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The role of AMP-activated protein kinase in the coordination of metabolic suppression in the common… Jibb, Lindsay A. 2008

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THE ROLE OF AMP-ACTIVATED PROTEIN KINASE IN THE COORDINATION OF METABOLIC SUPPRESSION IN THE COMMON GOLDFISH by LINDSAY A. JIBB B.Sc., Queen's University, Kingston Ontario, 2005 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE STUDIES (Zoology) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) April, 2008 © Lindsay A. Jibb, 2008 ABSTRACT Cell survival in conditions of severe oxygen deprivation depends on a wide variety of biochemical modifications, which result in a large-scale suppression of metabolism, preventing [ATP] from falling to fatally low levels. We investigated whether AMP-activated protein kinase (AMPK) has a role in the coordination of cellular modification during hypoxia, which leads to a regulated state of metabolic suppression in the goldfish (Carassius auratus). Energy charge, AMPK activity, protein and gene expression, as well as the translational capacity and phosphorylation state of a downstream target were measured in goldfish tissues during exposure to hypoxia (-0.3 mg 02/L) for up to 12 h. AMPK activity in the goldfish liver increased by 4- fold at 0.5 h hypoxia and was temporally associated with a —11-fold increase in calculated AMPfree/ATP. No change was observed in total AMPK protein or relative gene expression of identified AMPK isoforms. Changes in AMPK activity were also associated with a decreased rate of protein synthesis and an increase in the phosphorylated form of eukaryotic elongation factor-2 (eEF2; relative to total eEF2). Increases in AMPK activity were not seen in hypoxic goldfish muscle, brain, heart or gill, nor was a significant alteration in cellular energy charge seen in muscle. Still, the present study is the first to show that AMPK activity increases in liver in response to short-term severe hypoxia exposure in a hypoxia-tolerant fish. The decreased rates of protein synthesis, a well known component of metabolic suppression, combined with increased phosphorylation of eEF2, a downstream target of AMPK, potentially implicate the kinase in the cellular effort to suppress metabolism in hypoxia-tolerant species during oxygen deprivation. ii TABLE OF CONTENTS ABSTRACT^ ii TABLE OF CONTENTS^ iii LIST OF TABLES v LIST OF FIGURES^ vi LIST OF ABBREVIATIONS^ vii ACKNOWLEDGEMENTS ix CO-AUTHORSHIP STATEMENT^ x Chapter 1: GENERAL INTRODUCTION^ 1 1.1. Environmental Hypoxia^  1 1.2. Responses to Hypoxia  1 Problems associated with hypoxia^  1 Hypoxia-survival strategies of tolerant animals^ 2 Adjustments to metabolism in response to hypoxia exposure^ 2 1.3. AMP-activated protein kinase^ 5 1.4. Goldfish as a model organism 9 1.5. AMPK and its potential role in coordinating metabolic suppression in goldfish^ 9 1.6. References^  11 Chapter 2: THE ROLE OF AMP-ACTIVATED PROTEIN KINASE IN THE COORDINATION OF METABOLIC RATE SUPPRESSION IN THE COMMON GOLDFISH DURING HYPDXIA EXPOSURE^ 16 2.1. Introduction^  16 2.2. Materials and Methods^  19 Animal care^  19 Identification of AMPK subunits^  19 iii Tissue distribution of AMPK isoforms^ 21 Hypoxia exposure^ 22 Analytical procedures 24 Calculations and statistical analysis^  29 2.3. Results^ 30 AMPK isoforms^ 30 Responses to hypoxia 30 2.4. Discussion^ 33 2.5. References 61 Chapter 3: GENERAL DISCUSSION AND CONCLUSIONS^ 67 3.1. Overview^ 67 3.2. AMPK as a potential coordinator of hypoxic energy metabolism^ 67 3.3. Conclusions^ 70 3.4. Future Directions  71 3.5. References^ 72 iv LIST OF TABLES Table 2.1. Blood hemoglobin (Hb), hematocrit (Ht), mean cellular hemoglobin content (MCHC), and plasma [lactate] in goldfish exposed to normoxia (-9.5 mg 0 2/L) and during 12 h of hypoxia (-0.3 mg 02/L) exposure^ 56 Table 2.2. Liver free [creatine], [lactate], pHi, [ADPfree], [AMPfree], [ADPfree]/[ADP] and z\1G in goldfish exposed to normoxia (-9.5 mg 02/L) and during 12 h of hypoxia (-0.3 mg 02/L) exposure^ 57 Table 2.3. AMPKala, AMPKalb and AMPKI31 mRNA expression in goldfish liver in goldfish exposed to normoxia (-9.5 mg 02/L) and during 12 h of hypoxia (-0.3 mg 0 2/L) exposure.....58 Table 2.4. Muscle free [creatine], [lactate], pH., [ADPfree], [AMPfree], [ADPfree]/[ADP] and AtG in goldfish exposed to normoxia (-9.5 mg 02/L) and during 12 h of hypoxia (-0.3 mg 02/L) exposure^ 59 Table 2.5. AMPK activity in brain, gill and heart in goldfish exposed to normoxia (-9.5 mg 02/L) and at 0.5, 8 and 12 h of hypoxia (-0.3 mg 02/L) exposure^ .60 LIST OF FIGURES Figure 2.1. Distribution of AMPKala, AMPK alb and AMPKI31 mRNA in eye, heart, brain, kidney, liver, muscle, intestine and gill in goldfish. Expression of each gene is relative to the same standard sample and absolute expression is adjusted such that the tissue with the lowest expression for each gene has a expression quantity of 1 .44 Figure 2.2. Liver [ATP], [CrP] and calculated [AMP free] / [ATP] in goldfish exposed to— fre , normoxia (-9.5 mg 02/L) and during 12 h of hypoxia (-0.3 mg 0 2/L)^ ..46 Figure 2.3. Liver AMPK activity, protein and representative AMPKa western blot in normoxia (-9.5 mg 02/L) and during 12 h hypoxia (-0.3 mg 02/L) ^48 Figure 2.4. Liver phospho-eEF2, representative phosphoThr56-eEF2 and eEF2 Western blots and protein synthesis rate in goldfish exposed to normoxia (-9.5 mg 0 2/L) and during 12 h of hypoxia (-0.3 mg 02/L) 50 Figure 2.5. Muscle [ATP], [CrP] and calculated [AMPfree] / [ATP] in goldfish exposed to normoxia (-9.5 mg 02/L) and during 12 h of hypoxia (-0.3 mg 0 2/L)^ ..52 Figure 2.6. Muscle AMPK activity, AMPK protein expressed relative to total protein and representative AMPK Western blot in goldfish exposed to normoxia (-9.5 mg 02/L) and during 12 h of hypoxia (-0.3 mg 0 2/L)^ ..54 vi LIST OF ABBREVIATIONS ACC-1^acetyl-CoA carboxylase-1 ACC-2 acetyl-CoA carboxylase-2 AMPK^AMP-activated protein kinase ANOVA analysis of variance ATP^adenosine triphosphate AMP adenosine monophosphate AMPfree^free adenosine monophosphate ADP adenosine diphosphate ADPfree^free adenosine diphosphate °C degrees Celsius CaMKK^calmodulin-dependent protein kinase kinase cDNA complementary deoxyribonucleic acid Cr^free creatine CrP creatine phosphate DNA^deoxyribonucleic acid DTT dithiothreitol eEF2^eukaryotic elongation factor 2 eEF2-P phosphorylated eukaryotic elongation factor 2 EDTA^ethylene diamine tetraacetic acid EGTA ethylene glycol tetraacetic acid AiG'^effective Gibbs free energy change of ATP hydrolysis g gram GLUT-4^glucose transporter-4 GTP guanosine triphosphate h^ hour Hb hemoglobin (HMG)-CoA reductase 3-hydroxy-3-methylglutaryl-CoA reductase Ht^hematocrit 1 litre vii MCHC^mean cellular hemoglobin content min minute mol^mole MOPS 3-morpholinopropanesulfonic acid mRNA^messenger ribonucleic acid mg2+ magnesium 02^oxygen PCR polymerase chain reaction PEG^polyethylene glycol PFK phosphofructokinase PFK-2^phosphofructokinase-2 pHi intracellular pH PMSF^phenylmethanesulphonyl fluoride qPCR quantitative real-time polymerase chain reaction RNA^ribonucleic acid s second SE^standard error Ser serine TOR^target-of-rapamycin Thr threonine TTBS^Tween-20 tris-buffered saline viii ACKNOWLEDGEMENTS I would first like to extend a great deal of thanks to Dr. Jeffrey Richards. Were it not for his informative and insightful conversations and lessons, clever and helpful ideas, and unique ability to calm nerves at stressful times, this work would not have been possible. I am extremely indebted to him for these reasons and feel very privileged to have been able to complete my Master's degree under his supervision. I would also like to acknowledge and thank my fellow members of the Richards' Lab, Milica Mandic, Ben Speers-Roesch and Gigi Lau. They have been generous with their time, helping with both Sunday morning sampling sessions and assaying, and I am extremely appreciative of their efforts. Together we have had countless fun times in several Canadian cities, set a new standard for the minimum reasonable volume at which lab music should be played (to the chagrin of the lab post-doc), and had a large number of nerdy conversations. I feel very lucky to have made such great and I'm sure lasting friendships. There are also a number of people in the department who have brightened the course of my time at UBC with their friendships and I am extremely lucky to have been able to share time with them. Thanks are especially extended to Drs. Trish Schulte, Bill Milsom and Tony Farrell for their valuable suggestions concerning this project. My family has provided me with a great deal of support, encouragement and love throughout the whole of my education for which I am profoundly appreciative. Finally, I would like to thank Brandon Ash for his support and patience during the course of my thesis work. Nothing beats coming home from the lab at night to a delicious lasagna baking in the oven. ix CO-AUTHORSHIP STATEMENT Chapter 2 of this thesis is co-authored by Lindsay A. Jibb and Dr. Jeffrey G. Richards. The research program was identified and designed by J.G.R. The research and data analysis was conducted by L.A.J. under the supervision of J.G.R. The manuscript preparation was conducted by L.A.J. in consultation with J.G.R. x Chapter 1: GENERAL INTRODUCTION 1.1. Environmental Hypoxia Environmental hypoxia is common in several freshwater and marine aquatic ecosystems. These ecosystems are frequently subject to fluctuating oxygen (02) concentrations due to the relatively low capacitance of water for 02. Small lakes and ponds become ice-covered in the winter months, eliminating the diffusion of 02 from air to water and causing these water bodies to become anoxic for weeks to months at a time (Nilsson et al. 1993). Estuaries in temperate regions are very productive in the summer months, resulting in diel fluctuations in dissolved 02 and hypoxic waters at night (Cochran and Burnett 1996). These did 02 fluctuations also occur in the waters of the Amazon and in tidepools during low tide at night. Additionally, excessive anthropogenic loading of nutrients and organic matter into poorly circulated waters leads to eutrophication and has resulted in thousands of km 2 of hypoxic or anoxic marine water over the last several decades (Wu 2002). Still, nearly all aquatic environments, both those which are subject to fluctuating levels of dissolved 02 and those which are not, are inhabited by fish species. Not surprisingly, depending on the environment in which they reside, these fish vary greatly in their ability to tolerate 02 deprivation. 1.2. Responses to Hypoxia Problems associated with hypoxia The cells and tissues of hypoxia-intolerant fish are particularly sensitive to 02 deprivation. In situations of hypoxia or anoxia in these fish, the inability to produce large 1 amounts of ATP via oxidative phosphorylation sets in motion a chain of biochemical events whose end result can be necrotic cell death. Faced with the limited capacity to produce ATP via oxidative phosphorylation, hypoxia-sensitive cells react with large-scale increases in non- mitochondrial energy production, namely glycolysis (Boutilier and St-Pierre 2000). Glycolytic energy production however, is inefficient and quickly results in exhaustion of fermentable fuels followed by ATP depletion (Busk and Boutilier 2005). When the rate of production of cellular ATP can no longer meet the energetic demands of ionic and osmotic equilibrium, cell death results via membrane rupture due to uncontrolled Ca 2+ and H2O influx (Boutilier 2001, Busk and Boutillier 2005). This potentially fatal series of events can occur in hypoxia-sensitive animals within minutes to hours. Hypoxia-survival strategies of tolerant animals Those fishes tolerant of low 02 possess a diverse suite of traits allowing for an ability to avoid or postpone the above-mentioned catastrophic ATP depletion. These traits can be broadly grouped into two main classifications, (1) those traits that work to improve 02 extraction from the hypoxic environment and (2) those traits that come into play when a fish is unable to extract adequate 02 from the environment to maintain a routine metabolic rate. The second of these strategies is the subject of this thesis. Adjustments to metabolism in response to hypoxia exposure If 02 extraction and delivery capacities of a fish are insufficient to supply adequate 02 to the tissues for the maintenance of routine metabolic rate, hypoxia-tolerant fish rely on alterations of metabolic rate to extend survival time at low 02. These alterations are made in an effort to maintain cellular [ATP] thereby delaying the onset of a catastrophic energy loss that may lead to death. The ability of hypoxia-tolerant animals to maintain [ATP] in response to 2 hypoxia has been demonstrated