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Protein turnover and stability of the protein pool during metababolic arrest in turtle hepatocytes Land, Stephen C. 1995

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PROTEIN TURNOVER AND STABILITY OF THE PROTEIN POOL DURINGMETABOLIC ARREST IN TURTLE HEPATOCYTES.BYSTEPHEN C. LANDB.Sc. Hons. Zoology (Environmental Physiology)University of Aberdeen, Scotland, 1988.A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THEDEGREE OF DOCTOR OF PHILOSOPHYINTHE FACULTY OF GRADUATE STUDIES.(Department of Zoology).We accept this thesis as conformingto the required standard.THE UNIVERSITY OF BRITISH COLUMBIA.DECEMBER, 1994.© Stephen C. Land. 1994In presenting this thesis in partial fulfilment of the requirements for an advanceddegree at the University of British Columbia, I agree that the Library shall make itfreely available for reference and study. I further agree that permission for extensivecopying of this thesis for scholarly purposes may be granted by the head of mydepartment or by his or her representatives. It is understood that copying orpublication of this thesis for financial gain shall not be allowed without my writtenpermission.(Signature)______________________________Department of ZOOLOGYThe University of British ColumbiaVancouver, CanadaDate 15th December. 1994DE-6 (2188)UABSTRACT.Hepatocytes isolated from the western painted turtle (Chrysemys picta bellii) are capableof entering a period of metabolic suppression that is characterised by a coordinated and highlyregulated reduction in rates of ATP synthesis and ATP demand. To examine the demand sidethis relationship, the studies described here investigated the partitioning of energy usage inmetabolic suppression using protein turnover as an example of a highly regulated andenergetically expensive cell process. Absolute rates of protein synthesis fell by 92% during 12hof anoxia at 25°C. Using an empirically determined cost of 5.2 ATPs per peptide bond therelative cost of protein synthesis was determined to be 24.4mol ATP/g/h accounting for 28-36% of total ATP turnover. In anoxia, this fell to l.6mol ATP/g/h, constituting 25% of totalanoxic ATP turnover. The energy dependence of proteolysis was assessed in labile and stableprotein pools. During anoxia, labile protein half-lives increased from 24.7 to 34.4h, with stableprotein half-lives increasing from 55.6 to 109.6h. Inhibitors of energy metabolism revealed thata large proportion of whole cell proteolytic rates was ATP-independent, with the majority ofATP-dependent proteolysis appearing in the stable protein pool. Consequently, the combinedanoxic mean proteolytic suppression for both pools was 36%, but 93% of the ATP-dependentcomponent was suppressed. ATP demand for normoxic ATP-dependent proteolysis wasdetermined at 11. lmol ATP/g/h, accounting for 21.8% of total ATP- turnover. In anoxia, thiswas suppressed by 93% to O.73zmol ATP/g/h accounting for 12% of remaining ATP-turnover.Summation of anaerobic energy demand by proteolysis and protein synthesis accounts for about40% of remaining ATP-turnover in metabolic suppression.The final series of experiments tested the hypothesis that a heme-protein oxygen sensor isinvolved in modulating protein expression profiles. Cells incubated in anoxia consistently111expressed proteins of 83, 70.4, 42.5, 35.3 and 16. lkDa and suppressed proteins of 63.7, 48.2,36.9, 29.5 and 17.7kDa. Except for the 70.4 kDa protein, this expression was not found duringaerobic incubation with cyanide, used as a mimic of physiological anoxia. Incubation of cellswith factors that affect protoporphyrin conformation (Co2,Ni2, CO) produced predictablechanges in expression profiles for 42.5, 35.3, 17.7 and 16.1 kDa protein bands and incubationwith a heme-synthesis inhibitor abrogated the response. Remaining suppressed proteins inanoxia demonstrated a predictable sensitivity to Co2 and Ni2, but no effect with CO, possiblysuggesting control by a protoporphyrin with different 02 and CO binding kinetics. These resultsstrongly suggest that one or more heme-protein based oxygen sensor mechanisms are presentin turtle hepatocytes, which govern both positive and negative modulation of oxygen-sensitiveprotein profiles in anoxia.Overall, these results demonstrate coordinated reductions in ATP demand by proteinturnover, but also demonstrate that protein turnover rates constitute a large proportion ofremaining ATP-turnover in anoxic metabolic suppression. An important signal for positive andnegative changes in protein expression profiles on the anoxic transition appears to be oxygenitself, raising the suggestion that oxygen-sensing mechanisms may be important in themodulation certain cellular events during metabolic suppression.ivTABLE OF CONTENTS.Abstract iiTable of Contents. ivList of Tables viiiList of Figures ixAcknowledgements xiChapter 1: Principles and Mechanisms of Anoxia-tolerance.Introduction 1Design of Metabolism in Facultative Anaerobes 3Storage of Fermentable Substrate 4Optimising Anaerobic ATP and W Yields 4Minimising End-Product Accumulation 6Integrating Organ Systems in Metabolic Suppression. 9Signalling Mechanisms 12Mechanisms of Metabolic Suppression 16Reducing Enzyme Fluxes 17Reducing Membrane Ion Fluxes 23Anabolic Processes 29Protein Turnover and Gene Expression in Anoxia Intolerant Systems 31Kinetics of Protein Induction 31Effect of Hypoxia on Protein Turnover 35Oxygen Sensing Processes in Hypoxic Gene Expression 36Implications for Anoxia-tolerant Systems 39Consequences of Suppressing Metabolism: Remaining Viable 41Acute Mechanisms of Cell Death in Anoxia . 41Chronic Accumulation of Cell Damage 46Cell Damage on Recovery From Anoxia 47The Turtle Hepatocyte as a Model System for the Study ofAnoxia Tolerance 49Aims of the Research and Thesis Overview 51VChapter 2: Protein Synthesis During Anoxia and Recovery in Anoxia-tolerant Hepatocytesfrom the Western Painted Turtle, Chiysemys picta bellii.Preface 53Introduction 53Materials and MethodsChemicalsAnimalsSolutionsHepatocyte PreparationExperimental DesignPart A. - Rate of Protein SynthesisValidationIncorporation of Isotope intoOxygen ConsumptionExtraction and Measurement of Total Purine NucleotidesPart B. - Urea ProductionIn Vivo Labelling of Proteins During Recoveryfor Electrophoresis and AutoradiographyStatistical AnalysisResultsValidation for the Measurement of Fractional Ratesof Protein SynthesisRates of Protein SynthesisSpecific ATP Requirement for Protein SynthesisProportion of Metabolism Utilized by Protein SynthesisProportion of Metabolism Utilized by Urea ProductionPurine Nucleotide Phosphate ProfilesPatterns of Protein Synthesis During RecoveryProtein55555555565656565760616262636363646467677074Discussion 75viChapter 3: Protein Turnover During Metabolic Arrest: Role and Energy Dependence ofProteolysisPreface 80Introduction 80Materials and Methods 82Chemicals 82Primary Culture 82Measurement of Protein Degradation 83Assessment of Conditions for Measuring Protein Degradation 83Pre-labelling of Proteins 84Experimental Procedure 85Measurement of Normoxic and Anoxic Proteolytic ATP Turnover 86Data Handling and Statistics 87Results 88Culture Conditions 88Validity of the Technique 88Protein Degradation Rates in[3H]phe- and[‘4C]phe-labelled Proteins 89Effect of Anoxia and Recovery on Protein Half-life 89Proportion of ATP-dependent and ATP-independent proteolysis 94Energetic Cost of Proteolysis 96Effect of Cycloheximide, Emetine-HC1 and Protease Inhibitors 100Discussion 102Chapter 4: A Heme-Protein Based Oxygen Sensing Mechanism Controls the Expressionand Suppression of Multiple Proteins in Anoxia-tolerant Turtle Hepatocytes.Preface 108Introduction 108Materials and Methods 111Chemicals 111Culture Preparation 111Experimental Design 111viiPart A: