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The unusual energy metabolism of elasmobranch fishes


Comparative Biochemistry and Physiology, Part A 155 (2010) 417–434

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Comparative Biochemistry and Physiology, Part A
j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / c b p a

Review

The unusual energy metabolism of elasmobranch ?shes☆
Ben Speers-Roesch a,?, Jason R. Treberg b
a b

Department of Zoology, University of British Columbia, 6270 University Blvd., Vancouver, British Columbia, Canada V6T 1Z4 Ocean Sciences Centre, Memorial University of Newfoundland, St. John's, Newfoundland, Canada A1C 5S7

a r t i c l e

i n f o

a b s t r a c t
The unusual energy metabolism of elasmobranchs is characterized by limited or absent fatty acid oxidation in cardiac and skeletal muscle and a great reliance on ketone bodies and amino acids as oxidative fuels in these tissues. Other extrahepatic tissues in elasmobranchs rely on ketone bodies and amino acids for aerobic energy production but, unlike muscle, also appear to possess a signi?cant capacity to oxidize fatty acids. This organization of energy metabolism is re?ected by relatively low plasma levels of non-esteri?ed fatty acids (NEFA) and by plasma levels of the ketone body ?-hydroxybutyrate that are as high as those seen in fasted mammals. The preference for ketone body oxidation rather than fatty acid oxidation in muscle of elasmobranchs under routine conditions is opposite to the situation in teleosts and mammals. Carbohydrates appear to be utilized as a fuel source in elasmobranchs, similar to other vertebrates. Amino acid- and lipidfueled ketogenesis in the liver, the lipid storage site in elasmobranchs, sustains the demand for ketone bodies as oxidative fuels. The liver also appears to export NEFA and serves a buoyancy role. The regulation of energy metabolism in elasmobranchs and the effects of environmental factors remain poorly understood. The metabolic organization of elasmobranchs was likely present in the common ancestor of the Chondrichthyes ca. 400 million years ago and, speculatively, it may re?ect the ancestral metabolism of jawed vertebrates. We assess hypotheses for the evolution of the unusual energy metabolism of elasmobranchs and propose that the need to synthesize urea has in?uenced the utilization of ketone bodies and amino acids as oxidative fuels. ? 2009 Elsevier Inc. All rights reserved.

Article history: Received 5 August 2009 Received in revised form 28 September 2009 Accepted 29 September 2009 Available online 9 October 2009 Keywords: Fatty acid Ketone body Amino acid Carbohydrate Substrate oxidation Fuel preference Urea Evolution Fish Chondrichthyes Review

Contents 1. 2. Introduction . . . . . . . . . . . . . . . . . . . . . . . . Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Capacity for fatty acid oxidation . . . . . . . . . . . . 2.2. Fatty acid transport . . . . . . . . . . . . . . . . . . 2.3. Hepatic lipid storage. . . . . . . . . . . . . . . . . . 2.4. Physiological regulation . . . . . . . . . . . . . . . . Ketone bodies . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Ketone bodies as routine metabolic fuels . . . . . . . . 3.2. Physiological regulation . . . . . . . . . . . . . . . . Amino acids . . . . . . . . . . . . . . . . . . . . . . . . 4.1. Amino acid oxidation . . . . . . . . . . . . . . . . . 4.2. Role of amino acids in gluconeogenesis and ketogenesis . 4.3. Amino acid transport . . . . . . . . . . . . . . . . . 4.4. Physiological regulation . . . . . . . . . . . . . . . . 4.5. Lessons from whole animal physiology . . . . . . . . . Carbohydrates . . . . . . . . . . . . . . . . . . . . . . . Environmental in?uences . . . . . . . . . . . . . . . . . . 6.1. Salinity . . . . . . . . . . . . . . . . . . . . . . . . 6.2. Hypercapnia and hypoxia . . . . . . . . . . . . . . . 6.3. Temperature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 418 418 418 420 421 422 422 422 423 424 424 424 425 425 425 425 426 426 426 426

3.

4.

5. 6.

☆ This paper derives from a presentation at the session entitled ‘Biology of Elasmobranchs: from Genes to Ecophysiology and Behaviour’ at the Society for Experimental Biology's Annual Main Meeting, Glasgow, 28 June–1 July, 2009. ? Corresponding author. Tel.: +1 604 822 4201; fax: +1 604 822 2416. E-mail address: bensr@zoology.ubc.ca (B. Speers-Roesch). 1095-6433/$ – see front matter ? 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.cbpa.2009.09.031

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Energy metabolism and the incidence of cancer in elasmobranchs Why do elasmobranchs have an unusual energy metabolism? 8.1. Paleozoic atmospheric oxygen levels . . . . . . . . . 8.2. Effects of TMAO on fatty acid oxidation . . . . . . . . 8.3. Buoyancy role of the liver . . . . . . . . . . . . . . 8.4. The urea–albumin hypothesis. . . . . . . . . . . . . 8.5. Requirement for urea synthesis . . . . . . . . . . . . 8.6. Random loss of muscle fatty acid oxidation machinery . 9. Comparative and evolutionary considerations . . . . . . . . 9.1. Basal Actinoptergyii . . . . . . . . . . . . . . . . . 9.2. Sarcoptergyii . . . . . . . . . . . . . . . . . . . . 9.3. ‘Agnatha’: Myxini and Petromyzontida . . . . . . . . 9.4. Other Chondrichthyes . . . . . . . . . . . . . . . . 9.5. Energy metabolism of early vertebrates . . . . . . . . 10. General conclusions . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1. Introduction Elasmobranchs are an ancient group of carnivorous ?shes that ?rst appeared at least 400 million years ago (mya) (Grogan and Lund, 2004). They are found worldwide in both marine and freshwater environments, from the deep-sea and polar oceans to coral reefs and tropical rivers (Compagno, 1990). Elasmobranchs and their close relatives the Holocephali comprise the monophyletic class Chondrichthyes, which represent the basal extant gnathostomes (jawed vertebrates) (Grogan and Lund, 2004; Kikugawa et al., 2004). The Chondrichthyes have a fossil record that traces back to at least 450 mya (Miller et al., 2003). The ancient evolutionary history of elasmobranchs and their relatively simple body plan has long made them an attractive research subject in ?elds such as evolutionary developmental biology, comparative morphology, and comparative physiology. Certain metabolic and physiological peculiarities of elasmobranchs, including their unusual osmoregulatory strategy and organization of energy metabolism, have been of particular interest to physiologists. Marine elasmobranchs as well as holocephalans are osmoconforming ionoregulators that synthesize and accumulate relatively high levels of ureatohelpmaintainosmoticequilibriumbetweeninternalsolutelevels andthesurroundingseawaterwithouttheneedforhighlevelsofinorganic ions (Yancey, 2001). The perturbing effects of urea on macromolecular structures are thought to be counteracted to a certain extent by coaccumulation of trimethylamine oxide (TMAO) and other methylamines (Yancey, 2001). Whereas detailed investigations of the ureabased osmoregulation of elasmobranchs were underway as early as the 1930s (Smith, 1936), little was known about their energy metabolism until the 1970s when the prominent British metabolic biochemists V.A. Zammit and E.A. Newsholme provided the ?rst indication that unlike most other vertebrates, elasmobranchs show an increased reliance on ketone bodies rather than fatty acids as oxidative substrates in muscle (1979). Newsholme's group at Oxford at the time was investigating the metabolic biochemistry of muscle from a wide variety of animals, including insects, ?shes, and tetrapods, in order to learn about the diversity of fuels utilized for ATP production. After completing a doctorate on invertebrate anaerobiosis under Newsholme's supervision, Zammit stayed on to follow up on preliminary measurements of relatively low enzyme activities related to ketone body oxidation in teleosts. Working at Plymouth Marine Laboratory, Zammit found that the activity of β-hydroxybutyrate dehydrogenase, an important enzyme involved in ketone body oxidation, was low in teleost muscle and high in elasmobranch muscle. Conversely, the activity of carnitine

palmitoyltransferase, a key enzyme in mitochondrial long-chain fatty acid oxidation, was non-detectable in elasmobranch muscle but relatively high in teleost muscle. Subsequent work by comparative biochemists and physiologists, including but not limited to J.S. Ballantyne, W.R. Driedzic, T.W. Moon, and C.D. Moyes, greatly increased our knowledge of the organization of energy metabolism in elasmobranchs, provided evolutionary context via comparative studies on teleosts and other ?shes, and suggested possible explanations for its origin. J.S. Ballantyne synthesized much of this work in his comprehensive and seminal review of elasmobranch metabolism (Ballantyne, 1997). Since then, further discoveries have re?ned and enhanced our comprehension of this topic. In this review, we provide a broad survey of our current understanding of the energy metabolism of elasmobranchs and suggest future research directions. We have structured our review around the major oxidative fuels: lipids, ketone bodies, amino acids, and carbohydrates. In Sections 2–5 we provide a comprehensive overview on their utilization and relative importance in elasmobranchs. In Sections 6 and 7 we summarize what is known about environmental in?uences on the energy metabolism of elasmobranchs and brie?y discuss the potential link between energy metabolism and the incidence of cancer in elasmobranchs. In Section 8 we explore potential explanations and suggest new hypotheses for why the energy metabolism of elasmobranchs is unusual. In order to distinguish the metabolic organization of elasmobranchs, we provide comparisons to two other major vertebrate groups, teleosts and mammals, whose metabolisms are well known compared with elasmobranchs. In Section 9 we extend our comparisons to other groups of ?shes, including basal actinopterygians, lobe-?nned ?shes, jawless ?shes, and holocephalans, in order to put the energy metabolism of elasmobranchs into a broader evolutionary context. Finally, in Section 10 we provide general conclusions on the unusual energy metabolism of elasmobranchs. 2. Lipids 2.1. Capacity for fatty acid oxidation Carnitine palmitoyltransferase (CPT) is a mitochondrial enzyme that is essential for oxidation of long-chain fatty acids (12 to 22 carbons in length), a major energy source for oxidative phosphorylation (Kerner and Hoppel, 2000). CPT is present as two proteins, CPT-1 and CPT-2, which are localized to the outer and inner mitochondrial membranes, respectively, and facilitate the carnitine-dependent entry of long-chain

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fatty acids into the mitochondrial matrix, where β-oxidation occurs (McGarry and Brown, 1997; Kerner and Hoppel, 2000). CPT-1 is an important regulatory step in long-chain fatty acid oxidation due to allosteric inhibition by malonyl CoA. Malonyl CoA is a lipogenic intermediate formed from acetyl CoA in a reaction that is closely linked to systems of energy supply and demand (McClelland, 2004). CPT-1 is thus generally considered the rate-limiting step in mitochondrial β-oxidation of fatty acids (Kerner and Hoppel, 2000). On a technical note, many measurements of CPT activity re?ect total CPT activity of both isoforms in homogenates of freeze-thawed tissues. Pande et al. (1990) suggested that CPT activity measured in these preparations represents CPT-2 activity due to freezing-induced inactivation of the more labile CPT-1. However, no appreciable effect of freezethaw of tissues on CPT activity has been reported (e.g. Suarez et al., 1986a; Sidell et al., 1987) indicating that CPT activity measurements are appropriate even on frozen tissues. Moreover, the functional linkage between the two enzymes makes the measurement of ‘apparent CPT activity’ useful as an indicator of fatty acid oxidation capacity. CPT activity has been utilized in this manner in many comparative studies and CPT activity as a marker of long-chain fatty acid oxidation has been validated by other means such as measurements of mitochondrial fatty acid oxidation. The same cannot be said for 3-hydroxyacyl CoA dehydrogenase (HOAD), a β-oxidation enzyme commonly measured in comparative studies; we suggest that HOAD activity should not be considered equally reliable to CPT activity as a marker of fatty acid oxidation capacity (e.g. Ballantyne et al., 1981; Suarez et al., 1986a; Sidell et al., 1987; Crockett and Sidell, 1990; Maillet and Weber, 2007). The observation of Zammit and Newsholme (1979) of nondetectable CPT in the red muscle and heart of four elasmobranch species was con?rmed in some of the same as well as additional species by Sidell et al. (1987), Singer and Ballantyne (1989), and Moyes et al. (1990a) (Table 1), suggesting that long-chain fatty acid oxidation was absent in these tissues in elasmobranchs. In teleosts, however, CPT activity was found at relatively high levels in red muscle and heart, as well as in liver, gill, and kidney (Table 1, references therein). These data led researchers to question whether mitochondria from red muscle and heart of elasmobranchs could oxidize fatty acids in the apparent absence of CPT activity. The measurement of readily detectable HOAD activity in red muscle and heart from elasmobranchs suggested active mitochondrial β-oxidation of fatty acids (Moon and Mommsen, 1987; Singer and Ballantyne, 1989; Moyes et al., 1990a; Speers-Roesch et al., 2006a). Studies on isolated mitochondria showed conclusively, however, that palmitoylcarnitine is not oxidized under isosmotic conditions by mitochondria from red muscle of mako shark (Isurus oxyrinchus) (Ballantyne et al., 1992) or red muscle and heart of spiny dog?sh shark (Squalus acanthias) and little skate (Leucoraja [=Raja] erinacea) (Moyes et al., 1990a; Chamberlin and Ballantyne, 1992). In contrast, mitochondria from heart and red muscle of teleosts and mammals oxidize long-chain fatty acylcarnitines at relatively high rates (Moyes et al., 1989; Moyes et al., 1990b; Chamberlin et al., 1991; Kiessling and Kiessling, 1993; Ballantyne, 1997). Low levels of palmitoylcarnitine oxidation occur in S. acanthias red muscle and heart mitochondria under hypoosmotic incubation (Moyes et al., 1990a), although the physiological signi?cance of this is unclear. Low rates of oxidation of octanoylcarnitine, a medium-chain fatty acylcarnitine, have been detected in mitochondria from S. acanthias red muscle and heart (Moyes et al., 1990a). This led Moyes et al. (1990a) to speculate that medium-chain fatty acids (6-12 carbons in length), which can be produced by peroxisomal oxidation of very long-chain fatty acids (>22 carbons in length) and oxidized mitochondrially via carnitine acyltransferases speci?c for shorter chain fatty acylcarnitines (the activity of which would not be detected in assays of CPT activity), could be used to a certain degree as an energy source in elasmobranchs. Consistent with this idea, Speers-Roesch et al. (2006a) found that activity of carnitine octanoyltransferase is present at low levels in heart

