.05) and showed trends similar to the pattern observed for glucose turnoverrate.DISCUSSIONThe present study examined the role of circulatory glucose in the swimmingenergetics of carp. Glucose turnover increased with exercise, but the absolute difference influx between resting and swimming fish was dependent on plasma glucose concentration.Despite this concentration effect on turnover, glucose availability was apparently of lessimportance to the active locomotory muscle than to other tissues, like myocardium, sinceGUI 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 thefollowing 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 carpwere more reliant on circulatory glucose than were better aerobic performers like rainbowtrout, with the latter using glucose for an estimated < 10 % of total fuel utilization duringexercise at 80 % U.Oxygen consumption. Carp oxygen uptake in the present study compares well withprevious measurements. The extrapolation of active oxygen uptake to zero activityconcurred with other such procedures and with determinations of oxygen consumption ininactive carp at 15 °C (about 50 mgO2h1kg in 0.8 - 2.0 kg animals: Christiansen et a!.1982; Hughes et a!. 1983; Ultsch et a!. 1980). Maximum active oxygen consumption inthe present study was about 4-fold higher than the estimated resting rate, resulting in a574white muscle3 0- 2 •0110 00— 0o 0 I• I I IE 50-30-red muscle.— ‘, .10- 00(1•pp I fl I I40-— ..—.._.e-n - heart0c, 20oC.) .10- 0 Q00 00 I I I I0 5 10 15 20Plasma glucose (mM)FIGURE 7. Muscle glucose utilization index (GUI) inresting (hollow symbols) and swimming (filled symbols)carp in relation to plasma glucose concentration.58metabolic scope of 132 mg02h’kg1. This expansion of oxygen consumption from restto near maximal exercise is less than one-half of the metabolic scope in salmonids ofsimilar mass (Kiceniuk and Jones 1977). Although resting rates of oxygen consumptionare comparable in -1 kg carp and trout, maximal oxygen consumption is higher insalmonids as a result of their superior swimming capacity (Ut of 2 body lengthssec1;Kiceniuk and Jones 1977). Oxygen consumption in carp swimming at 80 % of U (130mg02h1kg)was about 3-fold higher than the predicted resting rate. This metabolicscope 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 beenmeasured previously for other teleosts (Garin et a!. 1978). This resulted largely fromhigher glucose levels in carp in the present study. The regression describing proportionalincreases in resting glucose turnover with glucose concentration allows us to predict that at2 - 4 mM glucose, whole-body glucose utilization would be within the range estimated inmost species of teleosts that have been studied (1.4 - 2.2 molmin1kg (Garin et at.1978)).The effect of exercise on glucose kinetics in carp was revealed upon inspection ofthe concentration-dependent glucose fluxes during rest and exercise (Figure 5). Over therange of 5 - 10 mM plasma glucose, exercise was clearly accompanied by a constant 2-foldstimulation of turnover. The absolute difference in turnover between resting andswimming fish was dependent on glucose concentration, as a result of the difference inslopes of the data regressions. However, proportionality in these relationships meant thatglucose clearance (turnover divided by glucose concentration) was independent of plasmaglucose and the doubling of clearance rate (Figure 5) confirmed the concentrationindependent effect of exercise on glucose kinetics, at least over the range of glucoseconcentrations measured.59The regression for swimming carp would be predicted to overlap that for the restingfish at 2 - 4 mM plasma glucose, suggesting that an exercise effect would not be detectedat low plasma glucose concentrations. The implication (discussed further in chapter 4) isthat glucose is used increasingly for swimming energetics depending on its plasmaconcentration. However, this would appear to not be the case since GUI in red muscle, themain locomotory muscle at 80 % was concentration-independent. The possibilitythat 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 incardiac GUI parallel concentration-dependent turnover rates in the whole-animal (Figure7).Muscle glucose uptake. The effect of exercise on red muscle glucose utilization wouldappear to be pertinent to the oxidative demands of this fiber mass. If, as a startingassumption, the metabolic scope for sustained swimming (87 mg O2h1kg)in carp isattributed solely to the increased aerobic functioning of the lateral red muscle, thenoxidative demand for glucose would have been equivalent to about 7400 nmol’min1kg.In turn, assuming about 70 g of red muscle per kg body mass (Bone 1978), the oxidativeglucose demand in active red muscle per gram of tissue mass is estimated at just over 100nmolmin1.This compares to an estimated red muscle GUR of about 28 nmolmin’g1 (excluding the one value for the glycogen-depleted fish and using a red muscle lumpedconstant of 0.40; West et a!. 1993). Therefore, a significant portion (25 - 30 %) of theoxidative energy demand in swimming carp could potentially have been derived fromcirculatory glucose. This estimate would increase in the likely event that other aerobicmetabolic costs also contributed to the change in whole-body oxygen consumption. Forinstance, electromyograms from previous work on swimming carp (40 cm length andacclimated to 15 °C) indicate that intermediate pink muscle fibers are recruited at 1.3 bodylengthssec (Johnston et cii. 1977), a speed comparable to 80 % Uj in the present study.60Given that sustained swimming was established for individual carp by visual inspection ofswimming behaviour (i.e., water speed was slowed until burst-and-glide swimmingceased), we cannot rule out the possibility of aerobic involvement of intermediate fibers inthe present study. White muscle recruitment, however, was probably minimal since suchrecruitment in carp is generally accompanied by rapid fatigue (Rome et al. 1984).Intramuscular glycogen could have augmented, or even dominated, thecarbohydrate 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 presentstudy were inconclusive because of the variability in resting glycogen levels and because ofthe potential intermittent use of muscle glycogen during step changes in swimming speed.Nevertheless, it is noteworthy that GUI increased markedly when red muscle glycogen hadbeen depleted to below 10 mol glucosyl unitsg1 in swimming fish (Figure 6). At themoment, it is not known whether glucose transporters in fish muscle are sensitive to eitherthe 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 notsaturated in the carp in this study. An interpretation that favors carbohydrate as thepreferred substrate in active carp red muscle is that this situation reflected reversal of theinhibitory effects of glycogenolysis on hexokinase activity (Williamson 1965) as glycogendepletion progressed. In humans with McArdle’s disease, the absence of musclephosphorylase and thus of glycogenolysis, is in a sense analogous since glucose utilizationis enhanced in patients exercising at intensities that normally induce muscle glycogenmobilization (Vissing et al. 1992). As has been argued for swimming trout, however, asimilar inhibition of red muscle glucose utilization could also occur if lipid-based fuelswere preferred for oxidation (West et al. 1993). In any case, it would seem that redmuscle glucose utilization was dependent less on the availability of glucose than on theavailability of other competing substrates. Given that the endurance of carp at high relativeswim speeds (at least 25 h can be sustained; Van Dijk et al. 1993) would over-burden61typical red muscle glycogen stores, the possibility of shifts in red muscle fuel sources(intramuscular to extramuscular) and types (carbohydrate versus lipid) throughout anendurance swim should be examined more closely.Unlike skeletal muscle, GUI in cardiac muscle showed defmite concentration-dependent uptake and an effect of exercise that paralled the responses seen in whole-bodyturnover. Complete oxidation of glucose at the utilization rates estimated in this studywould have accounted for 10 - 40 % of total myocardial fuel consumption in resting andswimming carp (assumes resting power output of about 1 mWg for slow swimmingspecies (Farrell 1984), exercising power output 4 x resting (A.P. Farrell, personalcommunication), 20 % cardiac efficiency (Graham and Farrell 1990), caloric equivalent of20.1 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 myocardialsubstrate use for energy provision. Fuels other than glucose would appear to dominatecardiac energetics at low glucose levels. Possible shifts toward greater reliance on glucoseoxidation at higher plasma glucose concentrations would be consistent with previousarguements (Sephton et a!. 1990, Sidell et a!. 1987) that the teleost myocardium seemspoised 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 duringsustained swimming in the carp (Cyprinus carpio). Am. J. Physiol. (submitted). I amgrateful to C.J. Brauner for helping with the swimming trials and for measuring oxygenconsumption in active carp.62CHAPTER 3. CARBOHYDRATE RECOVERY IN SKELETAL MUSCLE OFRAINBOW TROUT (Oncorhynchus myldss) AND SKIPJACK TUNA (Katsuwonuspelamis) AFTER BURST EXERCISE.Evaluating the role of hyperglycemia in post-exercise resynthesis of glycogen in troutskeletal muscle.INTRODUCTIONBurst swimming in teleosts causes dramatic changes in metabolite levels which canleave the white muscle depleted of fuel reserves at the end of an exercise bout (Milliganand Wood 1986b; Mommsen and Hochachka 1988; Schulte et a!. 1992). The resultingaccumulation of lactate occurs in stiochiometries that quantitatively reflect depletion ofintracellular glycogen. Subsequent post-exercise clearance/recovery of white musclemetabolites is generally a slow process with little metabolite exchange occuring betweenmuscle and plasma (Tang and Boutilier 1991). With respect to glycogen recovery,evidence suggests that the quantitative significance of glycogenesis from glucose is minimalbecause of the predominance of an, as yet undefined, intramuscular glyconeogenic pathwayfrom lactate (Arthur et a!. 1992; Moyes et a!. 1992; Schulte et a!. 1992; Pagnotta andMilligan 1991; Tang and Boutilier 1991). However, the importance of white muscle as asite of circulatory glucose disposal cannot be overlooked, if only because of the largerelative mass of the tissue.In contrast to the situation in white muscle, the stoichiometry of lactateaccumulation in red muscle does not appear to be matched with glycogen depletion at theend of exercise (Parkhouse et a!. 1987), which is perhaps indicative of greater transfer ofmetabolites between plasma and the well perfused red muscle mass. Post-exerciseglycogen repletion in red muscle of other species is very much dependent on the uptake ofsubstrate from the circulation, primarily glucose in mammals (Bonen et a!. 1990; Johnson63and 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 muscleare greatly depleted by high-intensity exercise (Parkhouse et a!. 1987), the relative utilityof lactate vs. glucose as a precursor for glycogen synthesis and the rate of fuel repletion isnot known for this tissue. Post-exercise hyperglycemia is sometimes seen in recoveringsalmonids (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 glucoseutilization 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 vivoskeletal muscle glycogenesis, particularly during a period of net glycogen synthesis(Kusunoki et