Metabolic Biochemistry, Volume 4 (Biochemistry and Molecular Biology of Fishes) (Vol 4)

Preface The idea of editing a series of volumes on The Biochemistry and Molecular Biology of Fishes was born out of the present-day lack of a forum for state-of-the-art review articles in this rapidly expanding field of research. On the one hand, researchers and students in this area always find themselves combing the literature on general (rat-dominated) biochemistry before discovering short and usually incomplete and disappointing coverage of the situation in the piscine setting. On the other hand, the rapidly expanding volume and quality of the primary literature in fish biochemistry and molecular biology supply convincing evidence for a maturing field. This discipline is no longer the younger sibling of rat or human biochemistry but has recently led to a number of major conceptual breakthroughs; for this reason, and because its activity domain is sometimes nonoverlapping with 'mainstream' biochemistry, the field is certainly ripe and ready for a review series of its own. Comparative biochemistry and molecular biology and comparative physiology as disciplines by definition use organisms as a special kind of experimental parameter for probing general mechanisms and principles of function. In theory this approach is relatively blind to phylogenetic boundaries, but in practise the realities of funding and availability of experimental material greatly narrow the field of play. As a result, two phylogenetic groups - - the insects and the fishes - - have over the last several decades provided the bulk of the experimental data base in these disciplines. Interestingly, although comparative biochemistry in many ways grew out of comparative physiology, the growth and development of these two activities in the insect field have to major extent proceeded along independent paths. By contrast, the comparative physiology and biochemistry of fishes have not been so independent of one another and the tendency has been for the former to envelope the latter. We believe that the current conceptual developments in the fields as well as the simple logistics of dealing with massive data bases make this the right time for the reality of independence to match the perception of independence, which we feel is another important rationale for this review series. Our goal is to provide researchers and students with a pertinent information source from theoretical and experimental angles. To be useful to students, theoreticians, and experimentalists alike, contributing authors are urged to emphasize concepts as well as to relate experimental results to the biology of the animals, to point out controversial issues, and todelineate as much as is possible directions for future research. Peter W. Hochachka Thomas P. Mommsen Vancouver and Victoria, B.C.

Contributors

Hiroki Abe, Department of Food Science and Nutrition, Kyoritsu Women's University, 1- 710 Motohachioji, Hachioji, Tokyo 193, Japan (Chapter 14) James S. Ballantyne, Department of Zoology, University of Guelph, Guelph, Ontario, Canada NI G 21/11 (Chapter 10) Andrew H. Bass, Section of Neurobiology and Behavior, Cornell University, Ithaca, New York 14853, USA (Chapter 12) Ralf Bastrop, Universiti~tRostock, Fachbereich Biologie, Zoologisches Institut, Universitiitsplatz 2, D-02500 Rostock 1, Germany (Chapter 7) Richard W. BriU, Southwest Fisheries Science Center, Honolulu Laboratory, National Marine Fisheries Service, National Oceanic and Atmospheric Administration, Honolulu, Hawaii 96822-2396, USA (Chapter 1) Stephen EJ. Brooks, Nutrition Research Division, Health Canada, Tunney's Pasture, Ottawa, Ontario, Canada K1A OL2 (Chapter 13) C.G. Carter, Department of Aquaculture, University of Tasmania, PO Box 1214, Lauceston, Tasmania 7250, Australia (Chapter 8) Nathan L. Collie, Department of Biological Sciences, Texas Tech University, Lubbock, Texas 79409-3131, USA (Chapter 9) Ronaldo P. Ferraris, Department of Physiology, University of Medicine and Dentistry of New Jersey,, New Jersey Medical School, Newark, New Jersey 07103-2714, USA (Chapter 9) Glen D. Foster, Department of Biology, University of Ottawa, 30 Marie Curie, Ottawa, Ontario, Canada KIN 6N5 (Chapter 4) Edward M. Goolish, National Oceanic and Atmospheric Administration, Southwest Fisheries Science Center, La Jolla, California 92038, USA and Scripps Institution of Oceanography, Center for Marine Biotechnology and Biomedicine, University of California, San Diego, La Jolla, California 92093, USA (Chapter 15) Joaquim Guti6rrez, Departament de Bioquimica i Fisiologia, Universitat de Barcelona, Unitat de Fisiologia Animal F, Av. Diagonal, 645, E-08071 Barcelona, Spain (Chapter 17) Peter W. Hochachka, Department of Zoology, University of British Columbia, Vancouver, British Columbia, Canada V6T 2A9 (Chapter 1)

viii

Contributors

D.E Houlihan, Department of Zoology, University of Aberdeen, Aberdeen, TiUydrone Avenue, Aberdeen AB9 2TN, Scotland, UK (Chapter 8) Karl Jiirss, Universiti~t Rostock, Fachbereich Biologie, Zoologisches Institut, Universitiitsplatz 2, 1)-02500 Rostock 1, Germany (Chapter 7) Odile Mathieu-Costello, Department of Medicine, Universityof California, San Diego, La Jolla, California 92093-0623, USA (Chapter 1)

I.D. McCarthy, Department of Zoology, University of Aberdeen, Aberdeen, Tillydrone Avenue, Aberdeen AB9 2TN, Scotland, UK (Chapter 8) Thomas P. Mommsen, Department of Biochemistry and Microbiology, University of

Victoria, P.O. Box 3055, Victoria, British Columbia, Canada VSW3P6 (Chapter 12) 9Thomas W. Moon, Department of Biology, University of Ottawa, 30 Marie Curie, Ottawa, Ontario, Canada KIN 6N5 (Chapter 4) Christopher D. Moyes, Department of Biology, Queen's University, Kingston, Ontario, Canada K7L 3N6 (Chapter 16) Isabel Navarro, Departament de Bioquimica i Fisiologia, Universitat de Barcelona,

Unitat de Fisiologia Animal F,, Av. Diagonal, 645, E-08071 Barcelona, Spain (Chapter 17) Bernd Pelster, lnstitut ffir Physiologie, Ruhr-Universitdt Bochum, D.44780 Bochum,

Germany (Chapter 5) Jean-Francois Rees, Laboratory of Animal Physiology, Catholic University of Louvain,

Croix du Sud 5, B-1348 Louvain-la-Neuve, Belgium (Chapter 18) Kenneth B. Storey, Departments of Biology and Chemistry, Carleton University,

Ottawa, Ontario, Canada KIS 5B6 (Chapter 13) Eric M. Thompson, Laboratoire de Biologie Cellulaire, Unit~ de Biologie du D~veloppement, Institut National de la Recherche Agronomique, F-78352 Jouy-en-Josas, France (Chapter 18) Guido van den Thillart, Institute of Evolutionary and Ecological Sciences, Animal

Physiology, Gorlaeus Laboratories, University of Leiden, PO Box 9502, 2300 RA Leiden, The Netherlands (Chapter 3) Douglas R. Tother, NERC Unit of Aquatic Biochemistry, School of Natural Sciences,

University of Stirlinb StirlingFK9 4LA, Scotland, UK (Chapter 6) Marcel van Raaij, Institute of Evolutionary and Ecological Sciences, Animal Physiology, Gorlaeus Laboratories, University of Leiden, PO Box 9502, 2300 RA Leiden, The Netherlands (Chapter 3) Patrick J. Walsh, Marine Biology and Fisheries Division, Rosenstiel School of Marine and Atmospheric Sciences, Universityof Miami, 4600 Rickenbacker Causeway, Miami, Florida 33149-1098, USA (Chapter 12)

Contributors

ix

Jean-Michel Weber, Biology Department, University of Ottawa, 30 Marie Curie, Ottawa, Ontario, Canada KIN 6N5 (Chapter 2) Timothy G. West, Department of Zoology, Cambridge University, Downing Street, Cambridge, CB2 EJ3, UK (Chapter 16) Harold H. Zakon, Department of Zoology, Patterson Laboratory, The University of Texas, Austin, Texas 78712, USA (Chapter 11) Georges Zwingelstein, Laboratoire Maritime de Physiologie, Institut Michel Pacha, Universit~ de Lyon, 1337 Corniche Michel Pacha, Tamaris, F-83500 La Seyne sur Mer, France (Chapter 2)

Abbreviations

Amino acid(s) Acetylcholine receptor Adrenocorticotropic hormone Alanine aminotransferase Aldolase Ammonia quotient Atlantic salmon cell line Aspartate aminotransferase Brushborder membrane vesicles Branched-chain amino acid aminotransferase BCKAD Branched-chain a-ketoacid dehydrogenase Bluegill fry cell line BF-2 Immunoglobulin binding protein BiP Basolateral membrane vesicles BLMV Y,5'-cyclic adenosine-monophosphate cAMP Cytochrome C oxidase CCO CHSE-214 Chinook salmon epithelium cell line Creatine phosphokinase CPK Carnitine palmitoyl transferase CPT Citrate synthase CS Diacylglycerol DAG 5a-Dihydrotestosterone DHT Dimethylformamide DMF Dimethylsulfoxide DMSO 17fl-Estradiol E2 EAA Essential amino acid(s) Ethylenediaminetetraacetic acid EDTA Electric organ EO Electric organ discharge EOD Erythropoietin EPO Free amino acid(s) FAA Fatty acid binding protein FABP Fructose 1,6-bisphosphatase FBPase Free fatty acid(s) FFA Fast glycolytic (muscle fiber) FG Fathead minnow cell line FHM Fast oxidative glycolytic (muscle fiber) FOG Glucose 6-phosphatase G6Pase Glucose 6-phosphate dehydrogenase G6PDH Gamma-aminobutyrate GABA GAPDH Glyceraldehyde 3-phosphate dehydrogenase Glutamate dehydrogenase GDH Glucagon-like peptide GLP Glycogen phosphorylase GPase oeGPDH ~-Glycerophosphate dehydrogenase Glycogen synthase GSase High density lipoproteins HDL 12-Hydroxyeicosapentaenoate HEPE 12-Hydroxyeicosatetraenoate HETE AA AChR ACI'H AIaAT ALD AQ AS AspAT BBMV BCAAT

HK HPLC HSP IDL LCAT LDH LDL LT LX ME MT NEAA NMJ NMR ODC PAF PC PCA PCr PDG 6PGDH PEPCK PFK-I PG PG I PGK PK PKA PKC PMN PtdA PtdCho PtdEtn Ptdlns PtdSer PUFA RQ RT-2 RTG SDA SO T3 TAG TF TPI TRH TX VHDL VLDL XDH XO

Hexokinase High performance liquid chromatography Heat-shock protein Intermediate density lipoproteins Lecithin:cholesterol acyl transferase Lactate dehydrogenase Low density lipoproteins Leukotrienes Lipoxins Malic enzyme 17oe-Methyltestosterone Non-essential amino acids Neuromuscular junction Nuclear magnetic resonance Ornithine decarboxylase Platelet activating factor Pyruvate carboxylase Perchloric acid Phosphocreatine Phosphate-dependent glutaminase 6-Phosphogluconate dehydrogenase Phosphenolpyruvate carboxykinase Phosphofructokinase- 1 Prostaglandins Phosphoglucose isomerase Phosphoglycerate kinase Pyruvate kinase Protein kinase A Protein kinase C Pacemaker nucleus Phosphatidic acid Phosphatidylcholine Phosphatidylethanolamine Phosphatidylinositol Phosphatidylserine Polyunsaturated fatty acids Respiratory quotient Rainbow trout germ cell line Rainbow trout gonad cell line Specific dynamic action Slow oxidative (muscle fiber) 3,5,3? -Triiodo-L-thyronine Triacylglycerol Turbot fin cell line Triosephosphate isomerase thyrotropin releasing hormone Thromboxanes Very high density lipoproteins Very low-density lipoprotein Xanthine dehydrogenase Xanthine oxidase

Hochachka and Mommsen (eds.), Biochemistry and molecular biology of fishes, vol. 4 9 1995 Elsevier Science B.V. All rights reserved. CHAPTER 1

Design for a high speed path for oxygen: tuna red muscle ultrastructure and vascularization ODILE MATHIEU-COSTELLO, RICHARD W. ]]RILL * AND PETER W. HOCHACHKA **

Department of Medicine, University of California, San Diego, La JoUa, CA 92093-0623, U.S.A., * Southwest Fisheries Science Center, Honolulu Laboratory, National Marine Fisheries Service, National Oceanic and Atmospheric Administration, Honolulu, HI 96822-2396, U.S.A. and **Department of Zoology, University of British Columbia, Vancouver, B.C., Canada V6T 2,49

I. Introduction II. Materials and methods 1. Animals 2. Tissue preparation 3. Morphometry III. Results and discussion Acknowledgements IV. References

I. Introduction Because it is one of the most aerobic muscles in fish, the red muscle of tuna is of particular interest to study strategies and constraints in structural designs for high 02 flux from capillary to muscle fiber mitochondria. Tuna can maintain extremely high aerobic metabolic rates and reach high swimming speeds 4. The tuna red muscle is well known to operate at higher than ambient water temperature by conserving heat via the central counter-current heat exchange (for review, see ref. 36), and white muscle lactate turnover rates after exercise are known to be closer to those found in mammals than in other fish 1'39. In this chapter, we summarize our morphometric findings on the three-dimensional arrangement of the capillary network and its relationships with fiber ultrastructure in red muscle of skipjack tuna, Katsuwonus pelamis, in comparison to highly aerobic skeletal muscles of birds and mammals. Muscles designed for high sustainable activity (hummingbird and bat flight muscles as well as the red muscle of tuna) are all composed of only one population of very highly aerobic fibers, instead of the mosaic of fiber types with different metabolic pattern found in the vast majority of skeletal muscles. This homogeneity allows one to specifically examine capillary-fiber geometrical relationships across species, in particular vascular supply in relation to muscle

2

O. Mathieu-CosteUo, R.W. Brilland RW. Hochachka

fiber aerobic capacity in cases of very high demand for 02 flux. As summarized further in this chapter, previous studies showed striking similarities in structural design for high 02 flux in hummingbird and bat flight muscles despite several differences in capillary-fiber geometry2s,29. In fish as in birds, red blood cells are nucleated and less deformable than mammalian red cells, but they can be larger than bird red cells, and fishes operate at different body temperature than both birds and mammals. Thus, it is of particular interest: (1) to examine capillary-fiber structural arrangement in the red muscle of one of the most athletic fishes known; and (2) to compare it with that in highly aerobic skeletal muscles of birds and mammals.

II. Materials and methods While the details of methods used here have been described elsewhere 22, it is important to briefly highlight aspects that are relevant to properly explain the results.

1. Animals Five Skipjack tuna (Katsuwonus pelamis); body mass 1.5-2 kg; fork length 43-44 cm) were purchased from local commercial fishermen and held in outdoor 10 m diameter holding tanks supplied with continuously flowing seawater (25 4- 1~ at the Kewalo Research Facility (National Marine Fisheries Service, Honolulu, Hawaii).

2. Tissuepreparation After the tunas had been netted and anesthetized, muscle peffusion fixation with glutaraldehyde fixative (four animals) or infusion with Batson's casting material (one animal) were performed following procedures and subsequent tissue processing described elsewhere in detail22. Transverse and longitudinal sections (1 /zm thick) of perfusion-fixed tissue were used for light microscopy morphometry of capillarity and fiber size. Ultrathin transverse sections (50-70 nm) were examined with a Zeiss 10 transmission electron microscope and sampled for morphometry of fiber ultrastructure. Samples injected with casting material were examined with a Stereoscan 360 scanning electron microscope (Cambridge Instrument).