Table 1 Carnitine palmitoyltransferase activities (μmol·min? 1·g ww? 1) in liver, kidney, elasmobranch rectal gland, heart, and red muscle of ?shes, including elasmobranchs, holocephalans, teleosts, basal actinopterygians, sarcopterygians, petromyzontidians, and myxines. Liver Elasmobranchii Potamotrygon hystrixa Potamotrygon magdalenaeb Potamotrygon motoroc Himantura signiferc Taeniura lymmac Chiloscyllium punctatumc Leucoraja erinacea Squalus acanthias Raja clavatag Scyliorhinus caniculag Mustelus asteriasg Holocephali Hydrolagus collieih Teleostei Lophius piscatoriuse Scomber scombrus Gaidropsarus vulgarise Morone saxatilise Hemitripterus americanuse Zoarces americanusi Makaira nigricansj Gadus morhuak Dicentrarchus labraxg Pleuronectes platessag Cyprinus carpiof Oncorhynchus mykiss Lepomis cyanellus Trematomus newnesin Notothenia gibberifronsn Myoxocephalus octodecimspinosusn Tautoga onitisn Salvelinus alpinuso Basal Actinopterygii Lepisosteus platyrhincusp Amia calvaq Acipenser fulvescensr Sarcopterygii Protopterus dollois Petromyzontida Petromyzon marinust Myxini Myxine glutinosa Eptatretus stoutiif Kidney Rectal Heart gland – ND 0.13 0.17 1.31 0.53 0.24d – – – – 0.076 – – – – – – – – – – – – – – – – – 0.17 – 0.011 ND 0.1 –
u

Red muscle – –

– 0.002 0.27 0.77 0.62 1.09 0.09d 0.09f – – – 0.066 – – – – – – – – – – 0.39 0.30f, 0.25l 0.1m – – – – 0.20 0.13 0.029 ND 0.1 0.09 0.18 0.077

– – – – 0.30 0.49 – – – – – – – – – – – – – – – – – – – – – – – – – – – – – – –

0.17 ND 0.033 0.072 0.085 0.010 0.10d, NDe <0.01f,g, NDe <0.01 <0.01 <0.01 ND 0.23 0.28e 0.41 0.52 0.16 0.21 0.39 0.44 0.60 0.20 0.17 0.28f – 0.29 0.18 0.14 0.15 0.29 0.57 ND 0.054 ND – 0.06 0.044
k

ND 0.02d <0.01f,g <0.01 <0.01 <0.01 – – 0.60g – – – – 0.29 – 0.75 0.07 0.32 0.51f – 0.73 0.34 0.16 0.34 0.21 ND 0.009 0.039 ND 0.06 ? – 0.022 ?

– –

CPT activity is an indicator of capacity for fatty acid oxidation. Superscript letters indicate references for data, which are provided below with the temperature at which each measurement was made. ND = not detectable. – = not measured. ? = lateral muscle (E. stoutii) or somatic muscle (P. marinus). a Driedzic and De Almeida-Val (1996), 25 °C. b Singer and Ballantyne (1989), 25 °C. c Speers-Roesch et al. (2006a), 25 °C. d J. Berges and J.S. Ballantyne, unpublished data (cited in Ballantyne, 1997), 10 °C. e Sidell et al. (1987), 15 °C. f Moyes et al. (1990a), 15 °C. g Zammit and Newsholme (1979), 10 °C. h Speers-Roesch et al. (2006b), 12 °C. i Driedzic and Stewart (1982), 10 °C. j Suarez et al. (1986b), 25 °C. k Hansen and Sidell (1983), 15 °C. l Gutières et al. (2003), 30 °C. m Londraville and Duvall (2002), 20 °C. n Crockett and Sidell (1990), 1 °C. o J.S. Bystriansky and J.S. Ballantyne, unpublished results, (mg protein? 1 data for liver and red muscle published in Bystriansky et al., 2007) 10 °C. p Frick et al. (2007), 25 °C. q Singer and Ballantyne (1991), 10 °C. r Singer et al. (1990), 10 °C. s Frick et al. (2008b), 25 °C. t LeBlanc et al. (1995), 10 °C. u Leary et al. (1997), 10 °C.

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of several elasmobranchs as well as red muscle of the shark Chiloscyllium punctatum. The ability to oxidize medium-chain fatty acids at low rates may partly explain the presence of detectable HOAD activity in red muscle and heart in elasmobranchs (Moyes et al., 1990a). Carnitineindependent oxidation of medium- or long-chain fatty acids, which could allow for fatty acid oxidation in the absence of CPT or other acyltransferases, does not occur in S. acanthias red muscle or heart mitochondria, whereas medium-chain fatty acids are utilized in this manner in mammals (Moyes et al., 1990a). Physiological studies support the notion that lipid oxidation in muscle of elasmobranchs is of minimal importance. Driedzic and Hart (1984) showed that isolated perfused hearts from the skate Leucoraja erinacea performed relatively poorly when provided with palmitate as an aerobic fuel, contrasting with the superior performance under these conditions by heart from a teleost, the sea raven (Hemitripterus americanus). Also, white muscle in S. acanthias does not appear to rely on lipids during exercise or recovery from exercise (Richards et al., 2003) whereas lipids are important in both these states in white muscle of rainbow trout (Richards et al., 2002). In contrast to the absence of CPT and fatty acid oxidation in cardiac and skeletal muscle of elasmobranchs, CPT activity was found in liver from S. acanthias at levels similar to those found in teleosts (Moyes et al., 1990a) (Table 1). Correspondingly, mitochondria from liver of the skate L. erinacea were found to readily oxidize palmitoylcarnitine and other fatty acylcarnitines (Ballantyne and Moon, 1986), probably in part to contribute acetyl CoA for ketogenesis (see Section 3.1) (Treberg et al., 2006a). Meanwhile, CPT activity was found to be absent in kidney and brain from the Amazonian stingray Potamotrygon magdalenae (Singer and Ballantyne 1989) (Table 1). Together, these studies led researchers to work with the understanding that fatty acid oxidation was virtually absent in extrahepatic tissues of elasmobranchs (Ballantyne, 1997; Watson and Dickson, 2001; Richards et al., 2003; Wood et al., 2005). The measurement of Driedzic and De Almeida-Val (1996) of a teleost-like level of CPT activity in the heart of the Amazonian stingray Potamotrygon hystrix (Table 1) suggested, however, that the story was not clear-cut. In fact, in a recent examination of the metabolic organization of elasmobranchs, including three stingrays and one shark, Speers-Roesch et al. (2006a) showed in all the presence of CPT activity in rectal gland and kidney at high levels similar to those found in the liver (Table 1). CPT activity was absent in red and white skeletal muscle, in agreement with previous studies (Table 1). The hearts of the stingrays but not the shark C. punctatum possessed relatively low but readily detectable levels of CPT (Speers-Roesch et al., 2006a), in agreement with the measurements of Driedzic and De Almeida-Val (1996) (Table 1). Although the physiological signi?cance of these heart CPT activities is unclear, it suggests that there may be a low capacity for cardiac fatty acid oxidation in some elasmobranch species. Based on these ?ndings, Speers-Roesch et al. (2006a) suggested that the original concept of absent extrahepatic fatty acid oxidation in elasmobranchs was an oversimplication. Instead, these authors suggested that fatty acid oxidation occurs in most tissues in elasmobranchs and is absent or limited only in skeletal and cardiac muscle due to the tissue-speci?c distribution of CPT. Studies on the ability of kidney and rectal gland mitochondria to oxidize fatty acids are needed to con?rm the apparent presence, based on CPT activities, of fatty acid oxidation in those tissues. Additionally, the capacity for fatty acid oxidation in other extrahepatic tissues such as intestine, spleen, and brain should be ascertained. Further exploration of the physiological importance of fatty acid oxidation for tissue and organ function in elasmobranchs is also warranted. Investigations into the determinants of the tissue-speci?c distribution of CPT in elasmobranchs, including transcriptional or translational regulation, are also needed. Transcriptional regulation may not be a primary explanation because mRNA transcripts of at least two CPT-1 isoforms are present at appreciable levels in red muscle and heart of

S. acanthias, despite the fact that CPT enzyme activity is not present (B. Speers-Roesch and J.G. Richards, unpublished data). 2.2. Fatty acid transport Plasma non-esteri?ed fatty acids (NEFA) are the most metabolically dynamic fraction of lipid in vertebrate blood and a reliable estimate of the importance of lipids in energy metabolism because NEFA transport in the blood helps sustain fatty acid oxidation in tissues (Henderson and Tocher, 1987). Plasma [NEFA] in elasmobranchs is 2- to 10-fold lower than that found in teleosts (Table 2, references therein; also see Larsson and F?nge, 1977; Zammit and Newsholme, 1979; Fellows et al., 1980), which is consistent with the limited fatty acid oxidation in muscle. In mammals and many other vertebrates, including some teleosts, plasma NEFA is carried to peripheral tissues by albumin, which is important for transport of the highly insoluble long-chain fatty acids (Peters,

Table 2 Total non-esteri?ed fatty acid (NEFA) concentrations in plasma of ?shes, including elasmobranchs, holocephalans, teleosts, basal actinopterygians, petromyzontidians, and myxines. [NEFA] (nmol mL? 1) Elasmobranchii Potamotrygon motoro Himantura signifer (freshwater) Himantura signifer (15 ppt seawater) Taeniura lymma Chiloscyllium punctatum Leucoraja erinacea Squalus acanthias Raja rhina Bathyraja sp. Isurus oxyrinchus Prionacea glauca Dasyatis americana Holocephali Hydrolagus colliei Teleostei Anguilla rostrata Myoxocephalus octodecimspinosus Myoxocephalus quadricornis Urophycis chuss Xiphias gladius Oncorhynchus nerka Salvelinus alpinus Basal Actinopterygii Lepisosteus platyrhincus Amia calva Acipenser fulvescens Acipenser brevirostrum Petromyzontida Petromyzon marinus Myxini Myxine glutinosa 106 123 154 216 172 573 416 399 167 252 203 193 466 652 874–1443 1183 2982 2374 1284 2316 (male) 3092 (female) 1215 607 758 2493 1060 Reference

Speers-Roesch et al. (2006a) " " " " Speers-Roesch et al. (2008) " Ballantyne et al. (1993) Speers-Roesch et al. (2008) " Ballantyne et al. (1993) " Semeniuk et al. (2007) Speers-Roesch et al. (2006b) Cottrill et al. (2001) Ballantyne et al. (1993) " " " Ballantyne et al. (1996) Bystriansky et al. (2007) Frick et al. (2007) Singer and Ballantyne (1991) Singer et al. (1990) Jarvis and Ballantyne (2003)

1136

LeBlanc et al. (1995)

551

P.J. LeBlanc, C. Hyndman, J.S. Ballantyne, unpublished results, cited in Speers-Roesch et al. (2006b)

Values are means and were obtained using the methylation procedure of Singer et al. (1990).