Oxygen-responsive Protein Expression 112Part B: Heme-group Conformation Experiments 113Part C. Heat Shock Response 114Detection of Radiolabelled Proteins 114Data Presentation and Statistics 116Results 116Cell Viability 116Oxygen-sensitive Protein Expression 116Oxygen-Sensing Mechanism 123Heat-Shock Response 128Discussion 135Chapter 5: General DiscussionPartitioning of ATP Demand in Metabolically Suppressed Tissues 141Importance of Protein Turnover in Metabolic Suppression 144Implications of Oxygen Sensing in Metabolic Suppression 150Perspective 156Literature Cited 158yinLIST OF TABLES.1. Metabolic characteristics of isolated turtle hepatocytes 502. Calculation of a specific ATP requirement for proteinsynthesis from cycloheximide inhibitable oxygen consumptionand absolute rates of protein synthesis 683. Percentage of normoxic and anoxic ATP turnover utilizedby protein synthesis 694. Percentage of normoxic and anoxic ATP turnover utilizedby urea synthesis 715. Purine nucleotide phosphate profiles during normoxia, anoxiaand cycloheximide administration 726. Viability criteria for plated cells under primary cultureconditions at 25°C from 0 to 55h post-plating 907. Logarithmic rate constants, half-lives and suppression ofproteolytic activity in[3HJphe and[14C]phe-labelled proteinsas influenced by anoxia and recovery 958. Cost of protein synthesis and ATP-dependent proteolysis undernormoxic and anoxic conditions determined by stepwise inhibitionof protein turnover 999. Oxygen-sensitive and -insensitive protein expression in anoxia 120ixLIST OF FIGURES.1. Relationship between oxygen uptake and oxygen concentrationin whole liver, isolated hepatocytes and mitochondria 132. Two-compartment model of protein expression . . . 323. Pathway of acute cell damage (cell death cascade) inanoxia-intolerant tissues, and mechanism for avoidancein anoxia-tolerant tissues 424. Validation of experimental conditions used to measurefractional rates of protein synthesis 655. Response of translation to anoxia and recovery 666. Protein profile on recovery from l2h anoxia in wholeturtle hepatocyte lysates 737. Validation of experimental conditions used to measureproteolytic rates from radiolabel release 918. Isotope liberation from[3H]phe-labelled and[14C]phe-labelledprotein pools expressed as percent of total radioactivity 939. Response of proteolysis to various metabolic inhibitors 9710. Rates of lactate production under anoxia, anoxia+cycloheximideand anoxia+cycloheximide+emetine 9811. Effect of various protease inhibitors and protein synthesisinhibitors on[3H]phe- and[14C]phe-labelled protein half-lives 10112. Representative autoradiograph demonstrating profile of proteinexpression and suppression in various states of anoxia 118x13. Oxygen sensitivity of protein expression during physiological anoxiaand environmental anoxia 12114. Response of oxygen-sensitive proteins to normoxia, cobalt,nickel and cobalt+dioxoheptanoic acid treatment 12415. Changes in protein expression for oxygen sensitive protein bandsduring treatment with cobalt, nickel and cobalt+dioxoheptanoic acid 12616. Response of oxygen and cobalt-sensitive protein expression totreatment with 10% carbon monoxide+anoxia 12917. Changes in oxygen and cobalt-sensitive protein expression as afunction of carbon monoxide treatment 13118. Response of turtle hepatocyte protein expression to heat-shockat 40°C for lh 13319. Partitioning of energy demand from normoxia to anoxia inturtle hepatocytes 14220. Control of anaerobic processes appears independent ofmitochondrial oxygen consumption and ATP concentrations 15121. The oxygen-sensing potential of the cGMP signalling pathway:feeding two messages into one second messenger system 155xiACKNOWLEDGEMENTS.The research for this thesis would not have been possible, or nearly as much fun, withoutthe input and interaction of a number of people. Special thanks go to my supervisor, PeterHochachka, whose overall philosophy in the gentle art of raising grad students, boundlessenergy and enthusiasm, and once vast financial resources were a central driving force behindthis work. Special thanks are also due to Drs. Les Buck and Tom Mommsen who helped in myearly attempts in isolating and experimenting with turtle hepatocytes. The experiments inChapter 4 were stimulated from a meeting held in Oct. 1993 at Woods Hole, MA (Oxygen asa Regulator of Cell Function) and I wish to thank the organisers for providing partial financialsupport and an excellent forum for interaction. Thanks also to Drs Jamie Piret and Chorng Hwafor instruction and use of a laser scanning densitometer.The Hochachka lab has always had a very open approach to discussion and collaborationand my time here was no exception. Interaction with Drs. Peter Arthur, Les Buck, GaryBurness, Chris Doll, Grant Mclelland, Mark Mossey, Petra Mottishaw, Chris Moyes, TrishSchulte, Jim Staples, Carole Stanley, Raul Suarez, Sheila Thornton and Tim West was moststimulating and something that I’m sure I’ll keep with me for a long time to come. I am gratefulfor excuses to get out of the lab, however, provided by Dr. Tom Carefoot, Barb Taylor andDeborah Donovan who ensured regular dive thps to Bamfield and an endless supply of sushi.Thanks also to Les, Chris and Petra for being reliable dive-buddies in the ‘tween-time, to Sheilafor raising Mytilus edulis to new and uncharted culinary heights and to Tim for ensuring therewas always a home, just, to come back to. Personal TLC and extensive financial aid came inthe form of Sandy Connors who is indirectly responsible for a large piece of this work. Irecommend anyone attempting a PhD to get a Sandy of their own.Formal financial aid was provided through a Canadian Commonwealth Scholarshipadministered by the Association of Canadian Colleges and Universities whom I gratefullyacknowledge for their co-operation, generosity and for providing me with an opportunity toview a new continent.1Chapter 1.Principles and Mechanisms of Anoxia-tolerance.Introduction.Living tissues have four sustaining requirements: energy (ATP), reducing power (NADH),a selectively permeable membrane, and precursors for biosynthesis. At the most basic level, theinter-relationships between these components defmes both the metabolic and physicalcharacteristics of tissues and also, the ability of tissues to adapt to severe changes in the externalenvironment (Atkinson, 1977).Oxygen acts as the terminal electron acceptor in the aerobic production of ATP and NADHin a reaction that proceeds from glucose as follows:1ONADH+1OH+2FADH+C6H120+ 602 + 38(36)ADP + 38(36)P1- 6CO2 + 6H20 + 38(36)ATP1ONAD+2FAD ++12H0The ratio of one mole of glucose to six moles of 02 and CO2 is fixed, as are the number ofreducing equivalents generated per mole of 02. The potentially variable factor within theseequations is the yield of ATP which is determined by evolution rather than chemical constraints(Atkinson, 1977). Nevertheless, in most biological systems, about 40% of the free energy2yielded from complete glucose oxidation remains conserved in the form of 38 ATPs.When oxygen is removed, the metabolism of glucose is entirely cytosolic and proceeds viaa fermentation pathway where there is no net change in oxidation state (cytosolic redox isbalanced). Lactate becomes the terminal electron acceptor in this pathway and the net reactionproceeds as:C6H120+ 2ADP + 2Pi -, 2lactate + 2ATP + 2H0The energy yield of this anaerobic pathway is drastically lower than when it is combined withoxidative phosphorylation, above. Complete aerobic oxidation of glucose has a free energychange (G°) of -686 kcal/mol. Lactate demonstrates a AG° change of -319.5 kcal/mol,therefore the net energy yield from anaerobic fermentation of glucose to 2 lactates is only -47kcal/mol. Assuming that ATP has a G° of -7.3 kcal/mol, then