in a variety of fish species. Perhaps the simplest way for a fish to decrease metabolic rate is to decrease locomotor activity, which has been described and quantified in the crucian carp (Carassius carassius L.). These fish display a near 50% decline in locomotion upon anoxia exposure (Nilsson et al. 1993). Fish exposed to hypoxia may also decrease metabolic rate by reducing food intake, which has been shown in a variety of fish species. Reproductive behaviours are additionally suppressed during hypoxia in an effort to decrease energy expenditure as demonstrated in the killifish, Fundulus grandis. Killifish display a delayed onset of spawning, a reduced daily egg production, and a decrease in circulating sex steroids when exposed to 30 days of hypoxia (Landry et al. 2007). An extremely important method by which hypoxia-tolerant fish may reduce their metabolic rate is through a controlled down-regulation of energy consuming processes in the cell. This down-regulation involves lowering energy expenditure via reductions in ATP- consuming processes without compromising the integrity of the cell, and may be coupled with increases in the efficiency of 02-independent energy production (Hochachka et al. 1996, Boutilier 2001, Hochachka and Somero 2002, Busk and Boutilier 2005, Bickler and Buck 2007). This coordinated down-regulation of ATP turnover has been described in isolated hepatocytes from the anoxia-tolerant painted turtle, Chrysemys picta bellii. Metabolic rate, assessed via both microcalorimetry, and ATP-turnover in these cells declined by —90% during exposure to hypoxia and this decline was completely reversible upon reoxygenation (Buck et al.1993a, Buck et al.1993b). The primary energy sinks of these cells during normoxia were found to be protein synthesis and Na+/K+ pumping, which constitute —35% and —28% of ATP demand respectively. Protein degradation, gluconeogenesis and ureagenesis also contribute to cellular ATP demand, although to a lesser degree (Hochachka et al. 1996). Upon exposure to 3 anoxia, the ATP demand by protein synthesis and Na±/K+ pumping are decreased by —90% and —75% respectively in hepatocytes (Hochachka et al. 1996). As such, the down-regulation of these processes can be interpreted to be a major contributing factor to the depression of ATP turnover at the cellular and whole animal level. The down-regulation of protein synthesis in hypoxia-tolerant fishes has been well documented both in isolated hepatocytes as well as in vivo. In crucian carp exposed to one week of anoxia, protein synthesis declines by 95% in liver, 48% in heart, 48 and 44% in red and white muscle respectively, but does not decline in brain (Smith et al. 1996). In the Amazonian cichlid exposed severe hypoxia, Astronotus ocellatus, the protein synthesis declined by 56% in liver, 60% in heart, 50% in gill and 27% in brain (Lewis et al. 2007). These studies demonstrate that the degree of down-regulation in response to 02 deprivation appears to be tissue-, species- and treatment-specific and may reflect the metabolic requirements of the tissue or species in question. In addition to the down-regulation of ATP-expenditure observed in hypoxia-tolerant organisms when exposed to low 02 , a coordinate induction of 0 2-independant ATP production is also often observed. During hypoxia exposure, ATP production shifts from that based in oxidative phosphorylation to substrate level phosphorylation, which involves a rapid hydrolysis of phosphocreatine and an activation of glycolysis to buffer declining [ATP] (van den Thillart and Smit 1984, Dalla Via et al. 1994, Virani and Rees 2000). Many studies have shown changes in expression and activity of glycolytic enzymes in response to hypoxia, which presumably alter the capacity of fish tissue for 'anaerobic' energy production (Martinez et al. 1996). In the hypoxia-tolerant goby (Gillichthys mirabilis) gene expression profiling revealed tissue-dependant changes in the regulation of several glycolytic genes (Gracey et al. 2000). 4 Expression of the glycolytic genes, lactate dehydrogenase A (LDH A), endolase (ENO), and triosephosphate isomerase was elevated in liver. In contrast, mRNA expression of ENO and glyceraldehydes-3-phosphate dehydrogenase decreased in muscle. These findings potentially reflect the differential metabolic roles of these tissues. Changes in the activity of glycolytic enzymes in response to hypoxia have also been demonstrated by several studies (Martinez et al. 1996, Kraemer and Schulte 2004) and similar to gene expression patterns, changes in enzyme activities during hypoxia are often enzyme-, tissue- and/or species-specific. Clearly, a large number of energy consuming and producing cellular processes are altered upon exposure to hypoxia and have been well documented in a variety of fishes. This ability to down-regulate the myriad of energy-consuming anabolic and energy-producing catabolic pathways is essential to maintaining balanced [ATP]. It is therefore essential that there be a well coordinated, temporally appropriate alteration of cellular processes to avoid cell damage or death. Still, the actual mechanisms responsible for this metabolic re-organization have not been studied to a great extent in hypoxia-tolerant fishes. One ideal candidate for this sort of coordination is the energy-sensitive sub-cellular protein, AMP-activated protein kinase (AMPK). 1.3. AMP-activated protein kinase AMP-activated protein kinase (AMPK) was first described as a regulator of lipid metabolism in mammalian cells nearly two decades ago (Hardie et al. 1989). Since that description, a great deal of research has centred on AMPK and its role in cellular and whole animal metabolism. AMPK is a heterotrimeric serine/threonine kinase composed of a catalytic a-subunit, and regulatory 13- and y-subunits (Hardie et al. 2006). Multiple genes code for each 5 of these subunits, and in mammals these genes give rise to two isoforms of the a-subunit (al and a2), two isoforms of the I3-subunit (01 and () and three isoforms of the 7-subunit (71, 72 and y3) (Hardie et al. 2006). The fact that multiple genes encode for each subunit combined with splice variation means that there is the potential for a large diversity in expressed AMPK protein. Obvious homologous sequence exists in the genome of all eukaryotic species tested, including the primitive protozoan Giardia lamblia suggesting the AMPK system arose early in the evolution of eukaryotes (Hardie 2003). The means by which AMPK is regulated makes the protein extremely sensitive to the energy status of the cell. An increase in AMP/ATP in the cell, indicating a cellular energy deficit, results in activation of the kinase, and conversely a decrease in the ratio prevents activation of the kinase, as an energy stress is not perceptible (Hardie 2007). The method by which AMP results in activation of AMPK is two-fold. Firstly, binding of AMP to the 7-subunit of the protein results in a conformational activating change (Suter et al. 2006). The second method through which AMP activates AMPK is through prevention of dephosphorylation. Two kinases have been identified as having phosphorylation/activation action on AMPK, LKB 1 (Sakamoto et al. 2005) and CaMKK (Witters et al. 2006). Both of these kinases phosphorylate mammalian AMPK at Thr 172 on the a-subunit, and this phosphorylation results in an increased ability of AMPK to, in turn, phosphorylate down-stream targets (Hawley et al. 2003). This phosphorylation appears to occur continuously, however under normal conditions, those where the cell is not faced with a energy deficit, this phosphate group is rapidly removed by cellular phosphatases. The binding of AMP to the 7-subunit of the protein attenuates this dephosphorylation (Suter et al. 2006). The combined effects of AMP binding to AMPK, namely this prevention of dephosphorylation and direct allosteric stimulation, can result in a 6 >1000-fold activation of AMPK (Hardie 2007). Importantly, ATP binds the same allosteric sites on AMPK as AMP although with a lower affinity (Scott et al. 2004), thus ATP acts as a competitive inhibitor of AMPK. Since ATP is capable of binding these sites, but upon binding causes none of the activating effects of AMP, high concentrations of ATP inhibit activation of AMPK by antagonizing AMP binding to AMPK (Hardie et al. 2006). This suggests that decreases in cellular [ATP] combined with increased [AMP] is a requisite for activation of AMPK. Once activated, AMPK coordinates cellular events in an effort to restore energy balance. This is accomplished via inhibition of ATP-consuming anabolic pathways and activation of ATP-producing catabolic pathways. Regarding the inhibition of ATP-consumption, AMPK was first identified as a kinase of acetyl-CoA carboxylase-1 (ACC-1; Hardie and Pan 2002) and 3- hydroxy-3-methylglutaryl (HMG)-CoA reductase (Hardie et al. 2006) and as such results in decreased fatty acid and cholesterol synthesis respectively in rat hepatocytes. AMPK activation also results in the inhibition of glycogen synthesis through inactivation of glycogen synthase (Nielsen et al. 2002) and inhibition of protein synthesis through phosphorylation of eukaryotic elongation factor 2 (eEF2; Horman et al. 2002) as well as via inhibition of the target-of- rapamycin (TOR) pathway (Kimura et al. 2003). Additionally, AMPK appears to cause G1 phase cell cycle arrest, preventing entry into the DNA replicating S phase of the cycle, another ATP costly process (Jones et al. 2005). With regard to the ATP-producing pathways, AMPK activation results in increased skeletal muscle hexokinase activity, glucose transporter GLUT-4 expression (Holmes et al. 1999) translocation of GLUT-4 to the membrane (Kurth-Kraczek et al. 1999) and increased phosphofructokinase-2 (PFK-2) activity in rat cardiomyocytes (Marsin et al. 2000). Regarding 7 mitochondrial energy production, AMPK decreases cellular malonyl-CoA through phosphorylation of ACC-2, relieving inhibition of uptake of fatty acids in the mitochondria via the carnitine shuttle and thereby stimulating oxidation of fatty acids (Kudo et al. 1995, Hardie et al. 2006). AMPK also up-regulates mitochondrial biogenesis in mammalian cells (Winder et al. 2000, Reznick and Shulman 2006) increasing the tissue capacity for aerobic energy production. Mitochondrial function, however, during 0 2-deprivation is markedly depressed in hypoxia- tolerant animals (St-Pierre et al. 2000). These mitochondria-specific actions of AMPK are likely not to be of consequence in hypoxia-tolerant animals. Still, many of the known actions of AMPK make it a very interesting candidate for coordinating hypoxia-induced metabolic suppression by delaying or preventing a catastrophic cellular ATP loss. As of yet, two studies have attempted to elucidate the potential role for AMPK during 02-deprivation in hypoxia-tolerant animals and these studies have produced conflicting results. The first of these studies showed an increase in the percent phosphorylation of AMPK, and thus activation, in the liver of the frog (Rana perezi) during severe hypoxia exposure (3.33 kPa; Bartrons et al., 2004). In contrast, a more recent study showed no alteration in AMPK activity in either the skeletal muscle or the liver of another ranoid frog (Rana sylvatica) in response to anoxia exposure (Rider et al., 2006). These contradictory results are intriguing and leave open the question as to whether or not AMPK is involved in metabolic organization during 02 deprivation. Additionally, the role of AMPK in metabolic rate suppression has not been investigated in any hypoxia-tolerant fishes. 8 1.4. Goldfish as a model organism The Carassisus sp. represent the champions of hypoxia tolerance among teleost fishes. For instance, the half-maximal survival time (LT 50) of the common goldfish (Carassius auratus) exposed to complete anoxia ranges from 22 h at 20°C (Bickler and Buck 2007) to weeks to even months at 5°C (van den Thillart et al. 1983, van Waarde et al. 1991). These animals survive low 02 conditions by employing a unique combined strategy. Firstly, goldfish possess extremely large muscle and liver glycogen stores (Nilsson 2000, Bickler and Buck 2007). This unusually large amount of glycolytic substrate extends the period of time that can be spent in hypoxia. Secondly, goldfish possess the ability to undergo a strong suppression of metabolic rate, which extends hypoxic survival time delaying the complete exhaustion of fermentable fuels. Specifically, during anoxia the goldfish heat production, declines to 30% of the normoxic level (van Waversveld et al. 1988, van Waversveld et al. 1989). Goldfish also possess a capacity producte ethanol and CO2 from lactate as 'anaerobic' metabolic end products (Shoubridge and Hochachka 1980). These are then excreted at the gill, thereby avoiding lactate accumulation. Together, these strategies help to allow for goldfish to survive 02 deprivation and make it an ideal model organism for study of the potential role of AMPK in coordinating metabolic suppression in hypoxia-tolerant fishes. 