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1996). The presence of albumin in elasmobranchs was ?rst questioned by Irisawa and Irisawa (1954) and has been debated ever since (see Metcalf and Gemmell, 2005). Using re?ned techniques for albumin identi?cation, Metcalf and Gemmell (2005) recently resolved the dispute via a comprehensive analysis of elasmobranch plasma proteins that showed conclusively that elasmobranchs lack albumin. Earlier reports of albumin in elasmobranchs were likely the result of the use of imprecise techniques for albumin detection (Metcalf and Gemmell, 2005). In elasmobranchs, plasma NEFA appear to be carried by lipoproteins. Metcalf and Gemmell (2005) showed that palmitate binds to elasmobranch plasma proteins that are probably low density lipoprotein (LDL) and very low density lipoprotein (VLDL). This result supports earlier work showing the importance of LDL and VLDL, but not high density lipoprotein (HDL), as NEFA carriers in elasmobranchs (Lauter et al., 1967; Mills et al., 1977; Babin and Vernier, 1989). Chylomicrons, which are lipoproteins that transport dietary lipids from the gut, also are enriched in NEFA (Lauter et al., 1967). Lipoproteins in elasmobranchs additionally carry other lipid fractions including triacylglycerols, hydrocarbons, and cholesterol (Lauter et al., 1967; Mills et al., 1977). Further details of the role of lipoproteins in lipid transport and turnover in elasmobranchs remain unknown. The absence of albumin in elasmobranchs has been suggested to be the primary cause of the comparatively low plasma [NEFA] seen in these ?shes (Fellows et al., 1980; Fellows and Hird, 1981; Ballantyne, 1997). However, many teleosts, including common carp (Cyprinus carpio), Antarctic tooth?sh (Dissostichus mawsoni), and eels (Anguilla spp.), lack albumin and transport fatty acids via HDL (De Smet et al., 1998; Metcalf et al., 1999a; Metcalf et al., 1999b). Furthermore, despite the lack of albumin, common carp and eels have plasma [NEFA] typical of other teleosts (van Raaij et al., 1996; Cottrill et al., 2001) as well as the capacity to oxidize fatty acids at relatively high rates in peripheral tissues including heart and red muscle (Moyes et al., 1989; West et al., 1999). The lower levels of plasma NEFA in elasmobranchs compared with teleosts may therefore simply be the consequence of the limited fatty acid oxidation in muscle of elasmobranchs, which may lessen the requirement for NEFA transport (Speers-Roesch et al., 2008). In mammals, at least, plasma [NEFA] and rates of muscle and whole body fatty acid oxidation are closely related (Ebeling and Koivisto, 1994; Piatti et al., 1996). Little is known about cellular uptake and transport of fatty acids and other lipids in elasmobranchs. Limited work shows that, like teleosts and other vertebrates, elasmobranch liver appears to possess fatty acid binding proteins (FABP) similar to the mammalian hearttype FABP, which transports lipids related to energy production, as well as to the mammalian liver-type FABP, which transports fatty acids involved in phospholipids synthesis (Bass et al., 1991; Medzihradszky et al., 1992; Córdoba et al., 1999). Consistent with these roles, the heart-type FABP found in liver of elasmobranchs have a higher af?nity for saturated and monounsaturated fatty acids and the liver-type FABP binds more readily to polyunsaturated fatty acids (Bass et al., 1991; Córdoba et al., 1999). Another FABP identi?ed from liver of the guitar?sh Platyrhinoides triseriata could not bind lipids (Sugiyama et al., 1982). It is unknown to what degree cellular fatty acid uptake and transport mechanisms, including FABP, have been modi?ed in muscle of elasmobranchs, where fatty acid oxidation is absent or low. 2.3. Hepatic lipid storage Lipid storage in elasmobranchs was reviewed comprehensively by Ballantyne (1997) but given its importance in energy metabolism we provide a new overview here. Unlike mammals and many teleosts, adipose tissue is absent in elasmobranchs and the liver is the main lipid storage site as well as a major site of lipid synthesis (Malins 1968; Bone and Roberts, 1969; Baldridge, 1972; Sargent et al., 1972; Ballantyne, 1997). Elasmobranchs are renowned for their large livers and high hepatosomatic indices (HSI) in comparison with teleosts and other

vertebrates. There is great interspeci?c variation in HSI values, however, ranging from <4% in some benthic species, similar to teleosts, to >25% in some deep-sea squaliformes and large pelagic sharks (Bone and Roberts, 1969; Corner et al., 1969; Baldridge, 1970; Rossouw, 1987; Treberg and Driedzic, 2007). Triacylglycerols (TAG), alkyldiacylglycerols (ADAG) and hydrocarbons (e.g. squalene, pristane) are the primary lipid classes stored in the liver. High concentrations of wax esters are present in some species (Ballantyne, 1997; Navarro-Garcia et al., 2000; Wetherbee and Nichols, 2000). The acylchains of ADAG, wax esters, and especially TAG are mobilized for utilization in peripheral tissues, being exported from the liver as NEFA or esteri?ed as TAG or, following hepatic β-oxidation and ketogenesis, as ketone bodies (see Section 3.1) (Sargent et al., 1972; Anderson, 1990; Treberg et al., 2006a). Squalene, as an intermediate in sterol biosynthesis, has limited metabolic fates. Ballantyne (1997) postulated that squalene might be a metabolic ‘dead-end’ in elasmobranchs, based in part on the observation that there is a proportional increase in hepatic squalene content with age in Centrophorus spp. sharks (Peyronel et al., 1984). Thus, those species that accumulate vast amounts of squalene, such as many deep-sea sharks, do not appear to be able to subsequently utilize that lipid carbon for energy metabolism (Ballantyne, 1997). The metabolically inert status of squalene and other hydrocarbons in the elasmobranch liver may re?ect their importance in buoyancy in many elasmobranchs. The major liver lipids accumulated by elasmobranchs are less dense than water: hydrocarbons such as squalene are least dense (0.86 g ml? 1), followed by ADAG (0.89 g ml? 1), and then TAG (0.9 g ml? 1) (sea water has a density of 1.03 g ml? 1). Thus, the hepatic accumulation of low density lipids, especially squalene and ADAG, has an important role in buoyancy in elasmobranchs by providing hydrostatic lift in a group of animals that lack a swimbladder (Bone and Roberts, 1969; Baldridge, 1972). Turnover of ADAG in S. acanthias is much lower than that of TAG, potentially re?ecting a conservation of ADAG for buoyancy (Sargent et al., 1972). In deep-sea sharks, the energetic savings of a lessened reliance on hydrodynamic lift may explain why they sequester large amounts of lipid as metabolically inert squalene in an oligotrophic environment (Bone and Roberts, 1969; Corner et al., 1969; Phleger, 1998). In a famous but never repeated experiment, Malins and Barone (1970) demonstrated a shift in the proportion of ADAG to total liver lipids and a subsequent decline in the relative TAG content when the density of S. acanthias was arti?cially increased with weights. There was no change in the percentage of lipid content of the liver or the HSI, rather only what appears to have been a shift from the relatively dense TAG to the less dense ADAG. Further research is needed to evaluate the intriguing possibility that control of whole animal density in elasmobranchs can be achieved via metabolic restructuring of hepatic lipids. The dual role of the elasmobranch liver as an energy store and as a buoyancy aid may in?uence the mobilization of storage lipids. A substantial decrease in the HSI of the winter skate (Leucoraja ocellata) occurs after 7 and 28 days of fasting (de?ned here as food deprivation for a duration of more than 2 days) (Treberg and Driedzic, 2007). This is consistent with the mobilization of liver lipid reserves in a benthic species where buoyancy presumably is not a major concern. On the other hand, in the pelagic S. acanthias HSI is relatively unaffected by several weeks of fasting (Kajimura et al., 2008). Lipid mobilization is still likely, however, because the liver does reduce in size roughly parallel with whole body mass decline (Kajimura et al., 2008). Seibel and Walsh (2002) suggested that lipid accumulation in the elasmobranch liver might be a result of the need to sustain high rates of TMAO synthesis, which may limit availability of choline, a TMAO precursor, for hepatic phosphatidylcholine production. In mammals, choline de?ciency leads to reduced phosphatidylcholine synthesis and accumulation of diacylglycerol and TAG, resulting in a form of non-alcoholic fatty liver disease (hepatic steatosis). Seibel and Walsh

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(2002) speculated that a similar situation presents in elasmobranchs, but their hypothesis relies on the assumption that relatively high rates of TMAO synthesis occur. In fact, many elasmobranchs, including S. acanthias, apparently lack the capacity for endogenous TMAO synthesis (Baker et al., 1963; Goldstein et al., 1967; Treberg et al., 2006b). Instead, maintenance of TMAO homeostasis is achieved via dietary input and low rates of loss (≤1% day? 1) to the environment (Treberg and Driedzic, 2006). Earlier measurements of higher TMAO losses (4.4–13.8% day? 1) (Goldstein and Palatt, 1974) were likely overestimates due to insuf?cient tracer equilibration (Treberg and Driedzic, 2006). Although the hypothesis of Seibel and Walsh (2002) is not supported by these data, investigations into how the pathogenesis of mammalian non-alcoholic fatty liver disease (Marra et al., 2008) is avoided in the lipid-rich elasmobranch liver may prove useful. 2.4. Physiological regulation The physiological regulation of lipid metabolism in elasmobranchs is not well understood. Most studies have focused on hormonal control. Administration in S. acanthias of a low dose of the thyroid hormone 3,5,3′-triiodothyronine, which has widespread stimulatory metabolic effects in other vertebrates, increased HOAD activity in liver but not kidney (Battersby et al., 1996). Corticosterone injection of S. acanthias caused a decrease in hepatic lipid content (Patent, 1970), although it is unclear whether this was due to stimulation of NEFA or ketone body mobilization, or both. Similarly, Lipshaw et al. (1972) demonstrated a decrease in liver lipid content of the nurse shark (Ginglymostoma cirratum) injected with epinephrine and norepinephrine. These results suggest that like other vertebrates, corticosteroids and catecholamines are important in mobilization of energy substrates in elasmobranchs. Conlon et al. (1994) investigated the effects of in vivo synthetic catshark Scyliorhinus canicula urotensin-II infusion on plasma [NEFA] in S. canicula, but they were unable to detect the low levels of plasma [NEFA] in this species using a commercially available enzymatic NEFA kit. Further studies using methods with a higher sensitivity of detection, such as gas chromatography, are needed to ascertain whether urotensin-II stimulates NEFA mobilization, as in teleosts (Sheridan, 1994). Conlon et al. (1994) did show that urotensin-II had no effect on plasma [TAG]. Further work is needed to better understand the hormonal regulation of lipid mobilization, cellular uptake, and oxidation in elasmobranchs, and how this may be in?uenced by the unusual tissue-speci?c organization of fatty acid oxidation and preference for ketone body oxidation. As in mammals and teleosts (Morash et al., 2008), malonyl CoA plays a role in regulation of mitochondrial fatty acid oxidation in the elasmobranch liver via inhibition of CPT-1 (Treberg et al., 2006a). Comparable to fasted mammals, during ketosis induced by several weeks of fasting the IC50 of malonyl CoA increased markedly in liver of S. acanthias, likely to facilitate greater ?ux of fatty acids into ?-oxidation and ketogenesis (Treberg et al., 2006a). Presumably, control of CPT-1 activity via malonyl CoA also occurs in other tissues, with the possible exception of muscle where fatty acid oxidation is limited. Nothing is known about enzymatic or hormonal regulation of malonyl CoA levels via acetyl-CoA carboxylase in elasmobranchs. Much remains unknown regarding the physiological regulation of lipid metabolism after feeding and during food deprivation. Fatty acid composition of plasma [NEFA] in the southern stingray (Dasyatis americana) re?ects the fatty acid composition of the diet but the physiological effects of this are unclear (Semeniuk et al., 2007). The effects of fasting on lipid storage were previously discussed (Section 2.3). Several weeks of fasting in the catshark S. canicula caused no increase in plasma [NEFA] (Zammit and Newsholme, 1979), whereas in S. acanthias plasma [NEFA] decreased during fasting over a similar time period, which was attributed to increased