3. Morphometry Sarcomere length was measured on longitudinal sections, after careful control of the angle of each section 19. Fiber cross-sectional area, capillary diameter and capillary number around a fiber were measured on transverse sections with an image analyzer. Capillary numbers per fiber sectional area in transverse and longitudinal sections were collected by point-counting, and the data were used to estimate

Design for a high speed path for oxygen: tuna red muscle ultrastructure and vascularization

3

the degree of orientation of capillaries and capillary length per fiber volume 18. Capillary-to-fiber ratio (i.e. capillary number per fiber number) was computed as the product of capillary density (i.e. number per fiber cross-sectional area) and mean fiber cross sectional area. Capillary surface per fiber volume was obtained by intersection-counting on vertical (i.e. longitudinal) sections using a cycloid grid 2. Capillary-to-fiber perimeter ratio in transverse section, which is an index of the size of the capillary-fiber interface 25 was measured by intersection-counting in transverse sections 21, and capillary surface per fiber surface estimated as the product of capillary-to-fiber perimeter ratio and an orientation coefficient c'(K',O) as described elsewhere 25. The volume of mitochondria per volume of muscle fiber was estimated by standard point-counting 22, and mitochondrial volume per/zm fiber length calculated as the product of mitochondrial volume density and fiber cross-sectional area. Where appropriate, data on fiber size and capillary density were normalized to sarcomere length, in order to compare morphological data between muscles, independent of the particular length at which each sample was fixed and therefore examined. A normalizing sarcomere length of 2.1/zm was chosen because it is in the mid-range of the sarcomere lengths where maximal tension is developed in skeletal muscles, and it is within the range of operating sarcomere lengths in hindlimb muscles of mammal during terrestrial locomotion (range, 1.7-2.7 ~m) 6, wing muscles of bird during wing beat cycle (1.7-2.3/zm) 5 and red muscle in fish during swimming at slow speed (1.9-2.2/zm) 35.

III. Results and discussion Figure la-c illustrates the high capillary density, small fiber size and high mitochondrial volume density previously reported in red muscle of tuna 3,1~ In longitudinal sections (Fig. lb), we found a large number of capillaries cut in transverse or oblique section, as well as branches running perpendicular to the muscle fiber axis. This suggested the presence of capillary manifolds in tuna red muscle, as previously found in the highly aerobic pectoralis muscle of pigeon 2~ Figure 2a,b illustrate the remarkable similarity between the appearance of capillary manifolds in tuna red muscle (Fig. 2a) and pigeon pectoralis muscle (Fig. 2b). In that study, Potter and coworkers 34 showed that these capillary branches oriented perpendicular to the muscle fiber axis are venular capillaries which form dense manifolds around groups of muscle fibers. The examination of microcorrosion casts of tuna red muscle also showed that capillaries form a dense envelope of blood around muscle fibers (Fig. 2c). The functional implications of the particular arrangement of venular capillaries in those muscles are not fully understood. Capillary manifolds could facilitate an increased vascular supply to and from the muscle fibers at the venular end of the network where substrates and 02 content are lowest and metabolite concentration highest. They could also be related to other functional aspects such as heat dissipation and/or the blood pumping action of the muscle during flight in

4

O. Mathieu-Costello, R.W. BriU and RW. Hochachka

Fig. 1. Fine structure of tuna red muscle, a and b: light micrographs of portions of muscle bundles in transverse and longitudinal sections, respectively, c: electron micrograph of transverse section of muscle fibers and adjacent capillaries (c). Capillaries are empty after the fixation by vascular perfusion. Note large capillary density and small fiber size (a-c), large number of capillary branches running perpendicular to the muscle fiber axis (b) and high density of mitochondria, M (c). From ref. 22.

birds. Interestingly, however, capillary manifolds were found in flight muscle of hummingbird ~, but not in bat 24,29. The fact that they were found in tuna red muscle also suggest possible rheological implications since in fish, as in bird, red blood cells are nucleated and less deformable than mammalian red cells. Another possibility in tuna is transfer of heat from the muscle at the venular end of the network, as it possibly favors heat removal in bird flight muscle 2~ Table 1 summarizes morphometric data on capillarity and fiber ultrastrueture in red muscle of tuna compared with tuna white muscle, and aerobic muscles of birds and mammals with large differences in aerobic capacities. In tuna red muscle, fiber cross-sectional area was small (~500/~m 2) but not as small as in ultimate cases of high aerobic capacity in bird and mammal. In hummingbird and bat flight muscles, average fiber cross-sectional area was ~200 and 300/~m 2, respectively, in tissues similarly prepared. Note that the number of capillaries per number of fibers was similar in tuna red muscle and hummingbird flight muscle (~1.6). However,

Design for a high speed path for oxygen: tuna red muscle ultrastructure and vascularization

5

Fig. 2. Examples of capillary manifolds, a: light micrograph in a longitudinal section of tuna red muscle. b and c: scanning electron micrographs of vascular corrosion casts examined perpendicular to the surface of the manifold in pigeon flight muscle (b) and in cross-section in tuna red muscle (c). Note the remarkable similarity between the appearance in tuna (a) and pigeon (b) muscles, and the dense envelope formed by capillaries around muscle fibers (c). Based on fiber dimensions, two muscle fibers (A and B) could be contained in the empty space in c. From refs. 22 (a,c) and 34 (b).

because of the difference in fiber size, there was a huge difference in capillary numerical density between the muscles. The number of capillaries per mm 2 fiber cross-sectional area at 2.1/zm sarcomere length was 3400 in tuna red muscle and 8000 in flight muscle of hummingbird. Capillary length density is an important estimate of capiUarization which accounts for capillary geometry, and determines capillary volume and surface area available for exchange per unit volume of fiber and mitochondria. Figure 3 shows estimates of the degree of capillary orientation, expressed as the percentage added

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Sarcomere length,/~m Fig. 3. Plot of the degree of orientation of capillaries (expressed as the percentage added to capillary length by tortuosity and branching, compared with straight, unbranched capillaries oriented parallel to the muscle fiber axis) against sarcomere length in red muscle of tuna (solid circle) compared with group mean values (:I:SE) in highly aerobic muscles of birds and mammals. From refs. 22 (tuna), 28 (hummingbird) and 29 (bat). Relationships in rat hindlimb (solid line) and pigeon pectoralis muscles (broken line) are from refs. 25 and 20, respectively. to capillary length by tortuosity and branching, against sarcomere length in the muscles listed in Table 1. It is now well established that both fiber size 7,11,26 and the degree of orientation of capillaries 19'27,33 are functions of sarcomere length. As a consequence, capillary density in transverse sections can underestimate capillary length per fiber volume by a different percentage (e.g. 10-70% in rat muscles; see solid line in Fig. 3) depending on the sarcomere length at which samples are fixed and therefore examined. Figure 3 also shows that the degree of orientation of capillaries can vary between muscles and/or animals, for example bat flight muscle compared with rat soleus, or bird compared with mammal. Therefore, it may not be appropriate to compare muscle capillarity based on capillary densities in transverse sections alone even if all samples are fixed at the same sarcomere length 2~ It is interesting to note that in tuna red muscle, the contribution of tortuosity and branching to capillary length was similar to that in rat muscles (Fig. 3) in spite of the different geometry. In contrast to tuna and bird muscles 22,28 skeletal muscles of mammals showed no evidence of capillary manifolds 29. The different contribution of capillary tortuosity and branching to capillary length in tuna red muscle compared with bird muscles (Fig. 3) may be related to differences in fiber size, yielding differences in length and/or number of branches in manifolds between the muscles. In addition, capillary length density in tuna red muscle and pigeon pectoralis muscle were remarkably similar (4100 mm/mm 3) in spite of the different degree of capillary orientation, capillary-to-fiber ratio and fiber size. It was less than half the capillary length density seen in ultimate cases such as the flight muscles of hummingbird and small bats (Table 1).

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Our measurements of fiber size in red and white muscle of skipjack tuna are within the range of values reported by others, although direct comparison is often difficult because of differences in tissue preparation or because sarcomere length is not reported. To our knowledge, capillary density (e.g. capillary-to-fiber ratio and number per fiber cross-sectional area) or geometry in tuna red muscle had never been reported prior to our studies. Comparison with data in red muscle of other fishes 8,17,3~ revealed that neither was fiber size the smallest, nor capillary number around a fiber or capillary density the highest in red muscle of tuna. Similarly, mitochondrial volume density in tuna red muscle (28.5-35%; this study and ref. 16) was high, but not the highest, for fish muscle. The highest mitochondrial volume density for fish (45.5%) has been reported in red muscle of anchovy 17. The comparison of capillary length per fiber volume at a given mitochondrial volume density showed that values in tuna red muscle were as great as in mammalian heart and about half those in highly aerobic muscles of bird and mammals (Fig. 4). For example, capillary length per fiber volume at 30% mitochondrial volume density was 4300 mm -2 in tuna red muscle 22 compared with 7600 mm -2 in bat and rat muscles 29. It is interesting to note that in flight muscle of bird (hummingbird and pigeon), capillary length per unit volume of mitochondria was similar to that in bat and rat hindlimb (Fig. 4). There were about 25 km capillaries per ml of mitochondria in those muscles compared to 14 km in tuna red muscle. The different capillary geometry does not account for the different relationship between capillary length per fiber volume and mitochondrial volume density in tuna compared with

Design for a high speed path for oxygen: tuna red muscle ultrastructure and vascularization

9

highly aerobic muscles of birds and mammals (capillary manifolds were found both in tuna red muscle and bird flight muscle). On average, about one third of fiber mitochondrial volume was subsarcolemmal in tuna red muscle. This fraction was less than in flight muscles and more than in rat soleus, where subsarcolemmal mitochondria represented about one half and less than one fifth of the fractional volume of mitochondria, respectively (Table 1). In other words, comparison of highly aerobic muscles in fish, bird and mammal shows that the proportion of subsarcolemmal mitochondria is not greater in muscle with greater fiber size. Rather the opposite is observed, bat and hummingbird flight muscles (with the smallest fiber size) showing the greatest relative proportion of subsarcolemmal mitochondria. Interestingly the red muscle of anchovy, with the greatest reported volume density of mitochondria for fish skeletal muscle (45.5%), also showed a much greater fiber cross-sectional area (1115 /tm2; ref. 17) than tuna and other highly aerobic muscles (Table 1). This also indicated that intrafiber diffusion distances to mitochondria are not necessarily reduced in highly aerobic muscles of fish. A relatively large proportion of subsarcolemmal mitochondria (25% of total mitochondrial fractional volume) was found in tuna white muscle (Table 1). It was similar to that in bat hindlimb and almost as large as in pigeon pectoralis and tuna red muscle, i.e. muscles with much smaller fiber size and much greater proportion of interfibrillar mitochondria than in white muscle of tuna. Thus, a great ratio of subsarcolemmal relative to interfibrillar mitochondria is not necessarily a characteristic of highly aerobic muscles. Another important parameter to consider when assessing the three-dimensional arrangement of capillaries relative to the muscle fibers and its impact on the geometry of blood-tissue exchange, is capillary-fiber surface. Traditionally, muscle potential for 02 flux had been viewed in terms of intercapillary and diffusion distances. In contrast, recent experimental and theoretical evidence (see ref. 13 for review) suggested an important role of the capillary-fiber interface in determining 02 flux rates in working red muscles. Cryomicrospectroscopy measurements of myoglobin saturation in quick-frozen red muscles have shown that the major pO2 drop from capillary into a cross-section through the muscle fiber occurs within a few microns subjacent to the capillary and further decline towards the center of the fiber is very shallow because of myoglobin facilitated diffusion 9. In this context, capillary-to-fiber surface, i.e. the size of the capillary-fiber interface, is an aspect of capillary-fiber structure which needs to be also considered when assessing muscle capacity for 02 flux from capillary to fiber mitochondria. As pointed out by Sullivan and Pittman 3s, matching 02 supply and demand in muscles can be achieved by nature via different strategies. It can change fiber size (which affects capillary surface per fiber volume) or capillary-fiber contact area (i.e. capillary-fiber surface) or both. Figures 5 and 6 show the relationships between capillary surface per fiber volume and mitochondrial volume density (Fig. 5) and capillary-fiber surface and mitochondrial volume per unit length of fiber (Fig. 6) in red muscle of tuna compared with highly aerobic muscles of bird and mammal. Capillary surface density at a given volume density of mitochondria was smaller in tuna red muscle

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250

fiber length

Fig. 6. Plot of capillary surface per fiber surface against mitochondrial volume per unit length of muscle fiber (i.e. mitochondrial volume density multiplied by fiber cross-sectional area) in tuna red muscle (solid circle) compared with rat soleus (open diamond), flight muscle of hummingbird (solid triangle) and bat (solid diamond) and group mean value (:I:SE) in bat hindlimb. From refs. 22 (tuna), 28 (hummingbird) and 29 (bat and rat).

Design ]'or a high speed path ]:or oxygen: tuna red muscle ultrastructure and vascularization

11

(Fig. 5). This was due to the smaller capillary length density in tuna (Fig. 4) while capillary diameter was similar ("4/xm) among muscle groups. It is also interesting to note the similar capillary surface per unit volume of mitochondria in highly aerobic flight muscles (bat and hummingbird) and in bat hindlimb and rat soleus muscles ('-'3400 cm 2 per ml of mitochondria). In comparison, the value in tuna red muscle was only "~1800 cm 2 (Fig. 5). In contrast, capillary surface per fiber surface at a given mitochondrial volume per unit length of fiber was similar in tuna red muscle and rat M. soleus and it was about half that in the flight muscles of bat and hummingbird (Fig. 6). Interestingly, the ratio between capillary-to-fiber surface and mitochondrial volume per unit length of fiber in the most highly aerobic muscle in fish, i.e. the red muscle of anchovy (calculated from ref. 17; see ref. 22) was also close to that in tuna red muscle and it was more than half those in flight muscles of bat and hummingbird. This suggests consistent differences in the size of the capillary-to-fiber interface relative to the mitochondrial volume to be supplied per unit length of fiber in extremely highly aerobic muscle of fish compared with bird and mammal. The greater capillary-fiber surface ratio in flight muscles at a given mitochondrial volume per unit length of fiber suggests an increased capacity for 02 flux. It is consistent with the greater respiratory rates of mitochondria in flying hummingbirds 37 (7-10 ml 02 per ml mitochondria per min) compared with locomotory muscles of mammals running at VO2max (ref. 15) (5 ml O2/ml mitochondria/min). It supports the idea of an important role of the capillary-fiber interface in determining 02 flux rates in working red muscles 9. In tuna red muscle, capillary-fiber surface at a given volume of mitochondria per unit length of fiber was similar to that in rat soleus (Fig. 6), in spite of the lower capillary surface per unit volume of mitochondria in tuna (Fig. 5). Measurements of maximal respiratory rates of tuna red muscle mitochondria in vitro 31, yielded estimates of maximal in vivo mitochondrial respiratory rates at least 3-5 times lower in tuna red muscle than in mammals 22. The reason for this difference is not fully understood. The differences in operating temperatures between the muscles could play a role, since accounting for plausible Q l0 values yield maximal respiratory rates in tuna close to those in mammal 22. However, other explanations are also possible including the up-regulation of protein and amino acid metabolism in fish muscle compared with other vertebrates 12 which may require greater mitochondrial volume densities for the enzymes of amino acid and protein turnover. Both substrate and heat transfer may also require an increased capillary-fiber surface in tuna independently of 02 transfer per se 22. In summary, examination of capillary-to-fiber geometry in tuna red muscle displays both similarities and differences with features found in the most highly aerobic muscles of birds and mammal. Three features seem prominent in the design for high flux paths for oxygen: (1) small fiber size; (2) high capillary density; and (3) high mitochondrial density, but in tuna these are not as pronounced as in hummingbird and bat flight muscles. Additionally, a particular arrangement of capillary manifolds seem required in birds and tuna but not in mammals. Perhaps because of constraints of function at different temperatures, capillary length per unit volume of mitochondria is substantially shorter in red muscle of tuna than in

12

O. Mathieu-Costello, R. W. Brill and R W. Hochachka

skeletal muscles of both bird and mammal over a wide range of aerobic capacities. Similarly, capillary-to-fiber surface appears to be systematically smaller in highly aerobic muscles of fish than in flight muscle of birds and mammals for the volume of mitochondria to be supplied per unit length of fiber. Whether those differences are related to differences in mitochondrial properties or capillary function or both, remains to be determined.

Acknowledgements. Supported by Grant 5PO1 HL-17731 from the National Institutes of Health, U.S.A.