1.5. AMPK and its potential role in coordinating metabolic suppression in goldfish The maintenance of cellular [ATP] is integral to cell survival and the ability of hypoxia- tolerant organisms to accomplish this feat in 0 2-limited circumstances has been of great interest to comparative physiologists for several decades. Still, little is known about the mechanisms 9 that coordinate cellular processes and ultimately result in metabolic suppression. The objective of my thesis was to investigate the role of AMPK in coordinating metabolic suppression in the common goldfish. As AMPK is a sensitive detector of cellular energy status, I have measured or calculated the concentrations of cellular adenylates in tissues of interest in goldfish exposed to hypoxia. Knowledge of [adenylate] provides information about the presence or absence of the cellular signals for AMPK activation as well as information about metabolic energy balance during hypoxia. I have also measured the activity of AMPK in goldfish, determining whether or not activation occurs during hypoxia in these animals. Following the observation of AMPK activation in liver, I investigated the phosphorylation state of AMPK's down-stream targets. Additionally, I assessed the rate of protein synthesis, the cellular process known to be affected by AMPK phosphorylation of eEF2 (Browne et al. 2004). Protein synthesis has been well established to be down-regulated in hypoxia-tolerant animals during metabolic suppression (Land et al. 1993, Smith et al. 2006, Bickler and Buck 2007, Lewis et al. 2007) and may help to implicate AMPK in the coordination of cellular energy metabolism if shown to be co-ordinately down-regulated. I have also investigated the level of regulation of AMPK. Specifically, whether any increase in activity occurs via simple and timely covalent modification, or whether up-regulation of AMPK mRNA or protein expression occurs in hypoxic goldfish. 10 1.6. 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Response of protein synthesis to anoxia and recovery in anoxia-tolerant hepatocytes. Am JPhysiol . 265: R41-R48, 1993. Landry CA, Steele SL, Manning S, and Cheek AO. Long term hypoxia suppresses reproductive capacity in the estuarine fish, Fundulus grandis. Comp Biochem Physiol A. 148: 317-323, 2007. Lewis JM, Costa I, Val AL, Almeida-Val VM, Gamper! AK, and Driedzic WR. Responses to hypoxia and recovery: repayment of oxygen debt is not associated with compensatory protein syntheis in the Amazonian cichlid, Astronotus ocellatus. J Exp Biol. 210: 1935-1943, 2007. Marsin AS, Bertrand L, Rider MH, Deprez J, Beauloye C, Vincent MF, van den Berghe G, Carling D, and Hue L. Phosphorylation and activation of heart PFK-2 by AMPK has a role in the stimulation of glycolysis during ischaemia. Curr Biol. 10: 1247-1255, 2000. Martinez ML, Landry C, Boehm R, Manning S, Cheek AO, and Rees BB. Effects of long- term hypoxia on enzymes of carbohydrate metabolism in the Gulf killifish, Fundulus grandis. 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Deficiency of LKB 1 in skeletal muscle prevents AMPK activation and glucose uptake during contraction. EMBO J. 24: 1810-1820, 2005. 13 Scott JW, Hawley SA, Green KA, Anis M, Stewart G, Scullion GA, Norman DG, and Hardie DG. CBS domains form energy-sensing modules whose binding of adenosine ligands is disrupted by disease mutations. J Clin Invest. 113: 274-284, 2004. Shoubridge EA and Hochachka PW. Ethanol: novel end product of vertebrate anerobic metabolism. Science. 4453: 308-309, 1980. Smith RW, Houlihan DF, Nilsson GE, and Brechin JG. Tissue-specific changes in protein synthesis in vivo during anoxia in the crucian carp. Am J Physiol. 271: R897-R904, 1996. St-Pierre J, Brand MD, and Boutilier RG. Mitochondria as ATP consumers: Cellular treason in anoxia. Proc Nat Sci Acad. 97: 8670-8674, 2000. Suter M, Riek U, Turek R, Schlattner U, Wallimann T, and Neumann D. Dissecting the role of 5'-AMP for allosteric activation, and deactivation of AMP-activated protein kinase. J Biol Chem. 281: 32207-32216, 2006. van den Thillart G, van Berge Henegouwen M, and Kesbeke F. Anaerobic metabolism of goldfish, Carassius auratus (L.): ethanol and CO2 excretion rates and anoxia tolerance at 20, 10, and 5°C. Comp Biochem Physiol. 76A: 295-300, 1983. van den Thillart G and Smit H. Carbohydrate metabolism of goldfish (Carassius auratus L.) Effects of long-term hypoxia-acclimation on enzyme patterns of red muscle, white muscle and liver. J Comp Physiol B. 154: 477-486, 1984. van Waarde A, de Graaff I, van den Thillart G, and Erkelens C. Acidosis (measured by nuclear magnetic resonance) and ethanol production in anoxic goldfish acclimated to 5 and 20°C. J Exp Biol. 387-405, 1991. van Waversveld J, Addink ADF, and van den Thillart G. Simultaneous direct and indirect calorimetry on normoxic and anoxic goldfish. J Exp Biol. 142: 325-335, 1989. vanWaversveld J, Addink ADF, van den Thillart G, and Smit H. Direct calorimetry on free swimming goldfish at different oxygen levels. J Therm Anal. 33: 1019-1026, 1988. Virani NA and Rees BB. Oxygen consumption, blood lactate and inter-individual variation in the gulf killifish, Fundulus grandis, during hypoxia and recovery. Comp Biochem Physiol A. 126: 397-405, 2000. Winder WW, Holmes BF, Rubick DS, Jensen EB, Chen M, and Holloszy JO. Activation of AMP-activated protein kinase increases mitochondrial enzymes in skeletal muscle. J Appl Physiol. 88: 2219-2226, 2000. Witters LA, Kemp BE, and Means AR. Chutes and Ladders: the search for protein kinases that act on AMPK. TRENDS Biochem Sci. 31: 13-16, 2006. 14 Wu RSS. Hypoxia: from molecular responses to ecosystem responses. Mar Poll Bull. 45: 35- 45, 2002. 15 Chapter 2: THE ROLE OF AMP-ACTIVATED PROTEIN KINASE IN THE COORDINATION OF METABOLIC RATE SUPPRESSION IN THE COMMON GOLDFISH DURING HYPDXIA EXPOSURE 2.1. Introduction Environmental hypoxia is a common, naturally occurring phenomenon in many aquatic ecosystems. Further, it is increasing in prevalence due to anthropogenic nutrient loading and eutrophication of both freshwater and marine environments. One of the inherent challenges for animal life in an 02-limited environment is an inability to produce large amounts of metabolic energy per unit fuel. Despite this, many fish species can tolerate this energetic stress and are frequently found living in these waters, and as a result, these species must have evolved coping mechanisms to facilitate life during 0 2 deprivation. These mechanisms involve both those that maximize the capacity to extract 02 from the environment and those that facilitate survival when the environmental 02 tensions are below that required to maintain a routine metabolic rate. Both of these mechanisms involve adaptations that can be broadly categorized as behavioural or physiological and biochemical. Behavioural and some physiological strategies work to maximize 02 extraction from the hypoxic environment and include responses such as aquatic surface respiration to access more 0 2 rich environments (Sloman et al. 2008) or increased 1-1b- 02 affinity (Weber and Lykkeboe 1978) to increase 02 extraction from the hypoxic environment. An additional suite of modifications to behaviour and physiology come into play A version of this chapter has been submitted for publication: Jibb LA and Richards JG. AMP-activated protein kinase coordinates metabolic rate suppression in the common goldfish during hypoxia exposure. 2008. 16 when ambient 02 becomes so low that the ability of a fish to maintain a normal rate of ATP production becomes compromised. Regardless of this ATP-production limitation, hypoxia-tolerant animals defend cellular [ATP] when faced with an 02 deprivation. From a metabolic perspective this is due, at least in part, to a large capacity for energy production via substrate level phosphorylation and the ability to undergo controlled metabolic rate suppression. The latter is accomplished through the controlled reduction of ATP utilization that extends the amount of time that can be spent in hypoxia by slowing the usage of on-board fuels and the accumulation of deleterious metabolic end products. During hypoxia, large-scale changes in protein synthesis rates and ion-motive force account for the majority of observed reductions in ATP utilization (Hochachka et al. 1996). For instance, in isolated hepatocytes from the anoxia-tolerant turtle Chyrsemys picta bellii, ATP-utilization during anoxia falls to —10% of normoxic rates (Buck et al. 1993) with reductions in protein synthesis making up the largest percentage of this decline (Land et al. 1993). Overall, in order to maintain cellular [ATP] during hypoxia exposure, the hallmark measure of a hypoxia-tolerant animal, there must be an initiation of substrate-level phosphorylation and metabolic rate suppression. This requires a timely and well-synchronized restructuring of a multitude of cellular processes; however, little research has focused on how the cellular metabolic response is coordinated during hypoxia. AMP-activated protein kinase (AMPK) represents an ideal candidate protein to coordinate many of the metabolic responses to hypoxia. AMPK is a heterotrimeric protein kinase comprised of a catalytic subunit (a) and two regulatory subunits ((3 and y; Carling 2004). Phosphorylation at Thr 1 72 in the a-subunit is essential for AMPK activation (Carling 2004) and this is brought about via the activity of upstream kinases. Two of these upstream kinases have 17 been identified, LKB 1 (Sakamoto et al. 2005) and CaMKK (Witters et al. 2006) in mammals. AMPK appears to be continuously phosphorylated, however the phosphate group is rapidly removed under normal conditions, returning AMPK to an inactive form (Hardie 2007). Binding of AMP to the AMPK induces a conformational change and prevents this dephosphorylation (Sanders et al. 2007). Upon activation, AMPK inhibits anabolic processes in the cell and activates catabolic processes (Hardie et al. 2006) thereby helping to maintain cellular [ATP]. Specifically, AMPK has been shown to inhibit protein synthesis through phosphorylation of eukaryotic elongation factor 2 (eEF2; Horman et al. 2002), decrease glycogen synthesis through inactivation of glycogen synthase (Nielsen et al. 2002) and decrease fatty acid synthesis rates through phosphorylation of acetyl-CoA carboxylase-1 (ACC-1; Hardie and Pan 2002), among other actions in rats. With regard to the ATP-producing pathways, AMPK activation results in increased skeletal muscle hexokinase activity, glucose transporter GLUT-4 expression (Holmes et al. 1999) translocation of GLUT-4 to the membrane (Kurth-Kraczek et al. 1999) and increased phosphofructokinase-2 (PFK-2) activity in rat cardiomyocytes (Marsin et al. 2000). Combined, these actions have led to AMPK being termed the cellular 'energy gauge' because of it critical role in maintaining cellular energy balance. However, the cellular consequences of AMPK activation have been studied mainly in exercise- and ischaemia-stressed mammals. At the extreme of hypoxia-tolerance in teleost fishes are the Carassius sp., which are capable of surviving months of anoxia at cold temperature. An important means by which members of the genus Carassius accomplish this feat is through a strong hypoxia-dependant suppression of metabolic rate. This has been described in the common goldfish, Carassius auratus, which depresses metabolic rate by —70% during anoxic bouts, as assessed via direct calorimetry (van Waversveld et al. 1989). Metabolic suppression during hypoxia is key for the 18 survival of the goldfish as it allows for the conservation of endogenous glycogen reserves; thereby extending the amount of time that can be spent under 02-limiting circumstances. Given that AMPK is sensitive to cellular energy status, that its activation leads to a general reduction in anabolic pathways and a stimulation of catabolic pathways and that it has been shown to play a key role in metabolic suppression under ischaemic conditions, we hypothesize that it may play a role in the coordination of cellular metabolic suppression observed in the goldfish during exposure to severe hypoxia. In the present study, we determined cellular energy status, activation pattern of AMPK, and its interactions with a well characterized down-stream target, protein synthesis, in skeletal muscle and liver of normoxic and hypoxic goldfish. This was carried out in an attempt to determine whether or not AMPK may play a role in coordinating metabolic suppression during hypoxia exposure in hypoxia-tolerant organisms. 