a!. 1993). In addition to determinations of muscle glucose uptake, glucoseturnover was estimated simultaneously in recovering rainbow trout (Oncorhynchus mykiss)using bolus-injections of6-3H-glucose. The results are discussed in terms of the pertinenceof possible changes in post-exercise glucose availability to glycogenesis in different skeletalmuscle types.MATERIALS AND METHODSAnimals. Rainbow trout (Oncorhynchus mykiss) of both sexes and weighing 300 - 600g were obtained from a local supplier and maintained in the aquarium facility at theDepartment 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 atleast 1 month prior to experimentation.Surgery. Procedures followed for cannulating rainbow trout were precisely asoutlined in chapter 1.2. A single 60 cm cannula (PE-50, Clay-Adams) was placed in the6415-I0E00Pr0 2 4 6 8 10exercise Time (mm post-exercise)FIGURE 8. Preliminary observations of plasma glucoseand lactate in trout during recovery from enforced burstswimming.o lactate• glucose1265dorsal aorta of each fish. The cannula was filled with Cortland saline (Wolf 1963)containing 5 i.u. mL1 heparin (Glaxo Canada, Ont.) and was flushed twice daily. Eachexperimental fish was allowed to recover for 48 h in a black perspex box supplied with acontinuous flow of aerated freshwater (10 - 15 °C).Exercise and recovery protocol. Fish were netted from the recovery boxes andtransferred to a Brett-type swim tunnel. A protocol similar to the one used by Schulte eta!. (1992) was followed to ensure recruitment of white muscle and reduction of glycogenstores. Briefly, water speed was increased slowly over a period of 2 - 3 mm to amaximum level, defined as the point where the fish first displayed burst and glideswimming behavior. This speed was maintained until the fish started to rest against therear screen of the swim space. Water velocity was immediately reduced to a point wherethe fish could be induced to swim again and then once more increased gradually to themaximum. The cycle was repeated for about 15 mm until burst swimming could no longerbe induced. At this point the fish was quickly removed to a holding box for recovery andeventual administration of radiolabels.Studies of white muscle metabolite status in fish following maximal exercise showthat 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 expectedbetween 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). Isotopeadministrations were designed to bracket a portion of this interval in recovery so thatglucose utilization could be related to changes in muscle metabolite levels. Glucoseturnover and uptake in muscle were determined in a total of 9 exercised fish with bolusinjections of isotope given at 2.5, 4 or 5 h (3 fish per group) post-exercise. Glucose fluxesand tissue glucose utilization rates were also determined in 6 unexercised animals.66Isotope injection and tissue sampling. Bolus injections of6-3H-glucose (15 ILCi) and14C-2-deoxyglucose (2-DG, 5 MCi), purchased from Amersham Canada, were used todetermine in vivo circulatory glucose turnover and muscle glucose utilization, respectively.For each preparation, the appropriate volume of each isotope was dried under a continuousstream of N2 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 radiolabelinjected while the remainder was taken up in a glass Hamilton syringe to verify the bolusvolume. The syringe was connected to a 3-way valve positioned on the free end of thedorsal 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 thecannula. 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 fromeach sample by centrifugation and the plasma retrieved was deproteinized with an equalvolume of 0.6 N perchloric acid (PCA). The samples were neutralized with 3 molL1K2C03 in 0.5 molL1 triethanolamine and kept frozen (-70 °C) until analysed formetabolites and radioactivity.At the end of the sampling period, a mixture of sodium pentobarbital (1 ml of 65mgmL1;MTC Pharmaceuticals, Ont) and d-tubocurarine chloride (0.1 mgmU disslovedin 1 mL of saline; Sigma Chemicals, Mo) was delivered through the cannula to immobilizethe fish. When opercular movements ceased (30 - 60 see) the spinal cord was severedimmediately posterior to the head. Red and white muscle was taken with a single pass of adouble-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 N2, andimmersed in liquid N2. Only 10 - 15 sec passed from the time the fish was killed to themoment that the tissues were frozen. The samples were kept frozen at -70 °C.67Metabolite assays. Muscle was prepared for homogenization by dissecting away the skinand separating red and white fiber masses with the steak in a liquid N2 bath. Pieces oftissue (0.5 - 1 g) were placed in pre-weighed tubes and quickly reweighed. Ice cold 0.6 NPCA (4 vol) was added while the tissues were still frozen. The tissues were immediatelyhomogenized with three 20 sec passes of an Ultra-turrax homogenizer. Aliquots (2x100L) 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 tissuelactate and glucosyl units and plasma glucose were adapted for use with microtitrationplates (0.3 mL) and a Titertek Multiskan plate reader.The tissue content of 14C-2-deoxyglucose-6-phosphate (14C-2-DGP) wasdetermined in neutralized tissue extracts either by separation of 14C-2-DGP from 14C-2-DG on columns of DEAE Sephadex A-125 (Sigma Chemicals, Mo) or by precipitationwith 0.3 N Ba(OH)2and 0.3 N Zn(S04), as described previously (Chapter 2).The 2-DG lumped constant. The lumped constant (LC) is a correction factorneeded in the calculation of tissue specific glucose utilization to account for different ratesof 2-DG and glucose phosphorylation in tissues (Sokoloff 1983). In the study of trout redmuscle and heart the LC was estimated to be 0.40 (West et al. 1993), a value that wasunaffected by changes tissue glycolytic activity. In the present study, estimation of the LCfor trout white muscle was made by determining simultaneous in vitro rates of glucoseutilization and 2-DGP formation in slices of white muscle as described previously for redmuscle (West et a!. 1993). Briefly, slices of white muscle were prepared and preincubatedin saline as described previously. Slices were then incubated individually for 40 mm with0.3 Ci5-3H-glucose (Amersham Canada) and 0.15 Ci 14C-2-DG in fresh saline (2.5mL). At the end of the incubation the slice was removed, weighed and analyzed for 14C-2-DGP content. 3H20 was separated from 5-3H-glucose by distillation of the PCA68deproteinized/neutralized incubation medium as described previously (West et a!. 1993).The LC was formulated as the rate of14C-2-DGP formation divided by the rate of glucoseutilized (calculated from 3H20production).Glucose turnover and muscle uptake. Aliquots (20 ,LL) of deproteinized/neutralizedplasma were dried under a continuous stream of nitrogen gas for 5 h to remove 3H20.14c.2.DG was determined and3H-glucose was estimated after reconstituting the driedplasma with 1 mL H20 and adding 10 mL aqueous scintillant (ACS II; AmershamCanada). Radioactivity was assayed using a dual counting protocol prepared on an LKBRackbeta scintillation counter. Analysis of tracer washout for estimation of glucoseturnover and muscle glucose utilization was conducted as described previously (Chapter 2).Statistics. Data is presented as means ± se. The significance of differences betweengroups was assessed using ANOVA and Tukey’s HSD.RESULTSGlucose concentration and turnover rate, Average plasma glucose concentrations andwashout curves for the injected radiolabels are depicted in Figure 9 (resting fish) andFigure 10 (recovering fish). Plasma glucose concentration varied considerably amongexperimental animals (ranging 6 - 38 mmolL’), but individual fish remained in steadystate for the period following bolus-injection of radiolabels.No differences in plasma glucose concentration were evident between experimentalgroups (Table 6). High variability and small sample sizes during recovery likely accountedfor the lack of differences between groups, even though plasma glucose at 4 - 5 h ofrecovery was comparatively low. Average glucose turnover rate was also depressed duringthe 4- 5 h interval, with no statistical differences between groups (Table 6). Turnover ratewas linearly associated with steady state glucose concentration for individual fish at rest69000FUF•0s 1500 -E200‘e 1200 -0150900- 100600020 30 40 60TIME (mm)FIGURE 9. Plasma glucose concentration (upper panel) andwashout curves (lower panel) for‘4C-2-DG and6-3H-glucosein resting trout.0 10 20 30 40 50 60TIME (mm)0 10 5070— I I I I I I I0C)—0c)E3002502001501005000- 25- 20E 15I) 100C)-‘ 1800C,,1500_0bf. 1200Cd)00o 600:1300E0’63FIGURE 10. Plasma glucose concentration (upper panel) andwashout curves (lower panel) for‘4C-2-DG and6-3H-glucosein trout during recovery from burst exercise.10 20 30 40 50 60TIME (mm)I I I I I0 10 20 30 40 50TIME (mm)71and throughout the 2.5- 6 h of recovery (Figure 11). Separate regressions for resting andrecovering fish were not different, suggesting that changes in plasma glucose status had adirect influence on glucose kinetics while effects of exercise-state of the fish were lessimportant. The plasma clearance of glucose, calculated as turnover rate divided by glucoseconcentration, was shown to be independent of plasma glucose level (Figure 11). Thismeans that the volume of plasma cleared of glucose was relatively constant at 0.5 - 1.5mLmin1kg,regardless of plasma glucose concentration.Muscle glucose utilization. Pre- and post-exercise rates of glucose utilization in whitemuscle were calculated using the lumped constant determined from muscle sliceincubations (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 ratebetween control and recovering fish (Table 6). Like turnover rate, glucose utilization inwhite muscle between 4- 5 h of recovery tended to be 1/3 to 1/2 of the rates estimated atother recovery times and at rest. Glucose uptake in white muscle varied within relativelynarrow limits (Figure 12) and plasma glucose concentration had a less noticable influenceon utilization rate compared to its effect on turnover rate. As indicated, the significance ofa regression through these data was strongly influenced by a single observation which,emphasizing that utilization was only marginally dependent on glucose availability (Figure12).As is suggested in Figure 13, glucose uptake in red muscle of recovering troutchanged with plasma glucose concentration only between 6 - 12 mmol1J1. Above 12mmoiU1 utilization seemed independent of concentration. Since all of the glucoseutilization values for resting fish had relatively high plasma glucose levels (> 12 mmolU1), the average resting utilization rate from 9 fish with lower plasma glucose levels (< 5mmolL1, from West et al. 1993) is included in Figure 13 for comparison. The datasuggest that the trends in glucose uptake were similar for both resting and recovering trout.TABLE6.Whole-bodyglucoseturnoverandglucoseutilizationinskeletalmuscleduringrecoveryfromenforcedmaximalexercise.Utilizationinredmusclewascalculatedusingalumpedconstantof0.4(Westetat.1993),whileforwhitemuscle0.51wasused.RECOVERYINTERVALPRE-EXERCISE2.5-3.5h4-5h5-6hPLASMAGLUCOSE19.3±2.620.2±8.98.1±1.218.2±5.9(mmoFL1) GLUCOSETURNOVER19.1±2.319.7±8.87.1±0.6a20.7±6.8(molkg’min1)GLUCOSEUTILIZATION(nmolg’min1)REDMUSCLE18.5±1.712.2±3.25.2±1.1”10.9±3.6WHITEMUSCLE1.5±0.32.1±0.90.8±0.62.0±0.3a.p=0.07forcomparisontopre-exerciseglucoseturnover.b.p=0.008forcomparisontopre-exerciseredmuscleglucoseutilization.734030E2o8102.01.5. V001. I I I0 10 20 30i 40PLASMA GLUCOSE (mmol.L)FIGURE 11. Glucose turnover in resting and recovering troutas a function of plasma glucose concentration (upper panel),turnover = 0.97[glucose] +0.57, r=0.93 (solid line). Separatelines for resting (long dash) and exercised (short dash) fish werenot different. Plasma clearance (lower panel) was concentration-independent.I o PRE-EXERCISELv 3.5h, • 5h, • 6h000074‘5’3O 5O3’..4O2O2O-Io____ ____________10IPREEXERCISE0 PRE-EXERCISE, 4 v 3.5h, • 5h, • 6h08i+ .p I I I0 10 20 30 40PLASMA GLUCOSE (mmol.L1)FIGURE 12. White muscle glycogen and lactate in resting andrecovering trout (upper panel). *different from pre-exercise (p <0.05); **different from pre-exercise and 6 h (p <0.001). Glucoselevel had only a minor affect on utilization rate (lower panel), withutilization= 0.07[glucose] + 0.31, r = 0.68). *Denotes an observation with large influence on any regression (slope is not significantwithout this datapoint, r=0.48). Data from 9 resting trout with lowglucose levels (from West et al. 1993) is also shown (cross).I I I•4.0 5.0 6.0RECOVERY (hrs.)75_l —p. C.)4_I20C..2 .10CE000 0C.)l0.c-)•15L10oI0‘—015*4.000VL // ‘J0PRE- 5.0 6.0EXERCISE RECOVERY (hrs.)