II4. References 1. Arthur, P.G., T.G. West, R.W. Brill, P.M. Schulte and P.W. Hochachka. Recovery metabolism of skipjack tuna (Katsuwonus pelamis) white muscle: rapid and parallel changes in lactate and phosphocreatine after exercise. Can. I. Zool. 70: 1230-1239, 1992. 2. Baddeley, A.J., H.J.G. Gundersen and L.M. Cruz-Orive. Estimation of surface area from vertical sections./. Microsc. 142: 259-276, 1986. 3. Bone, Q. Myotomal muscle fiber types in Scomber and Katsuwonus. In: The Physiological Ecolo~ of Tunas, edited by G.D. Sharp and A.E. Dizon, New York, Academic Press, pp. 183-205, 1978. 4. Bushnell, P.G. and R.W. Brill. Responses of swimming skipjack (Katsuwonus pelamis) and yellowfin (Thunnus albacares) tunas to acute hypoxia, and a model of their cardiorespiratory function. PhysioL Zool. 64: 787-811, 1991. 5. Cutts, A. Sarcomere length changes in the wing muscles during the wing beat cycle of two bird species, l. Zool. London (.4) 209: 183-185, 1986. 6. Dimery, N.J. Muscle and sarcomere lengths in the hind limb of the rabbit (Otyctolagus cuniculus) during a galloping stride, l. Zool. London (tl) 205: 373-383, 1985. 7. Dulhunty, A.E and C. Franzini-Armstrong. The relative contributions of the folds and caveolae to the surface membrane of frog skeletal muscle fibres at different sarcomere lengths. I. Physiol. (London) 250: 513-539, 1975. 8. Dunn, J.E, W. Davison, G.M.O. Maloiy, P.W. Hochachka and M. Guppy. An ultrastructural and histochemical study of the axial musculature in the African lungfish. Cell Tissue Res. 220: 599-609, 1981. 9. Gayeski, T.E.J. and C.R. Honig. O2 gradients from sarcolemma to cell interior in red muscle at maximal VO2. Am. I. Physiol. 251: H789-H799, 1986. 10. George, J.C. and E.D. Stevens. Fine structure and metabolic adaptation of red and white muscles in tuna. Env. Biol. Fish. 3: 185-191, 1978. 11. Gray, S.D. and E.M. Renkin. Microvascular supply in relation to fiber metabolic type in mixed skeletal muscles of rabbits. Microvasc. Res. 16: 406-425, 1978. 12. Hochachka, P.W. and G.N. Somero. Biochemical Adaptation, New Jersey, Princeton University Press, 1984. 13. Honig, C.R., Gayeski, T.E.J. and Groebe, IC Myoglobin and oxygen gradients. In: The Lung, edited by R.G. Crystal, J.B. West, P.J. Barnes, N.S. Cherniack and E.R. Weibel. New York: Raven Press, p. 1489-1496, 1991. 14. Hoppeler, H. and S.R. Kayar. Capillarity and oxidative capacity of muscles. News Physiol. Sci. 3: 113-116, 1988. 15. Hoppeler, H. and S.L. Lindstedt. Malleability of skeletal muscle in overcoming limitations: structural elements. J. Exp. BioL 115: 355-364, 1985. 16. Hulbert, W.C., M. Guppy, B. Murphy and P.W. Hochachka. Metabolic sources of heat and power in tuna muscles. I. Muscle fine structure, l. Exp. Biol. 82: 289-301, 1979. 17. Johnston, I.A. Quantitative analyses of ultrastructure and vascularization of the slow muscle fibres of the anchovy. Tissue Cell 14: 319-328, 1982. 18. Mathieu, O., L.M. Cruz-Orive, H. Hoppeler and E.R. Weibel. Estimating length density and quantifying anisotropy in skeletal muscle capillaries. I. Microsc. 131: 131-146, 1983.

Design for a high speed path for oxygen: tuna red muscle ultrastructure and vascularization

13

19. Mathieu-Costello, O. Capillary tortuosity and degree of contraction or extension of skeletal muscles. Microvasc. Res. 33: 98-117, 1987. 20. Mathieu-Costello, O. Morphometric analysis of capillary geometry in pigeon pectoralis muscle. Am. J. Anat. 191: 74-84, 1991. 21. Mathieu-Costello, O. Morphometry of the size of the capillary-to-fiber interface in muscles. Adv. Exp. Med. Biol, 345: 661-668, 1994. 22. Mathieu-Costello, O., P.J. Agey, R.B. Logemann, R.W. Brill and P.W. Hochachka. Capillary-fiber geometrical relationships in tuna red muscle. Can. J. Zool. 70: 1218-1229, 1992. 23. Mathieu-CosteUo, O., P.J. Agey, R.B. Logemann, M. FIorez-Duquet and M.H. Bernstein. Effect of flying activity on capillary-fiber geometry in pigeon flight muscle. Tissue Cell, 26: 57-73, 1994. 24. Mathieu-Costello, O., P.J. Agey and J.M. Szewczak. Capillary-fiber geometry in pectoralis muscles of one of the smallest bats. Respir. Physiol., 95: 155-169, 1994. 25. Mathieu-Costello, O., C.G. Ellis, R.E Potter, I.C. MacDonald and A.C. Groom. Muscle capillaryto-fiber perimeter ratio: morphometry. Am. J. Physiol. 261: H 1617-H 1625, 1991. 26. Mathieu-Costello, O., D.C. Poole and R.B. Logemann. Muscle fiber size and chronic exposure to hypoxJa.Adv. Exp. Med. Bio1248: 305-311, 1989. 27. Mathieu-Costello, O., R.E Potter, C.G. Ellis and A.C. Groom. Capillary configuration and fiber shortening in muscles of the rat hindlimb: correlation between corrosion casts and stereological measurements. Microvasc. Res. 36: 40-55, 1988. 28. Mathieu-Costello, O., R.K. Suarez and P.W. Hochachka. Capillary-to-fiber geometry and mitochondrial density in hummingbird flight muscle. Respir. Physiol. 89:113-132, 1992. 29. Mathieu-Costello, O., J.M. Szewczak, R.B. Logemann and P.J. Agey. Geometry of blood-tissue exchange in bat flight muscle compared with bat hindlimb and rat soleus muscle. Am. J. Physiol. 262: R955-R965, 1992. 30. Mosse, P.R.L. The distribution of capillaries in the somatic musculature of two vertebrate types with particular reference to teleost fish. Cell Tissue Res. 187: 281-303, 1978. 31. Moyes, C.D., O. Mathieu-Costello, R.W. Brill and P.W. Hochachka. Mitochondrial metabolism of cardiac and skeletal muscles from a fast (Katsuwonus pelamis) and a slow (Cyprinus carpio) fish. Can. J. Zool. 70: 1246-1253, 1992. 32. Poole, D.C. and O. Mathieu-Costello. Analysis of capillary geometry in rat sub-epicardium and sub-endocardium. Am. J. Physiol. 259: H204-H210, 1990. 33. Potter, R.E and A.C. Groom. Capillary diameter and geometry in cardiac and skeletal muscle studied by means of corrosion casts. Microvasc. Res. 25: 68-84, 1983. 34. Potter, R.E, O. Mathieu-Costello, H.H. Dietrich and A.C. Groom. Unusual capillary network geometry in a skeletal muscle, as seen in microcorrosion casts of M. pectoralis of pigeon. Microvasc. Res. 41: 126-132, 1991. 35. Rome, L.C. and A.A. Sosnicki. The influence of temperature on mechanics of red muscle in carp. J. Physiol. (London) 427: 151-169, 1990. 36. Stevens, E.D. and Neill, W.H. Body temperature relations of tunas, especially skipjack. In: Fish Physiology. Vol. VII, Locomotion, edited by W.S. Hoar and D.J. Randall, New York, Academic Press, p. 315-359, 1978. 37. Suarez, R.K., J.R.B. Lighton, G.S. Brown and O. Mathieu-Costello. Mitochondrial respiration in hummingbird flight muscles. Proc. Natl. Acad. Sci. USA 88: 4870-4873, 1991. 38. Sullivan, S.M. and R.N. Pittman. Relationship between mitochondrial volume density and capillarity in hamster muscles. Am. J. Physiol. 252: H149-H155, 1987. 39. Weber, J.-M., R.W. Brill and P.W. Hochachka. Mammalian metabolite flux rates in a teleost: lactate and glucose turnover in tuna. Am. J. Physiol. 250: R452-R458, 1986.

Hochachka and Mommsen (eds.), Biochemistryand molecularbiology of fishes, vol. 4 9 1995 Elsevier Science B.V. All rights reserved. CHAPTER 2

Circulatory substrate fluxes and their regulation JEAN-MICHEL WEBER AND GEORGES ZWINGELSTEIN *

Biology Department, University of Ottawa, 30 Marie Curie, Ottawa, Ontario, Canada K l N 6N5 and * Laboratoire Maritime de Physiologic, Institut Michel Pacha, Universit~ de Lyon, 1337 Comiche Michel Pacha, Tamaris, F-83500 La Seyne sur Mer, France

I. II. III. IV. V. VI. VII. VIII.

Introduction Why and how to measure metabolite fluxes in vivo? Basic regulatory mechanisms Lactate fluxes Glucose fluxes Amino acid fluxes Lipid fluxes References

I. Introduction Multicellular life can only be sustained if selected metabolic fuels, end-products, and anabolic precursors are transported between cells at the appropriate rates and times. In vertebrates, most inter-tissue metabolite exchange depends on the cardiovascular system and, consequently, the regulation of circulatory substrate fluxes plays a crucial role in achieving homeostasis. Fishes are no exception. They must constantly adjust rates of blood metabolite turnover to coordinate biochemical processes involved in maintenance, growth, reproduction, locomotion and various responses to environmental stresses. As a group of vertebrates, however, they must use distinct metabolic strategies mainly imposed by their aquatic environment and high protein intake 48,83. Both, proteins and lipids dominate fish energy metabolism because low amounts of carbohydrates are ingested and their absorption is rather limited 42. Surprisingly, most of the detailed information concerning metabolite fluxes of fish deals with carbohydrates even though they often represent a very small fraction of these organism's total energy budget. This bias can be explained by: (1) an imitation of mammalian studies where carbohydrates can play a major role; and (2) the relative simplicity of carbohydrate biochemistry compared with lipids and proteins. In this chapter, we examine the main metabolic substrates found in the systemic circulatory system of fish (Fig. 1) and review what is presently known about the modulation of their fluxes. Plasma concentrations of these major substrates are

16

J.-M. Weber and G. Zwingelstein TRANSPORT

EXCHANGE WITH ENVIRONMENT

RFtEE 90LUW3~" INTESTINE

[~GLYCEROL ACIOiS ~T~A~

GILLS SKIN

~DNEY

STORAGEAND TRANSFORMATION

ADIPOSE SKi~ETAL I~SCI, F.S

~iOTEIV.BOUNQ

FATTYACIDS 1 TRIACYLGLYCEROLS PHOSFtlOUPIOS STEROLESTERS " I I

HEART BRAIN

! GONADS

..... i

Fig. 1. Major soluble and protein-bound circulatory fuels in fish: sources and destinations.

summarized in Table 1. The secondary circulation47.119 is not discussed separately because no metabolite measurements from this compartment are yet available. Also, the fluxes of several systemic substrates have never been measured directly. In such cases, we suggest important avenues for future work and provide indirect estimates whenever possible.

II. Why and how to measure metabolite fluxes in vivo ? All body constituents are constantly produced and utilized 44, and circulatory metabolites are therefore kept in a dynamic state, undergoing constant turnover. For decades, however, changes in plasma concentration have been used to draw quantitative conclusions about rates of substrate release into the circulation and uptake therefrom. Such conclusions are often not valid because concentration changes only indicate an imbalance between release and uptake, and major variations in flux can potentially occur while concentration stays constant 12s. Fortunately, flux and concentration of individual metabolites usually vary in parallel and some qual. itative information about flux can be gained from the direction of concentration changes. Modem access to various metabolic tracers has opened the door for the direct measurement of fluxes in vivo on a routine basis and mammalian biology has greatly benefited from this approach. In contrast, relatively few whole organism turnover studies have been attempted in fish with the two major techniques presently available: bolus injection and continuous infusion. The terminology, experimental procedures and calculations necessary to carry out reliable flux measurements have

Circulatory substrate fluxes and their regulation

17 TABLE 1

Resting concentrations of major plasma substrates in fish Metabolite

g 1-1

/zmol m1-1

Directly available (no hydrolysis required) Glucose 123,131 Lactate 9,26,71,123 Amino acids 54,77,92,95 Fatty acids 59,64,131

0.20.010.3 0.1 -

2.5 0.2 2.3 1.5

1 -14 0.1- 2 2 -14 0.1- 5.4

Only available after hydrolysis Triacylglycerols64'94,131 Phospholipids 19,94 Total proteins 1~

1 4 28

-11 -10 -35

1 -12 7 -12 m

Molar concentrations were calculated using average molecular weights of 160 (amino acids), 280 (fatty acids), 880 (triacylglycerols), and 780 (phospholipids).

been described in detail by Hetenyi 44, Katz 51-53, Okajima 81 and their coworkers and by Wolfe 128, amongst others. The bolus injection technique has almost been used exclusively in fish studies because it only requires a single catheter for both, tracer injection and blood sampiing. In contrast, continuous infusion takes two catheters to allow simultaneous infusion and sampling, and the added difficulties associated with surgical placing and maintenance of two lines have encouraged fish biologists to opt for the simpler experimental design of bolus injection 123. This is unfortunate because much more information could be obtained from continuous infusion where consecutive measurements of flux are possible in a single experiment under steady or non-steady state conditions (i.e. even when metabolite concentration varies during the experiment). A more complete understanding of flux regulation in fish will require common use of continuous infusion and the development of easier double catheterization techniques should make this possible.

III. Basic regulatory mechanisms How does the organism alter metabolite turnover rate in response to different stresses? In a study on the regulation of plasma metabolite fluxes in exercising Thoroughbred horses, blood flow and plasma metabolite concentration were proposed as the coarse and fine control, respectively 125,126. There is no reason to believe that flux regulation follows different principles in teleosts. Blood flow is the coarse control because its changes will affect all plasma fuels to the same extent. Metabolite concentration represents the fine control because modifying it for individual substrates will allow the modulation of flux for each fuel independently. This way, the respective contribution of each substrate to total metabolism can be affected by its relative concentration in the circulation, and a positive correlation between circulating concentration and turnover rate has been demonstrated for a

18

J.-M. Weber and G. Zwingelstein

LACTATE

[•'• X =) _J It.

SWIMMING

c~,.s. I TROUT

I ~ST

IrLouNoER~

~,.o,, I

CONCENTRATION Fig. 2. Relationships between plasma lactate concentration and flux in several species of teleosts. Note the positive correlation between the slope of this relationship and cardiac output.

variety of metabolites in all species studied to date. In addition, the slope of the relationship between concentration and flux increases as cardiac output rises 126. The regulating roles of cardiac output and circulating metabolite concentration can be demonstrated for teleosts in the case of lactate. Figure 2 shows how the slope varies between species and experimental conditions. Slopes range between 0.7 and 3.6 in resting teleosts where changes in lactate concentration were elicited by hypoxia or previous heavy exercise21,29,71. In contrast, during exercise, rainbow trout (Oncorhynchus mykiss) show a slope of 5.2 (ref. 123): almost twice the value found during hypoxia29. This large difference can be explained by the fact that cardiac output is much higher in swimming animals than in resting hypoxic fish. Finally, it is not surprising to find the highest slope of 15.1 in skipjack tuna (Katsuwonus pelamis), a species with a cardiac output more than 7 times higher than rainbow trout 32. There is no doubt that the above analysis of flux regulation is still extremely primitive. Potentially important biochemical signals and direct neural effects have not even been mentioned here because their influence has not been investigated in fish. Presumably, some of these factors will affect fluxes indirectly by changing blood flow or circulating concentration of the metabolite of interest. Several hormones are bound to play important regulating roles and their investigation should be a priority in future research.