2.2. Materials and Methods Animal care Adult goldfish (Carassius auratus) weighing 36.0 + 1.4 g were purchased from a local supplier (Delta Aquatics, Richmond BC) and held in either flow-through or static renewal dechlorinated city of Vancouver tap water. Temperature during holding was maintained at 16°C and the fish were fed a daily with commercial goldfish flakes. Identification of AMPK subunits Tissue sampling Goldfish were sampled directly from a holding tank and sacrificed with an overdose of benzocaine (1 g/L). Brain, eye, heart, gill, intestine, liver, kidney and muscle were rapidly excised, flash-frozen in liquid N2, and stored at -80°C. 19 RNA extraction, reverse-transcriptase PCR amplification and fragment sequencing Total RNA was extracted from the tissue samples following the methods of Chomczynski (1993) using Tri Reagent (Sigma-Aldrich, Oakville ON). Following isolation, total RNA was quantified spectrophotometrically and integrity of the two ribosomal bands was assessed by electrophoresis using a 1.5% agarose gel. RNA was stored at -80°C. Reverse transcription reactions and PCR amplification of cDNA sequence were carried out following the methods outlined in Richards et al. (2003). Briefly, cDNA was synthesized from 4 lig total RNA with the use of RevertAid H Minus M-MuLV Reverse Transcriptase kit (Fermentas, Burlington ON) following the manufacturer's instructions. Partial AMPK subunit sequences were obtained using primers designed from the conserved regions of sequenced AMPK subunit isoforms (al, a2, (31,(32, yl, y2, and y3) using sequence information available in the GenBank, although only primers designed for al and f31 yielded successful PCR. Primers for AMPKala were: (forward) 5' — GGGCCAGCGTAAAACCTTCCT — 3' and (reverse) 5' — GGAGGGGAACTGTTTGATTATAT — 3', and PCR for this gene product consisted of 35 cycles: 1 min at 94°C, 1 min at 51°C and 2 min at 72°C. Primers for AMPKa1b were: (forward) 5' — GGAGGGGAGCTATTTGATTATAT — 3' and (reverse) 5' — GGGTTCTTCTTCGTACACG — 3' and PCR for this gene product consisted of 35 cycles: 1 min at 94°C, 1 min at 53°C and 2 min at 72°C. Primers for AMPK(31 were: (forward) 5' — GCCGGAAGGAGAGCATCAGTACAAGT — 3' and (reverse) 5' — GCGCTAAGAACCATCACGCCAT — 3' and PCR for this gene product consisted of 35 cycles: 1 min at 94°C, 1 min at 60°C and 2 min at 72°C. Primers were designed using GeneTool Lite software (www.biotool.com ). PCR products were electrophoresed on a 1.5% agarose gel to verify that a product of the appropriate size was amplified. PCR products were then ligated into 20 a plasmid vector using the pGEM-T EasyVector System II kit (Promega, Madison WI) following the manufacturer's instructions, and transformed into Escherichia coli JM109 heat- shock competent cells (Promega, Madison WI). Colonies were grown overnight at 37°C on LB- agar plates and several colonies containing the ligated insert were selected and grown in liquid culture. Following overnight growth, plasmid DNA was harvested from cultured cells using a GenElute Plasmid Miniprep kit (Sigma-Aldrich, Oakville ON) and sequenced on an Applied Biosystems PRISM 377 sequencer at the Nucleic Acid Protein Service Unit at the University of British Columbia (Vancouver BC). Tissue distribution of AMPK isoforms Tissue distribution of goldfish AMPK isoforms was estimated using quantitative real- time PCR (qPCR). Isoform-specific primers were designed using Primer Express software (Applied Biosystems Inc., Foster City CA). Primers for AMPKala (GenBank accession number EU583380) were: (forward) 5' — GCCAAGATCGCTGACTTTGG — 3' and (reverse) 5' —CGCAGCTCGTTCTCAGGAA — 3'. Primers for AMPKa1b (EU583381) were: (forward) 5' —TAAGGACGAGTTGCGGTTCTC — 3' and (reverse) 5' — GCCCTGCGTATAACCTTCCA — 3'. Primers for AMPK131 (EU580137) were: (forward) 5' — GCTGCAGGTGCTCCTCAAC — 3' and (reverse) 5' — GTTGAGCATCACATGGGTTGGT — 3'. Total RNA was extracted from brain, eye, heart, gill, intestine, liver, kidney and muscle from fish sampled directly from the holding tank and cDNA was prepared using the same methods as outlined above. Expression was quantified by real-time PCR on an ABI PRISM 7000 sequence detector (Applied Biosystems, Foster City CA). PCR reactions consisted of 2 ill cDNA, 4 pmol of each primer and Universal SYBR green master mix (Applied Biosystems, Foster City CA) in a total volume of 22 111. PCR conditions included initial incubations of 2 min at 50°C and 10 min 95°C, 21 followed by 40 cycles consisting of 15 sec at 95°C and 1 min at 60°C. Melt curve analysis was performed following each reaction to ensure that a single product was amplified. Additionally, random products were sequenced following the methods above to ensure the amplified product was indeed the product of interest. Hypoxia exposure Temperature acclimation Three weeks before experimentation, a group of —80 fish were transferred into a —375 L aquaria equipped with a canister filter and a cooling coil. Water temperature was then lowered in the tank at a rate of 2°C per day using a recirculating water-chiller until it reached 10°C, at which point temperature was maintained for at least 2 weeks prior to experimentation. Fish were fed commercial goldfish flakes daily throughout the acclimation period. Hypoxia exposure Thirty-six hours before experimentation, goldfish were transferred into individual closed chambers with small, cut-out perforations that allowed for complete aeration and water flow, and allowed fish to move freely. Following transfer into these perforated chambers fish were returned to the original experimental tank at 10°C. To obtain normoxic tissue samples, eight of these chambers were slid smoothly into basins that were slightly larger than the chambers and did not have perforations. This was done below the surface of the water in the experimental tank and allowed a chamber to be removed from the tank in water and without the fish becoming agitated. An overdose of benzocaine (1 g/L) was added to the water in the basin containing a random chamber. At complete anaesthesia, (-3 min following the addition of benzocaine) individual fish were removed, patted dry, and weighted to the nearest 0.1 g. Blood was sampled following caudal severance using Ht tubes. Following blood sampling, skeletal 22 muscle, liver, heart, brain and gill were rapidly excised, flash-frozen in liquid nitrogen and stored at -80°C. Following the sampling of normoxic fish, the water 02 concentration in the experimental tank was lowered over a 1 h period by bubbling N2-gas into the water, until it reached —0.3 mg/L. Water [02] was monitored throughout the course of hypoxia exposure using an Oakton DO 6 dissolved 02 meter (Cole Parmer, Montreal QC). Eight fish were sacrificed at each of six time points (0.5, 1, 2, 4, 8 and 12 h hypoxia exposure) in an identical manner to normoxic fish. During the experiment water temperature was maintained at 10°C. To obtain sufficient tissue for a complete biochemical analysis, the acclimation and experimental trials were performed twice. Fish from the first experiment were used for the determination of muscle intracellular pH (pH,) and metabolite concentration, and AMPK activity, protein content and mRNA expression levels. Fish from the second experiment were used for the determination of liver pH, and metabolites, haematology, plasma [lactate], eukaryotic elongation factor-2 (eEF2) and phospho Thr56 eEF2 protein expression and analysis of protein synthesis rates. Liver pH, was determined at all time points in both experiments and no significant difference was found between the two experiments (Two-way ANOVA, between experiments: df = 1, p = 0.691), therefore the two experiments gave consistent results concerning alterations in liver pH,. Additionally, during the first experiment a second, —50 L tank was maintained at an [02] of —9.8 mg/L and 10°C and eight fish were sampled from this tank in the abovementioned manner to obtain additional normoxic samples. 23 Analytical procedures Haematology Blood [Hb] was determined spectrophotometrically using Drabkin's reagent (Blaxhall and Daisley 1973). Hematocrit was determined by centrifugation of whole blood at 5 000 g for 3 min in sealed capillary tubes. Mean cellular [Hb] (MCHC) was calculated as [Hb]/Ht. Tissue processing and pH, determination Frozen muscle (-200 mg) was ground to a powder under liquid nitrogen and pH, was determinated on an aliquot following the methods of POrtner et al. (1990) using a thermostatted Radiometer BMS3 Mk2 capillary microelectrode with PHM84 pH meter (Copenhagen, Denmark). The remaining ground muscle tissue was lyophilized for 72 hr and stored above dessicant at 80°C. For pH; determination in liver, -50 mg liver was sonicated using a micro- sonicator (Kontes, Vineland NJ) at medium frequency for -3 sec in 0.2 ml ice-cold metabolic inhibitor (POrtner et al. 1990). Liver pH, was measured using an Accumet pH electrode (Cole Parmer, Montreal QC). Metabolite determination Tissue (-20 mg lyophilized skeletal muscle, -100 mg wet liver) was homogenized at maximum speed in ice-cold 8% HC1O4 for 30 s using a Polytron homogenizer. Homogenates were then centrifuged at 20 000 g for 5 min at 4°C, and the supernatant adjusted to -pH 7.6 with 3M K2CO3. Neutralized samples were centrifuged at 20 000 g for 5 min at 4°C and the supernatant was immediately frozen in liquid nitrogen, and stored at -80°C until use. These extracts were then used for the enzymatic determination of tissue [lactate], [ATP] and [CrP] (Bergmeyer, 1983). Total creatine (Cr) was determined by heating an aliquot of extract in sealed Eppendorf tubes for 20 min at 60°C and assaying for Cr enzymatically (Bergmeyer, 1983). Free 24 [Cr] was calculated on a per sample basis by subtracting [CrP] from total [Cr]. Plasma [lactate] was measured enzymatically on deproteinized plasma (20 ill 8% HC10 3 added to 20 pi of plasma). Western blotting Sample preparation, SDS-PAGE and western blotting were carried out according to the methods outlined by Todgham et al. (2005) with the following minor modifications. The amount of protein loaded into each acrylamide gel well was 20 lig. Blots for total AMPKa were blocked using Tween-20 tris-buffered saline (TTBS; 17.4 mM Tris-HCI, 2.64 mM Tris Base, 0.5 M NaC1, and 0.05% Tween-20 [v/v]) with 2% (w/v) non-fat powdered milk. Blots for eEF2 and phospho-Thr56 eEF2 were blocked using TTBS with 3% (w/v) bovine serum albumin. All membranes were incubated overnight at 4°C in a 1:1000 dilution of primary antibody (either rabbit IgG anti-AMPKa, rabbit IgG anti-eEF2, or rabbit IgG anti-phospho Thr56 eEF2; Cell- Signalling Technology, Danvers MA). Following washing in TTBS, membranes were incubated in 1:5000 IgG goat anti-rabbit (Sigma-Aldrich, Oakville ON) for 1 h. AMPK activity AMPK activity was determined following the methods described by Davies et al. (1989). Briefly, —150 mg frozen tissue (muscle or liver) was homogenized for 30 s at medium speed in approximately 3 volumes of ice-cold homogenization buffer containing 50 mM Tris-base, 250 mM mannitol, 1 mM ethylene glycol tetraacetic acid (EGTA), 1 mM ethylene diamine tetraacetic acid (EDTA), 50 mM NaF, 5 mM Na-pyrophosphate, 1 mM phenylmethanesulphonyl fluoride (PMSF), 4 µg/ml trypsin inhibitor, 1 mM benzamidine, 1 mM dithiothreitol (DTT). Samples were then centrifuged at 4°C for 20 min at 14 000 g and 360 ptl supernatant was transferred to a new micro-centrifuge tube, and 40 of 25% (w/v) 25 polyethylene glycol-6000 (PEG-6000) was added bringing the concentration in the tube to 2.5% PEG-6000. Sample tubes were then vortexed for 10 min at 4°C and subsequently centrifuged at 10 000 g for 10 min at 4°C. Following centrifugation, —320 pd supernatant was transferred to a new micro-centrifuge tube and —60 p1 of 25% PEG-6000 was added, bringing the concentration in the tube to 6% PEG-6000. Tubes were again vortexed for 10 min at 4°C and then centrifuged at 10 000 g for 10 min at 4°C. The supernatant was then removed and discarded and the pellet was washed with 300 pl of 6% PEG-6000 (prepared in homogenization buffer) before being centrifuged a final time at 10 000 g for 10 min at 4°C. Following centrifugation, the supernatant was removed and discarded and the pellet was resupended in 75 1.11 ice-cold resuspension buffer containing 50 mM Tris-base, 250 mM mannitol, 1 mM EGTA, 1 mM EDTA, 50 mM NaF, 5 mM Na-pyrophosphate, 10% w/v glycerol, 0.02% Na-azide, 1 mM PMSF, 4 mg/mL trypsin inhibitor, 1 mM benzamidine, 1 mM DTT. An aliquot of the purified resuspended protein solution was taken for determination of total protein by the Bradford protein assay (Sigma- Aldrich, Oakville ON; Bradford 1976). Aliquots of 50 pL of 1 mg/mL resuspended protein were prepared for each sample in 0.12% Triton X-100 (Sigma-Aldrich, Oakville ON) made up in resuspension buffer and immediately frozen at -80°C for no longer than 2 weeks before