- o PRE-EXERCISE3.5h, • 5h, • 6h- 0•V• • VI.— I I I I0 10 20 30 40PLASMA GLUCOSE (mmol.U)FIGURE 13. Red muscle glycogen and lactate in resting andrecovering trout (upper panel). *different from pre-exercisevalue (p <0.05); **different from pre-exercise and 6 h (p <0.01).Glucose utilization in red muscle (lower panel) appeared to havebeen bi-phasic with respect to plasma glucose availability, becoming concentration-independent only at higher (>10mM)glucose levels. Uptake in 9 resting trout with lower plasmaglucose levels (see West et al. 1993) are also shown (cross).76There were differences (ANOVA p = 0.04, glucose concentration was an insignificantcovariate) 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 therange where utilization is sensitive to plasma glucose level (Figure 13).Glycogen replenishment in skeletal muscle. Post-exercise recovery of white and redmuscle glycogen is illustrated in Figures 12 and 13, respectively. Red muscle glycogencontent was still reduced 3.5 h into the recovery period and had recovered to 60 % of thepre-exercise value by the end of 6 h. The data indicate a rate of replenishment of about 20nmol glucosyl unitsmin1gred muscle over the interval from 3.5 to 6 h post-exercise.Glucose utilization in red muscle during recovery was 5 - 12 nmolming (Table 6),indicating that plasma glucose possibly supported a large portion, 25 - 60 %, of glycogenreplenishment in this tissue. In comparison, although white muscle glycogen contentremained significantly reduced 6 h post-exercise, the rate of glycogen replenishmentbetween 2 and 6 h (about 40 nmolmin1g)was 20 - 40 times higher than the rate ofglucose phosphorylation (1 - 2 nmolming’, Table 6).DISCUSSIONAn unexpected result in this study was the range of plasma glucose concentrationsamong the experimental animals. This might suggest variable levels of stress (Barton andSchreck 1987), but identifying the nature of any stress is difficult. In previous studies thatused the same pre-experimental care, anesthetization, surgical and handling proceduresthere was no hyperglycemia evident (West et a!. 1993). In the present study, glucose washigher in resting fish than in those that were exercised, indicating that hyperglycemia wasnot a direct result of the exercise protocol. In turn, trends in glucose turnover wereexplained primarily by plasma glucose status rather than exercise state (rest or recovery).However, while hyperglycemia and concentration-dependent glucose turnover are77remarkable from a glucoregulatory perspective, these kinetics seemed dissociated from thecarbohydrate recovery pattern in trout white muscle. It is important to note that turnoverincreased proportionally with plasma glucose concentration, suggesting that whole-bodyutilization of glucose may have occurred predominantly via substrate-mediated disposal.Despite this, glucose utilization rate in white muscle, the largest homogenous tissue massin trout, was low and was only slightly sensitive to plasma glucose concentration. Thissituation emphasizes that glucose availability was pertinent to tissues other than whitemuscle and, further, that white muscle glycogen was resynthesized essentiallyindependently of glucose availability. The interpretation in red muscle is lessstraightforward since the pattern of glucose utilization in relation to glucose availabilitywas bi-phasic, with an initial concentration-dependent phase followed by concentration-independence at higher plasma glucose levels. It would seem that glycogen reformation inthis muscle type is dependent on both glucose availability and glyconeogenic flux. Post-exercise blood glucose status, which is moderately hyperglycemic in some studies(discussed later), could therefore have consi&rable influence on the relative importance ofglucose 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 ratesdetermined 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 energyprovision. In neither of these species was glucose turnover necessarily accompanied byhyperglycemia. In trout, blood glucose status was closely connected to glucose turnoverrate in both resting and recovering fish (Figure 11). Turnover rates in resting salmonidswith similar plasma glucose levels are not available for comparison. However, the overallregression does suggest that at plasma glucose levels below 10 mmolL1 turnover rate islikely to fall within 2 - 10 molminkg. This estimate is in agreement with previous78measurements in resting rainbow trout in which plasma glucose was < 10 mmolL1(Dunn and Hochachka 1987; Washburn et a!. 1992) and demonstrates the apparent utilityof plasma glucose concentration as a predictor of whole-body glucose flux.Beyond its predictive value, the relationship between plasma glucose concentrationand turnover rate implies that whole-body glucose disappearance in trout responds toglucose availability over a wide range. Proportional changes in turnover withconcentration accounted for the observation that a relatively constant blood volume wascleared of glucose, independent of glucose concentration (Figure 11, lower panel).Concentration-dependent turnover further suggests that glucose concentration itself mayhave been an important regulator of whole-body glucose disposal. This would beconsistent with the generalization that glucoregulatory hormones are slow to respond toelevated glucose levels in teleosts (Harmon et a!. 1991), but it is not known whatregulatory characteristics are specifically involved with enhanced glucose production invivo. With respect to glucose disposal, an important point is that, even at relatively lowturnover 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 < 15 % of the total glucose removed from thecirculation. This also holds for resting salmonids with lower plasma glucoseconcentrations (2 - 4 mM), based on total muscle glucose disposal of about 0.5 molmin1kg body mass (West et a!. 1993) and turnover rates of 2 - 5 molmin1kg(Lin et a!.1978; Washburn et a!. 1992 or predicted from Figure 12). Therefore, the largest tissuemass 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 particularlyinteresting to investigate in relation to glucoregulation during stress-inducedhyperglycemia.79White Muscle Recovery. The present study provides direct evidence that extramuscularpathways of glucose formation are of minor importance to white muscle glycogenrecovery. Two observations point to the relative unimportance of changes in glucoseavailability to the process of glycogen resynthesis in this muscle mass. Firstly, post-exercise changes in white muscle glycogen and lactate appeared unaffected byhyperglycemia and elevated glucose kinetics since recovery rates were comparable toprevious findings (25 - 50 nmol glucosyl unitsmin1gand 40 - 100 nmol lactatemin1g1, 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 spannedquite narrow limits despite the highly variable plasma glucose levels and utilization was notlinked in a discernible way to recovery interval. It can be calculated that, regardless ofplasma glucose availability, glucose phosphorylation in white muscle could have potentiallyaccounted for 5 - 10 % of the glycogen formed during recovery (assuming white muscle is60 % of body mass and that glucose utilization at 1 - 2 nmolmin1gis uniformthroughout the tissue). This estimate of white muscle glycogenesis is somewhat higherthan one other estimate made previously with different methodology (see Pagnotta andMulligan 1991). Accounting for the extent to which competing pathways utilize G-6-Pmight be expected to bring these different calculations in closer agreement. However, thelikelihood that muscle glucose is directed primarily toward glycogen storage duringexercise-recovery conditions (> 80 % in rats, Kusunoki et a!. 1993) suggests that thepresent 2-DG measurements of glucose utilization rate are reasonable estimates of in vivoglycogenesis.Red Muscle Recovery. Unlike glucose utilization in white muscle, utilization rate inred muscle was bi-phasic and could account for a relatively large proportion of glycogenrepleted (25- 60 %). The difference in the rate and concentration-dependence of glucoseutilization between the muscle-types may be related to a higher amount of glucose80transporters in red muscle, as in mammalian skeletal muscle (Marette et a!. 1992). In anycase, the significance of an increased capacity to phosphorylate glucose in red muscle is thegreater potential use of glucose for glycogen synthesis compared to white muscle.Furthermore, the concentration-dependence of glucose uptake suggests that post-exerciseblood glucose status could have influenced the rate of glucose incorporation into redmuscle glycogen. The present data indicate that glucose incorporation into red muscleglycogen is potentially adjustable over a limited range of plasma glucose concentration, upto 10 - 12 mmolL1. While hyperglycemia in the present study was not specificallyrelated 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 whereconcentration-dependent utilization in red muscle was observed in the present study. Wecan conclude that glucose availability is likely to influence the rate of glucose incorporationinto red muscle glycogen, but given that glycogen synthesis occured at a faster rate thancould be accounted for by glucose utilization alone, it is probable that both of the processesof 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 forglycogen 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 forglycogen synthesis, > 90 % and 30 - 50 % respectively (Bonen et a!. 1990; Johnson andBagby 1988; Pagliassotti and Donovan 1990). Lactate is expected to be the majorglyconeogenic precursor in vertebrate muscle and greater oxidative losses of lactate inmammals (Brooks 1986) than in fish may partly explain greater dependence onglycogenesis in rat muscle-types. It is evident, however, that rats use glucose almostexclusively in red muscle, even when lactate is made available for glyconeogenesis (Bonenet a!. 1990). In trout, the process of glycogen resynthesis in red muscle resembles the81situation in mammalian white muscle since glucose accounts substantially, yetincompletely, for glycogen synthesis. Glyconeogenesis is probably involved in the overallprocess, as it is in mammalian white muscle (Bonen et at. 1990), but the specific role forlactate in trout red muscle is unknown. Changes in red muscle lactate content afterexercise (Figure 13) provide few insights. Nevertheless, plasma lactate increasesmeasurably in salmonids between 2- 8 h post-exercise (Milligan and Wood 1986a,b;Mommsen and Hochachka 1988; Pagnotta and Milligan 1991; Schulte et at. 1992; Figure8), as does lactate turnover (Milligan and McDonald 1988). The relative importance ofglyconeogenesis in trout red muscle might be expected to peak when plasma lactateavailability and flux is maximal. A better understanding of the relative importance oflactate and glucose in this muscle-type might emerge from studies of lactate incorporationinto glycogen in relation to glycogen recovery rate in individual fish - combiningtechniques of tracer lactate incorporation into glycogen with 2-DG uptake and sequentialbiopsy 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 ofhyperglycemia in the post-exercise resynthesis of glycogen in trout skeletal muscle. J.