IV. Lactate fluxes Lactate has occupied a prominent position in studies of hypoxia and muscle metabolism for a very long time. In the last 20 years, tracer experiments have allowed to establish that its fluxes were much higher than for other plasma substrates, even in resting organisms, and that it could become an important metabolic fuel

Circulatory substrate fluxes and their regulation

19 TABLE 2

Lactate turnover rate in post-absorptive teleosts Species

Mass (kg)

Rt (/zmol kg -1 min -1)

Predicted Rt for mammal of same size

Predicted mammal Rt/ Measured fish Rt

Anguilla rostrata 26 Platichthys stellatus 71 Oncorhynchus kisutch 71 Ictalurus punctatus 21 Oncorhynchus mykiss 29 Oncorhynchus mykiss 123 Katsuwonus pelamis TM

O.180 0.335 0.275 0.800 0.350 0.322 1.420

0.50 0.76 1.33 2.25 2.80 4.41 112

145 112 122 78 110 114 61

290 147 92 35 39 26 0.54

Predicted values for mammals of equivalent body mass were calculated as follows: Rt -- 70.78 Mb 0'42, where Rt = lactate turnover rate in ttmol kg -I min -1, and Mb ffi body mass in kg (modified from reference 123).

for oxidative tissues in mammals 76,122. This new picture of lactate metabolism has attracted the attention of fish biologists, and resting lactate turnover rates have been measured in several teleost species (Table 2). Except for tuna, the lactate fluxes of fish range from 0.5 /~mol kg -1 rain -1 in eels (Anguilla sp.) to 4.4/tmol kg -1 min -1 in rainbow trout (Oncorhynchus mykiss). Enough information is available from mammals to derive an allometric equation expressing the relationship between resting lactate turnover rate and body mass in this vertebrate group 123. We have used this equation to compare lactate fluxes in teleosts and mammals of equivalent size (Table 2). This comparison shows that turnover rate is 26 to 290 times lower in fish than in mammals, but skipjack tuna (Katsuwonuspelamis), the only scombrid measured to date, stands out as a clear exception with fluxes exceeding those of mammals

TM.

Ratios between lactate turnover and oxygen consumption rates of teleosts and mammals of the same size are similar, suggesting that the metabolic role of lactate is equivalent in all resting vertebrates when the effect of body mass is taken into account 123. This conclusion may not hold during exercise because patterns of lactate exchange between skeletal muscle and the circulation are so strikingly different in fish and mammals. After strenuous swimming for example, lactate is released extremely slowly from fish white muscle 113 and this typical pattern of retention found in all teleosts is exaggerated in bottom-dwelling, sedentary s p e c i e s 114'121. Therefore, during exercise, lactate oxidation may account for different proportions of total VO2 in mammals, pelagic and benthic fish. Stresses of different kinds are known to stimulate lactate fluxes in mammals, but little information is available for fish. Nonetheless, fasting, hypoxia and exercise have all been shown to increase lactate turnover rate in some teleosts. In American eels, long-term fasting causes a 2.5-fold rise in turnover rate 26, but the effect of food restriction has never been quantified in other fish species. Similarly, the effect of hypoxia has only been measured in rainbow trout where turnover rate increased by

20

J.-M. Weberand G. Zwingelstein

seven-fold at an environmental p O 2 of 4 kPa (ref. 29). Exercise studies have been limited by the steady state assumption of the bolus injection technique. Two steady state situations have been investigated to date: prolonged aerobic swimming and recovery from strenuous, anaerobic exercise. During sustained exercise, circulatory lactate transport between tissues could be a convenient way to shuttle carbohydrate energy between body compartments 76, and white to red muscle lactate exchange was hypothesized as a potential mechanism to support energy metabolism in active red muscle of fish. However, lactate flux only increases by two-fold during sustainable swimming in trout, showing that such a mechanism does not play a significant role in this species 123. In recovery from exhaustive exercise, fluxes are also elevated to accelerate the disposal of large lactate loads accumulated during anaerobic work. After strenuous swimming, a 3- and 9-fold increase in turnover rate was measured in flounder and salmon, respectively71. In neither species were recovery fluxes sufficiently high to explain the time course of decrease in muscle lactate concentration, suggesting that a fraction of the total lactate load never leaves white muscle and is metabolized in situ.

V Glucosefluxes Rates of glucose turnover have been measured in several species at rest (Table 3). They were determined under steady state conditions and therefore represent both, rates of glucose production (Ra) and disappearance from the circulation (Rd) at the whole organism level. The liver accounts for most of the glucose produced, but fish kidneys can probably also make a significant contribution unlike their mammalian counterpart 55.7s,1~ The relative importance of liver and kidney has not been quantified in vivo, and is likely to depend upon species, diet, and level of activity. Measured rates of glucose t u r n o v e r (Rt) are upper estimates of glucose oxidation TABLE 3 Glucose turnover rate (Rt) in resting, post-absorptive teleosts Species

Rt

Dicentrarchus labrax3s Oncorhynchus mykiss 27'3s Oncorhynchus mykiss 2 Paralabrax sp. 14,15 Oncorhynchus kisutch 65 Hemitriptems ameticanus 118 Hoplias malabaricus67 Pleuronectes platessa 12 Katsuwonus pe/.am/s124 Anguilla rostrata26

0.6 1.0 1.1 2.1 2.2 3.6 3.9 5.7 15.3 56

Turnover rates are given in/~mol glucose kg -1 min -1.

Circulatory substrate fluxes and their regulation

21

(Rox) because not all glucose leaving the circulation is usually oxidized. Rox glucose has not been quantified directly in vivo, but comparative recovery of expired 14CO2 after bolus injection of different 14C-substrates shows that glucose is oxidized at much lower rates than fatty acids and amino acids except for glycine 37,115. Unfortunately, the experimental approach used in these studies does not provide absolute rates of oxidation. Continuous infusion of 14C-substrates after priming the CO2/bicarbonate pool, and monitoring 14CO2 production will be needed to measure such rates. Then, comparing R~,, t and Rox values will allow to determine what percentage of total glucose turnover is oxidized. Except for eel and tuna, glucose turnover rates of fish range between 0.6 and 5.7 /zmol kg -1 min -1 (Table 3). These values are 20-100 times lower than for resting mammals of equivalent size ~24. The lower body temperature and lower metabolic rate of fish may account for this difference. The high glucose fluxes of tuna can be explained by their 'mammalian' metabolic rates and greater reliance on carbohydrates for energy metabolism, but it remains unclear why eels should have the ability to support even higher turnover rates than tuna or mammals. Species showing high turnover rates appear to have a better ability to maintain steady blood glucose concentrations 1~ However, the main factors involved in the regulation of glucose fluxes have not been investigated thoroughly. The evidence available to date suggests that the regulatory mechanisms of fish operate very slowly (hours) compared with mammals (minutes). Hepatic glucose production only shuts down 1-2 h after glucose loading in Paralabrax 14. Also, indirect evidence from changes in circulating glucose concentration suggests that insulin45,1~~and glucagon ~11 take at least 30 min to start modifying fluxes and that their effect lasts for several hours. Elevated plasma cortisol has no effect on the glucose turnover of rainbow trout (cortisol injection) 2 and sea raven (high cortisol induced by chronic stress) 118. Similarly, subjecting trout to 3 h of low water pO2 (4 kPa) had no effect on their glucose flux29, mainly because elevated plasma catecholamines tend to abolish the inhibitory effect of hypoxia 129. In future work, quantifying the respective effects of circulating glucose, insulin, glucagon and other hormones will require the use of continuous infusion and, eventually, the 'glucose clamp' technique 44 should be adapted for fish experiments. Because several tissues rely exclusively on glucose for energy metabolism, some attention has been devoted to potentially limiting glucose fluxes during fasting. The most dramatic effect has been shown in American eels where a 10-fold decrease in glucose turnover was measured after 15 months of food deprivation 26. Shorter studies in other species provide conflicting results. A 30% reduction in glucose flux was observed in Paralabrax 14 and Hoplias67,but Hemitripterus 118 and Dicentrarchus 38 showed an 80 and 320% increase, respectively. These species differences are quite puzzling and a closer look at the combined effects of several factors including size, age, diet, locomotory habits, and temperature may provide an explanation. The effect of exercise on glucose turnover rate has not been investigated in fish. However, West et al. 127 have recently used deoxyglucose to quantify glucose uptake of individual tissues from the circulation. This exciting approach will allow to determine the relative contribution of different organs to whole-animal glucose turnover

22

J.-M. Weber and G. Zwingelstein

and it opens the door for a detailed investigation of fish glucose metabolism in vivo. In a first series of experiments on trout, these authors have shown that exercise causes a 28-fold increase in red muscle glucose utilization but has no effect on cardiac muscle. Interestingly, glucose utilization only accounts for less than 10% of the oxidative metabolism of these two tissues during swimming 127.

VI. A m i n o acid fluxes Proteins represent a very important source of energy in teleosts 16, and rates of nitrogen excretion have been used to quantify protein catabolism 117. Different studies have concluded that amino acid oxidation accounts for 14-85% of total 1(/IO2 depending on species, feeding status, and level of activity 18'56'57'116. Despite this well-known dependence on protein for energy metabolism, very few researchers have tried to measure rates of circulatory amino acid turnover and oxidation. Furthermore, reports to date are qualitative only, providing relative rates between substrates or experimental conditions. Borer and colleagues estimated that alanine, glutamate, and aspartate fluxes of Paralabrax were equivalent to mammalian values 15, confirming the much higher relative importance of amino acid catabolism to total MO2 in fish than in mammals, because of the large metabolic rate difference between these two groups of animals. The turnover rate of the three amino acids measured was not affected by 72 days of fasting 15. Measurements of 14CO2production after injection of t4C-substrates show that circulating glutamate, alanine, leucine, and phenylalanine are oxidized much more rapidly than glucose in resting fish37. This is also true in swimming trout where leucine becomes the preferred amino acid substrate for oxidation in active muscles nS. As expected, non-essential amino acids are generally favored over essential amino acids 120. A significant fraction of total flux is channeled through gluconeogenesis. Teleosts have evolved a relatively high capacity for converting amino acids to glucose and this has been interpreted as a strategy to synthesize enough mucopolysaccharides for mucus production in organisms with little dietary carbohydrates 15. Two very interesting situations where amino acid fluxes should be particularly high have not been investigated so far: elasmobranchs and migrating salmon. Elasmobranchs have no significant ability to oxidize lipids outside the livers.1~ Therefore, during sustained locomotion, they should derive most of their energy from amino acid oxidation as indicated by their high capacity to metabolize glutarnine in muscle mitochondria 22. Similarly, salmon is known to depend almost exclusively on protein catabolism in the last stages of long migrations, after carbohydrate and lipid reserves have been depleted 3.16. In sockeye salmon (Oncorhynchus nerka), several amino acids appear to be converted to alanine before inter-organ transport 77, suggesting that alanine fluxes are much higher in migrating than in non-migrating teleosts. In addition, amino acid fluxes should increase throughout migration as white muscle proteins are progressively catabolized via the indirect action of androgens 4,s, and proteolytic agents such as cathepsins 130.

23

Circulatory substratefluxes and their regulation

Finally, no amino acid flux measurement should be attempted in fish without considering the large concentration gradient between red cell and plasma. Amino acids are three 11 to over 200 times 34 more concentrated inside fish erythrocytes than in plasma, and, therefore, the specific activity measured in plasma or whole blood will be different, leading to the calculation of distinct flux rates. Each experimental situation should be considered individually before selecting plasma or whole blood because concentration gradients and red cell membrane transport kinetics are so variable between species and amino acids.

I~I. Lipid fluxes To our knowledge, plasma lipid fluxes have never been measured in fish at the whole organism level even though fat represents a critical source of ATP in these animals. The following analysis will focus on indirect and qualitative information to point out promising directions and potential difficulties for future research. The major circulatory lipids of f i s h - free fatty acids (FFA) and triacylglycerols (TAG) - and their sites of appearance and disappearance are summarized in Fig. 3. Also, many important aspects of circulatory lipid transport in fish have been reviewed by Sheridan 98. Lipid substrates are shuttled between tissues either as FFA (rapid delivery) or as TAG and phospholipids (slow delivery). Most of our discussion will deal with INTESTINE DIETARY UPIDS STORAGE

CATABOLIC TISSUES

i

TISSUES

TG+PL

_

..........

~'

F

~

FFA ~

TG + PL

UVER

Fig. 3. Source, destination and composition of plasma lipids. FABP ffi fatty acid binding protein; FFA = free fatty acids; HSL = hormone sensitive lipase; LPL = lipoprotein lipase; PL = phospholipids; TG = triacylglycerols; VLDL = very low density lipoproteins.

24

J..M. Weber and G. Zwingelstein

the relatively simple situation of FFA rather than with other lipids (TAG, glyceryl ethers, phospholipids, free and esterified cholesterol) whose complex circulatory transport involves ehylomierons, VLDL, HDL, and LDL9s. Even in mammals, the present understanding of these compounds' kinetics is still very limited. In fish, 20--30% of circulating FFA are unsaturated with chain lengths of 20 and 22 carbons 1~176 The major function of these polyunsaturated FFA is to act as precursors of membrane phospholipids 13,1n and eicosanoid compounds 6,66,88,while shorter chain fatty acids (C18 and less) are used primarily for energy metabolism. Therefore, C20 and C22 acids should have much lower turnover rates than the FFA involved in oxidative pathways. Also, one would expect that swimming will have a much more pronounced effect on the flux of 'short' acids than on C20 and longer FFA. The choice of an appropriate marker fatty acid for measuring FFA fluxes for different purposes and under different conditions should take the above considerations into account. Plasma FFA concentration is approximately one order of magnitude higher in teleosts than in elasmobranchs and holocephalans (see Table 4). In addition, and contrary to eyclostomes and teleosts28,4~ elasmobranchs lack albumin-like plasma proteins 33, and they are incapable of oxidizing fatty acids in other tissues than in liver7,22,1~ The FFA fluxes of sharks, skates and rays should therefore be significantly reduced in view of their remarkably limited capacity to transport and metabolize lipids. The high cardiac output of elasmobranehs (53 v e r s u s 17 ml kg-lmin -1 in resting dogfish and trout, respectively5~ can only partially compensate for their low plasma FFA (approximately 0.15 v e r s u s 1.2 mM). With the same relative extraction from plasma, dogfish would only be able to support less than half the FFA delivery rate of trout. TABLE 4 Total plasma free fatty acid concentration in teleosts, elasmobranchs, and a holocephalan Species

FFA (~mol m1-1)

Teleosts Oncorhynchus mykiss 43 Salvelinus alpinus 39 Dicentrarchus labrax 131 Mullus surmuletus 131 Scomber scombrus 131 Gadus aeglefinus 59 Gadus morhua 59

1.52 2.11 1.10 1.42 1.22 1.54 1.28

Elasmobranchs Scyliorhinus canicula 131 Squalus acanthias 131 Raja rad/ata 59 Etmoptems spinax59

0.15 0.15 0.09 0.29

Holoeephalan Chimaera monstrosa 59

0.17

25

Circulatory substrate fluxes and their regulation

T h e effect of exercise on p l a s m a FFA fluxes should be very different b e t w e e n species b e c a u s e teleosts show very diverse swimming abilities and lipid s t o r a g e s t r a t e g i e s 1~ S o m e species store m o s t of their T A G in l o c o m o t o r y muscles (e.g. herring, Clupea harengus), o t h e r s c o n c e n t r a t e T A G in liver (e.g. cod, G a d u s m o r h u a ) or in a d i p o s e tissue 98. Table 5 lists a few species to illustrate muscle/liver s t o r a g e TABLE 5 Total lipid content of liver and muscle (% lipid per g tissue wet weight) Species

Liver

Muscle

Oncorhynchus mykiss 1 Oncorhynchus nerka 17 Clupea harengus 17 Scomber scombrus 17 AnguiUa anguiUa70 Dicentrarchus labrax a Gadus morhua 36 Gadus aeglefinus 82

5 7 2 8 12 18 63 63

4 15 11 13 18 3 0.3 0.4

a

G6rard Brichon, unpublished results. TABLE 6 Effects of hormones on plasma FFA concentration indicating similar changes in FFA fluxes