anaerobic glycolysis transfersa mere 2.1% of the total free energy within a glucose molecule towards ATP, with 31% of thethe overall energy yielded from lactate production conserved as ATP. Fermentation fromglycogen enhances the conservation of energy yield to 47% by generating a further net gain ofone more ATP per glucosyl unit liberated and subsequently fermented.This overview of aerobic versus anaerobic energy production efficiencies serves to illustratea basic, but important point: because of the inefficiency of energy yield from anaerobicpathways, the limits to life without oxygen are defined by the way in which the organismpailitions energy supply against energy demand. The research presented in this thesisexamines this relationship by taking protein turnover as an example of a compartmented, highlyregulated and energetically expensive process, in an organism that displays a specialised abilityto withstand long periods of anoxia, the western painted turtle (Chrysemys picta bellii). Thisspecies deals with the loss of ambient oxygen by entering a period of metabolic suppression3which involves a large-scale change in the way that energy is produced, partitioned amongstATP-demanding processes, and subsequently utilised. To closely examine the mechanismsbehind metabolic suppression, a turtle hepatocyte preparation has been developed which servesas a useful model system for studies aimed towards understanding the sub-cellular characteristicsof anoxia-tolerance (Buck et a!., 1993b). The series of studies presented here utilise thispreparation to examine the re-organisation, and relative importance of cellular processes, andin particular protein turnover, in the anoxic metabolic response. From this perspective,questions are broached as to how individual ATP demanding processes respond during thetransition to anoxia, at what relative advantage to enhancing anoxic survival, and at whatpotential cost to cell function over long bouts of anoxia.To establish the physiological and biochemical framework, the following literature reviewdiscusses the principles and mechanisms of anoxia tolerance, and in particular, metabolicsuppression from the perspective of an organ-to-cell response. The aim is to demonstrate howan elegantly integrated series of innate adaptations, and changes in cellular organisation,combine to conserve all four of the essential components to sustaining life, but in the completeabsence of oxygen.Design of Metabolism in Facultative Anaerobes.There are four conserved features of metabolic design in anoxia tolerant organisms that areadvantageous to sustaining life without oxygen: 1) tissues possess large stores of fermentablesubstrate 2) the yield of ATP per mole of substrate is maximised, 3) the problems of metabolicend-product accumulation are circumvented and 4) organ systems require to integrate the anoxicmetabolic response to sustain the specific needs of each tissue.4Storage ofFermentable Substrate. Glycogen hydrolysis provides a source of glucosyl unitsfor fermentation through glycolysis. In vertebrate anaerobes, the principal storage site forglycogen is the liver which plays a central role in the provision of substrate for other tissuesduring periods of anoxia. In turtles, glycogen comprises 15% of the liver mass at aconcentration of 86Omol glucoselg (high millimolar concentrations; Clark & Miller, 1973;Hochachka, 1982); in goldfish, this reaches as much as 13OOmol glucose/g (molarconcentrations; Van den Thillart, 1982). This is in stark contrast to anoxia-intolerant speciessuch as trout and rat where liver glycogen concentration is 235 and 21Omol glucose/grespectively (Hochachka, 1982).Optimising Anaerobic ATP and H Yields. The fermentation of glycogen to 2 lactatesgenerates a net production of 3 ATP. However, the ATP yield of this pathways can besubstantially improved by coupling glycolytic fermentation with other anaerobic ATP generatingpathways. In parasitic helminths, fermentation of glucose to succinate increases the yield ofATP to 4 ATP per mole of glucose; further reduction to propionate increases the yield of thispathway to 6 ATP per mole of glucose. Stoichiometric coupling of this pathway to the reductionof aspartate to succinate, and leucine to isovalerate, generates a further 1 ATP each andenhances the classical glycolytic yield from glucose by over four fold (Hochachka, 1980).Bivalves also appear to possess the necessary metabolic machinery for this pathway but itappears to be principally utilised in the early stages of anoxia when aspartate concentrations arehigh (De Zwaan, 1983).Turtles do not appear to utilise alternative fermentation pathways to any significant extent.In turtle hepatocytes over lOh of anoxia, lactate accumulates to 50mM and together with glucosemobilisation, accounts for almost all of the carbon loss from glycogen (Buck et al., 1993b).5However, the 1G° of lactate production and glucose release accounted for only 36% of anoxicheat flux measured by microcalorimetry re-issuing the question of whether other fermentativepathways might be active in anoxia (Buck et al., 1993a). Subsequent measurement of succinaterevealed an accumulation to only 2% of lactate concentrations indicating that the fumaratereductase pathway does not account for a significant proportion of the exothermic gap and isnot an important alternative fermentative pathways in turtle liver. Microcalorimetry of turtlebrain cortical slices also points to the lack of alternative fermentation pathways in this tissue asthe exothermic gap was negligible during anoxia (Doll et a!., 1994).Optimising the number of ATP produced per proton released helps to slow the rate oftissue acidification. The source of H in anoxia is believed to result from the mismatch ofprotons consumed in fermentation versus protons produced by ATP hydrolysis (Hochachka andMommsen, 1983). The stoichiometry of this relationship (glycogen fermentation and ATPhydrolysis) is always the same regardless of the fermentative pathway, favouring the netproduction of 2H per mole of glucosyl unit used and appears to be inviolable for numerousanaerobic systems (Hochachka and Mommsen, 1983).The ratio of ATP synthesised per net W produced in fermentation can be optimised,however. The fermentation of glucose to lactate generates a ratio of 1:1 for ATP synthesisedto H produced. If glycogen is the principal substrate, this ratio is improved to 1.5:1 and in thepathway from glucose to succinate and proprionate, the ATP:H ratio increases to 2 and 3respectively, per mole of glucosyl unit utilised (Hochachka and Somero, 1984).Therefore, in situations where ATP demand is maintained during anaerobiosis, theutilisation of alternative fermentation pathways presents two principal advantages: 1) anincreased yield of ATP per mole of substrate and 2) an increase in the yield of ATP per mole6of end-product (Hj produced. Since the principal fermentative substrate in turtle tissues isglycogen, both ATP yield and overall ATP:H are improved 33% over glucose fermentationto 2 lactates. However, with increasing time in anoxia, circulating glucose becomes theprinciple substrate for tissues such as heart and brain suggesting that both ATP yield andATP:H likely demonstrates a tissue specific decrease.Minimising End-Product Accumulation. Despite potential improvements in the ratio ofH production to ATP turnover in anaerobic pathways, the overall stoichiometry of glycolyticW utilisation to ATP hydrolysis and H production still favours a net accumulation of protons(Hochachka and Mommsen, 1983). Therefore, in anoxia the accumulation of protons andlactate as metabolic end-products presents a problem since pathways allowing their completeoxidation, re-utilisation or excretion are02-dependent processes. Reducing potentially harmfulaccumulations of