&p. Blot. Thanks to P.M. Schulte for assistance with the swimming trials.82Lactate, glycogen and PCr profiles in the white muscle of skipjack tuna duringrecovery from burst swimming.INTRODUCTIONTuna can sustain relatively high speeds of 3-5 body lengthss1 indefinitely, yetthey are also capable of short term speeds of up to 20 body lengthsc’. This intriguingcombination of sprint and endurance abilities is supported by a metabolism with severalunusual features. For example, skipjack tuna (Katsuwonus pelamis) are able to fuelburst swimming through an exceptional glycolytic capacity in white muscle, which canproduce lactate concentrations of about 100 Lmolg1 (Guppy et al. 1979; Hulbert eta!. 1979). Removal of elevated plasma lactate and metabolically-produced protonsfollowing exhaustive exercise occurs very rapidly in tuna (Barrett and Connor 1964;Perry et a!. 1985). In fact, lactate turnover in post-exercise tuna resembles ratesobserved in mammals (Weber et a!. 1986), suggesting that the circulatory translocationof 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 duringexhaustive exercise within the white muscle mass and require relatively long periods forlactate clearance (Batty and Wardle 1979; Milligan and McDonald 1988).In studies of recovery from exercise in other teleosts, such as trout, the fish isexercised and terminal samples are collected by freeze clamping at selected times afterexercise. Muscular contractions during sampling can cause artefacts resulting fromchanges in the concentrations of intracellular metabolites such phosphocreatine andATP. Nevertheless, with care and rapid sampling it has been possible to follow thechanges 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 lengthss1 to keep from sinking and to ‘ram ventilate’the gills (Guppy et a!. 1979). The capture of free swimming tuna is not compatible83with the requirement to reliably collect tissue samples from tuna that have notstruggled. Recently Bushnell et a!. (1990) described a protocol for spinally-blockingtuna which permitted continuous measurement of cardiovascular parameters undercontrolled conditions. By using this procedure, and sampling muscle with a high speedbiopsy technique, it was possible to separately collect and rapidly freeze musclesamples from the same fish at different times after exercise. The advantage of thisapproach was that it provided a means to analyse detailed muscle metabolite changesfrom individual fish and consequently to use fewer fish overall to elucidate generalpatterns of recovery metabolism than would be necessary if a terminal samplingprocedure were employed.MATERIALS AND METHODSExperimental animals. Live skipjack tuna, Katsuwonus pelamis, (1 - 2 kg) werepurchased from a local fisherman and maintained in outdoor holding tanks at theKewalo Research Facility (Southwest Fisheries Center Honolulu Laboratory, NationalMarine Fisheries Service, National Oceanic and Atmospheric Administration). Thetanks were supplied continuously with aerated sea water at 25 ± 2 DC. Fish were notfed and were used for experiments within three days of capture.Exercise and biopsy protocol. In preliminary studies, terminal muscle sampleswere 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 blowon the head. A sample of epaxial white muscle was dissected from behind the dorsalfm of each fish, freeze-clamped between Wollenberger tongs and immersed in liquidnitrogen as in earlier studies (Guppy et al. 1979). Very little struggling occurredduring this procedure, and less than 20 sec elapsed from the time of capture to the pointof tissue immersion in liquid nitrogen. Three other fish were sampled in the same84manner after first being isolated individually in another holding tank and chased withcapture nets for 15 mm. All dissected muscles were stored frozen over dry ice prior tohomogenization and assay procedures. The two groups were designated unexercisedfreezed-clamped and exercised freeze-clamped, respectively.A serial biopsy technique was developed to assess in vivo metabolite changes intuna white muscle during recovery from exercise. Each fish was first exercised asdescribed above, then captured by net and guided into a plastic bag containing bufferedMS-222 (tricaine methane sulfonate, 1.0 gL’ with 1.0 gL4 NaHCO3)dissolved inoxygen saturated sea water. This group is referred to as exercised biopsied. Tunataken from the holding tank and anesthetized without being chased served asunexercised controls and are referred to as unexercised biopsied. Immediatelyfollowing anesthesia (about 2 mm), each fish was placed on an operating table and thegills were irrigated with recirculated water containing a low level of anesthetic (MS222, 0.1 gL4). At this time initial biopsy samples were obtained from the epaxialmuscle using a 3 mm bore, at 10 - 15 mm depth (ALKO Diagnostic Corporation). Thetissue sample was drawn from the fish by suction into a chamber cooled with liquidnitrogen and freezing occurred almost instantly. Bleeding was minimal and wasstopped easily by inserting a cotton applicator tip into the puncture. Skin was separatedfrom 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 theoperating table and the ventral aorta was cannulated following procedures describedpreviously (Jones et a!. 1986). The fish was then righted and a 20-gauge needle, 6 cmlong, was inserted through the dorsal musculature (just lateral to the second spine of thedorsal 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 theswimming muscles. The needle was left in place for periodic re-injections throughoutthe experiment. Once secured in a foam-lined brace that was supported in a perspex85holding box (see Bushnell et al. 1990), the fish was presented with continuouslyflowing 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 waslowered 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 cmposterior to the spinal needle insertion site. Samples of white muscle and blood (0.25ml) 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 deproteinizedwith one volume of 0.6 N perchloric acid and stored on dry ice. A terminal musclesample 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-turraxtissue homogenizer in 2 ml of 8 % perchloric acid in 40 % ethanol which had beenprecooled over dry ice. Homogenization temperature was maintained below -20°C. A50 d aliquot of homogenate was removed for measurement of muscle glycogen and theremainder 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 theneutralized sample was stored on dry ice.Total creatine, creatine (Cr) + phosphocreatine (PCr), content remains constantfor a given tissue (Connett 1988) and was used as initial reference for all musclemetabolites measured. This is a reliable normalization procedure (Sabina et a!. 1983)which avoids possible inaccuracies associated with weighing small, deep-frozen tissuesamples. Conversion to moFg4,for comparison with other studies, was made afterdetermining total creatine content in tuna white muscle samples of known weight. Afactor of 39 (Jhmol total creatineg4white muscle) was used for these conversions.86Muscle lactate, glucose, Cr and PCr were measured according to Bergmeyer(1985), modified for use with microtitration plates (0.3 ml) and a Titertek Multiskanplate spectrophotometer. Glycogen was measured (see Chapter 1.2) in aliquots (100j.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 plasmasamples.Data analysis. Linear regression was used to determine break points in thecurves describing the recovery of white muscle lactate and PCr. Break points wereused as estimates of recovery time for these metabolites and means from exercised andnon-exercised tuna were compared using the two-sample t-test. Comparisons of allother metabolite concentrations in the white muscle of the various groups of tuna wereassessed with Tukey’s multiple comparison. Dependent t-tests were used to comparemetabolite levels in biopsies taken at different times throughout the recovery period.RESULTSMuscle metabolites after exercise. Terminal muscle samples from the exercisedfreeze-clamped tuna show significantly elevated lactate levels and depressed glycogenand PCr levels (Table 7) consistent with the utilization of anaerobic glycolysis infueling intense white muscle activity. Similarly, the initial biopsy from the exercisedbiopsied fish had significantly elevated lactate and depleted PCr relative to unexercisedfreeze-clamped tuna (Table 7). Glycogen in the first biopsy was reduced, but notsignificantly compared to unexercised freeze-clamped fish (p = 0.09), which likelyreflects the large variation associated with these values.TABLE7.Metabolitesinwhitemuscleofexercisedandunexercisedtuna.Freeze-clamped,unexercisedfishweretakendirectlyfromtheholdingtank,whilethefreeze-clamped,exercisedfishwerechasedfor15mm.andthenkilledimmediately.Biopsiesweretakenfrombothexercisedandunexercised,spinally-blockedtuna.BIOPSIESFROMSPINALLY-BLOCKEDTUNAFREEZE-FREEZECLAMPED,INflALFINAL(180MIN)CLAMPED,UNEXERCISEDEXERCISEDUNEXERCISEDEXERCISEDUNEXERCISEDEXERCISED(N=11)(N=5)(N=4)(N=5)(N=4)(N=3)PCr74.6±3.454.5±9.6’20.1±8.7a,b75.7±3.276.5±8.511.5±4•73,i(%TOTALCr)LACTATE7.3±3.838.5±10.4a107.5±36.7a,b11.9±8.114.9±15.775.8±4•9i,b(mol/g)GLYCOGEN145.0±34.4127.3±56.988.6±32.4c145.6±40.1139.3±53.674.6±25.1”(jLmol/g)PYRUVATE0.05±0.010.39±0.14a0.79±0.30c10.05±0.010.19±0.190.14±0.04e(JLmol/g)(a)significantlydifferentfromfreeze-clampedcontrolsandfinalbiopsy(unexercised&exercised);(b)significantlydifferentfrominitialbiopsy,unexercised;(c)p=0.09forcomparisontofreeze-clampedcontrols,andp<0.05forcomparisontofinalbiopsy,exercised;(d)significantlydifferentfromfreeze-clampedcontrolsandfinalbiopsy,unexercised;(e)significantlydifferentfromallothergroupsexceptfinalbiopsy,-Jexercised.88An intermediate metabolite profile was observed in the initial biopsies of theunexercised, biopsied tuna (Table 7) with lactate and PCr levels significantly differentfrom both unexercised and exercised freeze-clamped fish. These changes wereprobably the consequence of the fish struggling prior to complete anesthesia, perhapscombined with a direct effect of the MS-222 anesthetic on muscle metabolism (see Vanden Tillart et a!. 1989).It should be noted that the relative amount of creatine in the phosphorylatedstate is very high in tuna white muscle. In mammals and other teleosts PCr levels of30 - 50 % of total creatine are typically measured. Van Waarde et al. (1990) suggestedthat, based on measurements with NMR, in vivo Cr was 80 % phosphorylated in teleostwhite muscle and assert that lower levels arise because of handling stress and/or lessthan instantaneous freezing of tissue samples. The comparable