the activity assays were run. At the time of assay, samples were thawed on ice, and 2.5 !Al of suspension was assayed for total AMPK activity in a final volume of 25 pl, containing 40 mM HEPES, 80 mM NaCl, 8% w/v glycerol, 0.8 mM EDTA, 0.2 mM SAMS peptide with the amino acid sequence HMRSAMSGLHLVKRR (GenScript, Piscataway NJ), 0.2 mM AMP, 0.8mM DTT, 200 µM ATP, 5 mM MgC12, and [32P]-ATP (3500 cpm/pmol). Negative controls, where sample was replaced with distilled H2O, were also run for each sample. After incubation for 5 26 min at 20°C, 151A1 aliquots were spotted onto 2 cm round phosphocellualase paper (p81,Whatman) and the phosphorylation reaction immediately stopped by submergence of the spotted papers into 200 ml of 150 mM H3PO4. Spotted papers were washed 10 times for 5 min each in the same volume of fresh 150 mM H3PO 4 . Ten washes were necessary to reduce non- specific binding to near background levels. Spotted papers were then washed once in 300 ml of acetone for 5 min and air-dried. The amount of bound 32P on the papers was assessed using scintillation counting. Cell-free protein translation assay Protein synthesis rates were determined following the methods outlined by Rider et al. (2006). Briefly, frozen liver was homogenized at 1:5 (w/v) in ice-cold extraction buffer (50 mM Hepes (pH 7.4), 250 mM sucrose, 20 mM NaF, 5 mM sodium pyrophosphate, 1 mM EDTA, 1 mM EGTA) and then clarified by centrifugation at 14 000 g for 15 mM at 4°C. The resulting supernatant was removed and stored at -80°C until analysis, which was performed within 2 weeks of extraction. On the day of analysis, Sephadex G-25 columns (GE Healthcare, Piscataway NJ) were equilibrated with buffer containing 50 mM Hepes (pH 7.4), 200 mM potassium acetate, 5 mM magnesium acetate, 1 mM DTT, 5 iig/mL leupeptin, 1 mM benzamidine, and 1mM PMSF as instructed by the column manufacturer. Clarified tissue extracts were thawed on ice and a volume (0.5 ml) was gravity filtered through columns to remove endogenous amino acids. Filtrate, containing cellular proteins, was collected and analysed for total protein using the Bradford assay as above. To determine protein synthesis rates, a 50 Ill aliquot of the filtrate was added to assay buffer containing, 50 mM 3- morpholinopropanesulfonic acid (MOPS; pH 7.1), 140 mM potassium acetate, 20 mM magnesium acetate, 2 mM DTT, 20 mM creatine phosphate, 20 mM creatine kinase, 1 mM 27 ATP, 0.5 mM GTP, 0.1 mM spermidine, 10 U RNaseOUT (Invitrogen, location), 50 ug/ml total RNA prepared from goldfish liver using the Tri-Reagent method as outlined above (Sigma- Aldrich, Oakville ON), and 20 mM each amino acid (except leucine) to a final volume of 100 pl. The reaction was started with the addition of 0.9 uL of 20 uM activated leucine stock containing L-[4,5 3H]-leucine (-300 cpm/pmol) and incubated at 25°C for 90 min. Negative controls, where clarified extract was replaced with distilled H2O were assayed for each sample. Following incubation the reaction was immediately stopped with the addition of 1 ml 10% (w/v) trichloroacetic acid and placed on ice for 10 min. Samples were then centrifuged at 10 000g for 5 min to collect precipitated proteins and the pellet was resuspended in 0.2 ml of 0.1 M NaOH and re-precipitated in 1 ml of 5% (w/v) trichloroacetic acid. After 10 min on ice, proteins were collected by centrifugation and subjected to an additional wash. Following this wash, proteins were solubized in 1 ml formic acid and 0.9 ml of the solubized protein solution was taken for counting in 10 ml of toluene-based scintillant on a LS 1801 liquid scintillation counter (Beckman Coulter, Mississauga ON). AMPK gene expression The expression of cloned AMPKala, AMPKalb, and AMPK131 mRNA in liver was determined using quantitative real-time PCR as outlined above. In addition to the above- mentioned protocol, data from single tissues were normalized against (3-actin expression as an internal control. Quantitative real-time PCR primers were designed for 13-actin using NCBI sequence from goldfish (Accession No. AB039726). Primers for (3-actin were: (forward) 5' — TGACCGAGCGTGGCTACAG — 3' and (reverse) 5' — TCTCCTTGATGTCACGGACAAT — 3'. Non-reverse transcribed controls were run for random samples in order to quantify the amount of genomic DNA being amplified in the PCR. Genomic DNA contamination was 28 present in all samples, but never consistuted more than 1:1024 starting copies for AMPKala, 1:32 starting copies for AMPKalb, or 1:524 288 starting copies for AMPK(31. Genomic DNA, therefore, represents a minor contribution to the total quantitative PCR signal. A random sample was used to create a standard curve of threshold cycle versus quantity of cDNA and all data are expressed relative to this curve. Calculations and statistical analysis Free cytosolic [ADP] and [AMP] were calculated according to published protocols (Golding et al. 1995, Teague et al. 1996). The equilibrium constants for creatine kinase (1CcK) and adenylate kinase (K AK) were corrected for experimental temperature, pH, and estimated free Mg2+ (assumed to be 1 mM; Golding et al. 1995, Teague et al. 1996). The Gibbs free energy of ATP hydrolysis (A1G'; kJ/mol) was determined using the following equation A f G' A f G' ° ATP ± RT1n [ADPf„,[Pi [ATP] where R is the universal gas constant (8.3145J IC I^T is temperature in K and AiG'ATp is the standard transformed Gibbs energy of ATP hydrolysis (AJG"'ATp = -RT1nICATP) at measured pH and temperature and estimated free [Mg2+]. Cytosolic free [Pi] was estimated as the inverse of the changes in PCr assuming a resting level of 1 lAmol/g wet weight. All data are presented as means ± SE. All muscle metabolite concentrations determined on lyophilized tissue were converted back into wet weight using a 4:1 wet:dry ratio which was determined through direct measurement of goldfish muscle water content (data not shown). Statistical analysis involved one-way ANOVA followed by Holm-Sidak post-hoc test to identify where statistical difference occurred. All data were tested for normality (Kolmogorov-Smirnov test) and homogeneity of variance (Levene median test). In cases where data sets did not meet 29 these assumptions, data were In transformed, and statistical analyses repeated. For those data sets that still did not meet assumptions following transformation, statistical analysis involved Kruskal Wallis one-way ANOVA on ranks followed by Dunn's post-hoc test. Differences were considered statistically significant at p < 0.05. 2.3. Results AMPK isoforms We identified two isoforms of the gene coding for AMPKa 1 and one isoform of the gene coding for AMPK(31. Alignment of goldfish isoforms with corresponding gene sequences deposited in GenBank revealed homologies of 69-95% for the isoforms of the al-catalytic subunit and 78-84% for the (31-regulatory subunit. Each of these AMPK genes were expressed in all goldfish tissues examined and each had a tissue-specific distribution pattern (Figure 2.1). AMPKa 1 a appeared to be expressed most highly in brain, kidney and intestine (Figure 2.1 A), AMPKalb appeared to be expressed most highly in brain, kidney and gill (Figure 2.1B) and AMPK pl appeared to be expressed at relatively constant levels across tissues with the highest expression in brain. Responses to hypoxia Whole animal and blood. Goldfish remained quiescent throughout the period of hypoxia exposure and no fish deaths were observed. Any goldfish showing signs of distress or loss of equilibrium during the hypoxia exposure were removed from the treatment and placed into a well-aerated tank for recovery and not included in the data analysis. There were no significant effects of hypoxia exposure on blood [hemoglobin] or hematocrit (Table 2.1). Mean cellular hemoglobin content, however, decreased significantly compared with normoxia at 1 h hypoxia 30 exposure and remained lower than normoxic values for the 12 h hypoxia exposure (Table 2.1). Plasma [lactate] increased by approximately 6-fold over the first 0.5 h of hypoxia and continued to rise, reaching an 11-fold increase at 12 h hypoxia (Table 2.1). Liver Adenylates, CrP, pH,, lactate and effective Gibbs free energy of ATP hydrolysis. Liver [ATP] decreased by nearly half within 0.5 h of hypoxia exposure and remained at this lower concentration for the duration of the exposure (Figure 2.2A). Over the same time frame, CrP decreased by nearly one-quarter of normoxic concentrations, and was constant at this level for the remainder of the exposure (Figure 2.2B). This decrease in liver [CrP] was mirrored by stoichiometric increases in liver free [Cr], which was elevated compared with normoxia at all sampling points (Table 2.2). Lactate concentrations in liver increased by about 4-fold at 2 h hypoxia and continued to rise to a 7-fold elevation over normoxia by 12 h (Table 2.2). Intracellular pH decreased rapidly within the first 0.5 h of hypoxia exposure and remained low for the duration of the treatment (Table 2.2). Calculated [ADP fre„] was elevated over normoxia by 1 h exposure and remained elevated for up to 4 h at which point it returned to values that were not different than normoxic controls. The calculated ratio of ADPfree/ATP and calculated [AMPfree] followed similar patterns to one another with values for both increasing significantly over the first 0.5 h of hypoxia exposure and remaining elevated through to and including 12 h hypoxia (Table 2.2). The calculated ratio of liver AMP free/ATP increased rapidly following the onset of hypoxia, with concentrations 11-fold higher than normoxic controls at 0.5 h (Figure 2.2C). Calculated [AMPfree] increased continuously following 0.5 h, reaching —45-fold increase at 4 h, then declined following 4 h, but remained at concentrations higher than normoxic values (Table 2.2). The effective Gibbs free energy change of ATP hydrolysis in goldfish liver fell by 31 0.5 h hypoxia and remained significantly lower than normoxic controls for the duration of hypoxia exposure (Table 2.2). AMPK activity, protein content and mRNA expression. AMP-activated protein kinase activity in liver increased in response to hypoxia exposure, with hypoxia causing a rapid increase in activity to levels 4-fold higher than the normoxic control by 0.5 h. Activity remained elevated throughout hypoxia exposure with intermittent points of non-significance at 2 h and 12 h (Figure 2.3A). No change was observed in AMPKa protein expression in liver in response to hypoxia (Figure 2.3B). Hypoxia had no or only a slight effect on mRNA expression of AMPK ala, AMPK a 1 b, or AMPK /31 in goldfish liver (Table 2.3). eEF2 phosphorylation (Thr56) and protein synthesis. During hypoxia exposure, there was a rapid initial —2-fold increase in phospho-eEF2 by 0.5 h, which was significantly different from normoxia at all time points except 4 h and 12 h hypoxia exposure (Figure 2.4A). Total liver eEF2 did not change in response to hypoxia (data not shown). In concordance with eEF2 phosphorylation, the rate of protein synthesis decreased in liver of hypoxic goldfish. This decrease was rapid and severe, with protein synthesis rates dropping to approximately one-tenth of normoxic values by 4 h hypoxia and remaining at this level for the duration of the exposure (Figure 2.4C). Muscle Adenylates, CrP, lactate, p1-1„ and effective Gibbs free energy of ATP hydrolysis. Muscle [ATP] did not change in response to hypoxia exposure (Figure 2.5A). Hypoxia exposure caused a significant drop in muscle [CrP] by 2 h hypoxia that remained lower than normoxic values for the duration of the treatment (Figure 2.5B). As in liver, this decrease in [CrP] was mirrored by a stoichiometric increase in muscle free [Cr] (Table 2.4). Muscle 32 [lactate] increased by approximately 4-fold by 1 h hypoxia and continued to rise significantly throughout the hypoxia exposure (Table 2.4). Muscle pH, fell by 1 h hypoxia exposure and remained significantly lower than normoxic controls for the duration of treatment (Table 2.4). Calculated [ADPf„„], however, remained unaltered by hypoxia at all time points (Table 2.4). Similarly, both ADPfree/ATP and AMPfree/ATP ratios were constant compared to the normoxic control across hypoxia exposure (Table 2.4; Figure 2.5C). Calculated [AMPfree] in muscle tissue, showed no change from control values until 12 h hypoxia, where concentrations increased by nearly 5.5-fold (Table 2.4). The effective Gibbs free energy of ATP hydrolysis in goldfish skeletal muscle fell at 12 h hypoxia, but was unaltered from normoxic controls at all other time points (Table 2.4). AMPK activity and protein amount. Unlike liver, AMPK activity in muscle was not affected by hypoxia exposure (Figure 2.6A). Hypoxia exposure also did not affect total AMPK protein (Figure 2.6B). Brain, gill and heart AMPK activity. There was no effect of hypoxia exposure on AMPK activity in the brain, gills, or heart (Table 2.5). 