Hormone

Effect on plasma FFA

Insulin 63,74

Decrease

Glucagon46 Glucagon23,60,91,109

Increase No effect

Catecholamines3~ Catecholamines31 Catecholamines86

Increase Decrease No effect

ACTH 75 ACTH 3~176

Increase No effect

Somatostatin99,101

Increase

Urotensin I199,1~

Increase

Arginine vasotocin49,68 Arginine vasotocin 68

Increase Decrease

Thyroid hormone8~176 Thyroid hormone96

Increase No effect

Cortisol 2~ Cortiso196

Increase No effect

Sex steroid analog 1~

Increase

Growth hormone69,75

Increase

Prolactin61,75

Increase

J..M. Weber and G. Zwingelstein

26

options. During swimming, teleosts favoring hepatic and adipose storage will have to supply most FFA to their working muscles v/a the circulation. Such species should therefore increase plasma FFA fluxes to a much larger extent than fish with considerable TAG reserves in their muscles. A variety of hormones are potentially involved in the regulation of plasma FFA fluxes. In mammals, flux and concentration are positively correlated 41,s4 and fish should be no different. The direction of hormonal effects on FFA concentration and flux are probably also identical in this group of vertebrates. Table 6 summarizes the potential effects of several hormones on the turnover rate of circulating FFA. It is interesting to note that some hormones will not regulate FFA fluxes in all species because their effects are tissue specific 9s. For example, catecholamines stimulate lipolysis in fish hepatocytes 97, but have no effect on their adipocytes 79. Consequently, catecholamines should increase FFA turnover in species storing TAG in the liver, but they should play no regulating role in species using mostly adipose tissue for lipid storage. Finally, the study of circulating triacylglycerol, phospholipids, and cholesterol promises to be extremely complex, but experimental difficulties should be overcome TABLE 7A

Total lipoprotein concentration and respective percent contribution of VLDL, LDL and HDL in plasma. All values were measured by ultracentrifugation methods Species Oncorhynchus mykiss juvenile 1~ adult 24,25.35

Oncorhynchus nerka93 Sardinops caerulae 62 Myxine glutinosa 72 Latimeria chalumnae 72 Scyliorhinus canicula 72 Centrophorus squamosus 73 Conger vulgaris72

Lipoproteins

VLDL

LDL

(gl -~)

(~)

(~)

13-17

18

64

18

23-26 7 8 29 14 2 7 7

7 26 13 57 77 14 61 67

36 38 15 24 14 75 34 33

57 37 72 19 9 11 6 -

HDL

TABLE 7B

Average lipid and protein composition of fish lipoproteins (% lipoprotein wet weight) Proteins

Phospho-

Free

lipids a

cholesterol

..... Cholesteryl

Triacylglycerols

esters ,,

Chylomicron VLDL LDL HDL

2 13 26 49

8

1

2

15 23 26

7 9 6

18 14 12

84 43 25

10

a Phospholipids (mostly phosphatidylcholine and sphingomyelin). In Latimeria, Scyliorhinus, and Conger, 13 tO 70% of the triacylglycerol fraction is made of alkyldiacylglycerols72,73. Values calculated from references 24, 25, 35, 62, 72, 73, 93, 100, and 105.

Circulatory substrate fluxes and their regulation

27

because, in certain species, these compounds can represent 10 to 40 times the energy stored in circulating FFA (see Table 1). They are first transported as chylomicrons before being stored in liver or adipose tissue where they can be converted to lipoproteins and released back in the circulation 98. Measuring the fluxes of these compounds will be a real challenge because the relative contribution of VLDL, LDL, and HDL to total plasma lipoproteins varies greatly between species (Table 7A), and each class of lipoproteins has a different composition (Table 7B). For these two major reasons, great care will have to be taken in choosing adequate lipid tracers and modes of administration to decipher specific aspects of plasma lipoprotein kinetics.

VIII. References 1. Abdul Malak, N., G. Zwingelstein, J. Jouanneteau and J. Koenig. Influence de certains facteurs nutritionnels sur la pigmentation de la truite arc en ciel par la cantaxanthine. Ann. Nutr. Alim. 29: 459-475, 1975. 2. Andersen, D.E., S.D. Reid, TW. Moon and S.E Perry. Metabolic effects associated with chronically elevated cortisol in rainbow trout (Oncorhynchus mykiss). Can. J. Fish. Aquat. Sci. 48: 1811-1817, 1991. 3. Ando, S., M. Hatano and K. Zama. Deterioration of chum salmon (Oncorhynchus keta) muscle during spawning migration -I. Changes in proximate composition of chum salmon muscle during spawning migration. Comp. Biochem. Physiol. 80B: 303-307, 1985. 4. Ando, S., M. Hatano and K. Zama. Deterioration of chum salmon muscle during spawning migration - VI. Changes in serum protease inhibitory activity during spawning migration of chum salmon (Oncorhynchus keta). Comp. Biochem. Physiol. 82B: 111-115, 1985. 5. Ando, S., E Yamazaki, M. Hatano and K. Zama. Deterioration of chum salmon (Oncorhynchus keta) muscle during spawning migration - III. Changes in protein composition and protease activity of juvenile chum salmon muscle upon treatment with sex steroids. Comp. Biochem. Physiol. 83B: 325-330, 1986. 6. Anonymous. Potential importance of arachidonate in gill function of marine fish. Nutrit. Rev. 45: 58-60, 1987. 7. Ballantyne, J.S., M.E. Chamberlin and TD. Singer. Oxidative metabolism in thermogenic tissues of the swordfish and mako shark. J. Exp. Zool. 261: 110-114, 1992. 8. Ballantyne, J.S., H.C. Glemet, M.E. Chamberlin and TD. Singer. Plasma nonesterified fatty acids of elasmobranch fishes. Mar. Biol., 166: 47-52, 1993. 9. Bange-Barnoux, R. Evolution de I'acide lactique et du glucose plasmatiques chez la Tanche lors de rasphyxie par confinement. C.R. S~ances Soc. Biol. 159: 400-403, 1965. 10. Bange-Barnoux, R., C. Bange, H. Vanel and J. Pottu. Influence du jeQne sur la protid6mie et les compos6s alpha-amin6s du sang de la tanche (Tinca tinca L.). Ann. Inst. Michel Pacha 4: 77-101, 1971. 11. Barnoud, R. and G. Peres. Recherches sur la protid6mie de la tanche soumise ~ r asphyxie par confinement. III Aminoacid6mie. J. Physiol. (Paris) 55: 193-194, 1963. 12. Batty, R.S. and C.S. Wardle. Restoration of glycogen from lactic acid in the anaerobic swimming muscle of plaice Pleuronectes platessa L.J. Fish Biol. 15: 509-519, 1979. 13. Bell, M.V., R.J. Henderson and J.R. Sargent. The role of polyunsaturated fatty acids in fish. Comp. Biochem. Physiol. 83B: 711-719, 1986. 14. Beret, K., M. Chenoweth and A. Dunn. Glucose turnover in kelp bass (Paralabrax sp.): in vivo studies with (6-3H, 6-14C)glucose.Am. J. Physiol. 232: R66-R72, 1977. 15. Beret, K., M. Chenoweth and A. Dunn. Amino acid gluconeogenesis and glucose turnover in kelp bass (Paralabrax sp.). Am. J. Physiol. 240: R246-R252, 1981. 16. Bilinski, E. Biochemical aspects of fish swimming. In: Biochemical and Biophysical Perspectives in Marine Biology, edited by D.C. Malins and J.R. Sargent, New York, Academic Press, pp. 239-288, 1974. 17. Braekken, O.R. Reports on Technological Research Concerning Norwegian Fish Industry. Norwegian

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l'AnguiUe EuropEenne. E/lets de l'Adaptation en Eau Douce ou en Eau de Mer r Deux TempEratures (12 et 22~ Doctorat d'~tat d~s sciences naturelles, Universit6 de Lyon, 1984. 20. Butler, D.C. Structure and function of the adrenal glands of fshes. Am. ZooL 13: 839-879, 1973. 21. Cameron, J.N. and J.J. Cech Jr. Lactate kinetics in exercised channel catfish, Ictalurus punctatus. Physiol. Zool. 63: 909-920, 1990. 22. Chamberlin, M.E. and J.S. BaUantyne. Glutamine metabolism in elasmobranchs and agnathan muscle. J. Exp. Zool. 264: 267-272, 1992. 23. Chan, D.K.O. and N.Y.S. Woo. Effect of glucagon on the metabolism of the eel, Anguilla japonica. Gen, Comp. Endocr. 35: 216-225, 1978. 24. Chapman, M.J., S. Goldstein, G.L. Mills, L. Fremont and C. Leger. Some aspects of the serum lipoproteins in the rainbow trout. In: Protides of the Biological Fluids, edited by H. Peeters, Oxford, Pergamon Press, pp. 489-490, 1979. 25. Chapman, M.J., S. Goldstein, G.L. Mills and C. Leger. Distribution and characterization of the serum lipoproteins and their apoproteins in the rainbow trout (Salmo gairdneri). Biochemistry 17: 4455-4464, 1978. 26. Cornish, I. and T.W. Moon. Glucose and lactate kinetics in American eel Anguilla rostrata. Am. J. Physiol. 249: R67-R72, 1985. 27. Cowey, C.B., H. de la Higuera and J.W. Adron. The effect of dietary composition and of insulin on gluconeogenesis in rainbow trout (Salmo gairdneri). Br. J. Nutr. 38: 385-395, 1977. 28. Davidson, W.S., S.E. Bartlett, T.P. Birt, V.L. Birt and J.M. Green. Identification and purification of serum albumin from rainbow trout (Salmo gairdneri). Comp. Biochem. Physiol. 93B: 5-9, 1989. 29. Dunn, J.E and P.W. Hochachka. Turnover rates of glucose and lactate in rainbow trout during acute hypoxia. Can, J. Zool. 65: 1144-1148, 1987. 30. Farkas, T. The effect of catecholamines and adrenocorticotropic hormone on blood and adipose tissue FFA levels in the fish Cyprinus carpio. Prog. Biochem. Pharmacol. 3: 314--319, 1967. 31. Farkas, T. Studies on the mobilization of fats in lower vertebrates. Acta Biochim. Biophys. Acad. Sci. (Hungary) 4: 237-249, 1969. 32. Farrell, A.P. From hagfish to tuna: a perspective on cardiac function in fish. Physiol. Zool. 64: 1137-1164, 1991. 33. Fellows, EC.I. and EJ.R. Hird. Fatty acid binding proteins in the serum of various animals. Comp. Biochem. Physiol. 68B: 83-87, 1981. 34. Fincham, D.A., M.W. Wolowyk and J.D. Young. Characterisation of amino acid transport in red blood cells of a primitive vertebrate, the Pacific hagfsh (Eptatretus stouti). J. E~. Biol. 154: 355370, 1990. 35. Fremont, L. and C. Leger. Le transport des lipides plasmatiques. In: Nutrition des Poissons, edited by M. Fontaine, Paris, Editions du CNRS, pp. 263-281, 1981. 36. Garcia, M.D., J.A. Lovern and J. Olley. The lipids of fish. 6. The lipids of cod flesh. Biochem. J. 62: 99-107, 1956. 37. Garin, D. and M. Mockry. Relation entre les mc~.tabolismes amin~., glucidique et lipidique chez le poisson, Dicentrarchus labrax. Influence de facteurs de renvironnement et de la composition de la ration. Rapport de contrat CNEXO, lnstitut Michel Pacha, 1983. 38. Garin, D., A. Rombaut and A. Fr~minet. Determination of glucose turnover in sea bass Dicentrarchus labrax. Comparative aspects of glucose utilization. Comp. Biochem. Physiol. 87B: 981-988, 1987. 39. Glemet, H.C., M.E Gerrits and J.S. Ballantyne. A comparison of plasma non-esterified fatty acids in anadromous and land-locked Arctic charr, Salvelinus alpinus. Unpublished data. 40. Gray, J.E. and R.E Doolittle. Characterization, primary structure and evolution of lamprey plasma albumin. Protein Science 1: 289-302, 1992. 41. Hagenfeldt, L. Turnover of individual free fatty acids in man. Fed. Proc. 34: 2236-2240, 1975. 42. Halver, J.E. Fish Nutrition. New York: Academic Press, 1972. 43. Harrington, A.J., K.A. Russell, T.D. Singer and J.S. Ballantyne. The effects of tricaine methanesulfonate (MS-222) on plasma nonesterified fatty acids in rainbow trout, Oncorhynchus mykiss. Lipids 26: 774--775, 1991. 44. Hetenyi, G.H., G. Perez and M. Vranic. Turnover and precursor-product relationships of non-lipid

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29

metabolites. Physiol. Rev. 63: 606-667, 1983. 45. Ince, B.W. and A. Thorpe. Effects of insulin and metabolite loading on blood metabolites in the European silver eel (Anguilla anguilla R.). Gen. Comp. Endocrinol. 23: 460-471, 1974. 46. Ince, B.W. and A. Thorpe. Hormonal and metabolic effects on plasma free fatty acids in the Northern pike, Esox lucius. Gen. Comp. Endocrinol. 27: 144-152, 1975. 47. Ishimatsu, A., G.K. lwama and N. Heisler. In vivo analysis of partitioning of cardiac output between systemic and central venous sinus circuits in rainbow trout: a new approach using chronic cannulation of the branchial vein. J. Exp. Biol. 137: 75-88, 1988. 48. Jayaram, M.G. and EW.H. Beamish. Influence of dietary protein and lipid on nitrogen and energy losses in lake trout, Salvelinus namaycush. Can. J. Fish. Aquat. Sci. 49(11): 2267-2272. 49. John, T.M., E. Thomas, J.C. George and EW.H. Beamish. Effect of vasotocin on plasma free fatty acid levels in the migrating anadromous sea lamprey. Arch. Int. Physiol. Biochim. 85: 865-870, 1977. 50. Jones, D.R. and D.J. Randall. The respiratory and circulatory systems during exercise. In: Fish Physiology, edited by W.S. Hoar and D.J. Randall, New York, Academic Press, pp. 425-501, 1978. 51. Katz, J. Use of isotopes for the study of glucose metabolism in vivo. Techn. Metab. Res. B207: 1-22, 1979. 52. Katz, J., E Okajima, M. Chenoweth and A. Dunn. The determination of lactate turnover in vivo with all- and 14C-labelled lactate. Biochem. J. 194: 513-524, 1981. 53. Katz, J., H. Rostami and A. Dunn. Evaluation of glucose turnover, body mass, and recycling with reversible and irreversible tracers. Biochem. J. 142: 161-170, 1974. 54. Kaushik, S.J. and P. Luquet. Influence of dietary amino acid patterns on the free amino acid contents of blood and muscle of rainbow trout (Salmo gairdnerii R). Comp. Biochem. Physiol. 64B: 175-180, 1979. 55. Knox, D., M.J. Walton and C.B. Cowey. Distribution of enzymes of glycolysis and gluconeogenesis in fish tissues. Mar. Biol. 56: 7-10, 1980. 56. Kutty, M.N. Respiratory quotient and ammonia excretion in Tilapia mossambica. Mar. Biol. 16: 126-133, 1972. 57. Kutty, M.N. and M. Peer Mohamed. Metabolic adaptations of mullet Rhinomugil corsula (Hamilton) with special reference to energy utilization. Aquaculture 5: 253-270, 1975. 58. Kuyas, C., M. Riley, J. Bubis and R.E Doolittle. Lamprey plasma albumin is a glycoprotein with a molecular weight of 175,000. Fed. Proc. 42: 2085, 1983. 59. Larsson, A. and R. F~inge. Cholesterol and free fatty acid (FFA) in the blood of marine fish. Comp. Biochem. Physiol. 57B: 191-196, 1977. 60. Larsson, A. and K. Lewander. Effects of glucagon administration to eels (Anguilla anguilla L.). Comp. Biochem. Physiol. 43A: 831-836, 1972. 61. Leatherland, J.E, B.A. McKeown and T.M. John. Circadian rhythm of plasma prolactin, growth hormone, glucose and free fatty acids in juvenile kokanee salmon, Oncorhynchus nerka. Comp. Biochem. Physiol. 47: 821-828, 1974. 62. Lee, R.E and D.L. Puppione. Serum lipoproteins of the Pacific sardine (Sardinops caerulea). Biochim. Biophys. Acta 270: 272-278, 1972. 63. Leibson, L.G., E.M. Plisetskaya and T.I. Mazina. The NEFA content in the blood of cyclostomata and fishes and the effect of epinephrine and insulin. Z. Evol. Biokhim. Fiziol. (Fish. Res. Bd. Can. Translation Series #1253) 4: 121-127, 1968. 64. Lewander, K., G. Dave, M.L. Johansson, A. Larsson and U. Lidman. Metabolic and hematological studies on the yellow and silver phases of the European eel, Anguilla anguilla L. -I. Carbohydrate, lipid, protein and inorganic ion metabolism. Comp. Biochem. Physiol. 47B: 571-581, 1974. 65. Lin, H., D.R. Romsos, P.I. Tack and G.A. Leveill6. Determination of glucose utilization in coho salmon (Oncorhynchus kisutch, Walbaum) with (6-3H) and (U-14C) glucose. Comp. Biochem. Physiol. 59A: 189-191, 1978. 66. Lowenstein, J. The effects of sea water adaptation on renal eicosanoid production in the goldfish (Carassius auratus). Comp. Biochem. Physiol. 98B: 389-396, 1991. 67. Machado, C.R., M.A.R. Garofalo, J.E.S. Roselino, I.C. Kettlehut and R.H. Migliorini. Effect of fasting on glucose turnover in a carnivorous fish (Hoplias sp.). Am. J. Physiol. 256: R612-R615, 1989. 68. McKeown, B.A., T.M. John and J.C. George. Effect of vasotocin on plasma GH, free fatty acids and glucose in coho salmon (Oncorhynchus kisutch). Endocrinol. Exp. 10: 45-51, 1976. 69. McKeown, B.A., J.E Leatherland and T.M. John. Effect of growth hormone and prolactin on the mobilization of free fatty acids and glucose in kokanee salmon, Oncorhynchus nerka. Comp.