toxic metabolic end-products can be achieved by 1) metabolising the end-product into an excretable form, 2) increasing tissue tolerances to accumulations of the end-product by improving tissue buffering capacity and 3) reducing the rate at which the end-product is formed by suppressing metabolism.Vertebrate anaerobes demonstrate all three mechanisms. Decarboxylation of lactate toethanol and CO2 (mechanism 1, above) has been characterised in goldfish and crucian carp(Carassius spp; Shoubridge & Hochachka, 1980; Johnston & Bernhardt, 1983). In goldfish,whole body + external lactate changes marginally from 0.2 to 6mol/g over a 12h anoxicepisode (Shoubridge & Hochachka, 1980). The apparent rate of lactate production accounts foronly 17% of the metabolic needs of the animal and is low despite high activities of LDH withinvarious tissues (Van den Thillart et a!., 1976). This apparent discrepancy is founded in theability of these species to decarboxylate lactate to ethanol + CO2 by alcohol dehydrogenase and7then excrete both products to the environment via the gills (Shoubridge & Hochachka, 1980).The advantage of this is clear: lactate accumulations are minimised over long periods of time,and the W production associated with ATP hydrolysis is balanced with W utilisation infermentation resulting in no net yield of protons (Hochachka and Mommsen, 1983). Thismechanism is so effective in goldfish that, despite possessing weak plasma buffering capacity,pH1 remains above neutrality for at least 3h of anoxia (Van den Thillart & Van Waarde, 1993).Turtles deal with the problem of W and lactate end-product accumulation by combiningeffective mechanisms for buffering H with metabolic suppression, exhibiting mechanisms 2)and 3) above. In anoxic dives lasting up to 6 months at 3°C, plasma lactate rises to 200mMindicating a significant metabolic acidosis, yet plasma pH falls gradually over this time frompH 7.9 to a range between pH 7.6-7.2 (Herbert & Jackson, 1985; Jackson & Heisler, 1983;Ultsch & Jackson, 1982). Despite possessing exceptionally high intrinsic concentrations ofplasma HCO3 (4Omequiv/l), the non-bicarbonate buffering capacity of turtle blood is nodifferent from anoxia-intolerant species and as such, the disappearance of HCO3 only accountsfor about 20% of the lactate load (Jackson & Ultsch, 1982). The shortfall in plasma bufferingcapacity is compensated by an increase in the concentrations of K, Ca2 and Mg2, which,together with decreasing anion concentrations (HCO3 and Ci), serve to balance the largeanionic lactate load (Jackson & Ultsch, 1982). On the cation side, the most remarkable changesoccur in plasma Ca2 and Mg2 concentrations which rise to 68 and 26mM respectively.Together, these compensate for more than 50% of the lactate charge imbalance (Jackson &Heisler, 1982). The source of these cations appears to be the shell and long-bones which arerich in Ca(HCO3)2and Mg(HCO3)2(Jackson, 1993). With bicarbonate as the accompanyinganion, mobilisation of Ca2 and Mg2 also serves to buffer W associated with lactate8production.Further enhancing tolerance to metabolic end-product accumulation, especially on recovery,is the ability to store toxic metabolic end-products in low tissue concentrations. Despite a largeplasma lactate accumulation in turtles, skeletal muscle and liver lactate only accumulates toabout 35% of that in the extracellular fluid (Jackson & Heisler, 1983). Turtles therefore appearto be capable of “storing” tissue derived lactate at high concentrations in blood plasma andminimise excessive intra-tissue accumulations. In addition, turtle bladder epithelium possessesan WATPase activity which operates to alkalinise or acidify the urine (Youmans & Barry,1989; Fritsche et a!., 1991). Whether this W sequestering activity is active during anoxia hasnot been determined but since the bladder can occupy as much as one third of the body cavityvolume when full, it represents a potentially important site for sequestering H.By far the most significant mechanism to sustaining life without oxygen is the ability tosuppress the rate of ATP demand and, in so doing, enter a period of metabolic suppression.This has the dual effect of slowing both the rate at which fermentable substrate is utilised andat which metabolic end-products accumulate. When metabolic suppression is combined withmetabolic design features that optimise ATP yield, minimise H production, tolerate endproduct accumulation, all in the presence of large stores of fermentable substrate, anoxicsurvival time can be extended many fold.Amongst vertebrates, the ability to suppress metabolism by various degrees is widespreadamongst ectothermic and endothermic species in extreme environments (reviewed by Hochachka& Guppy, 1987). However the most profound suppression in metabolic rate is found in variousspecies of freshwater turtle. In the red-eared slider (Pseudemys scripta elegans) heat outputmeasured by direct calorimetry falls 85% during an anoxic dive at 24°C (Jackson, 1968). In9the western painted turtle (Chrysemys picta bellii), biochemical measures of anoxic metabolicdepression demonstrate a significant temperature dependent component, ranging from 84% ofnormoxic metabolic rate at 15°C (Q10=3.7), to 91% at 3°C (Q10=9.7). Over a temperaturerange of 20-3°C, the combined effects of anoxia and increasing Q10 result in a 99.5% reductionin metabolic rate from normoxic metabolism at 20°C (Herbert & Jackson, 1985). In the northernrange of these species (lat 54°) where winter lasts from 4-6 months, turtles hibernate byburrowing into anoxic mud at the bottom of ice-frozen ponds, where ambient temperatures areconstant at 3-4°C (Ultsch, 1989). Therefore in the wild, the combined effects of metabolicsuppression and Q10 extend survival time several fold beyond what could be achieved throughincreased tolerance to end-product accumulation and sequestration of large stores of fermentablesubstrate alone. To this end, Hochachka (1986) has calculated that dealing with metabolicacidosis, optimising ATP yield and possessing large glycogen stores could account for a 3-4fold increase in survival time. However, by combining these factors with metabolicsuppression, survival time in turtles is extended by 60 fold.Integrating Organ Systems in Metabolic Suppression. The final requirement for survivalas an anaerobe, which is particularly relevant during metabolic suppression, is to establish anoptimal arrangement of remaining metabolism between organ systems. In whole turtle forcedsubmergence studies, the general tissue response on entry into anoxia appears to involve a briefactivation of glycolysis (Pasteur effect) followed by a decrease in glycolytic rate as on-boardoxygen supplies become completely exhausted (Clark & Miller, 1973; Kelly & Storey, 1988).The reversal of the Pasteur effect, coincident with tissue anoxia, marks the entry into truemetabolic suppression.Entry into anoxia requires a significant metabolic readjustment within organ systems which10favours reduced, but sustained tissue function during metabolic suppression. In turtles at 3°C,heart rate falls 80% from 1.8 to 0.4 beats/mm, contractility of atria and ventricles falls by 50%and there is a significant decrease in cardiac output (Wasser et a!., 1990; Jackson, 1987).There is also a redistribution of blood flow which favours perfusion of central organs, such asheart and especially brain (3.5 fold increase over normoxic controls) over splanchnic organs(liver, kidney, gut) and skeletal muscle (Davies, 1989). Since arterial pressure is maintained,the basis for this re-distribution of blood supply seems to be a local increase in vascularresistance. Potentially of great importance in this regard is the role that endothelial cells playin the release of autocoids (affect local vascular tone) and