values of 75 % in whitemuscle of resting and recovered tuna presented in this study (Table 7) attest to thereliability of the freezing methods used.Lactate, glucose and glycogen in recovery. Changes in white muscle lactate andglycogen during 180 mm. of recovery from exercise are illustrated in Figure 14. Inboth groups of biopsied fish the lactate recovery curves declined initially in a linearfashion and after 80 - 100 mm., remained constant at the pre-exercise level. Lactateconcentrations in the final biopsy taken from exercised and unexercised biopsied fishwere not significantly different from the resting level (Table 7). Biopsy glycogenconcentrations were more variable than lactate in recovery and significant differencesbetween initial and final tissue levels were detected in only the exercised group (Table7). However, most of the variation in recovering fish appeared to result fromdifferences between individual fish. Consequently, t-tests on paired data indicated thatglycogen concentrations in the white muscle of exercised biopsied fish was significantlyhigher in the final biopsy than at the start of the recovery period.89200• 150Oci00ct 50E150- 125-10oE751)250400EQ200150I I I I I10050015012510075502504010• 10I I I I I 00 50 100 150 200 0 50 100 150 200Time (mm) Time (mm)FIGURE 14. Glycogen, lactate and PCr in biopsies of tunawhite muscle, ° unexercised, • exercised. Statisticalanalysis is provided in table 7.I I I I I90FIGURE 15. Lactate and glucose levels in plasma ofexercised.I,5040302010050403020100I I I504030020,40E30 c2010—0200unexercised0 50TimeI I100(mm) 150exercised and unexercised skipjack tuna.91Glycogen concentration in the final biopsy was not different from unexercised freeze-clamped 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 JLmolg4, notsignificantly different from the fmal biopsy glycogen content).Plasma lactate concentration in the exercised biopsied tuna was essentiallysteady, 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 theunexercised biopsied fish, plasma lactate dropped steadily between 20 and 100 mmrecovery and remained constant thereafter at about 10 mM. Plasma was not obtainedfrom unexercised freeze-clamped fish, but the apparent resting steady state achieved inrecovery by the unexercised biopsied fish is comparable to previous measurements ofplasma lactate in resting skipjack tuna (Perry et a!. 1986).The plasma concentration of glucose remained in a constant steady-statethroughout the recovery period in biopsied fish (Figure 15). Glucose concentrationaveraged 12.9 ± 2.9 mM and 13.6 ± 3.5 mM, mean ± sd, for the control andexercised biopsied groups, respectively.Estimates of recovery time (lactate and PCr). Figure 16 depicts lactate and PCrconcentrations in tuna white muscle from representative exercised and unexercisedbiopsied fish during recovery. Every fish showed the same general recovery patternconsisting of an initial phase of rapid lactate and PCr change followed by a period inwhich these metabolites changed much more slowly. Both stages were linear, whichpermitted the estimation of recovery time by solving a set of regession equations foreach of the fish biopsied. This analysis assumes a homogeneous distribution of musclemetabolites, which appears to be valid on the basis of the consistent changes inlactate/PCr observed during recovery for each fish despite random selection of92150100Ec)ci()ci02005040__________30201000 50 100 150 200Time (mm)FIGURE 16. Lactate and PCr in white muscle ofrepresentative exercised (upper panel) and unexercised(lower panel) tuna, showing regression lines for calculating break points in metabolite recovery.50 100 150 200Time (mm)0 50 100 150Time (mm)1005000 50 100 150 200Time (mm)93Initial lactate level (mol.g4) °0 25 50 75 100 125 150 175 200I I I I I I I IFIGURE 17. Estimated recovery time in relation to lactateand PCr concentration in the initial biopsy of individualexercised and unexercised tuna. Points on the lower x-axisare mean lactate and PCr levels determined in 11 restingtuna (see Table 7)..I1209060300.0.+15.8, r=O.88r=O.93I I I0 20 40 60 80Initial PCr level (as % of total creatine)100.94biopsy sites. The variability in muscle glycogen during recovery prevented the use ofthe break-point determination procedure on these data.It became apparent that the experimental protocol produced a wide range ofrecovery times, rather than two discrete groupings, which reflected the degree ofmuscle lactate accumulation or PCr depletion. This essentially agrees with the earlierobservation that the unexercised biopsied fish exhibited a muscle metabolite profile thatwas intermediate to the resting and exercised freeze-clamped tuna, probably resultingfrom the uncontrolled activity during anesthetization. The recovery times overlappedsomewhat between exercised and unexercised biopsied fish and therefore it seemedmore appropriate to consider recovery time as a continuum with respect to exerciseintensity. This is emphasized in Figure 17 where recovery time is displayed as a linearfunction of the concentration of both lactate and PCr in the first biopsy obtained fromindividual fish. As a result of this overlap, the average lactate (and PCr) recovery timefor 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 timenecessary for lactate recovery, for either the exercised or non-exercised biopsied fish,was not significantly different from the respective PCr recovery time. Furthermore, ifindividual biopsied fish were once again considered, then there was a strikingsynchrony 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 theindividual sample biopsies (Figure 19). This suggests that, unlike other teleosts whichhave been studied, some regulatory mechanism seemingly operates to tightly couplemajor adjustments of these metabolites in tuna white muscle.95DISCUSSIONMuscle lactate and glycogen recovery. The disappearance of white muscle lactatelevels at a rate of 1.3 moFg’miif is up to 20 times faster than has been measuredfor trout (0.055-0.12 jmolgin). This unusually rapid clearance of lactate isintermediate between the rate of clearance for humans (0.66 molg’min’) and rats(about 2.5 molg1in)following maximal exercise (Hermansen and Vagge 1977;Meyer and Terjung 1979).Weber et a!. (1986) noted that despite high rates of whole-body lactate turnoverin tuna, lactate recovery apparently occurs too quickly in white muscle to result fromturnover alone, suggesting that some portion of the lactate remained in the whitemuscle as a substrate for post-exercise, in situ metabolism. In some teleosts, whererates of lactate turnover are greatly mismatched with actual clearance rates from whitemuscle during recovery from exercise (Milligan and McDonald 1988), retention andutilization of lactate within the white muscle mass is likely the dominant means oflactate disposal. Such a proposal of in situ metabolism of muscle lactate is prevalentthroughout the area of exercise-recovery metabolism of both mammals and ectotherms.Some evidence comes from Hermansen and Vaage (1977) who biopsied humanquadriceps muscle following maximal exercise and point out that the closely matchedrates of lactate and glycogen change in muscle, combined with a small arterio-venousdifference for lactate, suggest direct reconversion of lactate to glycogen. The enzymicsteps involved in the process are not understood well (see Bonen et a!. 1989 for apartial review and Chapter 4), but numerous studies on vertebrate muscle proposevarying degrees of in situ conversion of lactate to glycogen and demonstratedependence of the pathway on muscle fiber type, pH, arterial lactate concentration andan extramitochondrial pathway of glycogen resynthesis (Bonen et a!. 1990; Johnson andBagby 1988; Pagliassotti and Donovan 1990).96TABLE 8. Estimated recovery times for PCr and lactate in tuna white muscle. Withineach 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 UNEXERCISEDm=4 (N=5’iPCr 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.97y = 3.43 + 0.92xr2=0.99I I I I0 30 60 90 120PCr recovery time (mm)FIGURE 18. The relationship between estimatedrecovery times for white muscle lactate and PCr inexercised and unexercised tuna.120-60-I .30-098150x0xirr 0fJlVU xo 0E o0+CC..) X-iv + 00C X+ •Q.++ +.‘o•% 1’i00 5 10 15 20 25 30 35PCr (imo1.g-i)FIGURE 19. White muscle lactate in relation to PCr in biopsies fromexercised (large symbols; each symbol-type represents a different fish)and unexercised (small crosses; individuals not depicted) tuna. Alsoshown are measurements from muscle taken from 3 fish following 15mm of exercise (small filled circles).99Results of the present study support and extend the suggestion that tuna whitemuscle clears lactate in situ, since the nearly stoichiometric changes in the lactate andglycogen recovery profiles (i.e., 2 lactate removed: 1 glucosyl unit reformed) indicatemainly a glyconeogenic fate for most of the lactate produced during burst activity. Theextent to which this result approaches a quantitative representation of lactateincorporation into white muscle glycogen is reinforced by estimates of minimalglycogenesis from plasma glucose and low capacities for other tuna tissues to utilizelactate (discussed below).Plasma lactate and glucose.Plasma lactate concentration in the exercised tunaremained essentially constant for the period during which muscle lactate declinedsharply. In terms of the kinetics of lactate, the restrictive condition of a plasma steady-state means that lactate had to enter the plasma pool at the same rate which it wasremoved. Furthermore, 35 - 40 mM plasma lactate suggests, indirectly, a high rate offlux through the plasma pool in tuna (see Weber et al. 1986). In the post-exercisestate, white muscle is the probable source of plasma lactate which could be utilizedeither oxidatively or for gluconeogenesis in other tissues. In mammals, the primarilyoxidative disposal of lactate observed at rest and during sustained, aerobic activity haslead to the formation of the lactate-shuttle hypothesis (Brooks 1986) which proposes thetranslocation of lactate from producing to consuming muscle fibres.It is not likely that oxidative metabolism is reponsible for the removal of whitemuscle lactate in tuna. The oxygen consumption of skipjack tuna swimming at 2-5lengthss1 is calculated to be 0.68 molg1in’ (Gooding et a!. 1981) which is 4times the rate of a recovering trout (Mulligan and McDonald 1988). This rate ofoxygen consumption is still about 30 times too slow to account for the rate of lactatedisappearance by oxidation. It is possible that tuna have higher oxygen consumptionsduring recovery. However, it is unlikely that red muscle or the heart of tuna even have100the capacity to oxidize lactate at the rate required. Based on maximal estimates ofmitochondrial oxidation of pyruvate, tuna red muscle and heart ventricle couldmaximally oxidise 0.9 and 2.5 jmol lactateg1min respectively (Moyes et a!.1992). Thus, in a 1 kg animal, red muscle (assume 70 gkg1) could oxidize about 5mmol of lactate and the heart (4 gkg1, Farrell et a!. 1992) could oxidize about 0.8mmol of lactate in 80 mm. Combined oxidation in these tissues would thereforeaccount for the removal of less than 10% of an initial white muscle lactate load of 72mmoles (based on 108 molg1,table 7). If the temperature of tuna red muscle was10 °C higher than ambient (Hochachka et a!. 1978) then maximal oxidation would stillaccount for less than 15 % of the lactate clearance.The low capacity of tuna red muscle to oxidize lactate may mean that redmuscle is glyconeogenic