utilization (Wood et al., 2010-this issue). Studies on fatty acid turnover during food deprivation and following feeding in elasmobranchs are warranted. 3. Ketone bodies 3.1. Ketone bodies as routine metabolic fuels The initial discovery of a limited capacity for fatty acid oxidation in elasmobranch muscle (Zammit and Newsholme, 1979) was accompanied by evidence suggesting that ketone bodies are routinely utilized as an alternate aerobic fuel source, a condition likely unique among carnivorous vertebrates. Ketone bodies, including acetoacetate and D-βhydroxybutyrate (β-HB), are high-energy fuels produced primarily from acetyl CoA by liver mitochondria and then exported to peripheral tissues for oxidation (Ballantyne, 1997; Laffel, 1999). Zammit and Newsholme (1979) and subsequent studies showed that elasmobranch tissues possess relatively high levels of enzymes related to ketone body metabolism, especially D-β-hydroxybutyrate dehydrogenase (βHBDH) (Moon and Mommsen, 1987; Driedzic and De Almeida-Val, 1996; Watson and Dickson, 2001; Treberg et al., 2003; Speers-Roesch et al., 2006a). β-HBDH is a mitochondrial enzyme involved in the interconversion of β-HB and acetoacetate during the oxidation of β-HB in extrahepatic tissues and the production of β-HB in the liver (Nelson and Cox, 2004). Activities of β-HBDH generally are much lower in teleosts (Zammit and Newsholme, 1979; LeBlanc and Ballantyne, 1993; Soengas et al., 1996; 1998). In at least one elasmobranch (the stingray Taeniura lymma), however, the kidney has low β-HBDH activity suggesting there may be variation in ketone body utilization among elasmobranchs (Speers-Roesch et al., 2006a). Mitochondria from teleost and mammal red muscles oxidize ketone bodies relatively poorly (Moyes et al., 1989; Chamberlin et al., 1991; Ballantyne, 1997). Ketone body utilization by the teleost brain appears to be insigni?cant and isolated perfused teleost heart performs poorly in the presence of ketone bodies (Driedzic and Hart, 1984; Soengas et al., 1998). In elasmobranchs, however, ketone bodies are important fuels. β-HB and acetoacetate are oxidized by elasmobranch red muscle and heart mitochondria at a similar or higher rate compared with pyruvate (Moyes et al., 1990a; Ballantyne et al., 1992; Chamberlin and Ballantyne, 1992) and isolated perfused hearts perform best in the presence of ketone bodies (Driedzic and Hart, 1984). White muscle in S. acanthias appears to rely on ketone bodies during recovery from exercise (Richards et al., 2003). β-HB enhances the rate of glucose-dependent salt secretion in the rectal gland of S. acanthias (Walsh et al., 2006) and it is a good oxidative fuel for the kidney of the skate L. erinacea (Mommsen and Moon, 1987). Based on arteriovenous differences of [β-HB] in the blood supply to the brain in S. acanthias, deRoos (1994) suggested that β-HB is taken up and consumed by the brain at a greater rate compared with glucose, lactate, or alanine. Conversely, ketone bodies are poor oxidative fuels for isolated elasmobranch hepatocytes (Mommsen and Moon, 1987). Hepatic activity of the ketolytic enzyme succinyl CoA-ketotransferase (SKT, =3-oxoacid CoA transferase) is relatively low, preventing oxidation of ketone bodies (Zammit et al., 1979; Moon and Mommsen, 1987). These data likely re?ect the ketogenic role of the elasmobranch liver. Relatively high rates of ketogenesis in elasmobranch hepatocytes and liver mitochondria have been demonstrated from lipid precursors as well as from amino acids and pyruvate (Anderson, 1990; Mommsen and Moon, 1987). Thus, in elasmobranchs the lipid-rich liver is a ketogenic powerhouse that sustains the heavy reliance on ketone bodies as oxidative fuel in extrahepatic tissues. Ketone bodies are also important metabolic fuels in mammals, but they are primarily utilized as an alternate fuel in brain and other tissues to spare glucose and muscle protein only during starvation or carbohydrate limitation (Laffel, 1999; Veech, 2004). Elasmobranchs, on the other hand,

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appear to preferentially utilize ketone bodies as oxidative fuels under routine conditions, leading to a chronic mild ketosis. The concentration of plasma β-HB in recently fed elasmobranchs is similar to that seen in plasma of fasting or diabetic hyperketonemic (1 to 3 mM) or even ketoacidotic (>3 mM) mammals, in which the normal fed range is 0.05– 0.5 mM (Table 3, references therein; also see Laffel, 1999; Richards et al., 2003). Plasma [β-HB] of elasmobranchs is higher than found in most teleosts and some teleosts apparently lack β-HB altogether (Table 3, references therein). Acetoacetate levels in elasmobranchs are also higher than in fed teleosts and mammals, although the difference usually is less striking than it is for β-HB (Table 3, references therein). It is unclear, but worthwhile exploring, how elasmobranchs avoid the deleterious effects of ketosis seen in diabetic humans (Jain and McVie, 1999). 3.2. Physiological regulation Utilization of ketone bodies as aerobic fuels increases during fasting in elasmobranchs, similar to mammals but apparently not teleosts. In fasting elasmobranchs there are increases in plasma [β-HB] and, in some species, plasma [acetoacetate] (Table 3) (Zammit and Newsholme, 1979; deRoos et al., 1985; deRoos, 1994; Soengas et al., 1996; Segner et al., 1997; Treberg et al., 2006a; Kajimura et al., 2008; Wood et al., 2010-this issue). Increased β-HB uptake by the brain may occur during fasting in S. acanthias (deRoos, 1994). Increased ketogenesis in elasmobranchs fasted for several weeks is controlled via increases in fatty acid oxidation

due to relaxed malonyl CoA inhibition of CPT-1 as well as by an increase in activity of the ketogenic enzyme acetoacetyl-CoA thiolase (Zammit and Newsholme, 1979; Treberg et al., 2006a). Conversely, feeding of fasted S. acanthias causes plasma [β-HB] to decrease and activities of βHBDH to increase in rectal gland, brain, and liver, suggesting increased oxidation of β-HB in peripheral tissues but also increased hepatic ketogenesis (Walsh et al., 2006). Plasma [β-HB] varies throughout the year in captive S. canicula, potentially related to changes in reproductive status and feeding activity (Gutiérrez et al., 1988). The hormonal control of ketogenesis and ketone body oxidation in elasmobranchs is not well understood. Whereas insulin inhibits ketogenesis in mammals, infusion with bovine insulin did not affect ketone body levels in the catshark S. canicula (deRoos et al., 1985) nor did infusion with S. canicula insulin (Anderson et al., 2002). Administration of urotensin-II does not stimulate β-HB release from S. canicula hepatocytes nor does plasma [β-HB] change in S. canicula infused with this hormone, but epinephrine causes release of β-HB from hepatocytes in line with its role in mobilizing energy substrates (Conlon et al., 1994). Mammalian corticotropin (ACTH), which has stimulatory effects on substrate mobilization in mammals, had no effect on plasma [β-HB] in infused S. acanthias (deRoos and deRoos, 1992). More studies, preferably incorporating turnover measurements, are needed to better understand the regulation of ketone body production and utilization in elasmobranchs. The relationship between regulation of lipid and ketone body metabolism in

Table 3 ?-hydroxybutyrate and acetoacetate concentrations in plasma of elasmobranchs, teleosts, and mammals under various conditions. ?-hydroxybutyrate (mM) Elasmobranchii Squalus acanthias 4.91 1.18 0.72 1.59 10.7 16.9 0.2 2.2–3.8 3.12 0.71 1.11 3.21 0.06 1.61 ~ 3.0 ~ 0.5–~ 4 Acetoacetate (mM) 0.15–0.30 0.15–0.30 0.15–0.30 0.15–0.30 0.15–0.30 0.15–0.30 0.13 – – – – Condition Reference Wood et al. (2010-this issue); Kajimura et al. (2008) " " " " " Zammit and Newsholme (1979) Wood et al. (2005) deRoos et al. (1985) Treberg et al. (2006a) " " Zammit and Newsholme (1979) " Conlon et al. (1994) Gutiérrez et al. (1988) Zammit and Newsholme (1979) " Carrillo et al. (1982) Zammit and Newsholme (1979) " Segner et al. (1997) Segner et al. (1997) Morata et al. (1982) Soengas et al. (1996) Jain and McVie (1999) " Pan et al. (2000) " " Féry and Balasse (1983) " Owen et al. (1969) " " " " Keller et al. (1978) "

Scyliorhinus canicula

Fasted 7 days 6 h after feeding 2.5% ration 6 h after feeding 5.5% ration 60 h after feeding 5.5% ration 15 days after feeding 5.5% ration 35 days after feeding 5.5% ration 2–3 h after capture 2 h to 45 h after feeding Fasted 2–5 days 0–2 days after capture (no feeding) 4–9 days after capture (no feeding) 16–33 days after capture (no feeding) 0.07 2–3 h after capture 0.29 Fasted 40 days ~ 1.0 Fasted 40 days 0.03–0.55 (individual values) Fasted 1day, sampled throughout 1 year 2–3 h after capture Fasted 40 days Fed 2–3 h after capture 2–3 h after capture Fed Fasted 2 months Fed Fed Fed Hyperketonemic diabetic Fed Fasted 2 days Fasted 3 days Fasted 16 h Fasted 3–5 days Fed Fasted 3 days Fasted 10 days Fasted 17 days Fasted 31 days Fasted 48 h Diabetic

<0.001 <0.001 ND Mullus surmuletus <0.001 Scomber scombrus <0.001 Cyprinus carpio ND ND Oncorhynchus mykiss ND Salmo salar ~ 0.4 Mammalia Homo sapiens 0.12 1.4 0.03 1.67 3.15 0.22 4.52 0.07 1.21 4.3 5.10 5.84 Canis familiaris 0.099 1.09

Teleostei Dicentrarchus labrax

0.04 0.04 ~ 0.04 0.01 0.05 0.03 0.002 ~ 0.03 ~ 0.57 0.13 0.7 – – – 0.09 1.15 0.03 0.41 1.0 1.12 1.37 0.072 0.44

Values are means except where noted. ND = not detectable. – = not measured. ~ = estimated value based on graphical data.

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elasmobranchs also is of interest considering the similar origins of these fuels in the liver, the lipid storage organ. 4. Amino acids 4.1. Amino acid oxidation Besides their unusual organization of ketone body and lipid metabolism, another distinguishing characteristic of elasmobranchs is an apparently heavy reliance on amino acids, especially glutamine, as aerobic fuels. Ballantyne and colleagues were the ?rst to note this after detecting a higher rate of oxidation of glutamine compared with pyruvate in mitochondria isolated from elasmobranch red muscle (Ballantyne et al., 1992; Chamberlin and Ballantyne, 1992). Red muscle mitochondria from a teleost oxidize glutamine more slowly than pyruvate (Chamberlin et al., 1991), although amino acid metabolism is still clearly important in teleosts (Ballantyne, 2001). Very low levels of glutamine oxidation occur in mammalian red muscle and in general oxidation of amino acids in mammals and other tetrapods is primarily localized to the liver (Moyes et al., 1990b). Correspondingly, the activity in red muscle and other tissues of phosphate-dependent glutaminase (PDG), an enzyme involved in the catabolism of glutamine, is greatest in elasmobranchs compared with holosteans, teleosts, birds, and mammals (Chamberlin et al., 1991; Chamberlin and Ballantyne, 1992; Ballantyne, 1997). The oxidation of glutamine occurs by deamidation of glutamine to glutamate by PDG followed by deamination by glutamate dehydrogenase (GDH) of glutamate to α-ketoglutarate, which enters the Krebs cycle. Unlike the oxidation of glutamate, glutamine oxidation by elasmobranch muscle mitochondria is relatively insensitive to aminooxyacetate, an inhibitor of transamination (Chamberlin and Ballantyne, 1992). This suggests that the glutamate and α-ketoglutarate formed from glutamine

represents a different pool than that involved with cytosolic glutamate, which is preferentially involved with the malate-aspartate shuttle. This partitioning also is seen in the mammalian kidney, which under some conditions oxidizes glutamine as a preferred metabolic fuel (Schoolwerth et al., 1978). The elasmobranch liver is also an important site of amino acid oxidation, as it is in teleosts (Ballantyne, 2001). Liver mitochondria from the skate L. erinacea oxidize amino acids, with glutamate having the greatest rate and alanine, β-alanine, sarcosine, dimethylglycine, serine and glycine showing signi?cant rates of oxidation (Ballantyne et al., 1986; Moyes et al., 1986a,b). In isolated L. erinacea hepatocytes, rates of CO2 production from alanine, serine and leucine are higher than rates from other potential fuels including lactate, oleate, glycerol, and β-hydroxybutyrate (Mommsen and Moon, 1987). 4.2. Role of amino acids in gluconeogenesis and ketogenesis Amino acids in the liver of elasmobranchs are important substrates for gluconeogenesis, as in teleosts (Ballantyne, 2001). In hepatocytes from the skate L. erinacea, serine and alanine are comparable to or better than lactate as gluconeogenic substrates but both amino acids are poor substrates compared with glycerol (Mommsen and Moon, 1987; Ballatori and Boyer, 1988). Gluconeogenesis from alanine is likely via the phosphoenolpyruvate carboxykinase (PEPCK) pathway whereas carbon from serine is made available from transamination via serine-pyruvate transaminase (Mommsen and Moon, 1987). In elasmobranchs, amino acids, especially glutamine and aspartate, serve additional essential roles as nitrogen donors for urea synthesis in the liver and possibly in muscle (Ballantyne, 1997; Steele et al., 2005). Glutamine and aspartate both likely are regenerated using nitrogen from transamination of other amino acids as well as ammonia scavenging by GDH and glutamine synthetase (Fig. 1).