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Circulatory substrate fluxes and their regulation

95.

96. 97. 98. 99. 100. 101. 102. 103. 104. 105. 106. 107. 108. 109. 110. 111. 112.

113. 114. 115. 116. 117.

31

- comparative observations in serranides and sparides. Comp. Biochem. Physiol. 99B: 251-255, 1991. Schlisio, W. and B. Nicolai. Kinetic investigations on the behaviour of free amino acids in the plasma and of two aminotransferases in the liver of rainbow trout (Salmo gairdnerii Richardson) after feeding on a synthetic composition containing pure amino acids. Comp. Biochem. Physiol. 59B: 373-379, 1978. Sheridan, M.A. Effects of thyroxine, cortisol, growth hormone and prolactin on lipid metabolism of coho salmon, Oncorhynchus kisutch, during smoltification. Gen. Comp. Endocrinol. 64: 220-238, 1986. Sheridan, M.A. Effects of epinephrine and norepinephrine on lipid mobilization from coho salmon liver incubated in vitro. Endocrinology 120: 2234-2239, 1987. Sheridan, M.A. Lipid dynamics in fish: aspects of absorption, transportation, deposition and mobilization. Comp. Biochem. Physiol. 90B: 679-690, 1988. Sheridan, M.A. and H.A. Bern. Both somatostatin and the caudal neuropeptide, urotensin II, stimulate lipid mobilization from coho salmon liver incubated in vitro. Regul. Pept. 14: 333-344, 1986. Sheridan, M.A., J.K.L. Friedlander and W.V. Allen. Chylomicrons in the serum of postprandial steelhead trout (Salmo gairdneri). Comp. Biochem. Physiol. 81B: 281-284, 1985. Sheridan, M.A., E. Plisetskaya, H.A. Bern and A. Gorbman. Effects of somatostatin-25 and urotensin II on lipid and carbohydrate metabolism of coho salmon, Oncorhynchus kisutch. Gen. Comp. Endocrinol. 66: 405-414, 1987. Singer, TD. and J.S. Ballantyne. Absence of extrahepatic lipid oxidation in a freshwater elasmobranch, the dwarf stingray Potamotrygon magdalenae: evidence from enzyme activities. J. Exp. Zool. 251: 355-360, 1989. Singer, TD. and J.S. Ballantyne. Metabolic organization of a primitive fish, the bowfin (Amia calva). Can. J. Fish. Aquat. Sci. 48: 611-618, 1991. Singer, TD., V.G. Mahadevappa and J.S. Ballantyne. Aspects of the energy metabolism of lake sturgeon, Acipenserfulvescens, with special emphasis on lipid and ketone body metabolism. Can. J. Fish. Aquat. Sci. 47: 873-881, 1990. Skinner, E.R. and A. Rogie. The isolation and partial characterization of the serum lipoproteins and apolipoproteins of the rainbow trout. Biochem. J. 173: 507-520, 1978. Suarez, R.K. and T.P. Mommsen. Gluconeogenesis in teleost fish. Can. J. Zool. 65: 1869-1882, 1987. Takashima, E, T Habiya, N. Phan-Van and K. Aid. Endocrinological studies on lipid metabolism in rainbow trout. -II. Effects of sex steroids, thyroid powder, adrenocorticotropin on plasma lipid content. Bull. Jap. Soc. Sci. Fish. 38: 43-49, 1972. Tashima, L. and G.E Cahill. Fat metabolism in fish. In: Handbook of Physiology, Section 5: Adipose tissue, edited by A.E. Renold and G.E Cahill, Washington D.C., American Physiological Society, pp. 55-58, 1965. Tashima, L. and G.E Cahill. Effects of insulin in the toadfish Opsanus tau. Gen. Comp. Endocrinol. 11: 262-271, 1968. Thorpe, A. and B.W. Ince. Effects of pancreatic hormones, catecholamines, and glucose loading on blood metabolites in the Northern pike (Esox lucius L.). Gen. Comp. Endocrinol. 23: 29-44, 1974. Thorson, T The partitioning of body water in Osteichthyes: phylogenetic and ecological implications in aquatic vertebrates. Biol. Bull. 120: 238-254, 1961. Tocher, D.R. and J.R. Dick. Incorporation and metabolism of (n-3) and (n-6) polyunsaturated fatty acids in phospholipid classes in cultured Atlantic salmon (Salmo salar) cells. Comp. Biochem. Physiol. 96B: 73-79, 1990. Turner, J.D., C.M. Wood and D. Clark. Lactate and proton dynamics in the rainbow trout (Salmo gairdneri). J. Exp. Biol. 104: 247-268, 1983. Turner, J.D., C.M. Wood and H. H6be. Physiological consequences of severe exercise in the inactive benthic flathead sole (Hyppoglossoides elassodon): a comparison with the active pelagic rainbow trout (Salmo gairdneri). J. Exp. Biol. 104: 269-288, 1983. van den Thillart, G. Energy metabolism of swimming trout (Salmo gairdneri). J. Comp. Physiol. 156: 511-520, 1986. Van den Thillart, G. and E Kesbeke. Anaerobic production of carbon dioxide and ammonia by goldfish, Carassius auratus (L.). Comp. Biochem. Physiol. 59A: 393-400, 1978. Van Waarde, A. Aerobic and anaerobic ammonia production by fish. Comp. Biochem. Physiol. 74B:

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675--684, 1983. 118. Vijayan, M.M. and T.W. Moon. The stress response and the plasma disappearance of corticosteroid and glucose in a marine teleost, the sea raven. Can. I. ZooL 72: 379-386, 1994. 119. Vogel, W.O.P. Systemic vascular anastomoses, primary and secondary vessels in fish, and the phylogeny of lymphatics. In: Cardiovascular Shunts: Phylogenetic, Ontogenetic and Clinical Aspects, edited by K. Johansen and W. Burggren, Copenhagen, Munksgaard, pp. 143-159, 1985. 120. Walton, M.J. and C.B. Cowey. Aspects of intermediary metabolism in salmonid fish. Comp. Biochem, PhysioL 73B: 59-79, 1982. 121. Wardle, C.S. Non-release of lactic acid from anaerobic swimming muscle of plaice Pleuronectes platessa L.: a stress reaction. 3`. Exp. Biol. 77: 141-155, 1978. 122. Weber, J.-M. Design of exogenous fuel supply systems: adaptive strategies for endurance locomotion. Can. I. Zool. 66: 1116-1121, 1988. 123. Weber, J.-M. Effect of endurance swimming on the lactate kinetics of rainbow trout. 1. Exp. Biol. 158: 463-476, 1991. 124. Weber, J.-M., R.W. Brill and P.W. Hochachka. Mammalian metabolite flux rates in a teleost: lactate and glucose turnover in tuna. Am. 3`. Physiol. 250: R452-R458, 1986. 125. Weber, J.-M., G.E Dobson, W.S. Parkhouse, D. Wheeldon, J.C. Harman, D.H. Snow and P.W. Hochachka. Cardiac output and oxygen consumption in exercising Thoroughbred horses. Am, 3'. PhysioL 253: R890-R895, 1987. 126. Weber, J.-M., W.S. Parkhouse, G.P. Dobson, J.C. Harman, D.H. Snow and RW. Hochachka. Lactate kinetics in exercising Thoroughbred horses: regulation of turnover rate in plasma. Am. 3'. Physiol. 253: R896-R903, 1987. 127. West, T.G., P.G. Arthur, R.K. Suarez, C.J. Doll and P.W. Hochachka. In vivo utilization of glucose by heart and locomotory muscles of exercising rainbow trout (Oncorhynchus mykiss). I. Exp. Biol. 177: 63-79, 1993. 128. Wolfe, R.R. Tracers in Metabolic Research. Radioisotope and Stable Isotope~Mass Spectrometry Methods, New York, Alan R. Liss, 1984. 129. Wright, P.A., S.E Perry and T.W. Moon. Regulation of hepatic gluconeogenesis and glycogenolysis by catecholamines in rainbow trout during environmental hypoxia. 3`. Exp. Biol. 147: 169-188, 1989. 130. Yamashita, M. and S. Konagaya. High activities of cathepsins B, D, H and L in the white muscle of chum salmon in spawning migration. Comp. Biochem, Physiol. 95B: 149-152, 1990. 131. Zammit, V.A. and E.A. Newsholme. Activities of enzymes of fat and ketone-body metabolism and effects of starvation on blood concentrations of glucose and fat fuels in teleosts and elasmobranch fish. Biochem. I. 184: 313-322, 1979.

Hochachka and Mommsen (eds.), Biochemistryand molecular biologyof fishes, vol. 4 9 1995 Elsevier Science B.V. All rights reserved. CHAPTER 3

Endogenous fuels; non-invasive v e r s u s invasive approaches GUIDO VAN DEN THILLART AND MARCEL VAN RAAIJ

Institute of Evolutionary and Ecological Sciences, Animal Physiology, Gorlaeus Laboratories, University of Leiden, P.O. Box 9502, 2300 RA Leiden, The Netherlands

I. II.

Introduction Quantifying endogenous fuels: destructive methods 1. Handling stress 2. Tissue damage 3. Tissue extraction 4. Storage 5. Measurement III. Non-destructive approaches 1. Cannulation 2. Calorimetry 3. Nuclear magnetic resonance spectroscopy IV. Storage of endogenous fuels 1. High energy phosphates 2. Carbohydrates 3. Lipids 4. Proteins and amino acids V. Mobilization of endogenous fuels 1. Hypoxia and anoxia 2. Exercise 3. Starvation and migration VI. Summary VII. References

I. Introduction Life is a condition that requires non-equilibrium conditions, since all processes proceed only when free energy is converted into entropy. A state of non-equilibrium is kept at the expense of free energy: energy consumption for maintenance and activity ultimately result in heat production, since the organism itself hardly changes. The conversion of energy forms such as from chemical energy to kinetic energy are always coupled with increase of entropy, normally resulting in heat production. In the case of exercise on a hometrainer the conversion-efficiency from chemical to kinetic energy is about 25%, so 75% is lost as heat. Of course, kinetic energy dissipates in the end as heat, so all free energy is then lost as heat. For metabolic pathways the conservation of the free energy in the form of ATP has not always the same efficiency37. We can distinguish high and low efficiency pathways depending on the amount of free

34

G. ,,an den ThiUartand M. van Raaij

energy loss22. We may ask ourselves why nature did not develop only high efficiency pathways in order to minimize energy losses. The answer to this question is: reaction rate. The higher the energy loss, the faster the reaction can proceed. So, high efficiency processes are necessarily slow because they proceed near the equilibrium condition, and low efficiency processes are fast and are hardly influenced by changes in substrate and product levels because they operate far from equilibrium 37. In order to keep an organism in a state of non-equilibrium and to enable a large number of physiological processes, a constant energy input is needed. The sources for this energy input are the substrates for fermentation and oxidative processes. From a thermodynamic point of view, cells possess three types of reactions: endergonic, equilibrium and exergonic reactions. Most anabolic and homeostasis reactions are endergonic and require reactions coupled with ATP hydrolysis. While instead most exergonic reactions are coupled with ATP synthesis of which the most important are: the creatine kinase reaction; the pyruvate kinase and the 3-phosphoglycerate kinase reactions of the glycolysis; and the oxidative phosphorylation. The substrates for these three processes are respectively: creatine phosphate, glycogen (glucose) and NADH. Although the first two can be considered as fuels, the last substrate is in fact an intermediate occurring at fairly low concentrations normally below 10 ~M 31,n9 and can therefore hardly be viewed as a fuel. Instead, the substrates for the NADH generating processes should be considered as fuels for the oxidative phosphorylation. NADH is generated mainly by three processes: (1) the/~-oxidation of fatty acids; (2) the Krebs cycle; and (3) the glycolytic pathway. The substrates for these processes are lipids, proteins and sugars, and can therefore be considered as the fuels for the oxidative phosphorylation. When we define a fuel as a compound that acts as a substrate for an ATP producing pathway, and that can be stored to some extent, we should include anaerobic processes as well, and consider both ATP and PCr as fuels. Biochemically speaking we know only two types of ATP synthesis: (1) chemically driven reactions like the pyruvate kinase reaction (substrate phosphorylation); and (2) electrochemically driven reactions like the H+-driven ATP synthesis in the inner mitochondrial membrane. The latter is more important from a quantitative point of view, i.e. 18 times as much ATP is produced by the mitochondria than by glycolysis during complete degradation of glucose. However, the energy generating processes under anaerobic condition are for most animals crucial for survival, since oxygen shortage is a regularly occurring phenomenon either on the tissue level (due to ischemia or high consumption level) or particularly with fish on the organismal level (due to low environmental 02) 22,43,1~176 Therefore, although the total ATP production capacity of anaerobic processes is limited in comparison with that of oxidative processes, we feel compelled to discuss the fuels for both pathways separately.

II. Q u a n t i f y i n g e n d o g e n o u s fuels: destructive m e t h o d s

The quantification of endogenous fuels and metabolites especially for the purpose of describing physiological processes, is difficult because measurement always implies

35

Endogenous [uels; non-invasive versus invasive approaches

interference. It is therefore crucial for the interpretation of the data to know to what level the process under study is disturbed by the determination of a certain parameter. Thus far most measurements are based on destructive methods (chemical and enzymatic reactions), for which an extract is required. Few people realize the problems associated with tissue sampling, extraction, and metabolite measurement. Depending upon the metabolite in question, the applied method of sampling, and the method of extraction, the concentration may vary by more than an order of magnitude, which should make us at least very cautious with respect to the interpretation of the results. The number of different procedures indicate already how difficult it is to obtain reliable metabolite concentrations. Since most metabolite measurements are based on chemical reactions with and/or purification of the compound in question, a solution of the metabolite must be obtained from a previous tissue extraction. We can distinguish 3 phases in an extraction procedure, and within each phase a number of steps (Fig. 1). Phase I refers to the way the animal is handled: how it is taken out of its box and manipulated in order to obtain tissue samples. Animals may be killed by electrocution 76, by a blow on the head, by decapitation, by anesthesia, a combination of these or even by immersion in liquid nitrogen a9,136 (see also Table 1). The major problem in this phase is to prevent struggling of the animal, particularly when one is interested in resting values. Certainly electrocution is the poorest and anesthesia the best way to reach resting values. On the other hand, one should take into account that during the period necessary to reach anesthesia, animals (after exposure to anoxia or exercise) have time for recovery (5-10 min). Phase II is the sampling phase. In this phase samples are taken by a biopsy needle, or by dissection. This takes time, depending on the skill of the operator

I Precauti~ I I sampling I I freezing I I extracti~ 1

l~r+ ~AI

[~wder-~AI

/

I~I

1 , high speed mixer/centrifugation

I

Fig. 1. Steps in tissue extraction. At every step artifacts may develop, disturbing the final metabolic picture. Only very critical consideration of the procedures may result in an acceptable estimation.