2.4. Discussion In order to avoid necrotic cell death during hypoxia exposure, hypoxia-tolerant organisms and cells must coordinate cellular energy turnover to account for a limited capacity for 02-independent ATP generation. The present study is the first to show that AMPK activity increases in response to short-term, severe hypoxia exposure in a hypoxia-tolerant fish, the common goldfish, and that this activation is associated with a down-stream reduction in protein 33 synthesis rates, potentially mediated through the phosphorylation of eEF2. It has been speculated, but never experimentally determined (Bartrons et al. 2004, Bickler and Buck 2006, Rider et al. 2006), that AMPK may have an important role in organizing cellular metabolic suppression in tolerant animals during hypoxia exposure. Responses of AMPK to acute hypoxia exposure; however, were tissue specific, with responses observed only in the liver and not in other tissues. AMP-activated protein kinase has been termed the cellular energy gauge partly because of its acute sensitivity to [AMP]. Activation is mediated via increases in cellular [AMP free] ,e,' indicating a cellular energy deficit, and is accomplished through direct binding of AMP to the 7- subunit. This induces an activating conformational change in the protein and prevents deactivation by dephosphorylation (Suter et al. 2006). In addition, activation is greatly reduced in the absence of AMP, making its presence in elevated concentrations key to the AMPK response (Frederich and Balschi 2002, Suter et al. 2006). In the present study, there was a close temporal relationship between calculated increases in [AMPfree] and the activation of AMPK in the liver of goldfish (Table 2.2; Figure 2.3), further supporting the notion that a disruption of cellular energy charge is essential for activation of AMPK. It has been suggested that maintenance of a stable cellular [ATP] during hypoxia exposure is the hallmark measure of a hypoxia tolerant animal (Hochachka et al. 1996, Boutilier 2001). This however, may be an over simplification. In goldfish, hypoxia exposure caused liver [ATP] to decrease by nearly half during the first 0.5 h hypoxia, but following this initial drop, liver [ATP] stabilized for the duration of exposure (Figure 2.2). The fact that [ATP] is maintained in the liver at a new steady state, rather than falling to fatally low concentrations likely highlights the ability of hypoxia-tolerant animals to enter a state of metabolic rate 34 suppression. As a point of comparison, rat hepatocytes exposed to cyanide, which effectively mimics cellular hypoxia, lose 90% of intracellular [ATP] within 0.5 h of exposure and display a complete loss of viability after 2 h (Aw and Jones 1989). This decrease in [ATP] in fish liver, but not muscle, during hypoxia has been described previously in goldfish (van den Thillart et al. 1980) and sole (Solea solea; Dalla Via et al. 1994). These authors have postulated that the inherent low [CrP] in the liver results in an inability to adequately buffer [ATP] during the onset of hypoxia. This seems to be a reasonable explanation for findings in the present study, as there is a rapid drop in liver [CrP] and rise in free [Cr] in goldfish liver over the same 0.5 h period as this decline in [ATP]. (Figure 2.2; Table 2.2). The declining [ATP] results from its hydrolysis in the face of blunted oxidative phosphorylation and leads to the observed cellular accumulation of its breakdown products, ADPfree and AMPfree (Table 2.2). In accordance with this, ADPfree/ATP and AMPfree/ATP also increased significantly during the first 0.5 h, and remained elevated for the duration of the treatment (Figure 2.2; Table 2.2). The decline in [ATP] was also associated with a drop in pH„ which occurred over the same time period and was also unaltered from its original decline for the duration of hypoxia (Table 2.2). With specific regard to energy status of goldfish liver, following the initial decline by 0.5 h hypoxia, the effective Gibbs free energy of ATP hydrolysis remained unaltered for the duration of exposure (Table 2.2). The free energy of ATP hydrolysis calculated in goldfish liver following the initial decline was —54 kJ/mol, which is above values calculated as being required for the function of ATPases in rat myocardium (-49, -51, -53 and —45-50 kJ/mol for sarcolemmal Na+-K+-ATPases, Ca2+-ATPases, sarcoplasmic reticulum Ca2+-ATPases and actomyosin-ATPases respectively; Kammermeier 1987, Kammermeier 1993). Species differences in ATP free energy requirements do exist (POrtner et al. 1996), and the actual 35 requirements of goldfish ATPases are not known. Still, this would suggest that ATP free energy in goldfish liver during hypoxia exposure is maintained at a balanced level that continues to allow for the function of integral cellular processes, albeit at substantially reduced levels. Clearly, by some mechanism, the liver is capable of readjusting metabolism after a period of transition. There are a number of reasons as to why the [ATP] drop is attenuated following the initial decline. For instance, the rate of ATP hydrolysis may have slowed in goldfish liver due to decreased energy expenditure and metabolic rate depression. Secondly, an increased flux through the glycolytic pathway could also explain the attenuated [ATP] decline. Enzymes that participate in energy generating metabolic pathways can be acutely sensitive to adenylate concentrations. The glycolytic enzyme phosphofructokinase (PFK) for instance, is allosterically activated by AMP and ADP and is inhibited by ATP. The change in cellular adenylates in goldfish liver may be sufficient to activate PFK and increase flux through glycolysis. The observed activation of AMPK in the present study may play into either of these hypotheses. AMPK activation decreases rates of cellular anabolism and also upregulates PFK-2 activity in rat cardiomyocytes (Marsin et al. 2000) and increases GLUT-4 transcription and transport (Holmes et al. 1999, Kurth-Kraczek et al. 1999) to the cell membrane in rat skeletal muscle. The lack of increase in liver [lactate] until after 2 h exposure to hypoxia (Table 2.2) would tend to argue against an increased flux through glycolysis, however plasma [lactate] is increased by 0.5 h (Table 2.1) and the liver may be shuttling lactate to the muscle for conversion to ethanol. The observed decline in [ATP] may play another important role in cellular signalling in the liver as it may be a requisite for the observed AMPK activation. Several authors have reported that ATP binds the same allosteric domains as AMP on AMPK (Scott et al. 2004, 36 Hardie 2007). ATP binding to these sites causes none of the activating effects of AMP; high cellular [ATP] therefore results in competitive inhibition of AMPK (Corton et al. 1995, Hardie et al. 2006). In the present study, we show an activation of AMPK in liver, which is closely associated with both an increase in [AMP 1 and a decrease in [ATP], suggesting that both offreeJ these adenlyates may have to change in concentration in order to activate the AMPK. The likely mechanism for the observed AMPK activation in goldfish liver is post- translational modification through phosphorylation. As mentioned, AMPK is activated largely via phosphorylation of the a-subunit at Thr172 by upstream kinases and several studies have shown an association between kinase activation in a variety of situations and increased phosphorylation of this residue (Beauloye et al. 2001, Rider et al. 2006). In the present study, we screened for the phospho Thr172 form of AMPK using antibodies raised against the residue in rabbits, but found the antibodies unable to detect the phosphorylated protein in any goldfish tissue tested, suggesting a difference in amino acid sequence in this region of AMPK. Regardless of this, the large-scale changes in activation state of liver AMPK observed very early during hypoxia exposure (0.5 h) would appear to be the result of simple post-translational modification. No change in total AMPKa protein was observed during the 12 h hypoxia exposure (Figure 2.3B), therefore up-regulation of protein expression cannot explain AMPK activation. Consistent with the lack of change in the expression pattern of total AMPKa protein, no change was observed in the mRNA expression of either of the AMPKal isoforms, and only a small change was observed in AMPKI31 expression (Table 2.3). These data agree with previously published findings from human glioblastoma cells, which show no change in AMPK 37 al mRNA or protein expression in response to hypoxia and an up-regulation of both measures in the AMPK a2 isoform only after 24 h hypoxia (Neurath et al. 2006). Regardless of expression pattern, it has been well established that activation of AMPK results in decreased rates of protein synthesis in the cell. Further, it has been shown that one of the mechanisms by which AMPK affects this change is through direct phosphoryation of eukaryotic elongation factor-2 kinase (eEF2K) at Ser398, resulting in its activation (Horman et al. 2002, Browne et al. 2004). Upon phosphorylation-activation, eEF2K in turn phosphorylates eEF2 at Thr56 and modification at this site is exclusive to eEF2K (Browne and Proud 2002). This modification renders eEF2 unable to bind ribosomes and therefore inactive, and results in an ultimate decline in rates of protein synthesis. In the present study, we demonstrate significant increases in phosphorylation of eEF2 at Thr56 in hypoxic livers occurring by 0.5 h hypoxia (Figure 2.4A). This increase in phosphorylation is associated temporally with a significant decline in protein synthesis rates, which fall to —70% of normoxia by 0.5 h hypoxia exposure and continue to fall to —93% of normoxia by 4 h hypoxia exposure (Figure 2.4B). Interestingly, both the increase in phosphorylation of eEF2 and the decline in protein synthesis occur within the same timescale (0.5h hypoxia) as increases in AMPK activity in the goldfish liver (Figure 2.3), suggesting that AMPK may be important in decreasing ATP expenditure due to protein synthesis. Decreasing rates of ATP consumption is key to survival during 02 deprivation, as it prevents the catastrophic loss of fuel and energy. Protein synthesis, for its part, accounts for a 20-30% fraction of total ATP-coupled 0 2 demand (Bickler and Buck 2007) and has been shown to decline by —90% in anoxia-tolerant hepatocyte cultures (Land et al. 1993) and by 56-95% in liver of crucian carp (Carassius carassius; Smith et al. 1996) and Amazonian cichlid, 38 (Astronotus ocellatus; Lewis et al. 2007). These declines represent considerable energy savings and the association of this savings with increases in AMPK activity may implicate the protein in the cellular effort to coordinate a myriad of metabolic processes during hypoxia, leading to the regulated state of metabolic suppression in the liver of hypoxia-tolerant animals. Because it appears that AMPK activation can affect expensive cellular processes like protein synthesis in hypoxia-tolerant fish and facilitate metabolic rate suppression, it is of interest to consider other proteins or pathways, which are important in metabolic rate suppression and may be activated by AMPK. For instance, second to protein synthesis, iono- regulation is the largest energy sink in the cell, comprising -20% of ATP demand in hypoxia- tolerant hepatocytes of the western painted turtle, Chrysemys pieta belli (Hochachka et al. 1996). In goldfish hepatocytes specifically, Na+ pump activity and K+ leak pathways are down- regulated in a coordinated manner during chemical anoxia for energy conservation purposes (Krumschnabel et al. 1996). This ability of hypoxia-tolerant cells to manipulate ion regulatory processes contributes to a large degree to metabolic rate suppression, and represents an appealing target for regulation by AMPK. Interestingly, epithelial Na+ channel currents in Xenopus oocytes and mouse collecting duct cells are inhibited in an AMPK-dependant manner (Carattino et al. 2005), showing that some iono-regulatory action of AMPK is known. This again shows parallelisms between the cellular consequences of metabolic suppression in hypoxia-tolerant animals and the actions of AMPK, and makes it an appealing candidate protein for assisting in coordinating metabolic suppression. The observation that AMPK activation during hypoxia exposure is a liver-exclusive phenomenon may be a result of liver's high tissue specific metabolic rate. In normoxia, the liver has high mass specific rates of [ATP] turnover, and along with brain and heart, has been shown 39 to be one of the most metabolically active tissues in humans (Elia 1992). This high intrinsic metabolic rate of the liver means that as a tissue it is potentially a large