factors that alter the capillaryproliferation. The release of these substances is regulated by changes in local oxygenconcentrations through a number of “oxygen sensing” mechanisms (discussed later). This haslead to the conceptual development of the vascular system as a systemic oxygen sensor (Pohi,1990).Brain activity is also reduced: EEG and evoked potential activity fall by 50-80% (Feng eta!., 1988; Pérez-Pinzón et al., 1992), action potential thresholds increase for Ca2 and Na andpostsynaptic transmission is depressed (Pérez-Pinzón et a!., 1992). The overall effect is toreduce, but not inhibit, the ATP demanding activity of the brain, a phenomenon that has beentermed spike arrest (Sick et a!., 1993). Skeletal and smooth muscle also become quiescent.Entry into anoxia is associated with the cessation of movement (Ultsch, 1989) and althoughthere have been no studies of gut motility in turtles during metabolic arrest, the accompanyingstarvation is probably associated with a reduction in gut motility and possibly gut length (notedin other reptiles- Secor & Diamond, 1994 & pers. comm.).The systemic response to anoxia is largely based on fuel economy and during metabolic11suppression, the liver plays a central role ensuring a supply of plasma glucose that meetsdemand. In turtles, entry into anoxia is associated with a 50 and 60 fold increase in the plasmaconcentrations of adrenaline (Ad) and noradrenaline (NA) respectively. Subsequent activationof hepatic fl-adrenergic receptors leads to an increase in total glycogen phosphorylase activityand the percentage in the active “a” form. The resulting mobilisation of glucose from glycogenincreases plasma glucose 6 fold over controls in a 4h period of anoxia (Keiver & Hochachka,1991). Corticosterone levels remain low during anoxia but increase dramatically on reoxygenation and are suggested to pay a role in enhancing glycogen deposition and lactatemetabolism after a bout of anoxia (Keiver et a!., 1992).Organ-wide metabolic suppression, mobilisation of hepatic glucose and a redistribution insystemic blood flow are important factors in glucose sparing for fermentation by vital organssuch as brain and heart and probably aids in sustaining glycogen concentrations within thesetissues over the anoxic period. The importance of liver glucose mobilisation to sustained heartfunction has been demonstrated in studies which show that depleted cardiac glycogen suppliesdo not affect heart function so long as liver glycogen stores are present (Daw et a!., 1967).Therefore, vital organs possess a 3-way back-up system: metabolic suppression (80% in brain),large intrinsic stores of glycogen, and liver-derived plasma glucose. Glucose delivery is assuredby relative increases in the perfusion of these organs which also serves to remove metabolicend-products.A final and remarkable feature of anoxic metabolism that is demonstrated by all tissues,is the maintenance of ATP concentrations and cellular energy charge on entry into, and duringanoxic metabolic suppression (Kelly & Storey, 1988). As discussed later, the maintenance ofATP concentrations are central to sustaining cell function and avoiding acute and lethal forms12of cell damage. To achieve this, the metabolic changes that occur during the transition toanoxia and metabolic suppression must be balanced and highly coordinated by a singlesignalling event. The next section discusses signalling mechanisms that may be at the root oftransducing a change in environmental 02 to a cellular response.Signaffing Mechanisms.The signal that links changes in 02 concentrations to events in metabolic suppression mustact through a mechanism that can achieve a global and synchronous change in ATP demand andATP synthesis, and that can also coordinate this change between organ systems. Hormones canbe dismissed as the primary signal in this response since anoxic metabolic suppression has beendemonstrated in numerous isolated systems without the input of exogenous effectors. AlthoughpH is a potent modulator of cellular metabolism in invertebrate systems (Artemia embryos forexample), in vertebrates, it can also be dismissed as the primary signalling mechanism sinceintracellular pH (pH1) falls on a different time-scale to metabolic changes. Metabolicsuppression is also associated with little alteration in [ATP], [ADP] or [AMP] and changes inmetabolite concentrations are not sufficient to cause a 90% drop in enzyme flux along apathway (cf-Kelly & Storey, 1988). What then, relays changes in environmental 02 to invokea systemic and controlled metabolic suppression?Figure 1 demonstrates the change in oxygen uptake versus oxygen concentration in theliver. The iç of this curve is about 170MM 02 and is more than 170 times the K. for 02 atcytochrome c oxidase in liver tissue (<1PM02;Longmuir, 1957). Inset is the relationshipbetween oxygen concentration and oxygen uptake in isolated hepatocytes and isolatedmitochondria (Yoshihara et a!., 1988). Most significant is the observation that in intact13150 Whole Liver100- Hepatocytes-Mitochondria50 j’ 2*” h4”<40-204-0 0o 400 800 —Oxygen Concentration (FM)0200 400 600Mean Ilepatic 02 Concentration(1LM)FIGURE 1. Relationship between oxygen uptake and oxygen concentration inwhole liver, isolated hepatocytes and isolated mitochondria. Figure is adaptedfrom Yoshihara et al., (1988). Abbreviation: P - protein.14hepatocytes, V02 changes by more than 50% over an oxygen concentration range where isolatedmitochondrial V02 remains constant. Assuming that the gradient of oxygen from the exteriorof the cell to the mitochondrion is small (about 6PM; Jones et a!., 1990) this suggests thathypoxia-associated changes in metabolism occur whilst 02 is still saturating at themitochondrion. In isolated, contracting dog gracilis preparations, evidence presented suggeststhat during hypoxia, 02 modulates ATP supply and demand in such a way that ATP synthesisand ATP demand remain balanced and coordinated with one-another (Hogan et a!., 1992;Arthur et al., 1992). ATP turnover rates also show a direct, linear correlation with 02 supplywhen 02 is still well above limiting concentrations at the mitochondrion (reviewed byHochachka, 1994). Evidence such as this strongly suggests that 02 itself acts as the globalsignalling mechanism on entry into anoxia.There are four principal mechanisms for cellular02-sensing that have been presented inrecent years. These include 1) a nitric oxide (NO) model where hypoxia-induced increases in[Ca2]1 activate NO-synthase with a subsequent increase in cGMP and protein-levelphosphorylation (King et a!., 1993), 2) an H20 generating NAD(P)H oxidase model wherechanges in the redox state of glutathione, or H2O effects on guanylate cyclase activation leadto changes in channel efflux and gene expression (Acker et a!., 1994), 3) a high km oxygenasemodel where the activity of a rate limiting oxygenase is modulated through its high sensitivityto [02] (Thurman et a!., 1993), and 4) a heme protein oxygen receptor model (discussed later)where changes in the02-dependent conformation of a membrane bound heme-protein transducechanges in extracellular 02 to a cellular response through a second messenger pathway(Goldberg et a!., 1988). The exact nature of the heme-protein receptor is currently vague butan interesting consideration is the potential for its association with other cellular signalling15mechanisms. For instance the cytosolic form of guanylate cyclase, the enzyme responsible forcGMP synthesis, possesses a protoporphyrin sub-unit which is activated and modulated by NO(Ignarro, 1989).The most significant feature of these kinds of 02 signalling mechanisms is that by virtue oftheir extreme sensitivity to changes in 02 concentrations, they demonstrate the potential to bringabout subsequent changes in