Fig. 1. Proposed functional coupling between alanine and ammonia release from skeletal muscle and the hepatic synthesis of urea and ketone bodies in elasmobranchs. Some substrates and cofactors have been omitted for clarity. Dotted lines represent two or more complex enzymatic pathways. Abbreviations: α-KG, alpha-ketoglutarate; β-HB, betaHydroxybutyrate; OUC, ornithine urea cycle; OXA, oxaloacetate. Enzyme/protein legend: 1, aminotransferases (general); 2, alanine aminotransferase; 3, aspartate aminotransferase; 4, glutamate aspartate transporter; 5, pyruvate dehydrogenase; 6, phosphate-dependent glutaminase; 7, glutamate dehydrogenase.

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Amino acids are important substrates for ketogenesis in elasmobranch liver. The rate of ketogenesis from leucine is greater than from oleate in hepatocytes of the skate L. erinacea (Mommsen and Moon, 1987). Remarkably, in S. acanthias liver mitochondria alanine and its transamination product pyruvate are preferentially converted to ketone bodies rather than to phosphoenolpyruvate for gluconeogenesis (Anderson, 1990). In contrast, an elevation in extracellular alanine has a hepatic antiketogenic effect in mammals, possibly via additional oxaloacetate provision that may prevent the intramitochondrial accumulation of acetyl-CoA probably necessary for ketogenesis (Nosadini et al., 1980). The conversion of pyruvate to acetyl CoA, which can be used in ketogenesis, requires catalysis by pyruvate dehydrogenase (PDH), which itself is inhibited by acetyl CoA. It is unclear how ?ux through PDH in elasmobranch liver is maintained under the conditions of relatively high [acetyl CoA] that persist in the ketogenic mitochondria. Elasmobranch liver PDH, or alternately the mitochondrial ketogenic pathway, should be characterized in an effort to explain this unusual carbon handling characteristic of elasmobranch liver mitochondria. Alanine and ammonia are released from muscle and this increases during food deprivation in S. acanthias (Leech et al., 1979), which raises the possibility of the existence of an alanine–ketone body cycle (Fig. 1) analogous to the alanine–glucose shuttle used in mammals for nitrogen shuttling from the muscle to the liver (Goldstein and Newsholme, 1976). In this scenario (Fig. 1), alanine and ammonium released from muscle may act as major nitrogen carriers to the liver to help sustain high rates of urea synthesis. [Ammonium] is kept low in elasmobranch tissues and in the liver glutamine synthetase likely plays a major role in scavenging free ammonia that may have come from the muscle. In the same sequence (Fig. 1), alanine taken up by the liver can be deaminated in the mitochondrion or the cytosol to pyruvate, which is preferentially converted into ketone bodies in mitochondria (Anderson, 1990). The ketone bodies are then released to the plasma for return to peripheral tissues for oxidation, including to the muscle to help replenish carbon lost to amino acid export. An alanine–glucose shuttle, as described for mammals, may operate alongside this carbon cycling through ketone bodies. Thus, we propose that in elasmobranchs there may be a link between urea synthesis and ketogenesis and ketone body oxidation, although further evidence is needed (see also Section 8.5). 4.3. Amino acid transport Amino acid transport does not appear to be markedly different between elasmobranchs and other vertebrates, with free amino acids being delivered from the gut or sites of proteolysis (e.g., muscle) to other tissue via the blood (Ballantyne, 2001). The level of total amino acids in elasmobranch plasma is similar to other vertebrates (Ballantyne, 2001). Ballantyne (1997) noted that the glutamine level in the plasma of elasmobranchs is lower than in other vertebrates, ostensibly because of substantial utilization as an oxidative fuel in muscle and as a nitrogen donor for hepatic urea synthesis. Subsequent measurements on other species of sharks showed higher plasma levels (Ballantyne 2001), however, suggesting that plasma [glutamine] may vary depending on species, recent feeding activity, and other factors. 4.4. Physiological regulation Few data exist on physiological regulation of amino acid oxidation in elasmobranchs. In S. acanthias, feeding causes a modest increase of alanine aminotransferase activity in rectal gland (Walsh et al., 2006) and fasting causes no major changes in the total amount of plasma free amino acids (Kajimura et al., 2008), although alanine export from muscle increases (Leech et al., 1979). Studies on hormonal control of amino acid metabolism have focused largely on the role of amino acids as gluconeogenic precursors. In S. acanthias, insulin causes decreases in plasma alanine and ACTH causes

increases in plasma [alanine] and this was attributed to increased and decreased gluconeogenesis, respectively (deRoos et al., 1985; deRoos and deRoos, 1992). In another study on S. acanthias, GDH increased in liver but not other tissues in animals injected with 3,5,3′-triiodothyronine (Battersby et al., 1996). Further studies are needed to better understand the regulation of amino acid oxidation in elasmobranchs. 4.5. Lessons from whole animal physiology Measurement of respiratory quotients (RQ) in elasmobranchs would be bene?cial in elucidating whole animal metabolic fuel preference, but there are few data available on RQ or whole animal CO2 production by elasmobranchs. The singular exception is the determination of Ogden (1945) of an RQ between 0.85 and 0.95 for the dusky smooth-hound shark (Mustelus californicus). These values are very close to the ‘classic’ mammalian value of 0.8–0.9 for protein/amino acid carbon being the predominant carbon source for aerobic metabolism. Alternatively, the nitrogen quotient (NQ) re?ects the ratio of the amount of nitrogen released to oxygen consumed and an NQ of 0.27 suggests 100% of aerobic respiration is fueled by protein catabolism (van den Thillart and Kesbeke, 1978; Wood 2001). Wood et al. (2007) determined NQ values of 0.19 and 0.51 for fed and 5-day fasted S. acanthias, further supporting the importance of amino acid oxidation in whole body metabolism of elasmobranchs. The > 100% NQ in fasted S. acanthias deserves brief discussion. Wood et al. (2007) interpreted this value as indicating a negative nitrogen balance, where more nitrogen was lost than could be supplied via protein oxidation. However, the relatively stable plasma [urea] found in fasted S. acanthias argues against negative balance (Kajimura et al., 2008). Meanwhile, urea losses, which are substantial in elasmobranchs, remained constant over 56 days of food deprivation (Kajimura et al., 2008). Using values from Kajimura et al. (2008), food deprived S. acanthias lose nitrogen at approximately 0.6 mmol N kg? 1 h? 1 or 14.4 mmol N kg? 1 day? 1. Based on the rate of loss of body mass recorded by Kajimura et al. (2008), the amount of nitrogen made available for urea replacement in S. acanthias is remarkably similar to the rate of nitrogen losses (primarily as urea) (Appendix A). Our calculations are somewhat equivocal, however, because the lower 0.3% day? 1 value suggests nitrogen de?cit whereas with only a 0.2 % increase (=0.5% day? 1), nitrogen balance or excess nitrogen availability would be expected (Appendix A). Moreover, using the greater rate of body mass loss (~1%day? 1) found by Leech et al. (1979) over the ?rst 25 days of fasting in S. acanthias, nitrogen production from protein catabolism would far exceed the nitrogen losses. Further measurements are needed to better ascertain the role of amino acid oxidation in energy production and urea synthesis during fasting in elasmobranchs. Even if the available NQ values are overestimates of ‘normal’ conditions, or the animals are in fact in nitrogen de?cit, the data still support a substantial reliance on protein catabolism in food deprived elasmobranchs. This contrasts with the situation in fasting teleosts, where the order of aerobic fuel utilization is lipid > carbohydrate> protein (Wood, 2001). In elasmobranchs, unlike most teleosts, the reliance on protein catabolism and amino acid oxidation even in fed animals is likely not simply a case of providing aerobic fuels but also for providing substrates for urea synthesis in order to match urea losses to the environment. The potential consequences of this for the organization of energy metabolism of elasmobranchs are considered in Section 8.5. 5. Carbohydrates Carbohydrates are utilized as a fuel source in elasmobranch tissues but the overall contribution to energy metabolism is unclear. In elasmobranch heart, and to a lesser extent, red muscle, brain and rectal gland, signi?cant hexokinase activity is present, suggesting that circulating glucose may be an important oxidative fuel in these tissues (Moon and Mommsen, 1987; Sidell et al., 1987; Walsh et al., 2006). Isolated mitochondria from

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elasmobranch red muscle and heart oxidize pyruvate at relatively high rates, providing further support for carbohydrates as signi?cant aerobic fuels in these tissues (Chamberlin and Ballantyne, 1992; Ballantyne et al., 1992; Moyes et al., 1990a). Arteriovenous differences across the brain of S. acanthias indicate lactate production, but the lack of glucose uptake suggests that endogenous glycogen is the carbon source for this lactate (deRoos, 1994). Isolated perfused heart from the skate L. erinacea maintained performance via glycolysis, but rapidly failed following inhibition with iodoacetate (Driedzic and Hart, 1984). Glucose supports active chloride secretion by isolated perfused rectal gland and is needed for effective utilization of ketone bodies as a substrate for secretion (Walsh et al., 2006). Interestingly, severe hypoglycaemia induced by insulin administration was tolerated for several days in S. acanthias (deRoos and deRoos, 1979), potentially aided by the chronic ketosis of elasmobranchs. The reliance on carbohydrates as aerobic fuels appears to be broadly similar in elasmobranchs and carnivorous teleosts (Driedzic and Hart, 1984; Sidell et al., 1987; Moyes et al., 1989; Chamberlin et al., 1991). Also like teleosts, glycogen is an important fuel source in locomotory muscle of elasmobranchs. Following creatine phosphate depletion, glycogen is the primary source of anaerobic ATP generation in exercising muscle of S. acanthias, resulting in a marked accumulation of lactate (Richards et al., 2003). The activity of pyruvate dehydrogenase in white muscle is very low and does not respond to exhaustive exercise or recovery, indicating aerobic carbohydrate oxidation is of minor importance under these conditions (Richards et al., 2003). Much of the formed lactate was retained within the muscle and was converted back to glycogen in situ, although the mechanism of intramuscular glycogenesis is not clear. Although Crabtree et al. (1972) detected the gluconeogenic enzyme phosphoenolpyruvate carboxykinase (PEPCK) in elasmobranch muscle, their measurements were obtained using an assay now known to be unreliable for tissue extracts (Ballantyne, 1997). In fact, PEPCK activity is undetectable in elasmobranch muscle and the intramuscular recycling of lactate to glycogen may occur via the hypothesized reversal of pyruvate kinase proposed to occur in muscle of ?shes (Moon and Mommsen, 1987; Suarez et al., 1986b). Despite the apparent utilization of glucose as a metabolic fuel, levels of circulating glucose appear to be maintained relatively constant even during long-term fasting (Zammit and Newsholme, 1979; deRoos et al., 1985) or after feeding (Walsh et al., 2006), although Kajimura et al. (2008) reported an increase in plasma [glucose] during several weeks of fasting in S. acanthias (Kajimura et al., 2008). Studies are needed on glucose turnover in elasmobranchs. Hepatic gluconeogensis occurs in elasmobranchs but the rate is much lower than that found in teleosts, and, unlike teleosts, the kidney in elasmobranchs may not be an important site of gluconeogenesis (Moon et al., 1985; Mommsen and Moon, 1987). Alanine, serine and lactate are gluconeogenic substrates in elasmobranchs, as they are in other vertebrates (Mommsen and Moon, 1987; Anderson, 1990). Research on the regulation of carbohydrate metabolism in elasmobranchs has largely focused on hormonal control of glucose mobilization. Similar to other vertebrates, catecholamines and cortisol cause a rapid glucose mobilization (Patent, 1970; deRoos and deRoos, 1978). ACTH induces slower increases in plasma [glucose], probably indirectly via stimulation of corticoid release and mobilization of gluconeogenic substrates (deRoos and deRoos, 1992). In contrast, both elasmobranch and mammalian insulins have a slow hypoglycaemic effect in elasmobranchs (Patent, 1970; deRoos and deRoos, 1979; deRoos et al., 1985; Anderson et al., 2002). Studies are needed to ascertain the physiological role of hormonal control of carbohydrate oxidation in elasmobranchs, including during feeding and fasting. 6. Environmental in?uences The impact of environmental factors such as salinity, carbon dioxide, oxygen, and temperature on the unusual energy metabolism of elasmobranchs is understudied.