sink for fermentable fuel use during hypoxia. Many of the actions of the liver however, for instance protein synthesis and gluconeogenesis, are known to be down-regulated during 02-deprivation in hypoxia-tolerant hepatocytes (Hochachka et al. 1996). In contrast, the brain and heart must maintain at least partial function during hypoxia exposure. This suggests that several actions of the liver are not necessarily critical to cell and whole-animal survival during the severe stress of hypoxia. As such, they do not need to be maintained at the same level as during normoxia. The normoxic mRNA expression of AMPK genes was generally higher than that of liver in other tissues tested (Figure 2.1). This however did not translate into detected differences in AMPK activation in these tissues during 0 2 -deprivation. Unlike results demonstrated in the goldfish liver, no activation of AMPK was observed in muscle, brain, heart or gill during hypoxia exposure (Figure 2.6; Table 2.5). These results are in contrast with those obtained in hypoxia-sensitive models for muscle, brain and heart (Kudo et al. 1995, Mu et al. 2001, McCullough et al. 2005), where AMPK was activated in response to hypoxia exposure. There are a number of potential explanations for this tissue-specific activation. For instance, each tissue may not be metabolically down-regulated to the same degree in the goldfish, as such, AMPK may not be activated in these tissues. This type of tissue-specific differentiation in decreased metabolic rate has been previously described in Australian desert frogs (Neobatrachus pelobatoides) during aestivation (Flanigan et al. 1991). Additionally, goldfish were exposed to severe hypoxia rather than complete anoxia and this may explain why the brain, heart and gill displayed no activation of AMPK. Upon hypoxia exposure in fish, these tissues receive increased blood flow, and therefore 02-delivery (Booth, 1979, Gamperl et al. 1995, 40 Solengas and Aldegunde, 2002) and consequently may not experience a metabolic stress of the same degree as liver. Related to this, brain and heart of both killifish (Fundulus grandis) and trout (Salmo gairdneri) showed fewer signs of metabolic stress when exposed to hypoxia than did the skeletal muscle or liver (Dunn and Hochachka 1986, Martinez et al. 2006) showing differential responses potentially owing to selective blood flow. In the case of muscle, the lack of activation of AMPK is probably related to the muscle's inherent ability to dramatically reduce energy utilization through decreased contraction during hypoxia. Indeed, in the present study goldfish were noted to minimize movement and remain quiescent in response to hypoxia. Crucian carp (Carassius carassius L.) have been shown to decrease spontaneous locomotor activity (swimming distance) by 50% in response to anoxia (Nilsson et al. 1993), which would contribute substantially to decreasing energy demands and negate the need to employ additional mechanisms to suppress metabolism. Similar reasoning has been used to explain decreases in metabolic enzyme activities in Killifish (Fundulus grandis) muscle when exposed to long-term hypoxia. Killifish showed a strong suppression of glycolytic enzyme activity in muscle contrasted by increased activity in liver, potentially reflecting the decreased energy requirements of muscle associated with decreased locomotion and showing tissue-specific effects of hypoxia depending on the metabolic demands of tissues (Martinez et al. 2006). In accordance with the lack of activation of AMPK in muscle, there was no observed increase in the cellular signal, calculated [AMP 1 ' until 12 h hypoxia (Table 2.4).free, Additionally, there was also no change in the effective Gibbs free energy of ATP hydrolysis until 12 h hypoxia (Table 2.4), suggesting that only at this point might ATP free energy become limiting. It is possible therefore that during longer duration exposure to hypoxia (> 12 h 41 hypoxia), goldfish muscle may experience an energy stress great enough to result in activation of AMPK. However, observations in this study and other studies of hypoxia-tolerant fishes show their marked ability to maintain constant muscle [ATP] in low ambient 02 conditions (Figure 2.5A; van Ginneken et al. 1995, Zhou et al. 2000, Richards et al. 2007). This may in fact impede AMPK activation since, as mentioned, ATP competes with AMP for the same activation domains but does not result in increased AMPK activity. High concentrations of ATP are therefore inhibitory to AMPK activation. Indeed, in goldfish muscle AMP /ATP ratios - f free were unaltered by hypoxia at all sampling times during exposure, as were [AD Pf ree] and ADPfree/ATP measurements (Figure 2.5C; Table 2.4). [ATP] in hypoxia-tolerant animals remains constant partly due to the buffering action of CrP hydrolysis, which declines steadily in goldfish muscle (Figure 2.5B) and is stochiometrically matched to increased Cr (Table 2.4). There is also an increase in muscle [lactate] during hypoxia (Table 2.4) that would tend to indicate an increased flux through glycolysis, however as goldfish muscle is the site of ethanol production (Shoubridge and Hochachka 1980), it is unclear whether this lactate is indicative of glycolytic flux or if it is accumulated from other tissues. AMPK has been proposed as an appealing candidate for coordination of the many cellular metabolic processes during hypoxia in tolerant organisms (Bickler and Buck 2006, Bartrons et al. 2004, Rider et al. 2006). Indeed, AMPK activity increased in liver in response to hypoxia exposure and the characteristic interactions between AMPK and the down-regulation of protein synthesis were in place and responded to hypoxia exposure. However the responses were tissues specific, and activation of AMPK was not observed in brain, gill, heart or muscle during 12 h severe hypoxia. In the present study, AMPK activation was closely associated with increased [AMPfree] and decreased [ATP], suggesting that the ratio of these adenylates may have 42 been important for activation. The decreased rates of protein synthesis, a well known component of metabolic suppression, combined with the phosphorylation of eEF2, a downstream target of AMPK, potentially implicate AMPK in the cellular effort to suppress metabolism in tolerant species exposed to hypoxia. 43 Figure 2.1. Distribution of AMPKala, AMPK alb and AMPKI31 mRNA in eye, heart, brain, kidney, liver, muscle, intestine and gill in goldfish. Expression of each gene is relative to the same standard sample and absolute expression is adjusted such that the tissue with the lowest expression for each gene has an expression quantity of 1. 44 AMPKI mRNA Expression^AMPKal b roRNA Expression AMPKala mRNA Expression "Jo a}^CS CZ I.I..^I^I. = tv, .^I^.^I^. eye_ heart - brain - kid.' - liver - in ode — intestine - gill Figure 2.2. Liver [ATP] (A), [CrP] (B) and calculated [AMP^/ [ATP] (C) in goldfish[  free] exposed to normoxia (-9.5 mg 02/L; open squares) and during 12 h of hypoxia (-0.3 mg 0 2/L; closed squares). Horizontal dashed lines through normoxia are shown as a reference. Data are means ± SE (n 5 to 8). *Significant difference from normoxia, p<0.05. 46 A 1P ATP (x NZ ATP (pawl • g-I wet tissue) CrP (lanai • g -I wet tissue) 'Jo^ZN Figure 2.3. Liver AMPK activity (A), AMPKa protein (B) and representative AMPKa western blot (C) in normoxia (-9.5 mg 02/L; open squares) and during 12 h hypoxia (-0.3 mg 02/L; closed squares). Horizontal dashed lines through normoxia are shown as a reference. All AMPKa protein samples are relative to normoxia, which was set to equal 1. Data are means ± SE (n = 5 to 11). * Significant difference from normoxia, p<0.05. 48 8N 1 2^4 Time in hypoxia (h) N 0.5 1 2 4 8 12 2.5 Te 2.0 4" o ▪ 1:14 *r1^at 1.5 d° 1:14^1.0e ei - 0.5 0.0 1.5 0.0 12 C AMPKa 49 Figure 2.4. Liver phospho-eEF2 (A), representative phosphoThr56-eEF2 and eEF2 Western blots (B) and protein synthesis rate (C) in goldfish exposed to normoxia (-9.5 mg 02/L; open squares) and during 12 h of hypoxia (-0.3 mg 02/L; closed squares). Horizontal dashed lines through normoxia are shown as a reference. All phospho-eEF2 protein samples are relative to normoxia, which was set to equal 1. Data are means ± SE (n = 5 to 8). *Significant difference from normoxia, p<0.05. 50 phospho-eEF2 at Thr56 (normalized to total eEF2) Protein Synthesis Rate (pmol leucine • mg total protein4 • 11 -1 ) (11 Figure 2.5. Muscle [ATP] (A), [CrP] (B) and calculated [AMPrree] / [ATP] (C) in goldfish exposed to normoxia (-9.5 mg 02/L; open squares) and during 12 h of hypoxia (-0.3 mg 0 2/L; closed squares). Horizontal dashed lines through normoxia are shown as a reference. Data are means + SE (n 5 to 15). *Significant difference from normoxia, p<0.05. 52 AMP free / ATP (x 104) CrP (gmol • g-1 wet tissue) ATP (limo' • g-1 wet tissue) 0 A. CT 0 S (7: IT4 A. C' 00 S Figure 2.6. Muscle AMPK activity (A), AMPKa protein expressed relative to total protein (B) and representative AMPKa Western blot (C) in goldfish exposed to normoxia (-9.5 mg 0 2/L; open squares) and during 12 h of hypoxia (-0.3 mg 0 2/L; closed squares). Horizontal dashed lines through normoxia are shown as a reference. All AMPKa protein samples are relative to normoxia, which was set to equal 1. Data are means ± SE (n = 6 to 12). *Significant difference from normoxia, p<0.05. 54 r. 4 0 3,0 4 ^AMPKa Protein^AMPK Activity (relative to total protein)^(nmol • mg protein -1 • min -1 ) CA 0 ^r- Table 2.1. Blood hemoglobin (Hb), hematocrit (Ht), mean cellular hemoglobin content (MCHC), and plasma [lactate] in goldfish exposed to normoxia (-9.5 mg 021L) and during 12 h of hypoxia (-0.3 mg 021L) exposure. Time Measure Normoxia 0.5 h 1 h 2 h 4 h 8 h 12 h Hb 1.18±0.09 1.21±0.12 1.24±0.04 1.23±0.09 1.04±0.13 1.20±0.07 1.06±0.11 Ht 30.2±1.8 35.1±2.9 39.6±3.3 38.2±3.0 35.4±2.3 36.1±1.8 31.2±3.0 MCHC 3.9±0.1 3.4±0.2 3.2±0.2* 3.2±0.1* 3.2±0.1* 3.3±0.1* 3.4±0.1* Plasma Lactate 1.23±0.12 7.71±0.94* 9.03±0.35* 10.40±0.33* 11.73±0.40* 12.05±0.44* 13.44±0.37* Data are means ± SE (n = 5 to 8). Hemoglobin (Hb) is expressed in mM; hematocrit (Ht) is expressed in %; mean cellular hemoglobin content (MCHC) is expressed as ([Hb]/Ht); plasma lactate is expressed as gmol/g wet tissue. *Significant difference from normoxia, p<0.05. Table 2.2. Liver free [creatine], [lactate], pH„ [ADPfree], [AMPfree], [ADPfree]/[ADP] and AJG' in goldfish exposed to normoxia (-9.5 mg 021L) and during 12 h of hypoxia (-0.3 mg 02/L) exposure. Time Measure Normoxia 0.5 h 1 h 2h 4h 8h 12h Cr 1.82±0.80 3.71±0.87* 4.02±0.57* 4.81±0.67* 5.39±0.84* 4.13±0.65* 5.01±0.99* Lactate 1.29±0.20 2.66±0.33 4.27±0.67 5.76±0.64* 5.76±0.21 * 6.36±0.61* 9.90±0.82* pH, 7.01±0.03 6.89±0.03* 6.90±0.01 * 6.88±0.03* 6.87±0.02* 6.86±0.02* 6.85±0.01* ADPfree 6.15±2.65 14.08±4.50 19.22±4.33* 25.69±3.60* 32.19±4.45* 17.93±4.06 17.78±4.47 AMPfree 0.02±0.01 0.20±0.08 * 0.27±0.09* 0.46±0.15* 0.78±0.19* 0.30±0.11* 0.37±0.15* ADPfree/ATP 2.18±0.81 13.52±4.20* 13.62±2.81* 20.20±4.10* 27.63±3.78* 16.61±3.18* 19.63±4.55* Ap' -63.93±1.29 -55 .72±1.11* -55.80±1.01* -53.75±0.72* -52.49±0.53* -54.01±0.78* -54.11±0•87* Data are means ± SE (n = 5 to 8). Cr, free creatine; pH,, intracellular pH; ADPfree, free ADP; AMPfree, free AMP; A1G', effective Gibbs free energy change of ATP hydrolysis. Cr and lactate are expressed in j.imolig wet tissue; ADPf ree and AMPfree are expressed in nmol/g wet tissue; Ap' is expressed in kJ/mol; ADPfree/ATP is x10 -3 . *Significant difference from normoxia p<0.05. Table 2.3. AMPKala, AMPKalb and AMPK(31 mRNA expression in goldfish liver in goldfish exposed to normoxia (-9.5 mg 02/L) and during 12 h of hypoxia (-0.3 mg 02/L) exposure. Time Gene Normoxia 0.5 h 1 h 2 h 4 h 8 h 12 h AMPKa1a 1.00±0.14 0.81±0.22 1.09±0.29 0.84±0.20 0.91±0.29 1.15±0.32 1.13±0.30 AMPKa1b 1.00±0.17 0.96±0.30 0.65±0.13 0.70±0.17 0.80±0.22 1.19±0.32 0.62±0.15 AMPK(31 1.00±0.14 1.49±0.12* 0.65±0.20* 0.59±0.09* 0.72±0.12* 0.75±0.12* 0.58±0.14* AMPKala, AMPKalb and AMPK(31 mRNA expression is relative to (3-actin expression, and all hypoxia data are relative to normoxic liver samples. *Significant difference from normoxia p<0.05. Table 2.4. Muscle free [creatine], [lactate], pH,, [ADPfree], [AMPfree]„[AD_Pfree]/[ADP] and AEG' in goldfish exposed to normoxia (-.9.5 mg 02/L) and during 12 h of hypoxia (-0.3 mg 02/L) exposure. Time Measure Normoxia 0.5 h 1 h 2h 4h 8h 12h Cr 13.39+0.92 17.50+0.91* 18.59±1.34* 18.46+1.00* 18.19+1.69* 20.01+1.64* 21.23+1.50* Lactate 1.02+0.11 3.51+0.68 4.28+0.35* 5.30+0.57* 6.96+0.73* 8.45+0.89* 10.76+0.85* pH, 7.36+0.02 7.29+0.02 7.27+0.04* 7.28+0.02* 7.23+0.04* 7.15+0.02* 7.15+0.03* ADP free 24.67+3.70 47.76+9.21 49.46+6.07 34.97+21.20 37.76+12.36 44.77+18.83 61.22+18.36 AMPfree 0.28+0.05 0.56+0.13 0.60+0.11 0.54+0.38 0.58+0.24 0.74+0.37 1.27+0.46* ADPfree/ATP 1.27+0.16 1.49+0.37 1.40+0.22 1.33+0.39 1.53+0.20 1.50+0.35 2.13+0.23 -60.54+0.38 -59.65+0.77 -59.91+0.75 -60.71+2.47 -59.01+0.77 -58.82+1.76 -56.96+0.49* Data are means ± SE (n = 5 to 15). Cr, free creatine; pH„ intracellular pH; ADPf„ e, free ADP; AMP free, free AMP; Ap', effective Gibbs free energy change of ATP hydrolysis. Cr and lactate are expressed in gmol/g wet tissue; ADPfree and AMPfree are expressed in nmol/g wet tissue; AJG' is expressed in kJ/mol; ADPfr ee/ATP is x10 -2 . *Significant difference from normoxia p<0.05. Table 2.5. AMPK activity in brain, gill and heart in goldfish exposed to normoxia (-9.5 mg 0 2/L) and at 0.5, 8 and 12 h of hypoxia (-0.3 mg 021L) exposure. Time Tissue Normoxia 0.5 h 8 h 12 h Brain 0.32+0.04 0.49+0.07 0.51+0.09 0.49+0.07 Gill 0.40+0.05 0.35+0.11 0.59+0.16 0.61+0.15 Heart 0.16+0.04 0.29+0.03 0.23+0.07 0.22+0.02 Data are means ± SE (n = 5 to 8). Values are expressed in nmol • mini • mg-1. 