to some degree, using lactate derived from white muscle in amanner similar to the version of a “lactate shuttle” proposed for reptiles (Gleeson andDalessio 1990). The glycogen content of tuna red muscle is reduced followingexhaustive swimming activity (Hulbert et a!. 1979), but the quantitative importance oflactate incorporation into red muscle glycogen is expected to be low because of therelative masses of red and white muscle. Integration of a shuttling mechanism with thelactate and glycogen changes observed in tuna must accommodate the apparentconservation of lactate carbon within white muscle mass.Another way for simultaneous glycogen recovery/lactate disappearance to occuris through the delivery of plasma glucose to muscle for glycogenesis. However, thecapacity for skipjack tuna liver to utilize lactate as either a gluconeogenic or anoxidizable substrate is low (Buck et a!. 1992) and in the general context of post-exercise lactate removal, contributes negligibly. Weber et a!. (1986) came to a similarconclusion by noting that Cori cycle activity is probably minimal in tuna recoveringfrom exercise. While liver glycogenolysis and gluconeogenesis from amino acids arealso potential sources of plasma glucose in teleost fishes (Suarez and Mommsen 1987),101it remains unlikely that turnover from all of these sources is high enough to supplyglucose 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 oftenindependent 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 thanan order of magnitude higher than rates measured in tuna by Weber et a!. (1986) toreflect just 50 % repletion of the white muscle glycogen by plasma glucose.The implication remains that much of the lactate formed during exercise stays inthe white muscle, or is retrieved from the plasma, and is the principle substrate forglycogen resynthesis. It is interesting that plasma lactate in the exercised tuna declinedmarkedly only after lactate and glycogen recovery in white muscle approachedcompletion. High plasma lactate likely produces a concentration gradient favorable forlactate uptake and utilization by the tissues mentioned above, but may also help toattenuate diffusive efflux of lactate from white muscle. In this regard, tuna resemblethe mammals in which prolonged elevation of plasma lactate seems necessary to sustainin situ muscle glyconeogenesis (Johnson and Bagby 1988; Stevenson et a!. 1987). Thissituation is different from other teleosts where plasma lactate is elevated onlytransiently relative to the prolonged period required for white muscle lactate andglycogen recovery.Tuna need to swim continuously and this may mean that muscular contractionsin free swimming fish influence changes in lactate, and other muscle metabolites,compared to spinally-blocked fish. There are no studies of lactate removal from whitemuscle of free swimming tuna for comparison of methodologies, but the rate ofglycogen replenishment in skipjack tuna measured by terminal sampling in recoveryfrom exercise (Barrett and Connor 1964) is comparable to changes in lactate andglycogen measured in the present study. In addition, while swimming after intenseexercise might change both the rate of lactate clearance from white muscle and whole-102body turnover, it is still likely that the principal fate of lactate in tuna is as we haveproposed since the potential to oxidize lactate is low and the proportion of red musclerelative to total muscle mass is small.Lactate and PCr. The extraordinary inverse correlation between the concentrationof PCr and the concentration of lactate has not, to our knowledge, been previouslyreported in muscle. Meyer and Terjung (1979) appeared to find a similar relationshipbetween between concentrations of PCr and lactate in the recovering gastrocnemius ofrats. However, they suggested that the recovery of phosphocreatine was biphasic,whereas the recovery of lactate was fitted best by a single exponential. A comparablerelationship between PCr and lactate has also been noted in hypoxic turtle brains (Lutzet a!. 1984).We suggest that recovery of pHi, dependent on the clearance of lactate, causesthe apparent linkage of phosphocreatine to lactate during recovery from exercise intuna. If, as proposed, lactate is a glyconeogenic substrate then there would besimultaneous consumption of protons within the white muscle. This suggestion issupported by the finding of Tang and Boutilier (1991) who found that protons producedin rainbow trout during intense exercise were most likely cleared by metabolicprocesses within the white muscle compartment.As discussed earlier, net efflux seems to account minimally for clearance oflactate from tuna white muscle. Thus an imbalance, if any, between the clearance oflactate and protons would have only a minimal effect on the relationship between pHiand lactate. Nevertheless, the efflux of lactate from tuna white muscle may also belinked with proton transport. Lactate and proton movements in muscle appear to becoupled in a carrier-mediated transport process (Mason and Thomas 1988; Juel andWibrand 1989). Furthermore, lactate transport across the mammalian sarcolemma ishighly sensitive to a pH-gradient (Roth and Brooks 1990). It is not clear to what extent103these findings can be generalized as Wiseman et al. (1989) found that the efflux ofproton and lactate equivalents was not tightly coupled in molluscan muscle tissue. Intuna there is an excess of blood lactate over blood metabolic protons after exercise, butPerry et al. (1985) noted that more efficient removal of protons from plasma, ratherthan differential lactate/proton release from white muscle, probably accounted for thisdifference. Directly measuring pHi and lactate in white muscle would resolve therelationship between the concentrations of protons and lactate. Nevertheless, theevidence is consistent with the suggestion that changes in pHi are linked with theconcentration of lactate.Phosphocreatine is linked to pHi through the near equilibrium reaction catalysedby creatine kinase where:PCr+ADP+H<---->ATP+CrIn addition to pHi, the ratio of ATP/ADP will also affect the final concentration ofPCr. Work on turtle brain and fish white muscle indicates a clear relationship betweenacidosis, or glycolysis, and PCr depletion induced by hypoxia (Lutz et a!. 1984; VanWaarde et a!. 1990). During recovery from anoxia the rephosphorylation of PCr infish muscle becomes dissociated from recovery of pHi, and presumably lactateclearance. However, during recovery from anoxia there were substantial increases inthe ratio of ATP/ADP which would dissociate pHi from PCr. Similarly, adenylateratios seem to influence PCr in rainbow trout white muscle where, in recovery fromexhaustive exercise, lactate is cleared at a much slower rate than PCr repletion (Schulteet a!. 1992). The close relationship between lactate and PCr in tuna suggests thatchanges in pHi determine the extent of phosphorylation of creatine during recovery.PUBLICATION STATUS: The preceding study has been published. See Arthur eta!. 1992 in the reference list. Thanks to Dr. P.G. Arthur for measuring PCr and forconceptual and practical collaboration in this study.104CHAPTER 4. GENERAL DISCUSSION.Glucose as a fuel for swimming. A comparison of glucose utilization in trout and carpis interesting in two respects. Firstly, as hypothesized from observations of superior fatutilization capacity in faster swimming fish (Moyes et a!. 1992b), it is evident that the slowswimming carp utilizes circulatory glucose for muscle energetics to a greater extent than dorainbow trout. As will be discussed later this interpretation assumes that red muscleaccounts for most of the increased metabolic demands at 80 % U11.j, which are lower incarp than in trout. Glucose dependency may be related largely to differences in absoluteaerobic performance capacity rather than differences in muscle glucose utilization rate.Secondly, these data, particularly the concentration-dependence of glucose turnover in carpand 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 beregulated in the whole-animal and in various muscle-types. In addition to the energeticimportance of glucose, the following discussion focuses on the significance of muscletissue 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 comparisonof 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 glucoseutilization, but also on the relative masses of the muscles involved. Muscle-typedifferences in utilization rate are in accord with observations in trout and carp indicatinghigher hexokinase (HK) activity in heart and red muscle compared white muscle (at 15 °C,approximately 5 - 10 molmingin heart and 1 - 2 molmin1gin red muscle versus0.1- 0.5 molmin1gin white muscle, Johnston 1977; Dnedzic 1987). It is likely that105muscle-type differences in glucose utilization capacity parallel differences in transportcapacity, suggesting that fish may be similar to mammals, in which the relative capacityfor 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 utilizationin fish white muscle, this muscle is a potentially significant site for circulatory glucosedisposal 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 incarp that is about 10 % of the total muscle (Johnston 1977; Johnston et al. 1977). Simplybecause of its large mass, white muscle at rest is estimated to account for a large portion ofthe glucose disposal in the carp (see table 9). In total, heart, red and white skeletal muscleare estimated to account for 50 - 70 % of glucose diappearance in resting animals. Therelative importance of each skeletal muscle type is similar because of the differences inrelative glucose uptake and relative mass of the two tissues. However, glucose disposal inred muscle was somewhat higher when turnover was low and increased in both low andhigh turnover conditions during exercise. Note that the one glycogen depleted fish(discussed earlier) was eliminated from this analysis because of the apparent dependency ofglucose uptake on muscle glycogen below levels of 10 mol glucosyl unitsg1 (Figure 6).In the more general situation, red muscle GUI was stimulated to a relatively constant levelin individual swimming carp (Figure 7), despite the concentration-dependence of whole-body turnover. Concentration effects on glucose flux in swimming carp therefore hadmore relevance to tissues other than the active red muscle. White muscle, given itspreference for carbohydrate fuels (Driedzic and Hochachka 1975; Moyes et a!. 1989),might be expected to be one such tissue. However, exercise- and concentration-effects onGUI in white muscle were not significant, even though trends (Figure 7) looked similar tothose observed for whole-body turnover (Figure 5). On the other hand, while GUI in heartmuscle was concentration-dependent (Figure 7), its small relative mass makes it an106TABLE 9. Total glucose disposal in the cardiac and skeletal muscle of carp. It isassumed that carp body mass is comprised of 66 % white muscle, 7 % red muscle andthat ventricle mass is 0.76 gkg1 (Bone 1978; Sidell et a!. 1987). The contribution ofspecific muscles to the whole-body disappearance of glucose assumes a 2-DG lumpedconstant (see text) of 0.40 for red muscle and heart (West et a!. 1993) and 0.51 forwhite 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 ofturnover rate - 5 molmin1kg’ was chosen arbitrarily to designate carp as havinglow turnover (< 5) or high turnover (5 and above). The one glycogen depleted fishwas excluded.Wet weight Glucose GUR Precentage ofof Carp Turnover