6.1. Salinity The effect of changes in environmental salinity on the energy metabolism of elasmobranchs is not well understood. During salinity acclimation in the moderately euryhaline freshwater stingray Himantura signifer, a decrease was observed in the ratio of n?3/n?6 plasma NEFA, which may re?ect utilization of n?3 fatty acids during osmoregulatory acclimation (Speers-Roesch et al., 2008). Few other metabolic effects were observed: total plasma [NEFA] was unaffected and activities of enzymes of energy metabolism were largely unchanged in liver, kidney, or heart (Speers-Roesch et al., 2006a; Speers-Roesch et al., 2008). Salinity acclimation in teleosts also has little effect on lipid metabolism, including plasma [NEFA], but causes more substantial changes in amino acid metabolism (Bystriansky et al., 2007). In elasmobranchs, oxidation and resultant depletion of intracellular osmolytes, including the amino acid derivatives ?-alanine and sarcosine, may be important during whole animal, cellular, and mitochondrial hypoosmotic stress (King et al., 1980; Ballantyne et al., 1986). Further studies are needed to evaluate the role of lipids and amino acids in salinity acclimation in elasmobranchs. 6.2. Hypercapnia and hypoxia How hypercapnia affects the energy metabolism of elasmobranchs is unknown, but some data are available on the effects of hypoxia. Exposure to hypoxia is associated with a depression of oxygen uptake in the hypoxia-tolerant epaulette shark (Hemiscyllium ocellatum) (Routley et al., 2002). Reliance on anaerobic glycolysis is indicated by accumulation of lactate, but there is no increase in plasma [glucose], unlike what is typically seen in hypoxia-tolerant teleosts (Routley et al., 2002). Plasma [glucose] also does not change in S. acanthias exposed to 1 mg O2/L for 5 h (B. Speers-Roesch and J.G. Richards, unpublished results). How lipid, amino acid, or ketone body metabolism is affected by hypoxia exposure in elasmobranchs is unknown. Ketone body metabolism, in particular, potentially presents an interesting avenue of research because neuroprotective effects of ketone bodies have been demonstrated in mammalian ischaemia models (Suzuki et al., 2001). Additionally, ketone bodies have an ATP yield per mol O2 that is similar to glucose, making them a good alternative aerobic fuel in hypoxia when minute yet important rates of substrate oxidation still occur (Hochachka and Somero, 1984; Veech, 2004). 6.3. Temperature Temperature is one of the most pervasive abiotic factors in?uencing the physiology of aquatic animals but there are very few studies on its effects on the energy metabolism of elasmobranchs. The typical ectothermic pattern of increased metabolic rate (oxygen uptake) at warmer acclimation temperature and decreased metabolic rate at cooler acclimation temperature is observed in elasmobranchs (e.g. Butler and Taylor, 1975; Tullis and Baillie, 2005). In an attempt to investigate in elasmobranchs the concept of metabolic compensation, Tullis and Baillie (2005) found no changes in the activities of citrate synthase, a marker of aerobic metabolism, and lactate dehydrogenase, a marker of anaerobic potential, in white muscle of the tropical bamboo shark Chiloscyllium plagiosum during acclimation to 15 °C compared with 25 °C controls. Further studies are needed on the metabolic responses to temperature in elasmobranchs. It would be particularly interesting to investigate how pathways of substrate oxidation are affected by cold temperature acclimation and adaptation in elasmobranchs. In cold-adapted and cold-acclimated teleosts and invertebrates, fatty acids are preferred oxidative fuels in muscle and other tissues and lipid storage is enhanced (P?rtner et al., 2005). This preference for lipids is probably in part due to increased mitochondrial densities and related biochemical consequences as well as ecological factors such as metabolic requirements for overwintering (P?rtner et al., 2005). Because elasmobranch muscle does

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not rely on fatty acid oxidation, instead it is possible that pathways of ketone body and amino acid oxidation are enhanced in polar or coldacclimated elasmobranchs. Alternatively, elasmobranchs provide a model to investigate whether an inability to effectively utilize fatty acids as an energy source in muscle constrains cold tolerance and adaptation. Compared with teleosts, few elasmobranchs routinely inhabit polar waters (M?ller et al., 2005; Verde et al., 2005), even though their isoosmotic solute system would seem to pre-adapt them to sub-zero water: due to colligative properties, the freezing point of elasmobranch blood is similar to or below the surounding seawater (Watts and Watts, 1974). Only 20 elasmobranch species commonly occur in polar waters, 17 of which are rajid skates, the most speciose elasmobranch family (M?ller et al., 2005). The paucity of elasmobranchs in polar waters does not appear to simply re?ect their lower worldwide species richness compared with teleosts. The number of rajid species inhabiting polar waters represent ~ 7% of the total number of known rajid species (M?ller et al., 2005). Among some of the more diverse teleost families found at the poles (and ignoring the endemic Antarctic notothenioid radiation), the percentage of species found in polar waters is ~ 38%, ~ 24%, and ~ 57% for snail?shes (Liparidae), eelpouts (Zoarcidae), and cods (Gadidae) (M?ller et al., 2005). Similarly, only 3 families of elasmobranchs are represented routinely in polar waters (~ 6% of total families), vs. 64 families of teleosts (~ 14% of total families) (M?ller et al., 2005; Nelson 2006). Studies on the energy metabolism of polar elasmobranchs and cold-acclimated elasmobranchs are warranted to assess whether the inability to utilize fatty acid oxidation in muscle may have been one constraint on polar invasion by elasmobranchs. 7. Energy metabolism and the incidence of cancer in elasmobranchs Ballantyne (1997) speculated that the comparatively low levels of glutamine and [NEFA] found in plasma of elasmobranchs could be related to their reported low incidence of cancer, because glutamine and NEFA are important fuels for tumor growth. Ostrander et al. (2004) have argued, however, that the scarcity of records of cancer in elasmobranchs may be an artifact of observer bias, a lack of systematic surveys of cancer rates in elasmobranchs (in contrast to the many such studies done on teleosts), and reduced exposure to environmental contaminants among pelagic sharks. Ostrander et al. (2004) concede, however, that in the absence of more detailed analyses, a relatively low incidence of cancer in elasmobranchs remains a possibility. The possible physiological explanations suggested by Ballantyne (1997) deserve further attention. What is clear is that the promotion and use of shark cartilage and other crude by-products as anti-cancer ‘medicines’ is exploitative and unsupported by scienti?c evidence (Ballantyne, 1997; Ostrander et al., 2004). 8. Why do elasmobranchs have an unusual energy metabolism? A major challenge is to identify and empirically test hypotheses for the ultimate causes of the unusual metabolic organization of elasmobranchs, in which ketone bodies and amino acids are favored over fatty acids as oxidative substrates in muscle. Here we consider the evidence for and against previously suggested hypotheses as well as propose new hypotheses. 8.1. Paleozoic atmospheric oxygen levels Moyes et al. (1990b) speculated that the relatively low atmospheric oxygen levels that existed during the Silurian–Devonian period when elasmobranchs arose could have led to a metabolism that did not emphasize lipids because of the known inef?ciency and deleterious by-products of fatty acid oxidation in hypoxic conditions. Although this proposal remains untested, the assumption of low

atmospheric oxygen levels during this period may be incorrect based on more recent estimates showing values closer to those today (Berner et al., 2007). 8.2. Effects of TMAO on fatty acid oxidation Ballantyne and Moon (1986) and Ballantyne (1997) suggested that fatty acid oxidation may be restricted to the liver of elasmobranchs because it minimizes overall the disruptive effects of TMAO on fatty acid oxidation. They based this hypothesis on the observation of lower [TMAO] in liver than in muscle and on the discovery that TMAO inhibits the oxidation of long-chain fatty acylcarnitines by liver mitochondria from the skate L. erinacea, which was attributed to the similarity in structure between TMAO and carnitine, itself a trimethylamine (Ballantyne and Moon 1986). Thus, centralization of fatty acid oxidation to the liver may be a more ef?cient metabolic organization than possessing adaptations to the effects of TMAO in all tissues. As a corollary, maintenance of lower hepatic [TMAO] is a compromise between the needs to maintain fatty acid oxidation while counteracting the effects of urea (Ballantyne and Moon, 1986). However, TMAO does not affect the apparent Km of CPT-1 in S. acanthias liver mitochondria, which is inconsistent with the notion that TMAO competes with carnitine for access to CPT-1 (Treberg et al., 2006a). 8.3. Buoyancy role of the liver The large, lipid-rich liver that affords buoyancy in elasmobranchs is conspicuous as a potential cause of the de-emphasis on muscle fatty acid oxidation. Perhaps, the sequestration of lipids in the liver to achieve buoyancy constrained the extrahepatic utilization of fatty acids as fuels. As discussed previously (Section 2.3), the buoyancy role of the liver is apparently conserved even in energy-limited situations such as fasting and the deep-sea environment, underscoring the importance of lipids for hydrostatic lift rather than simply for energy production (Baldridge, 1972; Ballantyne, 1997). This hypothesis lacks an obvious explanation for why fatty acid oxidation would be limited in muscle only, but because muscle is the largest component of body mass, reductions in fatty acid oxidation and enhanced amino acid oxidation there might substantially conserve lipids for hepatic storage. Interestingly, in rays but not sharks examined to date, there appears to be a limited capacity for cardiac fatty acid oxidation (Speers-Roesch et al., 2006a), perhaps because increased extrahepatic fatty acid utilization in benthic rays is possible due to the relaxed buoyancy role of liver. This speculative hypothesis could be evaluated by studying the capacity for fatty acid oxidation in teleosts that accumulate lipids as their primary source of buoyancy (Phleger, 1998) as well as in muscles of pelagic rays. 8.4. The urea–albumin hypothesis The potential explanation that has been most thoroughly evaluated is the proposal that the adoption in elasmobranchs of urea-based osmoregulation compromised fatty acid transport needed to sustain oxidation in extrahepatic tissues (Ballantyne and Moon, 1986; Ballantyne et al., 1987). The reasoning behind this ‘urea–albumin hypothesis’ involves the perturbing effect of urea on hydrophobic interactions of proteins. Albumin, an important fatty acid binding protein in many vertebrates but absent in elasmobranchs (Metcalf and Gemmell, 2005), relies on hydrophobic interactions not only for maintenance of tertiary structure but also for fatty acid binding (Scheider et al., 1976; Peters, 1996). Ballantyne and Moon (1986) and Ballantyne et al. (1987) suggested that the high levels of urea that accompanied urea-based osmoregulation in elasmobranchs may have limited fatty acid binding by albumin, leading to impaired fatty acid transport and the evolutionary loss of albumin and extrahepatic fatty acid oxidation. Instead, highly soluble