2.5. References Aw TY, and Jones DP. Cyanide toxicity in hepatocytes under aerobic and anaerobic. Am J Physiol Cell Physiol. 257: C435-C441, 1989. Bartrons M, Ortega E, Obach M, Calvo MN, Navarro-Sabate A, and Bartrons R. 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Respiratory adaptations in carp blood influences of hypoxia, red cell organic phosphates, divalent cations and CO2 on hemaglobin-oxygen affinity. J Comp Physiol. 128: 127-137, 1978. Witters LA, Kemp BE, and Means AR. Chutes and Ladders: the search for protein kinases that act on AMPK. TRENDS Biochem Sci. 31: 13-16, 2006. Zhou BS, Wu RSS, Randall DJ, Lam PKS, IP YK, and Chew SF. Metabolic adjustments in the common carp during prolonged hypoxia. J Fish Biol. 57: 1160-1171, 2000 66 Chapter 3: GENERAL DISCUSSION AND CONCLUSIONS 3.1. Overview The goal of the research presented in this thesis was to determine whether AMP- activated protein kinase (AMPK) was activated during hypoxia exposure in a hypoxia-tolerant animal, suggesting that the protein may play a role in coordinating the suppression of metabolism observed in these animals during hypoxia. Given that AMPK is a sensitive detector of cellular energy status and that its known actions in mammalian models are similar to cellular changes observed during metabolic suppression, it has been previously suggested that AMPK may be important in coordinating cellular processes during low 02 exposure in tolerant animals (Bartrons et al. 2004, Rider et al. 2006, Bickler and Buck 2007). This thesis represents the first work lending credence to this theory by showing that AMPK is activated in a hypoxia-tolerant organism, the common goldfish (Carassius auratus), during short-term severe hypoxia. I show however, that tissue-heterogeneity exists in the goldfish with respect to the phenomenon and AMPK activation is observed exclusively in the liver, and not in the brain, heart, gill or skeletal muscle. 3.2. AMPK as a potential coordinator of hypoxic energy metabolism Hypoxia-tolerant organisms are known to undergo a strong suppression of metabolic rate in response to hypoxia, with this suppression being key in preventing the rapid decline in cellular [ATP] that may result in necrotic cell death. This metabolic suppression has been shown directly and indirectly by several groups in both isolated cells and whole animal studies 67 (van Waversveld et al. 1988, van Waversveld et al. 1989, Buck et al. 1993a, Buck et al. 1993b). Importantly to this thesis, metabolic suppression has been shown through direct and indirect means to occur in the common goldfish in response to 02 deprivation. The metabolic suppression in goldfish is severe, with heat production falling to 30% of normoxic values during complete anoxia (van Waversveld et al. 1988, van Waversveld et al. 1989). Although the concept of metabolic suppression has received a great deal of attention, the cellular mechanisms through which hypoxia-tolerant animals reduce their metabolic rate during 02 deprivation are poorly understood. AMP-activated protein kinase has emerged from studies of exercised-stressed mammalian models as a potent detector of cellular energy status that may coordinate cellular processes in hypoxia-tolerant animals during 02 deprivation. In many hypoxia-tolerant animals including teleost fishes, large-scale decreases in energy-consuming processes, such as protein synthesis and ion-pumping, and increases in the capacity of cells to produce energy anaerobically have been widely demonstrated (van den Thillart and Smit 1984, Dalla Via et al. 1994, Smith et al. 1996,Virani and Rees 2000, Martinez et al. 2006, Lewis et al. 2007). These cellular processes mirror many of the known actions of AMPK, which has been shown to decrease synthesis rates of several macromolecules (Hardie and Pan 2002, Horman et al. 2002, Nielsen et al. 2002, Hardie et al. 2006) as well as to increase rates of 02-independent energy protection (Kurth-Kraczek et al. 1999, Marsin et al. 2000). In this thesis, I have shown that AMPK is activated in response to hypoxia exposure in the tolerant goldfish, with the liver being the sole tissue where activation was detected. The observation that activation of AMPK occurs in a tissue-specific fashion during hypoxia exposure is interesting and potentially reflects differences in the degree to which various tissues can undergo metabolic rate suppression during hypoxia exposure. Tissue-specific differences in 68 rates of metabolic suppression have been previously demonstrated in aestivating desert frogs (Flanigan et al. 1991). Should goldfish liver undergo a more dramatic metabolic suppression than skeletal muscle, heart, brain or gill during 12 h hypoxia, if AMPK has a role in coordinating metabolic rate during hypoxia, it may be expected to be activated in this tissue solely . Alternatively, preferential blood-flow to the brain, heart and gills has been previously described in teleosts (Booth 1979, Gamperl et al. 1995, Solengas and Aldegunde 2002) and may result in these tissues not undergoing a metabolic stress as great as that of the liver. With regard to skeletal muscle, several studies, including the present work, have observed decreased locomotor activity in teleosts during hypoxia exposure and this phenomenon has been directly quantified by Nilsson et al. (1993) in crucian carp (Carassius carassius). A simple depression of locomotion in goldfish muscle may therefore be great enough to prevent the requirement for a biochemical suppression of metabolism in the tissue. As AMPK is acutely sensitive to changes in cellular energy charge, I have also measured or calculated the concentrations of cellular adenlyates prior to and during hypoxia exposure in goldfish liver and muscle. The pattern of change for cellular metabolites, including adenlyates, during hypoxia is markedly different between the two tissues. The liver showed a rapid decline (>0.5 h) in intracellular pH, [creatine phosphate] and interestingly [ATP] in response to hypoxia, which results in an increase in calculated [AMP] over the same time period. The concentration of ATP in liver fell by —50% during the 0.5 h period but stabilizes thereafter. Goldfish skeletal muscle showed a more gradual decline in intracellular pH and [CrP] during hypoxia and no significant changes in [ATP], resulting in calculated [AMP] increasing only at 12 h. The hallmark of hypoxia-tolerant animals is often considered to be a distinct capacity to maintain [ATP] in 02-limited circumstances (Hochachka et al. 1996, Hochachka and Somero 69 2002). The data presented in this thesis illustrate that this is not necessarily the rule, and in liver tissue at least, [ATP] does decline during hypoxia. The stabilization of [ATP] after the initial drop shows however that by some physiological mechanism the liver is capable of re-adjusting metabolism. Interestingly, the stabilization occurs over the same time period as AMPK activity in the tissue is increased, and it is tempting to propose that AMPK may have a role in this re- balancing of cellular energy status. One of the key means by which AMPK maintains balanced cellular energy in mammals is through a down-regulation of protein synthesis (Carling 2004). This down-regulation can occur via AMPK phosphorylation of eukaryotic elongation factor-2 kinase (eEF2K). Once phosphorylated and activated eEF2K phosphorylates eEF2 at an exclusive site, Thr56, which renders eEF2 inactive and halts protein elongation (Horman et al. 2002). In this thesis, I have shown that the phosphorylated form of eEF2 increases relative to total eEF2 protein by 0.5 h hypoxia in liver and protein synthesis rates in the tissue concordantly decline by —90%. These changes also occur over the same time period as increases in liver AMPK activity are observed. This suggests that AMPK may be activated by cellular energy disturbances and react by down- regulating energetically expensive processes like protein synthesis, helping to stabilize [ATP] and preventing cell damage or death. 3.3. Conclusions Through this thesis, I have shown that AMPK is activated during hypoxia in a hypoxia- tolerant fish, with this activation being tissue-specific and observed solely in the liver. I have additionally shown that in the liver during hypoxia, a known down-stream target of AMPK, eEF2, is phosphorylated, likely helping to result in observed declines in protein synthesis rates 70 in the tissue. Decreases in protein synthesis rates may then play into the observed ability of the liver to alter metabolism and attenuate the initial decline in [ATP], preventing a catastrophic energy loss ending in necrotic cell death. Though it remains unclear whether AMPK activation is directly implicated in metabolic suppression in goldfish during 02 deprivation, my thesis work makes tempting the suggestion that AMPK is involved in a rapid re-organization of cellular processes and an impediment of energy utilization, in liver tissue at least. These two items are central to the concept of metabolic suppression. My thesis therefore offers further weight to the tenet that AMPK may have an important function in organizing cellular processes upon low 02 exposure in hypoxia-tolerant animals (Bartrons et al. 2004, Rider et al. 2006, Bickler and Buck 2007). 3.4. Future Directions The findings in my thesis have better demonstrated a potential role for AMPK in coordinating metabolic suppression in hypoxia-tolerant animals. Testing in a more direct manner the implications of AMPK as a metabolic energy coordinator will expand on the data in this thesis. The development of hypoxia-tolerant primary hepatocyte cultures would be extremely useful in this regard. Several studies have used cultured goldfish hepatocytes and demonstrated the ability of these cells to shut down ATP-consuming functions in response to environmental anoxia (Krumschnabel et al. 1994, Krumschnabel et al. 2000). The use of pharmacological agents (e.g AICAR (5-aminoimidazole-4-carboxamide riboside), an activator of AMPK) to manipulate AMPK in primary hepatocytes would be an interesting first step in more directly studying its role in metabolic suppression. 71 3.5. References Bartrons M, Ortega E, Obach M, Calvo MN, Navarro-Sabatè A, and Bartrons R. Activation of AMP-dependant protein kinase by hypoxia and hypothermia in the liver of frog Rana perezi. Cryobiol. 49: 190-194, 2004. Bickler PE, and Buck LT. Hypoxia tolerance in reptiles, amphibians, and fishes: life with variable oxygen availability. Annu Rev Physiol. 69: 145-170, 2007. Booth J. The effects of oxygen supply, epinephrine and acetylcholine on the distribution of blood flow in trout gills. JExp Biol. 83: 31-39, 1979. Buck LT, Land SC and Hochachka PW. Anoxia-tolerant hepatocytes: model system for study of reversible metabolic suppression. Am JPhysiol. 265: R49-56, 1993a. Buck LT, Hochachka PW, Schon A and Gnaiger E. Microcalorimetric measurement of reversible metabolic suppression induced by anoxia in isolated hepatocytes. Am JPhysiol. 34: R1014-1019, 1993b. Carling D. The AMP-activated protein kinase cascade—a unifying system for energy control. Trends Biochem. Sci. 29: 18-24, 2004. Dalla Via J, van den Thillart G, Cattani 0, and de Zwaan A. Influence of long-term hypoxia exposure on the energy metabolism of Solea solea. II. Intermediary metabolism in blood, liver and muscle. Mar Ecol Prog Ser. 111: 17-27, 1994. Flanigan JE, Withers PC, and Guppy M. In vitro metabolic depression of tissues from the aestivating frog Neobatrachus pelobatoides. JExp Biol. 161: 273-283, 1991. Gamperl AK, Axelsson M, Farrell AP. Effects of swimming and environmental hypoxia on coronary blood flow in rainbow trout. Am JPhysiol. 40: R1403-R1414, 1996. Hardie DG and Pan DA. Regulation of fatty acid synthesis and oxidation by the AMP- activated protein kinase. Biochem Soc Trans. 30: 1064-1070, 2002. 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