cellular functions well before 02 is truly limiting to cell function.Therefore, the potential exists for scaling metabolic events with decreasing changes in 02. Allof these signalling mechanisms could have a potential role in coordinating the disappearance of02 with metabolic suppression and since hypoxia-anoxia occurs in all tissues at approximatelythe same time, they provide a means for systemic signalling.Whatever the mechanism, there is mounting evidence for wide scale protein-levelphosphorylation in metabolic suppression (see next section). Therefore transduction of external02 changes likely involves a receptor mechanism at some level.As mentioned above, a characteristic feature of tissues capable of arresting metabolism isthe long-term maintenance of steady ATP concentrations. However, a number of studies havenoted that in response to declining POz there is a small and transient reduction in ATPconcentrations that appears universal amongst tissues (Kelly & Storey, 1988; Nilsson & Lutz,1992; Land et a!., 1993) and results in the catabolism of a small amount of adenine nucleotidebeyond AMP. Studies on turtle brain slices find a transient increase in extracellular adenosinefrom 1 to 21 M over 100 mm of anoxia (Nilsson & Lutz, 1992) which coincides with therelease of inhibitory neurotransmitters and the time of down-regulation of metabolic rate foundfor the whole animal. Although these changes in adenosine concentrations are acute (half-lifeof adenosine in blood is a few seconds), there could be a role for this metabolite in effecting16local metabolic changes during the transition to metabolic suppression. As discussed below,there is strong evidence to suggest that enzyme activity can be modulated throughphosphorylation/dephosphorylation reactions during metabolic suppression. A current workinghypothesis for the induction of this phosphorylation involves the binding of adenosine to theadenosine A2 receptor to activate protein kinase C (PKC) through the diacyiglycerol secondmessenger system. PKC directly, or indirectly through increased [Ca2]1,could phosphorylateglycolytic enzymes and membrane associated proteins bringing about a coordinated suppressionof metabolism. However, the means by which metabolism stays suppressed once adenosineconcentrations have abated remain unknown.Whether the link between environmental anoxia and metabolic suppression lies in 02 itselfor a combination of effects such as those found with adenosine remains unclear. Whatever themechanism, there are a number of well defined cellular responses that ensue which arediscussed in the next section.Mechanisms of Metabolic Suppression.There are three principal components of metabolic suppression that are highly conservedbetween forms (i.e. caused by differing environmental conditions) and species in which this kindof dormancy occurs. Firstly, ATP synthesis and ATP demand are coordinately reduced with theeffect that ATP concentrations ([ATP]) and cellular energy charge remain high and constant.Secondly, ion gradients and the plasma membrane potential are conserved. Lastly, tissues aremaintained functionally viable during long periods of metabolic suppression such that theirphenotypic characteristics and active function are not compromised on re-oxygenation andrecovery. Successful survival of anoxia by metabolic suppression is therefore a balance of17conflicting demands: reduced ATPase and synthase activity versus the need to expend energyin the stabiisation of the membrane potential and the turnover of functionally importantproteins. This section discusses current concepts of how energy supply, membrane functionsand anabolic processes are controlled to achieve a reversible suppression in activity in hand witha balance between energy supply and demand.Reducing Enzyme Fluxes. The first conserved feature of metabolic suppression, reducedATP supply and demand, is associated with a coordinated inhibition of rate controlling,regulatory proteins. Slowing the rate at which ATP is both synthesised and hydrolysed requiresa coordinated, large scale depression of enzyme activity, with remaining flux through enzymepathways finely tuned to the specific requirements of the tissue. The rate at which substrate (S)fluxes to a product (P) through an enzyme, is expressed as V=lcat.eo, where V, is themaximum rate of S—’P flux through an enzyme, keat is the turnover number of S-.P per catalyticsite and e0 is the enzyme concentration. This relationship states that the principal enzyme fluxcontrol mechanisms affect changes in V, either through changes in enzyme catalytic efficiency(kcat) or by altering the absolute concentration of active catalytic sites (e0).On entry into metabolic suppression, the principal routes for changing keat are via covalentmodification and allosteric regulation. Covalent modification of enzyme activity occurs by thebinding of phosphate and directly influences enzyme catalytic efficiency. Becausephosphorylation is controlled by signal transduction pathways associated with cAMP and cGMPdependent protein kinases, this also has the potential to be influenced by exogenous signals suchas hormones.In anoxia-tolerant tissues, anoxia generally results in significant, tissue-specific changes inthe kinetic constants for rate limiting glycolytic enzymes, pyruvate kinase (PK),18phosphofructokinase (PFK), and also for glycogen mobilisation through glycogen phosphorylase(GPase). In turtle liver, PFK demonstrates a 1.5 fold increase in Km for ATP and a 2.9 folddecrease in inhibitory constant (‘50) for citrate. This tissue also demonstrates a 5 fold increasein the amount of GPase in the active “a” form (Keiver and Hochachka, 1991). Heart shows theopposite pattern with a 2 fold decrease in K,,, for ATP and a 1.7 fold increase in I for citrate(Brooks and Storey, 1989). PK also demonstrates the same general trend for both tissues.Changes in glycolytic intermediates during anoxia in turtle tissues demonstrate that alteredenzyme kinetic characteristics are correlated with a tissue specific glycolytic response: livertissue exhibits enhanced glycogenolysis and glucose release, and heart and brain show anincreased capacity for fermentation of exogenous glucose (Kelly & Storey, 1988; Brooks &Storey, 1989).Evidence that covalent modification by enzyme phosphorylation may be at the root of thiskind of response comes from in vitro comparisons of kinetic constants from purified aerobic andanoxic forms of PFK and PK. In the goldfish, significant shifts in the isoelectric focusing points(p1) of PK and PFK isolated from liver, red, and white muscle have been detected indicatinga larger charge is present in the anoxic enzyme forms. However, the increase in p1 is onlyassociated with changes in kinetic properties of PK and PFK from the liver (Rahman & Storey,1988). Subsequent in vitro treatment of the purified enzymes with protein kinase or alkalinephosphatase demonstrates same-direction, and similar magnitudes of change in the kineticconstants for the purified enzymes and the anoxic enzymes (Rahman & Storey, 1988). In turtles,PFK and PK-phosphorylation events are yet to be investigated, however it seems that there isan increase in 32P-protein interactions during anoxia in liver and brain (Brooks & Storey,1993a).19The most compelling evidence for phosphorylation-mediated covalent modification duringmetabolic suppression in vivo comes from studies on PK from the channelled whelk(Busycolypus canaliculatum). In anoxia, there is a 50% increase in 32P bound to total cellularprotein. Purification of PK from these crude extracts reveals a phosphorylated anoxic form ofthe enzyme which demonstrates a 34% lower V, 90% increase in the K1, forphosphoenolpyruvate, and increased inhibitory constants for alanine and ATP (Plaxton &Storey, 1984a and b). Incubation