Turnover(g) (nmolmin1) (nmolrnin1) Accounted forResting 1360 - 1830 3000 - 11400white 800 - 4250 26- 37red 800 -1250 26-11heart 12-56 <1Swimming 1175 - 1525 5750 - 15300white 1600-5200 28-34red 2340-2700 40-17heart 34-80 <1107insignificant disposal site for circulatory glucose (Table 9). Pink skeletal muscle was notsampled in the present study, but given its prominent mass, its glycolytic disposition andits probable recruitment at 1.3 body lengthssec1 (Johnston et a!. 1977) pink fibers couldwell have accounted for a large portion of glucose disposal in carp.Turnover rate was not measured in resting rainbow trout. However, glucose uptakein the total red muscle and heart masses (about 70 g and 1 g respecitvely) of a resting 1 kgtrout requires a rate of circulatory glucose delivery of about 80 nmolmin1 in fish at rest.The total white muscle mass (about 660 gkg1 body mass) is estimated to demandconsiderably more circulatory glucose at rest (about 300 nmolmin1,based on utilizationrates in Table 4). However, we cannot be certain what proportion of resting whole-bodyglucose turnover is accounted for by combined utilization in these tissues. Estimates ofglucose turnover range from 1 - 10 molminkgin trout with resting glucose levels of2- 5 mM (Dunn and Hochachka 1987; Garin et a!. 1987; Washburn et a!. 1992; orestimated from Figure 12, chapter 2.1). Uptake in muscle tissue at rest would thereforeaccount for 3 - 30 % of circulatory glucose turnover, suggesting that the bulk of glucosedisposal occurs in other tissues in trout. During exercise the significant increase in redmuscle glucose utilization would raise this estimate of disposal, but it is not known whetherturnover rate changes in swimming trout. The uncertainty in designating trout glucosedisposal at rest as low compared to carp arises from the possibility that glucose utilizationin trout skeletal muscle may display limited sensitivity to plasma glucose concentrationwhich would bring estimates of muscle glucose disposal in closer agreement with theestimations made for carp.Maximal glucose flux. As was discussed earlier, HK activity provides a guide to therelative capacity for glucose flux in different muscle-types. The general pattern of higherglucose utilization rates in more oxidative muscles is consistent with the trends seen in HKactivity in carp and trout muscle. Interestingly, an apparent excess in flux capacity,108relative to in vivo utilization rates, exists in all muscle-types examined. The reasons forexcess capacity to phosphorylate glucose are not clear, although such observations havebeen made before. In isolated rat hearts, }{K has been measured at 5-fold in excess ofpeak glucose oxidation rates (England and Randle 1967). It may be that these capacitiesare required for situations in which alternative fuels- those that may be inhibiting glucoseflux - are limiting, but it would appear that flux capacity is in excess of even the normalaerobic metabolic requirements of some tissues. The trout heart is one such system andprovides good example for discussion since glucose flux data is available from in vitro andin vivo experiments.Measurements of trout heart hexokinase in vitro indicate a potential glucose flux ofup to 8300 nmolg1’ in (see Driedzic et a!. 1987), more than 1000 times greater thanobserved in vivo fluxes. It is very likely however, that actual maximal in vivo fluxes arelower since cellular substrate concentrations are usually less than saturating and possiblestructural roles for enzymes may influence availability for maximal flux compared to invitro estimates (Wright and Albe 1990). It may be that the peak glucose utilization seen inthe NaCN heart perfusions (i.e., -500 nmolgmin1)more closely resembles fluxcapacity in vivo. The surprising observation is that in vivo rates of glucose utilization areso low (-5 nmolg1min’) - considerably lower, in fact, than even rates of uptake innormoxic isolated hearts (30 - 120 nmolgmin). Seemingly, both endogenous (eg.,preference for intracellular fat) and circulatory (hormone effects and circulatory fuelsupply) factors contribute to nearly complete suppression of glucose utilization in normalpost-absorptive trout. From another perspective however, the in vitro results suggest atleast the potential to expand myocardial reliance on glucose in vivo. It is relevant to askunder what conditions the apparent capacity for glucose flux might be approached in thewhole animal.Hypoxic-stress is one situation in which myocardial glucose utilization is expectedto be increased in vivo, with greater energy provision from glycolysis allowing the heart to109sustain performance in the absence of adequate oxidative ATP production. Increasedglucose utilization in the NaCN perfusions of isolated hearts is analogous to the glycolyticresponse expected, although during complete inhibition of oxidative metabolism in trouthearts can sustain only very low, subphysiological, work rates. The response toprogressive hypoxia has been examined in previous studies. In sea raven hearts, forexample, power output remains steady through the early stages of progressive hypoxia eventhough oxygen consumption declines steadily (Farrell et a!. 1985). Cardiac performancemay be protected to some extent by glycolytic stimulation until the oxygen level fallsbelow a critical point. The responses of trout heart to hypoxia are less clear, althoughthere 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 oxygenconsumption (see Boutilier et a!. 1988). A decline in heart metabolism in this case maymean that the heart is able to function aerobically and shows enhanced glycolytic activityonly at very low arterial oxygen levels. Support for this alternative possibility comes fromwhole animal responses to hypoxia in which metabolic depression occurs earlier in ahypoxic 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 ofglycolysis could greatly increase phosphorylation of glucose in an effort to support in vivocardiac function during periods of low oxygen availability.One way to increase normoxic glucose flux in vivo is to increase the availability ofglucose 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 turnoverrate (discussed next section). Interestingly, glucose uptake in carp red muscle, whileresponsive to exercise, was concentration-independent. However, it would seem unlikelythat the glucose phosphorylation process was somehow saturated in active red muscle sinceflux was able to increase considearbly when muscle glycogen content was low. Glucoseuptake and phosphorylation would seem to be regulated differently in the two muscle-110types, but the results indicate that glucose flux is indeed flexible in both muscles underdifferent in vivo conditions. Normoxic glucose fluxes probably never match in vitro fluxcapacity, perhaps because of inhibition of flux in vivo (England and Randle 1967) orintracellular compartmentation of the enzyme or reduced in vivo catalytic potential becauseof possible structural roles for enzymes (Wright and Albe 1990). HK activities that areseemingly in excess might be viewed more appropriately as reserve capacities (SeeDiamond and Hammond 1992) that are exploited for shorter term protection of cellularenergetics when oxidative metabolism is compromised.Regulation of glucose utilization. Among resting carp, blood glucose level variedconsiderably (3 - 17 mM) and glucose turnover increased with plasma glucoseconcentration. These observations suggest, firstly, that carp are typical of teleosts in notregulating resting blood glucose concentration to within rigid in vivo limits (Harmon et a!.1991; Mommsen and Plisetskaya 1991; Nagai and Shizunori 1973; Palmer and Ryman1972) and, secondly, that glucose availability itself may have been important in mediatinguptake in peripheral tissues. Substrate-mediated glucose disposal is evident in mammalianmodels that display ineffective (diabetics) or underdeveloped (neonate dogs) hormonalglucoregulation (Doberne et al. 1982; Hulman et a!. 1988; Verdonk et a!. 1981). Thismode of plasma glucose removal is consistent with a recent suggestion that amongsalmonids insulin secretion itself may be influenced by interactions with other circulatinghormones. While it is evident that insulin serves an important anabolic function in fish(Mommsen and Plisetskaya 1991), potential interactions with other glucose stimulatedhormones delay the insulin response, thereby, contributing to the apparent glucoseintolerance of salmonids (Harmon et a!. 1991). It is not known to what extent hormonalinteractions can be generalized among teleosts, nor is it clear that all species demonstratethe inability to regulate glucose within narrow limits, as evident in carp and in trout.However, proportional changes in glucose utilization with increasing glucose concentration111in both species tends to support a possibly dominant role for glucose availability in theregulation of tissue glucose disposal rate, regardless of activity state.The concentration-dependence of glucose turnover rate in carp has furtherimplications for glucoregulation in swimming fish. The difference in regression slopes forresting and swimming carp suggests that it would be effective for individual fish to becomehyperglycemic during a rest-to-work transition in order to enhance delivery of glucose tothose tissues which have an increased demand for oxidative substrate. We have no directevidence 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 wasunchanged from the pre-exercise level throughout exercise. This implies that stimulationof glucose flux in the present study most likely occurred in the absence of a direct effect ofexercise on plasma glucose concentration, supposedly via parallel up-regulation of the ratesof glucose appearance and disappearance (Wasserman et a!. 1989). Thus, potentialexpansion of glucose use in swimming carp would seem to be largely fortuitous, dependingvery much on the prevailing blood glucose status of individual fish prior to exercise. Thissituation is also consistent with having poor control over plasma glucose concentration andwith possible substrate-mediation of glucose uptake. However, the regulatory featureswhich would allow up-regulation of hepatic glucose release concomitant with increasedglucose disposal require further investigation.Exercise energetics in carp and trout. Some differences in the kinetics and utilizationof glucose between carp and rainbow trout imply that glucose is somewhat more importantfor 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 restingmetabolism. Skeletal muscle accounted for about 50 % of glucose disappearance in restingcarp (plasma glucose, 3 - 17 mM), compared to estimates of less than 20 in trout112(plasma glucose, 5 - 40 mM). Glucose in resting skeletal muscle is expected to dividedamong biosynthetic (lipid/glycogen storage) and biodegradative (oxidation/lactateproduction) pathways. While higher disposal in carp musculature suggests, at leastqualitatively, increased availability of intracellular glucose-carbon for resting muscleenergetics, the uncertainty in assigning any quantitative significance to species differencesin resting glucose utilization arises from the lack of satisfactory estimates of in vivomaintenance costs in red and white muscle.During exercise, cardiac energetics are difficult to compare directly between thetwo 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 mMand, unlike carp, an exercise-effect was not detected. Glucose accounted for < 1 % ofcardiac energetics in trout swimming at 80 % Uj (West et al. 1993). However, if