36

G. van den Thillart and M. van Raaij

TABLE 1 Muscle lactate under 'resting' conditions Species

Lactate (/zmol g-l)

lmmobilisation

Sampling

Trout49 Trout/carp 95 Trout98 Trout1~ Trout25 Tuna 5 Cod16 Perch76 Human 1~ Eel 124 Goldfish112 Dog 17

15 14 13 1 6 7 5 3.5 3 2.4 1.5 1.2

Liquid N2 A + blow a Blow A Decapitation A + blow Blow Electrocution A + biopsy A + curare A A + isol. prep.

Excision at-20~ Tissue in liquid N2 Tissue in liquid N2 Freeze clamp Freeze clamp Freeze clamp Freeze clamp Tissue in liquid N2 Liquid N2, freeze dry Freeze clamp Freeze clamp Freeze clamp

a A = anaesthetic.

and the number and kind of tissues that have to be sampled. To reduce the loss of precious time in this phase, dissection is sometimes carried out on frozen tissue. Normally the sampling phase is terminated by freeze-clamping the tissue at liquid nitrogen temperature (-195.8~ 1~ Sometimes this step is left out, and the tissue is extracted immediately in cold perchloric acid (PCA). Freezing is however the best way to 'freeze' metabolism and bring it within a few milliseconds to a complete stop 138. Obviously this is important when one is dealing with processes that have high reaction rates. Phase III includes extraction and denaturation of the sample. In order to extract and denaturate the sample properly, it is necessary to pulverize the frozen sample together with the extraction medium to a fine powder 1~2,~5. This way the time for denaturation is minimized, and total surface area for extraction is maximized. This step can be left out only when slow metabolic processes are studied. Although acid will eventually hydrolyze compounds like ATP and PCr, the rate of hydrolysis is only a few percent per day, and therefore in most cases negligible. Extraction and denaturation occur during thawing, therefore it is obligatory to mix the powdered tissue with the PCA during thawing thoroughly. At each step artifacts may be introduced, sometimes leading to spurious results. We can distinguish five conditions where artifacts are likely to develop: handling stress, tissue excision, tissue extraction, tissue and extract storage, and metabolite measurement. 1. Handling stress

Except for blood sampling from cannulated fish, tissue sampling leads to handling stress, since few animals will voluntarily give up a part of their body for scientific

Endogenous fuels; non-invasive versus invasive approaches

37

inspection. So the animals have to be anesthetized and/or killed quickly in order to prevent extreme struggling which will otherwise certainly lead to significant changes of the metabolite profile. Large animals offer the possibility to use biopsy needles, although this type of sampling has its restrictions, too, because of local tissue damage (see below). Handling stress is a behavioral type of stress, a stress reaction initiated by the central nervous system either via direct stimulation or indirectly via hormones (adrenaline and cortisol). A large number of physiological reactions are activated under these conditions, all aimed at preparing the animal for an outburst of activity by redirecting bloodflow to the muscles, stimulating heartbeat and ventilation, increasing muscle tone, etc. All these activities have their effects on tissue metabolism, the more so if the animal is already engaged in struggling. Handling stress can be overcome only if the animal is not able to respond to the sampling; this can be reached either by surprise, by anesthesia, or by a probing technique that is not sensed by the animal. Animals are not easily surprised, certainly not on the level of tissue sampling. The fastest method is needle biopsy, this technique is often used with experiments on humans and larger mammals 1~ but also on fish this technique has been applied 5. It can be carried out fast enough to reduce handling stress, certainly if a local anesthetic is applied, although due to local tissue damage sampling-artifacts cannot be completely prevented (see below). The major problem with anesthesia is the delay; it takes time for the anesthetic to take effect. During this delay recovery processes may take place, especially when they are fast, the original metabolic picture may change completely during a delay of 5-10 minutes. Besides this delay, anesthesia has also side-effects on metabolism such as erythrocyte swelling 96. Repeated anesthesia also changes markedly the levels of blood borne metabolites 12. In the case of resting metabolism anesthesia is the best approach, since the disturbance will be minimal, and no recovery processes are to be expected. In the case of exercise, a fast blow, followed by decapitation, should be preferred in order to prevent recovery during the delay of the anesthetic. The delay problem may be overcome by infusion of an anesthetic via a cannula inserted in the aorta. Cannulation of the dorsal aorta is widely used for acid-base and blood-gas studies, its application for metabolic studies is restricted since only blood can be sampled. As far as we know cannulation has never been used in order to reach a very fast anesthesia after exercise. In some papers on fish metabolism, the experimental fish were inactivated/killed with electrical shock, obviously this method has its limitations, since it will likely stimulate the whole animal and particularly the muscles, thus leading to significant changes in the metabolite profiles. It has been described several times that premortem stress not only reduces the levels of glycogen, PCr and ATP, but also dramatically accelerates the postmortem degradation rate in comparison with anesthetized fish 29'103. The best way to overcome handling stress is to use non-invasive and nondestructive (physical) probing techniques which are not sensed by the animal. The technique currently available is in vivo NMR (nuclear magnetic resonance), which will be discussed separately.

38

G. van den Thillart and M. van Raaij

2. Tissue damage To acquire a piece of tissue, it must be excised, which obviously causes damage to the cells. In addition to local damage, nerves are cut and/or damaged, leading to stimulation of the adjacent tissue via spinal reflexes, thus resulting in general activation of the excised tissue. The metabolic rate of certain tissues can thus be increased enormously, therefore one should employ two different strategies: (1) suppress activation as much as possible; and (2) inactivate the sample as fast as possible. Activation particularly of muscle and neural tissue can be reduced significantly by anesthesia, and muscle relaxants. For example, muscle from anesthetized eel (Anguilla anguilla) responds immediately to incision, only by intracardial injection of curare the spinal reflexes can be suppressed 124. Recently, Arthur and collaborators 5 applied lidocaine at the spinal cord of skipjack tuna (Katsuwonus pelamis) - after previous MS 222 anesthesia - to suppress spinal reflexes during biopsy sampling. To inactivate the sample, freeze-clamping is the ultimate procedure. The aim of sampling is to have a momentary view of the metabolite levels. Using aluminium blocks cooled by liquid nitrogen (-195.8~ Wollenberger and colleagues 138 demonstrated that a piece of tissue can be metabolically put to a standstill within a few msec. This seems fast enough, especially since muscle contractions are slower. The dimensions of the tongs determine the sample size, normally 0.1-2.0 g. With enlarged tongs even whole animals can be freeze-clamped to a weight of about 6 g (refs. 2, 60, 87, 136). Freeze-clamping is not always used, some authors immerse the samples 9s or even whole animals in liquid nitrogen 36.49. This technique is inferior to the clamping method, because the time needed to completely deep-freeze a sample may take several minutes or longer depending on the size. Temperature equilibration depends on distance, temperature-difference and heat-transfer capacity. The equilibration time is exponentially related to both distance and heat transfer capacity, so the distance should be minimal (7 BL s-1) the fish were rapidly exhausted and specific losses were observed for the fatty acids 18:2, 20:4 and 22:6. It is now well known that these fatty acids are associated with phospholipid hydrolysis and since these lipid classes arc normally hardly mobilized 41, this observation suggests that some tissue damage did occur. Nevertheless, it was estimated that during submaximal exercise, coho salmon would derive 45% of its energy from lipid catabolism whereas this value was decreased to about 15% during exhaustive exercise. The latter value however, may be of less significance because the authors did not account for glycogen utilization which was certainly present during exhaustive exercise. In mackerel (Trachuna symme~cus), it was observed that red muscle used endogenous triglycerides for sustained swimmings2 while this was not noted in white muscle. In resting rainbow trout, lipid oxidation may contribute about 20% to the energy metabolism whereas this value was about 10% during sustained swimming ~~ From the kinetics of arterially infused radiolabeled substrates, Van den Thillart 1~ concluded that a preferential oxidation of endogenous substrates occurred during the first hours of sustained aerobic exercise. Recently, S~ingerss demonstrated that the lipid content and the fine structure of red muscle showed considerable species-specific variation. The red muscle of Danube bleak, a cyprinid which is known to be a sustained swimmer, contained more then 10% lipid on a wet weight basis whereas in the Asp, a piscivorous predator which performs mainly burst type exercise, lipid contents of only 2% were observed. From these observations, S~ingerss proposed that these variations in lipid content are associated with a difference in swimming behavior. Interestingly, a positive statistical correlation between the amount of red muscle and mobility was reported ~. During intensive exhausting exercise or during burst type exercise, as can occur frequently in the white muscle of active pelagic or piscivorous species, the pattern of substrate utilization is different. Since oxygen supply is inadequate during these conditions, the changes in fuel mobilization resemble those during hypoxia with the result that metabolic flux is increased by several-fold. Thus, endogenous PCr and glycogen are the preferred substratcs during intensive exercise (see Hypoxia section). However, also during submaximal exercise, the white muscle may use endogenous glycogen as a substrate for anaerobic glycolysis resulting in the production of lactate. This area of fish physiology is well-documented 5'2~176176176 and we will only give an outline describing the general findings. The degree of glycogen depletion and the accumulation of lactate arc positively correlated with the intensity and the duration of the exercise24. Mobilization of glycogen may be extremely rapid and values of about 40 ~mol g-1 s-1 have been reported 24. Glycogenolysis in the muscle tissues is probably initiated by increased levels of Ca 2+ which is released from the sarcoplasmatic reticulum. These ions act upon protein kinase which subsequently activates glycogen phosphorylasc 44. The lactate formed in the white muscle diffuses to some extent to the blood, although the degree of this process seems to be species specific. Active pelagic fish, are sometimes called 'lactate-releasers' since their blood lactate levels increase significantly during exercise while in benthic inactive species blood lactate is elevated only modestly. The latter species have therefore been called , lactate-non-releasers ,79 ' 104 . As stated

Endogenous fuels; non-invasive versus invasive approaches

55

above, lactate released into the circulation is a preferred substrate for aerobic metabolism in other tissues or for gluconeogenesis in the liver ~~

3. Starvation and migration A number of fish species will encounter prolonged periods of starvation during their life, often seasonally dependent and associated with migration and reproduction. The utilization of endogenous fuels is dependent on the length of the starvation period and the species under investigation. A considerable number of papers have been published on this area during the last 30 years. In this section we will describe the generalities that have arisen from this research emphasizing the importance of mobilizing endogenous fuels for energy metabolism. With respect to the mobilization of glycogen stores, the fish species studied so far may be divided in two categories: species which do or do not mobilize glycogen during the initial phase of starvation. Glycogen utilization during the first stage of starvation was found in common carp (C. carpio), roach (R. rutilus), killifish (Fundulus heteroclitus), rainbow trout (O. mykiss) and brown trout (Salmo trutta). During wintering of common carp, Takeuchi and Ishii 1~176 found that liver and (white) muscle glycogen levels were reduced by 65 and 80% respectively. Glycogen contents of the carcass of juvenile roach were decreased from 5.1 to about 1.6 /zmol g-1 after about three weeks and remained at this level during the remainder of the starvation period (50 days) 67. Similarly, whole body glycogen was depleted by 50% after 5 days of starvation in juvenile rainbow trout ss while glucose availability was significantly enhanced. The liver glycogen pool of killifish was depleted by over 90% after fasting for five weeks 62. Recently, Navarro et al. 75 demonstrated glycogen depletion in the liver and muscle of brown trout of about 80 and 20% respectively already after eight days of starvation, although after 4 weeks the glycogen contents were partly restored (from gluconeogenesis). In common carp, starved for about one month, no significant mobilization of liver glycogen is observed whereas after three months, these stores were depleted to 20% of the initial value 73. Thus carp may display an intermediate response. Representatives of the second category are migrating sockeye salmon (Oncorhynchus nerka) and fasting European (Anguilla anguiUa) and American (A. rostrata) eels. During their migration to the spawning grounds, the liver and muscle glycogen levels of salmon are not significantly changed and are in fact increased just prior to spawning 3~ The glycogen content of liver and red muscle of American eel was not affected after six months of starvation while white muscle glycogen was decreased by 40% (ref. 69). Similarly, Larsson and Lewander 61 showed that liver glycogen of European eels was not decreased during the first three months of starvation but was decreased by 40% after about five months. Muscle glycogen was not changed at all. However, these observations do not rule out the possibility of carbohydrate catabolism since liver glycogen may be continuously replaced by gluconeogenesis as was proposed by Larsson and Lewander 61. Increased gluconeogenetic activity from lactate and alanine was observed in migrating salmon 3~ Liver glycogen contents of fasting cod were extremely low 54 (Table 2) and although these levels were reduced during

56

G. van den ThiUan and M. van Raaij

starvation, liver glycogen is probably of minor quantitative importance in this species. In the second category, glycogen is only used as energy source at prolonged starvation when the availability of other fuels (e.g. lipids) is reduced. In the first category, glycogen utilization may be of transient importance during the first stage of starvation until the mechanisms for utilization of other substrates are activated. Lipids and protein are then the major fuels for energy metabolism. Lipid mobilization is a common strategy in fasting fish although there appears to be some variation with respect to target tissues. Visceral lipid depots are easily mobilized and decline almost immediately after cessation of feeding 4s. Especially in salmonids, the mesenteric fat deposits are the first to be depleted 4s,75. The visceral index of brown trout decreased from 8% of total body weight to about 5% after starvation for one week 7s. A similar mechanism may be operative in fasting American eel (tt. rostrata) as well since a significantly increased [FFA] was observed in the plasma while liver and muscle lipids were not affected 69. Similarly, increased plasma [FFA] was also found in fasting plaice (Pleuronectus platessa) 13s. Lipid mobilization from liver and red muscle was observed in European eel (,4. anguilla) 21'61, rainbow trout 48,84, and carp 19,73. The lipid depletion of these tissues is much more gradual then of visceral depots and will contribute to the energy metabolism during the whole starvation period. In general, there appears to a preferential mobilization of saturated fatty acids from visceral depots while saturates and monounsaturates are derived from liver and muscle lipids. These fatty acids indicate the mobilization of triglycerides, the polyunsaturated fatty acids of the phospholipid fraction are usually retained 41'134. It is generally believed that during prolonged starvation, proteins are the major source of energy. The nitrogen loss of several tissues of common carp was found to decrease in the following order: muscle > spleen > kidney > liver > intestine 19. In particular the large mass of white muscle is believed to be a huge reservoir of energy equivalents. Protein utilization was observed in the muscles of starving eel (A. rostrata) 69 and in migrating sockeye salmon (O. nerka) 68. In both species, the 'insoluble' myofibrillar proteins were mostly affected, whereas the 'soluble' fraction was relatively unaffected. Based on changes in enzyme activities (especially alanine aminotransferase) M6ndez and Wieser 67 concluded that protein catabolism was enhanced also in fasting juvenile roach (Rutilus rutilus). Thus, amino acids may be mobilized from tissues proteins, especially in white muscle, and serve as substrates for energy metabolism either directly via catabolism in situ or via gluconeogenesis. The latter process requires inter-organ transport, since the highest gluconeogenetic activities are found in the liver. It has indeed be observed that the white muscle of migrating salmon releases relatively large amounts of alanine 68 and, moreover, gluconeogenesis from alanine was increased in hepatocytes from migrating salmon 3~ as well as from fasted rainbow trout 14. Increased protein catabolism during starvation is enabled by increased activities of proteolytic enzymes like cathepsin and other acid and neutral proteinases during migration of salmonids 3'4'68. In addition, it was found that the rate of protein synthesis in the white muscles of fasting rainbow trout and carp was decreasedl 1,53.

Endogenous fuels; non-invasive versus invasive approaches

57

In conclusion, the energy metabolism in fasting and migrating fish is fuelled primarily by amino acids and fatty acids. However, glycogen may be an additional substrate either during the first stage of starvation or only during prolonged periods of starvation.