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ketone bodies and amino acids, whose transport does not require carriers, were favored. Researchers have looked to freshwater elasmobranchs, which accumulate little urea (Thorson et al., 1967; Treberg et al., 2006b), as a test of the urea–albumin hypothesis, reasoning that the evolution of teleost-like osmoregulation in these species may have been attended by an adoption of a metabolic organization similar to teleosts, where fatty acid oxidation plays a central role. The early studies examined the enzymatic capacity for fatty acid oxidation in tissues from Amazonian freshwater stingrays (Potamotrygon spp.), which accumulate no urea and lack the ability to synthesize urea (Thorson et al., 1967). The results were contradictory. Singer and Ballantyne (1989) detected very low to undetectable levels of enzymes involved in fatty acid oxidation (including CPT) in heart, kidney, brain, and muscle from captive juvenile . magdalenae (Table 1). Driedzic and De Almeida-Val (1996), however, found relatively high levels of CPT in heart (other tissues were not examined) from wild-caught adult P. hystrix (Table 1). Speers-Roesch et al. (2006a) attempted to resolve this discrepancy and provide a more comprehensive test of the urea–albumin hypothesis by examining the capacity for fatty acid oxidation, as indicated by tissue CPT activities and total plasma [NEFA], in a variety of marine and freshwater elasmobranchs possessing a wide range of urea levels: virtually none in the stingray Potamotrygon motoro, medium levels (~40–80 mM) in the moderately euryhaline stingray H. signifer in freshwater and acclimated to 50% seawater, and high levels in the marine C. punctatum (a shark) and T. lymma (a stingray) (>200 mM) (Treberg et al., 2006b). There was no indication of a relationship between urea content and the capacity for fatty acid oxidation or transport; the freshwater forms did not possess enhanced capacity compared with the marine forms (Speers-Roesch et al., 2006a) (Table 1; Table 2). The capacity for ketone body oxidation also was similar across all species (Speers-Roesch et al., 2006a). Speers-Roesch et al. (2006a) concluded that these results are evidence against the urea–albumin hypothesis. A potential criticism of this conclusion is that it assumes that the loss of high urea content in freshwater rays would be accompanied by an adoption of enhanced fatty acid oxidation if the urea–albumin hypothesis were correct. However, even if high urea levels accompanying the evolution of urea-based osmoregulation were the initial cause of the loss of muscle fatty acid oxidation capacity via impaired fatty acid transport, ketone bodies may have been an adequate alternative fuel such that adoption of fatty acid oxidation was not selected for in freshwater rays. In addition, the loss of molecular and biochemical machinery of fatty acid oxidation in muscle may have constrained the reevolution of this pathway. Nevertheless, the ?nding of Speers-Roesch et al. (2006a) of the capacity for fatty acid oxidation, as measured by CPT activity, in rectal gland and kidney begs the question of why the loss of effective fatty acid transport, as posited by the urea–albumin hypothesis, would have caused loss of fatty acid oxidation in muscle but not other extrahepatic tissues. Furthermore, as mentioned previously (Section 2.2), there are a number of teleosts with relatively high capacities for peripheral fatty acid oxidation that lack albumin, and like elasmobranchs they utilize lipoproteins to transport fatty acids (Moyes et al., 1989; De Smet et al., 1998; Metcalf et al., 1999a,b; West et al., 1999). Thus, there is no a priori reason to hypothesize that lacking albumin is a death knell for effective transport of plasma NEFA and peripheral fatty acid oxidation. Finally, both albumin-mediated NEFA transport and fatty acid oxidation occur substantially in the mammalian kidney, where relatively high levels of urea exist (Guder et al., 1986; Trimble, 1993). Considering these points and the results of Speers-Roesch et al. (2006a), it is reasonable to suggest that the urea–albumin hypothesis does not explain the organization of energy metabolism in elasmobranchs. However, further studies on the capacity for fatty acid oxidation in other urea-retaining animals, especially ureosmotic species such as the coelacanth (Latimeria spp.) (Yancey, 2001), may prove useful to further test this hypothesis.

8.5. Requirement for urea synthesis The evolution of urea-based osmoregulation in elasmobranchs may have had other consequences for the energy metabolism of elasmobranchs. Even during fasting the high levels of urea in elasmobranchs are maintained for osmoregulation in the face of signi?cant urea losses to the environment (see Sections 4.2 and 4.5). We propose that the need to replace this urea places unique constraints on the energy metabolism of elasmobranchs that may have been important in the evolution of their unusual pattern of fuel selection. During fasting, catabolism of endogenous proteins is likely the most signi?cant source of nitrogen for replacing urea. Periodic but great reliance on protein catabolism during food deprivation may have favored the utilization of glutamine and other amino acids as a major muscle oxidative fuel. Ballantyne (1997) ?rst noted the possible relationship between the need for amino acids for urea synthesis and the utilization of amino acids as oxidative fuels. We also suggest that alanine release from muscle and uptake in the liver for urea synthesis (Leech et al., 1979; Fig. 1) may have helped shift alanine from an ‘antiketogenic’ to a ketogenic precursor, following our proposal of an elasmobranch alanine– ketone body cycle (Fig. 1; see Section 4.2). This in turn could have emphasized the production and utilization of ketone bodies in elasmobranchs and this could have led to relaxed selection for and eventually loss of fatty acid oxidation in muscle. The muscle-speci?city could be explained by preferential cycling of carbon as ketone bodies to muscle to help replenish losses from protein catabolism (Fig. 1). The proposed functional coupling between amino acid release from muscle and ketogenesis should be evaluated in marine and freshwater elasmobranchs where the demands on urea replacement are expected to differ. 8.6. Random loss of muscle fatty acid oxidation machinery While the urea–albumin hypothesis posits that a loss of effective fatty acid transport resulted in reduced extrahepatic fatty acid oxidation and enhanced ketone body oxidation, it is potentially more parsimonious that an initial loss of fatty acid oxidation speci?cally in muscle of elasmobranchs was followed by a reduced capacity for fatty acid transport and an emphasis on ketone body oxidation. Possibly, the cause of the unusual energy metabolism of elasmobranchs could simply have been a random loss- or impairment-of-function mutation in a fatty acid oxidation protein (e.g. CPT-1, CPT-2) in muscle that impaired this pathway locally. Rat liver CPT-1 can be inactivated by a single amino acid substitution at a highly conserved residue (Napal et al., 2003); CPT-1 de?ciency disease is known for the liver isoform of mammals, but not for the muscle isoform, limiting its applicability to the elasmobranch question (Bonnefont et al., 2004). Speculatively, neutral or positive selection potentially combined with an ancient genetic bottleneck could have ensured that the impaired muscle fatty acid oxidation phenotype prevailed in elasmobranchs. An emphasis on ketone body and amino acid oxidation in part could have been achieved as described in Section 8.5 and may have even preceded and lessened the consequences of the random loss of fatty acid oxidation. Molecular and genetic analyses of the proteins of the fatty acid oxidation pathway in elasmobranch muscle are needed to assess this ‘random loss’ hypothesis. Preliminary analysis of the partial cDNA sequences of two isoforms of CPT-1 expressed at relatively high mRNA levels in heart and skeletal muscle of S. acanthias showed conservation of amino acid residues with known catalytic and regulatory importance within this region of the CPT-1 gene in mammals (B. Speers-Roesch and J.G. Richards, unpublished results). However, cloning and functional analyses of the full-length sequences and the promoter and untranslated regions of CPT-1 genes in elasmobranchs are needed to fully address the hypothesis and to evaluate the possibility that these isoforms are ‘fossilized’ pseudogenes. Identifying the proximate causes of the limited fatty acid oxidation in muscle of elasmobranchs will be important in evaluating ultimate explanations for its origin.

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Most explanations for the unusual energy metabolism of elasmobranchs considered here rely on the assumption that fatty acid oxidation in muscle was lost within this lineage. Another possibility is that the ancestral metabolic state of ?shes involves negligible fatty acid oxidation in muscle, which we consider in the next section. 9. Comparative and evolutionary considerations We have shown that elasmobranchs rely heavily on ketone bodies rather than fatty acids as oxidative fuels in muscle, whereas the opposite is true in teleosts and, under routine conditions, in mammals. Here we will compare the energy metabolism of elasmobranchs with those of some of the other major groups of ?shes in order to gain insight into the patterns, origins, and evolution of energy metabolism in ‘lower’ vertebrates. 9.1. Basal Actinoptergyii Energy metabolism has been investigated in some basal actinopterygians, including the more ancestral chondrostean sturgeons (Acipenser spp.) as well as the neopterygian bow?n (Amia calva) and Florida gar (Lepisosteus platyrhincus), which are thought to be more closely related to teleosts (Kikugawa et al., 2004; Nelson, 2006). Although the extant species possess certain derived anatomical characteristics, the origins of these groups trace back to the early to middle Devonian period (400– 370 mya) and later led to the evolution and diversi?cation of teleosts in the Jurassic and Cretaceous periods (200–65 mya) (Pough et al., 1999; Kikugawa et al., 2004; Zhu et al., 2009). Bow?n and sturgeon show detectable but very low CPT activity in most tissues, with the exception of heart in bow?n and liver and kidney in lake sturgeon (Acipenser fulvescens) where CPT is undetectable (Singer and Ballantyne, 1991; Singer et al., 1990) (Table 1). The low CPT activities in tissues may be explained in part by comparatively low mitochondrial densities as indicated by citrate synthase activities, especially considering that Chamberlin et al. (1991) subsequently demonstrated that isolated mitochondria from bow?n red muscle oxidize long-chain fatty acylcarnitines at relatively high rates. In the Florida gar, CPT activity is high in liver and heart but, similar to elasmobranchs, is undetectable in red muscle (Frick et al., 2007) (Table 1). Sturgeons have high plasma [NEFA] compared with elasmobranchs, whereas gar and bow?n have lower elasmobranchlike levels (Table 2) (Singer et al., 1990; Jarvis and Ballantyne, 2003; Frick et al., 2007; Speers-Roesch et al., 2008). Our understanding of the capacity of fatty acid oxidation in basal actinopterygians is unclear, although it may be lower overall compared with teleosts. In tissues from sturgeon, bow?n, and gar, activities of β-HBDH and SKT are detectable but relatively low (Singer et al., 1990; Singer and Ballantyne, 1991; Frick et al., 2007). However, in some of these studies the detergent Triton-X 100 was included in the homogenizing buffers, which is now known to potentially cause loss of activity of β-HBDH, at least in elasmobranchs (Treberg et al., 2006a; Speers-Roesch et al., 2006a). Nonetheless, bow?n red muscle mitochondria oxidize ketone bodies relatively poorly in comparison with those from elasmobranchs, possibly due in part to lower β-HBDH activity (Chamberlin et al., 1991; Singer and Ballantyne, 1991). These data indicate that basal actinopterygians possess a relatively low capacity for ketone body oxidation. Although red muscle mitochondria from bow?n show lower rates of oxidation of glutamine in comparison with pyruvate (Chamberlin et al., 1991), amino acid oxidation is still clearly signi?cant in this species, as in elasmobranchs (Singer and Ballantyne, 1991). Enzyme activities support the importance of amino acid oxidation in sturgeon (Singer et al., 1990), as well as gar, although glutamine utilization in gar may be limited (Frick et al., 2007). The metabolic organization of basal actinopterygians has been described as intermediate between elasmobranchs and teleosts with

regard to their utilization of fatty acids and ketone bodies as oxidative fuels (Singer et al., 1990; Singer and Ballantyne, 1991). While the available data are not inconsistent with this hypothesis, more information is needed on the fuel preferenda of mitochondria and turnover of oxidative fuels in basal actinopterygians. Future studies should be extended to other species of basal actinopterygians, including the bichirs. 9.2. Sarcoptergyii The Sarcopterygii are the sister taxon to the Actinopterygii and include the ancient lobe-?nned ?shes (lung?shes and coelacanths) as well as tetrapods (Brinkmann et al., 2004). The earliest lobe-?nned ?shes appeared at least 418 mya and share characteristics with early actinopterygians, chondrichthyans, and, possibly, the common gnathostome ancestor (Zhu et al., 2009). The extant lung?shes and coelacanths therefore are of particular interest with regards to the evolution of energy metabolism in early vertebrates. Unfortunately, the organization of energy metabolism has been examined only in the African lung?sh (Protopterus dolloi). Interestingly, it has a tissue-speci?c distribution of CPT activity comparable to elasmobranchs, with relatively high levels in kidney and liver and undetectable levels in heart and muscle (Frick et al., 2008b) (Table 1). P. dolloi also possess detectable but relatively low activities of D-β-HBDH in all tissues (Frick et al., 2008b), but use of Triton-X 100 may have caused underestimates of activities. Relatively high levels of SKT further suggest the utilization of ketone bodies (Frick et al., 2008b). Carbohydrates and amino acids both appear to be important aerobic fuels in lung?sh (Frick et al., 2008a). Plasma [NEFA] have not been measured in lobe-?nned ?shes, but the Australian lung?sh (Neoceratodus forsteri) possesses an albumin that is tetrapod-like rather than teleost-like (Metcalf et al., 2007). 9.3. ‘Agnatha’: Myxini and Petromyzontida The jawless hag?shes (Myxini) and lampreys (Petromyzontida) are ancient ?shes whose origins, interrelationships, and relationships to gnathostomes are unclear (Nelson, 2006; Janvier, 2007). Hag?shes are generally considered to be the most ‘primitive’ of ?shes, in part based on their status as isoosmotic osmoconformers (Grif?th, 1991). The heart and lateral muscle as well as the liver of hag?shes (Myxine glutinosa and Eptatretus stoutii) possess detectable CPT activity but the levels are very low even when corrected for citrate synthase activity and are similar to values seen in elasmobranchs (Table 1) (Hansen and Sidell, 1983; Moyes et al., 1990a; Leary et al., 1997). Plasma [NEFA] in M. glutinosa is similar to that found in elasmobranchs (Table 2) (P.J. LeBlanc, C. Hyndman, and J.S. Ballantyne, unpublished results, cited in Speers-Roesch et al., 2006b). The heart of M. glutinosa appears to preferentially utilize glucose rather than fatty acids and cannot function when exogenous palmitate is the only available fuel (Hansen and Sidell, 1983; Sidell, 1983). The heart of M. glutinosa lacks β-HBDH activity (Hansen and Sidell, 1983), but muscle in hag?sh has glutaminase activity that suggests that amino acid oxidation occurs (Chamberlin and Ballantyne, 1992). Overall, these data suggest that carbohydrates are the preferred metabolic fuel of hag?shes (Sidell, 1983). Lampreys, which represent the only other extant group of jawless ?shes and osmoregulate in a similar manner as teleosts (Yancey, 2001), have a teleost-like metabolic organization. Petromyzon marinus and Geotria australis possess readily detectable CPT activities in liver and muscle (Table 1) and mitochondria from skeletal muscle of P. marinus oxidize long-chain fatty acylcarnitines at relatively high rates (Power et al., 1993; LeBlanc et al., 1995). Lampreys have albumin and relatively high plasma [NEFA] (Table 2) (LeBlanc et al., 1995; Peters, 1996). β-HBDH and SKT activities are present, but relatively low (LeBlanc et al., 1995). Mitochondrial oxidation of β-HB occurs in muscle and liver, although the rates are low compared with fatty acid