of the anoxic form of PK with alkaline phosphatase revertsthe kinetic properties of the enzyme back to the aerobic state (Plaxton & Storey, 1984b).Recently, the protein kinase responsible for this effect was identified as a cGMP-dependentprotein kinase G (Brooks & Storey, 1990) supporting the possibility of receptor-mediatedcontrol of metabolic suppression.Allosteric modulation of enzyme activity occurs by the association of a ligand with aspecific, modulatory binding site which is separate from the enzyme catalytic site. This affectsa change in enzyme tertiary and quaternary structure which influences the properties of theenzyme catalytic site. The role of fructose 2,6-bisphosphate (F-2,6P2)as an allosteric regulatorhas been extensively examined during metabolic suppression in whelk tissues. PFK-1 catalysesthe glycolytic reaction between fructose 6-phosphate and fructose 1,6-diphosphate bytransphosphorylation using ATP as a substrate. This is a non-equilibrium reaction which iscatalysed in the gluconeogenic direction by fructose 1 ,6-bisphosphatase (Fl ,6P2ase). Thecontrol of the rate of flux through glycolysis versus gluconeogenesis in the upper part of thepathway is controlled by the relative states of activation of these two enzymes (Hers & Hue,1983). F2,6P is a potent activator of PFK-1, and synergistically inhibits Fl ,6P2ase with AMP,and as such, enhances the potential for glycolytic flux over gluconeogenesis at this locus (Hue20and Rider, 1987). Concentrations of F2,6P are regulated by a bifunctional enzyme which isgoverned by phosphorylation through a cAMP-dependent protein kinase to favour either thesynthesis of F2,6P when dephosphorylated (PFK-2 activity) or its catalysis to F6P whenphosphorylated (F2,6Pase activity). In anoxic whelk tissues, there is a 4-fold drop in PFK-2activity with resulting, significant tissue-specific reductions in F2,6P concentrations (Bosca &Storey, 1991). This together with an anoxic phosphorylated form of PFK-1 that is lessresponsive to F2,6P, ensures that glycolytic activation by F2,6P is removed at the PFK locus.In turtle and goldfish, the change in F2,6P is tissue-specific but inconsistent between thesame tissues from these two species. Goldfish show a significant increase in brain and heartF2,6P and large reductions in liver, kidney and muscle (Storey, 1988). In turtles however, redmuscle shows a large increase in F2,6P whereas in heart, it decreases (Brooks and Storey,1989). Interpreting these results is difficult, however the authors suggest that increases inF2,6P are correlated with the relative activation of glycolysis in more metabolically activetissues (Storey, 1988).The final mechanism for modulating enzyme activity during metabolic suppression involvesthe binding of enzymes to sub-cellular structures. In a review by Srere (1987), the cellularmilieu is envisaged as a relatively organised environment where enzymes along pathways bindto sub-cellular structures and are arranged in close proximity to each other. Hypothetically,within this ordered system, substrates could be channelled from one enzyme to anotherincreasing the efficiency of catalysis along a pathway by removing the randomness of substratediffusion. Another possibility suggested by this model is that enzyme pathways could be locatedlocally to where their product is in highest demand.There is good evidence to suggest that glycolytic enzymes do reversibly bind to cytoskeletal21structures in a number of tissues, and that the reversible nature of this binding correlates withchanges in glycolytic rate. In trout white muscle for example, exercise to exhaustion producedan associated increase in PFK, aldolase, GAPDH and phosphoglycerate kinase (PGK) bindingto the particulate fraction (Brooks & Storey, 1988). However, in anoxia tolerant organisms, theevidence for enzyme binding in the control of glycolysis is inconsistent between systems. Inthe channelled whelk ventricle, activities of hexokinase (HK), PFK, aldolase, glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and PK all exhibit a reduction in binding to the particulatefraction during anoxia (Plaxton & Storey, 1986), but this is not the general rule for all tissues.In whelk foot muscle, HK and PFK exhibit increases in binding whereas all other enzymesmeasured exhibit a decrease (Plaxton & Storey, 1986). In turtle brain, the activities of HK,PFK, GAPDH, aldolase, phosphoglycerate kinase and PK all exhibited significant increases inbinding according to the assay method used (Duncan & Storey, 1992). Based on studies whichassociate changes in glycolytic flux with altered enzyme binding profiles, these authors concludethat binding enhances the rate of substrate passage through glycolysis. Therefore duringmetabolic suppression, increased enzyme binding in turtle brain would seem rather anomalousand is explained on the basis of situating the glycolytic pathway close to the source of ATPdemand in the cell.Although changes in enzyme binding to the particulate fraction have been demonstrated inassociation with anoxia, no study to date has assigned any real, measurable, metabolicsignificance to this phenomenon. Studies examining the effects of binding interactions betweenpurified glycolytic enzymes and F-actin on enzyme kinetics have been examined in vitro andtend to show inconsistent results. PFK activity is enhanced with binding according to itsphosphorylation state whereas aldolase, GAPDH and PK all showed an inhibition on binding22(Reviewed by Brooks & Storey, 1993c). However, the concept of substrate channelling throughbound enzymes is also weakened by the fact that enzymes are rarely at equilibrium wherechannelling would work best. Differences in the equilibrium constants between enzymes wouldresult in accumulations of products that would lead to product inhibition at points along thepathway. Whether this truly is a significant mechanism involved in modulating metabolic fluxduring metabolic suppression remains vague.Where enzyme binding to cell structures may have some true significance is in themodulation of effective enzyme concentration. Changing e0 requires a system that canaccommodate the rapid and reversible masking of enzyme catalytic sites in response to anexternal stimulus such as phosphorylation. A good example of such a system is found in theerythrocyte transmembrane anion transporter known as band 3 protein (reviewed by Low,1986). This protein possesses a rod-shaped cytoplasmic domain that binds, and in so doinginhibits, a number of glycolytic enzymes, notably GAPDH, aldolase and PFK. Theseglycolytic enzymes are released through the phosphorylation of a tyrosine group near the N-terminus of the band-3 cytoplasmic domain with a resulting increase in e0 and glycolytic rate(Harrison et a!., 1991). Other means for increasing e0 can occur through mechanisms such asligand-induced association-dissociation of enzyme subunits (Kurganov, 1983) or synthesis ordegradation of existing protein (Hargrove & Schmidt, 1989). However, for affecting rapidchanges in metabolic flux rates, when energy consumption must be conserved, the latter modelsrepresent less attractive means for changing effective enzyme concentration.A significant problem with the entire analysis above is the lack of demonstrated cause andeffect. The timing, magnitude and collective effect of phosphorylation events, changes inallosteric effector concentrations and alterations in enzyme binding have not been demonstrated23to be causative in bringing about the coordinated metabolic suppression of the magnitudedescribed in vivo. A recent theory put forward by Hochachka and Matheson (1992) providesa framework for effecting large scale changes in metabolic rate that may prove useful in futurestudies of regulation in metabolic suppression and activation. In