VI. Summary Fuels are compounds that act as substrates for ATP producing pathways and can be stored to a certain extent. Fuels for anaerobic processes are ATP, PCr and glycogen. These anaerobic processes have a limited capacity, however, their survival value is very high, since they allow either a very high energy flux (exercise) for a short period, or a low energy flux during hypoxia/anoxia for a long period. Fuels for aerobic processes are sugars, lipids and proteins. During catabolism they produce NADH and FADH2 which are the ultimate substrates for the electron transport chain and mitochondrial ATP synthesis. The quantitation of fuels is fundamental to the understanding of energy metabolism. Most methods are of a destructive nature and cause as such a series of possible artifacts: stress due to handling and sampling, tissue damage, and incomplete metabolite extraction and denaturation of proteins. Most problems occur with those pathways where a high energy flux can be generated such as in muscle, i.e. it is very difficult to obtain low levels for lactate, and a high PCr/total creatine ratio. Another source of interference is where low metabolite levels occur together with high fluxes; this is found with FFA: low levels are easily disturbed due to lipolysis. Non-destructive techniques preclude most but not all problems. The most promising technique is in vivo NMR, which allows metabolite measurements without invasive or destructive actions. Particularly important is the finding that under resting conditions the phosphorylation potential of muscle is very high, resulting in >90% phosphorylation of creatine. Furthermore a review is given of the range of different fuels occurring in different tissues and different species. Obviously glycogen is quantitatively of minor importance in fish. The major energy source is protein, although lipids are in some species of equal importance. Fuel mobilization under hypoxia, exercise and starvation is discussed. Under conditions where anaerobic metabolism is activated, PCr and glycogen are the major fuels, while for long term exercise and starvation, both lipids and proteins are the predominant source of energy.

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128. Van Waversveld, J., A.D.F Addink and G. Van den Thillart. Simultaneous direct and indirect calorimetry on normoxic and anoxic goldfish. J. Exp. Biol. 142: 325-335, 1989. 129. Veech, R.L., J.W.R Lawson, N.W. Cornell and H.A. Krebs. Cytosolic phosphorylation potential. I. Biol. Chem. 254: 6538-6547, 1979. 130. Von der Decken, A. and E. Lied. Dietary protein levels affect growth and protein metabolism in trunk muscle of cod, Gadus morhua. J. Comp. Physiol. 162B: 351-357, 1992. 131. Waiwood, B.A., K. Haya and L. Van Eeckhaute. Energy metabolism of hatchery-reared juvenile salmon (Salmo Salar) exposed to low pH. Comp. Biochem. Physiol. 101C: 49-56, 1992. 132. Walker, R.M. and P.H. Johansen. Anaerobic metabolism in goldfish (Carassius auratus). Can. J. Zool. 55: 1304-1311, 1977. 133. Walton, M.J. and C.B. Cowey. Aspects of intermediary metabolism in salmonid fish. Comp. Biochem. Physiol. 73B: 59-79, 1982. 134. Watanabe, T Lipid nutrition in fish. Comp. Biochem. Physiol. 73: 3-15, 1982. 135. White, A. and TC. Fletcher. Serum cortisol, glucose and lipids in plaice (Pleuronectus platessa L.) exposed to starvation and aquarium stress. Comp. Biochem. Physiol. 84: 649-653, 1986. 136. Wieser, W., U. Platzer and S. Hinterleitner. Anaerobic and aerobic energy production of young rainbow trout (Salmo gairdneri) during and after bursts of activity. J. Comp. Physiol. 155B: 485-492, 1985. 137. Williamson, J.R. and B. Corkey. Assays of intermediates in the citric acid cycle and related compounds by fluorometric enzyme methods. In: Meth. Enzymol., edited by J.H. Lowenstein, New York, Academic Press, pp. 435-512, 1969. 138. Wollenberger, A., O. Ristau and G. Schoffa. Eine einfache Technik der extrem schnellen Abk0hlung gr6sserer Gewebestiicke. Pfluegers Arch. Eur. I. Physiol. 270: 399-412, 1960.

Hochachka and Mommsen (eds.), Biochemistryand molecularbiology of fishes, vol. 4 9 1995 Elsevier Science B.V. All rights reserved. CHAPTER 4

Tissue carbohydrate metabolism, gluconeogenesis and hormonal and environmental influences THOMAS W. MOON AND GLEN D. FOSTER

Department of Biology, Ottawa-Carleton Institute of Biology, University of Ottawa, Ottawa, Ontario KIN 6N5, Canada

I. II.

Introduction Liver 1. The general organization of hepatic metabolism 2. Environmental adaptations 2.1. Fasting 2.2. Hypoxia 2.3. Temperature 2.4. Stress 2.5. Seasonality 3. Hormone signal transduction pathways III. Kidney 1. The organization of kidney metabolism 2. Environmental adaptations IV. Skeletal muscle and heart 1. Myotomal muscle 2. Cardiac muscle V. Brain VI. Red blood cells VII. Conclusions VIII. References

I. Introduction Carbohydrates are key to the metabolism of all vertebrates, including fish species. The diversity of lifestyles and habitats selected by fish has resulted in significant differences in the way species handle and partition dietary carbohydrates within their bodies. Few generalities can be presented, and the reader is directed to Love 9~ for a review of some of the issues which may be involved. Early studies by Leibson 87, Plisetskaya and Kuz'mina 134, and Palmer and Ryman 123 introduced the ideas of motor activities affecting carbohydrate disposition and glucose intolerance, but papers since have generally lost the comparative perspective which is critical to our understanding these species differences. The idea that at least carnivorous fishes (e.g. salmonids) have a poor tolerance to carbohydrate has been revisited

66

T.W. Moon and G.D. Foster

recently 133,177. It is clear that this is an issue in the metabolic biochemistry of fishes, but more information is needed. In particular, the sensitivities of hormonal release to circulating carbohydrate and the specific tissue hormone receptors and species with non-carnivorous life styles need to be examined. Although this review will mention a variety of species, many have similar nutrient requirements and a strong comparative approach is needed before an understanding of this issue will be possible. It remains unclear whether any fish tissues require or use preferentially carbohydrate as a source of energy. The red blood cell and kidney cortex of mammals have a strict glucose requirement based upon their anaerobic metabolic profile. Fish red cells are nucleated and may contain a few mitochondria, and it is reported that 90% of the resting nucleoside triphosphate is produced aerobically 33. Red cells of the sea raven (Hemitripterus americanus) also have an aerobic metabolism fueled by exogenous glucose x46. Lamprey (Petromyzon marinus) brain tissue minces do utilize glucose 4s and the trout brain is thought to use primarily exogenous glucose to supplement its low endogenous glycogen reserves 27. Glucose/glycogen is critical for the maintenance of cardiac performance in the Atlantic hagfish (Myxfne glutinosa) 151. Yet in no case is the quantitative significance of carbohydrates in overall tissue metabolism clearly defined. The ability of tissues to utilize a particular nutrient is dependent on the permeability of the cell to it and its cellular metabolism to ensure adequate membrane gradients. Carbohydrates such as glucose and lactate poorly penetrate the hydrophobic cell membrane, and transport is generally carrier-mediated. Fish tissues such as red cells and liver, with few exceptions, show non-saturable uptake kinetics with respect to these metabolites (red cells 16~ liver171). Hexokinase (HK) activities, the enzyme responsible for glucose phosphorylation and the maintenance of the membrane gradient for glucose, are generally low in fish tissues (see Tables 2-8), and although there is some evidence for its modulation by diet 37, there is no evidence for a mammalian-type glucokinase in fish. Unfortunately, our knowledge of glucose transporters in fish tissues is poor, with the possible exception of red cells 3s,lss, an area which could help identify the mechanism of 'glucose intolerance' in fish. Fish tissues do contain glycogen in varying quantities (Table 1) indicating that the inability of tissues to take-up glucose or maintain plasma glucose content is compensated for by other metabolic pathways. Gluconeogenesis is the pathway responsible for de novo glucose and glycogen synthesis (glyconeogenesis) from precursors including lactate, amino acids, glycerol, and fructose 1~ The importance of this pathway to tissue carbohydrate homeostasis in fish has been the subject of several recent reviews97,1~ The high dietary protein requirements of fish9~ should provide adequate substrate for this pathway and there is extensive evidence for the modulation of gluconeogenic rates by intrinsic and extrinsic factors. The prevalence of proteins in the carnivorous diet and their use as an energy source together with the generally lowered energy demands of fish n4, may have alleviated the strict need for carbohydrate as a key energy source to many fish species. The

Tissue carbohydrate metabolism, gluconeogenesis and hormonal and environmental influences

67

TABLE 1 Glycogen contents of various fish tissues Species/Condition

Liver

Hagfish fed fasted; 1 month fasted; 4 months anoxia

15.039 1.8 4.7 -

Skate

6311~

Crucian carp winter/normoxic summer Catfish winter/fed fasted; 60 days spring anoxic

Kidney

3039 0.72 36

Red muscle -

Heart

Brain

2257 0.9557

9.011~

150066,117 5566 600118 8012~ 67 31

White muscle

167117 22 _ 3.7 2.4 n

19118 4.2120 1.7 3.3 D

140118 12.912o 7.3 7.9 _

211118 4212o 30 23 _

1627 _

627

Perch fed fasted; 7 weeks Flounder normoxic hypoxic Tuna American eel fed fasted, 6 months Rainbow trout normoxic anoxic

62145 182 6476 18 4178 49 l~ 53 3526, 2000 u

15.345 10.8 45 TM

16TM 4

36

93178

24178

121o6 7

141o6 11

5.126 _

_

_

_

_

_

10136 3.027 1.527

Values are given in/zmol glucosyl units g-I tissue. These are representative values and not intended to be comprehensive. Superscript numbers represent reference numbers. If values are not followed by superscript numbers, reference is identical to that located above in the same column.

role of carbohydrates may, therefore, be for short-term responses to acute stress situations 19 and/or as a last resort. The purpose of this review, therefore, is to provide an overview of recent studies on carbohydrate metabolism in fish using a tissue approach. An overriding thesis in the review is that carbohydrate metabolism has a significant role in fish, and that this function becomes more understandable when considered in light of perturbations of the system. The question of whole fish carbohydrate homeostasis will be examined only in terms of its importance to the individual tissue. An attempt will be made to update those previous reviews on fish metabolism (e.g. refs. 19, 61, 176), and by doing so, to identify those areas which need further research.

68

T.W.Moon and G.D. Foster

II. Liver The liver is a central organ of metabolism and is key to the regulation of carbohydrate metabolism in all vertebrate classes. A huge literature is available and it is not our intent to be all-inclusive, but to select those areas of recent interest.

1. The general organization of hepatic metabolism The ability of fish to metabolize carbohydrate and the enzymes required for this purpose are clearly understood. A general listing of recently published enzyme activities in fish livers is provided on Table 2. A complete enzyme profile is not available for each fish, but as methods become more available, such holes will disappear. The overall importance of carbohydrate metabolism in the fish liver as well as the relative importance of glycolysis and gluconeogenesis, however, are less well defined. For instance, hepatectomy in the fiver lamprey (Lampetra fluviatilis) s6 and the Pacific hagfish (Eptatretus stouti) 69 affected neither survival nor basal blood glucose concentrations, leading to the suggestion that liver carbohydrate metabolism may not be critical to whole body carbohydrate homeostasis. These studies, however, did not consider the ability of these species to tolerate environmental stress, such as hypoxia, exercise, fasting, or temperature changes, nor do they explain the ubiquity of carbohydrate-metabolizing pathways in fish livers. We suggest that while the importance of carbohydrate metabolism in the whole animal energy budget may be less than in other vertebrate classes, carbohydrate metabolism is critical and becomes so during adaptive responses to the environment. Liver glycogen contents are extremely variable in fish (Table 1). Agnathans generally have levels below 20/zmol g-l, while elasmobraneh and teleost values range from 20 to 2000/zmol g-1. Measured glycogen values for fed rainbow trout (Oncorhynchus mykiss) range from 35 (ref. 26) to 2000 (ref. 84)/zmol g-1. This variability may reflect sampling procedures, strain and/or life history differences, or even the method of analysis. The Crucian carp (Carassius carassius) has liver glycogen concentrations above 1500/zmol g-1 liver, and during periods of high glycogen content, the liver may reach 15% of body weight66. This gives a total liver carbohydrate store of over 20,000/zmol 100 g-1 body weight, or 4% of body weight! A similar analysis using other teleosts gives values no higher than 1500/zmol g-1 body weight (0.3%). While other teleosts do not demonstrate these high amounts of liver glycogen, the liver still contains as much as 50% of the whole animal glycogen stores, assuming skeletal muscle contains the rest. The hormones glucagon, glucagon-like peptide (GLP), and catecholamines all increase gluconeogenesis and glycogenolysis in a variety of species by inhibition of pyruvate kinase (PK) and phosphofructokinase-1 (PFK-1) activities and increasing phosphoenolpyruvate earboxykinase (PEPCK) and glycogen phosphorylase (GPase) activities 23'42'44'47'56'98'100'102'103'118'119'130'167.These enzymes have all been reported in most species studied (Table 2), and most are controlled by reversible phosphorylation-dephosphorylation mechanisms. The vasoactive peptides vasotocin and isotocin also increase glycogenolysis and glueoneogenesis in three species

Tissue carbohydrate metabolism, gluconeogenesis and hormonal and environmental influences

69

where they have been tested, with the effects very pronounced in the American eel (Anguilla rostrata) 111. Elevated plasma cortisol concentrations stimulate gluconeogenesis in some teleosts, possibly by increasing the availability of gluconeogenic substrates (amino acids) and activation of specific liver enzymes 1~8. It is not clear to this point whether this is a direct affect of cortisol or it is mediated by other glucoregulatory hormones 169. Insulin, the only hypoglycemic hormone in mammals, acts to counteract glucagonstimulated gluconeogenesis and glucose production 65, with little or no effect on basal (no hormone) rates. This situation with insulin-counteracting the effects of glucagon on gluconeogenesis and glycogenolysis has been reported for American eel 42 and sea raven 41'46 hepatocytes. Petersen et al. 130, however, reported an inhibition of basal (no hormone) gluconeogenic flux by insulin in trout hepatocytes, with a concurrent inhibition of PK activity. This inhibitory effect of insulin in the absence of glucagon has been found in no other teleost species nor in mammals where PK is unaffected 65. In fact, insulin actually increased gluconeogenic flux in hepatocytes isolated from sea raven 41,46, American eel 42, and hagfish 49. These actions of insulin occurred through an enhanced inhibitory effect of ATP on PFK-1 in all three species. No effects of insulin on PK and PEPCK were found in sea raven hepatocytes. Insulin also decreased glucagon-stimulated glycogenolysis and decreased glycogenolysis below basal (no hormone) levels in both sea raven and American eel hepatocytes 41,42. It is apparent that the actions of insulin differ between species and with respect to the carbohydrate pathways in the liver. Mammalian hepatocytes show metabolic heterogeneity such that periportal cells catalyze glucose release and gluconeogenesis while perivenous cells preferentially take up glucose for glycogenesis and lipogenesis 77. No evidence for this mammalian form of heterogeneity has been found in the catfish 121 or the trout 99 liver. However, density gradient separation of cell types demonstrated more oxidative and gluconeogenic scope existed in the less dense cells of trout liver 99, and greater overall enzyme activities in the less dense cells of the Gulf toadfish (Opsanus beta) 1~ An interesting finding in the toadfish study was that although enzyme activities were higher in the less dense cells, the heavier cells exhibited 2.5-4 times more metabolic activity than the less dense population. Similarly, the distinctively large lobe of the hagfish liver was found to be more metabolically active than the small lobe 49. Metabolic heterogeneity may exist in fish livers, although rather than a segregation of metabolic pathways existing, as in the mammal, the heterogeneity may be more subtle or simply reflect differential overall metabolic activities of the cells. A review of substrate utilization has been published previously 1~ and the general principles will not be covered here. This section will concentrate on recent findings that relate to an understanding of the fate of certain substrates, as well take a comparative look at some interesting differences found in some species. Lactate is generally the most readily utilized substrate in fish liver 1~ leading to the suggestion that the liver is important in the Cori cycle 157 (Cori cycle is: muscle lactate ~ liver glucose ~ muscle glycogen). However, in starry flounder (Platichthys stellatus) 96 , American eel 21 and skipjack tuna (Katsuwonus

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