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carnitine and especially pyruvate (LeBlanc et al., 1995). Rates of mitochondrial amino acid oxidation are also low in muscle from lamprey compared with other ?shes (LeBlanc et al., 1995). 9.4. Other Chondrichthyes Elasmobranchs diverged from their closest relatives, the holocephalans (chimaeras), at least 400 mya (Grogan and Lund, 2004), yet the metabolic organization of the holocephalan Hydrolagus colliei is virtually identical to that of elasmobranchs (Speers-Roesch et al., 2006b). H. colliei has low plasma [NEFA] (Table 2), a high capacity for fatty acid oxidation in kidney and liver, and a low capacity for fatty acid oxidation in heart and skeletal muscle as indicated by CPT activities (Table 1). Enzymes related to ketone body oxidation are relatively high in all aerobic tissues (Speers-Roesch et al., 2006b). Thus, the unusual energy metabolism of elasmobranchs probably occurs in all chondrichthyans. Since all marine chondrichthyans are urea-based osmoregulators, our proposal regarding the effects of urea synthesis on the energy metabolism of elasmobranchs (Section 8.5) can be extended to the Chondrichthyes. 9.5. Energy metabolism of early vertebrates Because the divergence and relationship between holocephalans and elasmobranchs is well supported by paleontological, molecular, and anatomical data (Grogan and Lund, 2004), Speers-Roesch et al. (2006b) concluded that the unusual metabolic organization of chondrichthyans was also present in the common ancestor of all chondrichthyan ?shes, which lived 450–400 mya, near the time of the origin of jawed vertebrates. This is an example of ‘paleophysiology’, which includes not only physiological analyses of fossil remains but also the comparative study of the physiologies of at least two extant lineages whose divergence extends deep into geological time and whose phylogenetic relationship is supported by robust evidence. One goal of ‘paleophysiology’ is to achieve a reliable indication of the physiologic state of the common ancestor that is likely now extinct and may or may not be known from the fossil record. Based on the apparently ancient origin of the metabolic organization of chondrichthyans, Speers-Roesch et al. (2006b) speculated that it might re?ect the ancestral metabolism found in the earliest jawed vertebrates, including chondrichthyans and, potentially, the now extinct placoderms and acanthodians (Brazeau, 2009). Pointing to the preference for carbohydrates and potentially amino acids rather than fatty acids as oxidative fuels in muscle of hag?shes as the ancestral craniate and ancestral vertebrate metabolic state, SpeersRoesch et al. (2006b) suggested that ketone bodies and amino acids became important aerobic fuels in muscle of the earliest jawed ?shes, including chondrichthyans, possibly to help sustain higher activity levels. The discovery of an elasmobranch-like energy metabolism in lung?shes (Frick et al., 2007) is consistent with this view, as the sarcopterygians appeared during the same time period (Zhu et al., 2009). As described in Section 8.5, the need to synthesize urea could have contributed to the selection of ketone bodies and amino acids as aerobic fuels, especially considering that early jawed vertebrates may have possessed the urea-based osmoregulation found in chondrichthyans (Grif?th, 1991). Later in jawed vertebrate evolution fatty acids may have been adopted as an alternative fuel in muscle by basal actinopterygians, and then as a primary fuel in teleosts, at the expense of ketone body utilization (Speers-Roesch et al., 2006b). A similar scenario also may have occurred in the evolution of sarcopterygians leading to tetrapods. Why fatty acid oxidation might have become enhanced in teleosts and tetrapods is unclear, but one possibility is the loss in most species of the need for urea synthesis for osmoconformation and its potential enhancing effects on ketogenesis (Section 8.5). While highly speculative, this proposal presents another hypothesis for the unusual energy metabolism of elasmobranchs that should be

evaluated by undertaking a detailed, systematic molecular and biochemical examination of the pathways of fatty acid and ketone body oxidation in basal chordates, such as urochordates and cephalochordates, to the derived teleosts and tetrapods. This ancestral hypothesis typically has been refuted by pointing out that lampreys have a metabolic organization more similar to teleosts (Ballantyne, 1997). This remains a potentially valid criticism, but may be explained by the complex life history of lampreys, which involves energy-intensive spawning migrations (LeBlanc et al., 1995) that may have led to the adoption of enhanced lipid metabolism (Speers-Roesch et al., 2006b). Similarly, lampreys also are unusual in possessing a more ‘advanced’ hypoosmotic osmoregulatory strategy (Yancey, 2001). An interesting question for future research is why teleosts and lampreys share certain major physiological traits. Such work may help evaluate the hypothesis advanced above. 10. General conclusions The unusual energy metabolism of elasmobranchs is characterized by limited or absent fatty acid oxidation in cardiac and skeletal muscle and a great reliance on ketone bodies and amino acids as oxidative fuels in these tissues. Other extrahepatic tissues in elasmobranchs rely on ketone bodies and amino acids for aerobic energy production but, unlike muscle, also appear to possess a signi?cant capacity to oxidize fatty acids. This is an important distinction: it is likely inaccurate to refer to elasmobranchs as lacking extrahepatic fatty acid oxidation. Rather, only muscle appears to lack fatty acid oxidation and in some species the heart may even have a low capacity to utilize fatty acids as oxidative fuels. The apparent presence based on CPT activities of substantial fatty acid oxidation in rectal gland and kidney of elasmobranchs, and lesser rates in heart of some rays, needs to be con?rmed. The preference for ketone body oxidation rather than fatty acid oxidation in muscle of elasmobranchs under routine conditions is opposite to the situation in teleosts and mammals. Carbohydrates appear to be utilized as a fuel source in most tissues in elasmobranchs, comparable to teleosts and mammals. In elasmobranchs, amino acids serve multiple important roles as oxidative fuels, important ketogenic precursors, and essential nitrogen donors for high rates of urea synthesis. Elasmobranchs possess a large, distinctive liver that serves as a storage depot for lipids utilized as energy sources and for buoyancy. Hepatic lipids appear to be mobilized as oxidative fuels in the form of NEFA and TAG as well as ketone bodies via ketogenesis. Unlike mammals but like many teleosts, elasmobranchs lack albumin and instead NEFA and other lipids are carried by lipoproteins. Potentially related to the reduced fatty acid oxidation in muscle, NEFA levels are relatively low in blood of elasmobranchs compared with teleosts. Ketone bodies, especially β-HB, are present at relatively high concentrations even under fed conditions and consequently elasmobranchs experience a chronic ketosis like that of fasting mammals. Further studies are needed on whole animal and tissue-level turnover, utilization, and physiological roles of fatty acids, ketone bodies, amino acids, and carbohydrates in elasmobranchs. Broad similarities are apparent between physiological regulation of energy metabolism in elasmobranchs, teleosts, and mammals, especially regarding carbohydrate metabolism and the response of ketone body metabolism to fasting. On the other hand, differences in the hormonal regulation of ketone body and lipid metabolism appear to exist between these vertebrate groups. Almost nothing is known about physiological regulation of amino acid metabolism in elasmobranchs, despite their important physiological roles. Similarly, our understanding of the in?uences of environmental factors on the energy metabolism of elasmobranchs is cursory and further studies are warranted. The urea-based osmoregulation of elasmobranchs has major consequences for the function and potentially the evolution of their energy metabolism, as emphasized by Ballantyne (1997). Although the urea–albumin hypothesis relating to urea's presumed perturbation

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of fatty acid transport does not appear to be supported by available evidence, we suggest that the need for urea synthesis especially under fasted conditions may have favored the utilization of amino acids and, via an alanine–ketone body cycle, ketone bodies as oxidative fuels instead of fatty acids (see Section 8.5). More empirical data are needed to better understand why the energy metabolism of elasmobranchs is unusual among vertebrates. Like elasmobranchs, holocephalans show a preference for ketone bodies rather than fatty acids as oxidative fuels in muscle. This suggests that the common ancestor of all chondrichthyan ?shes, which lived at least 400 mya, also had this metabolic organization. Studies are needed to assess speculation that the metabolic organization of elasmobranchs and other chondrichthyans re?ects, at least in part, the ancestral energy metabolism of jawed vertebrates. The initial observations of Zammit and Newsholme (1979) on the aerobic fuel preferences of elasmobranchs were followed by a substantial body of research, reviewed here and by Ballantyne (1997), which has afforded us a good general understanding of the unusual energy metabolism of this fascinating and ancient group of ?shes. Nonetheless, many unanswered questions remain. Future work on the energy metabolism of elasmobranchs will undoubtedly provide many interesting discoveries and likely will shed light on major questions related to the evolution, organization, and regulation of energy metabolism in ?shes and other vertebrates. Acknowledgements We thank our former supervisors, Jim Ballantyne (BSR) and Bill Driedzic (JRT), for their mentorship and for affording us independence in the pursuit of our interests in elasmobranch physiology. BSR thanks his current supervisor Jeff Richards for his support and interest in this ongoing work. Victor Zammit kindly shared with us his memories of his research on the metabolic biochemistry of elasmobranchs. Joy Stacey provided helpful comments on an early draft. BSR was supported by an NSERC Canada Graduate Scholarship as well as a Paci?c Century Graduate Scholarship from the University of British Columbia and the Province of British Columbia. Research by BSR was supported by an NSERC Discovery grant to James S. Ballantyne and an NSERC Discovery grant to Jeffrey G. Richards. JRT was supported during the majority of his elasmobranch research by an NSERC Post-graduate scholarship and an NSERC Discovery grant to William R. Driedzic. Appendix A Our calculations for estimated nitrogen loss vs. nitrogen availability from tissue catabolism in fasted Squalus acanthias are based on the following data. Losses of 14.4 mmol N kg? 1 day? 1 (primarily urea) are based on Kajimura et al. (2008). To calculate nitrogen availability, we assumed the following: a) b) c) d) e) Mass loss = 0.3–0.5 % day? 1 Protein content = 0.2 g g wet mass? 1 N content of protein = 11.4 mmol g? 1 Urea–N content of tissue = 0.66 mmol g wet mass? 1 Metabolically available N-(amino acids and betaine)=0.15 mmol g wet mass? 1

Using assumptions a–e we calculated the nitrogen available for urea per day from catabolism for a 1 kg dog?sh as: = 1kg??a?b?c? + 1kg??a?d? + 1kg??a?e? ?1 ?1 = 9:27 ? 15:5mmol N kg day ?A:1?

The lower value is obtained utilizing the 0.3% day? 1 mass loss estimate and the higher value is obtained using the 0.5% day? 1 mass loss estimate. References
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Data for assumptions a–e are from: a, Leech et al. (1979) and Kajimura et al. (2008); b, JRT (unpublished results) of approximate tissue protein content in elasmobranchs; c, cited in Kajimura et al. (2008); d, the urea content of most marine elasmobranch tissues is approximately 300– 350 mmol kg? 1 wet mass? 1 (Robertson, 1975; Bedford, 1983; Treberg and Driedzic, 2002; Treberg et al., 2006b), so assuming urea content is 330 mmol kg? 1 wet mass? 1, then 0.66 mmol urea–N g? 1; e, the approximate sum of α-amino acids and betaine in S. acanthias muscle (Robertson, 1975; Bedford, 1983).

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