theconcentration-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 todetect. Moreover, the maximal proportion of glucose that could account for estimatedheart oxygen consumption of active carp would approach estimates for the trout heart.In contrast to heart, carp red muscle glucose uptake was independent of plasmaglucose concentration and estimated glucose utilization was stimulated to similar levels incarp and trout (20- 30 nmolmin’g1). However, the smaller metabolic scope forsustained swimming in the less active carp accounted for a higher estimate of glucose usefor energy provision in carp red muscle (25 - 30 % in carp, < 10 % in trout) whileswimming at 80 % U. Obviously, comparison of glucose utilization in these species ismost useful if the assumption holds that red muscle accounts for the bulk of the activeoxygen consumption. If, in particular, aerobic recruitment of white muscle is an importantmetabolic cost during exercise, then this would have to be accounted for in calculations ofred muscle dependence on glucose. Simply stated, the nearly 2.5-fold higher aerobiccapacity of trout white muscle compared to carp (based on citrate synthase activity and113mitochondrial protein content per gram red muscle, Moyes et a!. 1989; 1992b) maycontribute 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 tendto bring estimates of glucose oxidation in closer agreement in these species. While theother 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 redmuscle, this would have minimal effect on the estimate of the use of glucose for swimmingenergetics of trout. Greater use of glucose in the less active carp would be consistent withthe generalization that aerobic exercise capacity influences the type of substrate used insupport of maximal oxidative energy provision.Post-exercise recovery of muscle glycogen. Comparison of the rate of lactateclearance, or glycogen repletion, in trout and tuna indicates a greater rate of recovery, by10- 30 times, in tuna white muscle. However, the balance between lactate clearance andglycogen 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 ratebetween trout and tuna, but clearance rate across temperate water species correlates with anumber of enzymatic indicators of tissue metabolic capacities, including glycolytic capacity(eg., suggested by pyruvate kinase (PK) activities), marker enzymes for muscleglyconeogenesis (eg., malic enzyme (ME) activities) and tissue aerobic capacity (eg.,citrate synthase activities), with the highest lactate fluxes evident in species with greatermetabolic capacities (Moyes et al. 1993). The tunas demonstrate superior capacities, evenwhen 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 lactateclearance. The relationship with aerobic capacity is unlikely to be indicatative of enhancedoxidative removal of lactate, since conservation of lactate carbon for glyconeogenesis114seems consistent across species, including the highly aerobic tuna white muscle.Interestingly, the mitochondrial capacity to produce ATP, although inhibited in vivo byhigh ATP:ADPfr ratios, is far in excess of the energetic demand for conversion of whitemuscle lactate to glycogen (Moyes et a!. 1992b). Thus, aerobic capacity would appear notto 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 tothe capacity to convert lactate to glycogen. PK and ME activities may have implicationsfor the route, and rate, of glyconeogenic flux in white muscle (next section), but it has yetto be demonstrated experimentally that the capacity for flux through these pathways limitsrecovery in vivo.Despite differences in recovery rates, teleost white muscle across species can beregarded as almost a self-contained metabolic system with respect to carbohydrate statusduring burst exercise and recovery. The dependence of white muscle on glycolysis and onintramuscular reconversion of lactate to glycogen allows gross level lactate/glycogenchanges 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 lactateclearance and the role of glucose are somewhat more difficult to place into a quantitativeframework. Apparent deficits in white muscle glycogen in trout range 0 - 50 %, troutrelative to resting control glycogen levels (Mulligan and Wood 1986b; Mommsen andHochachka 1988; Pagnotta and Milligan 1991; Pearson et al. 1990; Schulte et a!. 1992 andthis 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 tospare lactate that is retained in white muscle for primarily glyconeogenesis. Most analysessuggest 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 oxidation115(Scarabello et a!. 1991; Moyes et a!. 1992) or hepatic gluconeogenesis. Similarly,maximal losses of lactate in tuna white muscle, sampled with sequential biopsy, areexpected to be < 10 %, based on the capacities for oxidation and gluconeogenesis. Invivo rates of lactate oxidation in heart and red muscle and of possible red muscleglyconeogenesis 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 theseprocesses might explain minor mismatches between white muscle glycogen replenishmentand lactate clearance rates that are sometimes observed. In turn, low glucose uptake, asdetermined in trout, would be most useful for post-exercise glycogen restoration over thelong-term. A better understanding of the role of glucose uptake in white muscle mayemerge from studies that include specific examinations of mismatches in lactate/glycogenstoichiometries and of apparent deficits in glycogen recovery. It is expected, however, thatcirculatory glucose serves mainly as a supplement to the normal turnover of intramuscularglycogen reserves and perhaps in an energetic role for resting muscle metabolism.Muscle glyconeogenesis. These studies, like most examinations of white muscleglyconeogenesis in vertebrates, provide evidence for an intramuscular pathway of lactatereconversion to glycogen. The major gap in our understanding of this process is that thepathway 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 eliminatemitochondrial conversion of pyruvate to oxaloacetate (OAA) and subsequent shuttling ofOAA to the cytoplasm for decarboxylation to PEP since pyruvate carboxylase has not beendetected in muscle extracts (Opie and Newsholm 1967). Furthermore, there is no apparentrandomization of 14C 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 C02, GTP and the116presence 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 reduceslactate incorporation into glycogen. Conversly, the data of Pagliasotti and Donovan (1990)points to glycogen repletion independent of PEPCK activity in rabbit muscles. Thesituation in fish seems less ambiguous since PEPCK activity has not been detected in whitemuscle of species for which lactate clearance rates are known (Moyes et al. 1993),including tuna muscle where rapid carbohydrate recovery might be expected to beassociated with relatively high levels of PEPCK. The only known extramitochondrialpathway that is independent of PEPCK activity is reversal of PK. It is not known whethermass action ratios in recovering fish white muscle favor flux from pyruvate to PEP throughPK. 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 introut, Schulte et a!. 1992), would favor PK reversal. Conditions may well be similar intuna (pyruvate increases 20-fold over resting levels, for example) and it is evident thatreverse flux rates in tuna white muscle would have to equal about 0.1 % of maximal PKactivities (about 1300 molmin1g,Guppy et a!. 1979) to completely account for ratesof 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 molmin1g, Moon and Johnston 1980b) would account for lactate clearance in white muscle(Moyes et a!. 1992b). Generally, species with lower lactate clearance rates also displayreduced PK activities, thereby maintaining the need for PK reversal at 0.03 - 0.1 % of themaximal reaction velocity (Moyes and West, in press).It is not known by what mechanism trout red muscle may convert lactate carbon toPEP, although observations in rabbit muscles (Pagliassotti and Donovan 1990) indicate that117glyconeogenesis occurs by a general pathway in all muscle types. However, the capacityfor glyconeogenesis is related to oxidative capacity, with red fibers displaying the lowestlevels of lactate incorporation. In trout, the same tendency is suggested indirectly from theincreased potential of red muscle to use glucose for glycogen resynthesis after exercise.Red muscle, should be examined in comparative manner, similar to previous studies onwhite muscle, to identify regualtory features that help to distinguish lactate dynamics inrelation to aerobic capacity, glycolytic potential and fiber-type across species.Glucose and lactate in fish white muscle. Regardless of species differences in the rate ofwhite muscle lactate clearance, post-exercise lactate turnover and white muscle aerobiccapacity, the fmal fate of lactate seems restricted to glycogen restoration in fishes. Lowrates of glucose utilization in trout, demonstrate that flux is insufficient to support glycogensynthesis rates observed. In tuna, previous estimates of recovery glucose turnover, even inthe unrealistic situation of exclusive delivery to white muscle, are too low (about 15molmin’kg’, plasma glucose 2- 6 mmolL1;Weber 1986) for glycogenesis to accountfor glycogen repletion in the present study (requiring > 350 molmin1kg,assuming660 g white muscle per kg body weight).The stoichiometry between white muscle lactate and glycogen argues againstsignificant rerouting of lactate away from a glyconeogenic fate. The apparentinconsistency is that tuna display relatively high lactate turnover rates, as measured withradiolabelled lactate (Weber et al. 1986). In trout, although plasma lactate kinetics appearunrelated to white muscle glycogen replenishment, there is evidence that an initial outwardflux of lactate from white muscle is followed by inward transport to favor metabolism insitu (Turner and Wood 1983). A similar kinetic pattern in tuna would mean that whole-body turnover reflects lactate uptake from plasma by the lactate-producing as well asconsuming tissues. Simultaneous lactate extraction and removal has been observed inexercising human leg muscle, in which the extraction of tracer lactate from plasma occurs118despite net lactate release by muscle (Stanley et al. 1986). High plasma concentrations oflactate and mammal-like turnover rates would, therefore, seem to be indicative of theplasma compartment acting as a temporary storage space for lactate that is eventually takenup by mainly the white muscle mass. This kinetic pattern cannot be confirmed in thepresent tuna study, but low capacities for lactate use in other tissues and the observationthat plasma lactate remained elevated until the late stages of white muscle clearanceprovides some indirect evidence.In both trout and tuna the significance of an exercise-recovery cycle of glycogenlactate-glycogen is to conserve carbohydrate status of the white muscle. Glycogen reservesrepresent a short term fuel reserved for burst swimming and that by conserving lactate, it ispossible to conserve glycogen. The advantage, given that dependence on glycogenesisalone would greatly lengthen recovery time, is that carbohydrate recovery can occurindependently 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 energystore that can also be rapidly replenished after exercise in both trout and tuna. Rapidmetabolite recovery ensures that fish are ready to engage in another burst of high speedswimming as quickly as possible. 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