systems that undergo large andrapid, positive or negative changes in metabolic rate (eg hummingbird flight muscle ormetabolically suppressed tissues) changes in 1cat such as those described above, have beenhypothesised to be insufficient in bringing about the orders of magnitude change in metabolicactivity required. Furthermore in each case, and especially in metabolically suppressedsystems, the change in metabolic activity occurs whilst concentrations of regulatory metabolitesand allosteric effectors are relatively constant (Hochachka et al., 1991; Hogan et al., 1992).This prompted Hochachka and Matheson (1992) to suggest that a primary mechanism for largescale changes in flux through a pathway, in response to a change in oxygen availability,primarily involves a change in the concentration of enzyme catalytic sites (e0) rather than achange in the turnover number of S to P per catalytic site (U. Within this scenario, largechanges in metabolic flux are accommodated primarily through changes in e0, with allostericand covalent modulation performing the fine tuning of flux through the changed number ofcatalytic sites. As a framework for studying mechanisms of reducing metabolic flux in tissuesof anoxia tolerant organisms, Hochachka and Matheson’s theory is particularly attractive sinceit proposes a mechanism where large positive and negative changes in metabolic flux can beaccommodated, then finely tuned to the specific needs of the tissue.Reducing Membrane Ion Fluxes. The membrane potential is generated by the activepumping of charged ions against their electrochemical gradient and provides the kinetic energyfor cellular processes such as solute transport, amino acid and carbohydrate uptake, ATP24synthesis, cellular communication and so forth (reviewed Harrison & Lunt, 1980). A loss ofthis charge imbalance across the membrane results in a depolarisation of the cell membraneresulting in the rapid opening of voltage-gated Ca2 channels, activation of phospholipases A1and A2, rupture of the cell membrane and ultimately cell death (see later).The pumping of ions against an electrochemical gradient is an energetically demandingprocesses. In rat hepatocytes Na/K ATPase activity accounts for 16% of cellular ATPdemand, and in combination with Ca2 and H ATPases, this is extended up to 40% (Schneideret al., 1990). Na/K ATPase activity alone utilises 60% of total ATP demand in themammalian brain, and in kidney accounts for as much as 80% (Erecinska & Silver, 1989;Brezis et al., 1984). Clearly, ion pump ATPases constitute a significant ATP sink within thecell. Therefore, metabolic suppression in hand with stable ATP concentrations requires thatrates of ATP turnover should be reduced by lowering ion pumping activity. This represents onlyhalf of the solution, however. Maintenance of the cell membrane potential demands that reducedion pumping rates must be balanced at all times with opposing rates of ion flux as ions migratealong their respective electrochemical gradients. This flux of ions is controlled by a gatingmechanism within trans-membrane channel proteins that are specific for individual ion species.The gating mechanism can seal the channel against further ionic conductance, however this isfar from an all or nothing process. In inactive excitable tissues, inefficiencies in channelclosure result in a statistical probability of a certain percentage of channels being open at anygiven time (Hille, 1984). This background conductance is countered by pump-ATPase activitywhich maintains a polarised membrane potential over the cell membrane. Therefore duringmetabolic suppression, without a mechanism to coordinate the reduction in ion pumping rateswith opposing passive ion flux through channels, the cellular membrane potential would slowly25dissipate.The channel arrest hypothesis (Lutz et al., 1985; Hochachka, 1986) predicts that duringmetabolic suppression in anoxia-tolerant tissues, the membrane potential is maintained bydecreased ion-pump ATPase activity with coordinated reductions in channel ion flux. Thisresults in an overall reduction in the permeability of the cell membrane with sustained, butgreatly diminished rates of ion flux. If the predictions of the channel arrest theory hold true,then anoxia should be associated with a reduction in the conductance of specific ion speciesacross the plasma membrane with a resulting increase in membrane resistance and a steadymembrane potential. Studies conducted on isolated turtle cerebellum demonstrate that thecellular transmembrane potential does indeed remain constant at -60 to -85 mV during an anoxicmetabolic suppression (Pérez-Pinzón et a!., 1992a) which is estimated to be on the order of80% in turtle brain (Lutz et a!., 1984; Chih et al., 1989a). These studies also demonstratedthat input resistance fell by 36% during anoxia suggesting that cell membranes in anoxicpreparations are generally less leaky to ions. This has also been demonstrated in slicesincubated with ouabain where K leakage was significantly slowed in anoxic brains versusnormoxic controls (Chih et a!., 1989b). Ca2 flux through Ca2-channels also appears to bereduced during at least 5h of anoxia (Bickler, 1992). The evidence for channel arrest is notclear-cut for all brain regions, however. Pyramidal neurons from the turtle cortex demonstrateno apparent change in membrane resistance with anoxia (Doll et a!., 1991). Although thesecells can survive at least 120mm of anoxia, they appear to exhibit only a moderate 42%suppression in calorimetric heat-flux (Doll et al., 1994) which may factor into a specificmembrane and metabolic response of this brain region to anoxia.Experiments in turtle hepatocytes also present evidence for channel arrest in metabolic26suppression. By using 86Rb as a potassium analogue, it has been estimated that Na/KATPase activity is diminished by 75% during anoxia (Buck & Hochachka, 1994). Under thesame conditions, hepatocyte membrane potential remains constant at -3lmV and is apparentlymaintained independently of 02 availability. Therefore, despite a large reduction in the ATPturnover specific to Na/K ATPase activity, the membrane potential remains constant, anobservation that is consistent with the channel arrest hypothesis.A unifying observation from all of these studies is that despite a large metabolic suppressionassociated with constant [ATP] (Brain - Lutz et al., 1984; Doll et al., 1994, Kelly and Storey,1988; Hepatocytes - Buck et a!., 1993), preservation of the membrane potential is dependenton energy provision through glycolysis. In each case there is a rapid depolarisation onadministration of the GAPDH inhibitor, iodoacetate, and in the case of turtle brain, this alsooccurs during incubation without exogenous glucose (Doll et al., 1991; Pérez-Pinzón et a!.,1992a and 1992b). Therefore, although the membrane potential is maintained during metabolicsuppression, there is still a requirement for continued pump ATPase activity which is vital tothe continued survival of the tissue.How coordinated changes in rates of ion flux through channels and pump ATPases mightoccur remains a mystery. From the equation Vmax=kcat.eo discussed earlier, it appears that areduction in ion flux could occur either through an effective concentration change of thefunctional proteins involved or through a change in their “catalytic” efficiency (efficiency ofgating). There is some evidence to suggest that modulation of channel e0 occurs during anoxiain turtle brain. The density of both active and inactive Na channels decreases by 42% onexposure of turtle brain slices to anoxia (Pérez-Pinzón et a!., 1992c). Moreover, there is a goodpossibility that phosphorylation-mediated inhibition of channel activity may be involved here27also. In turtle brain, anoxia results in an increase in the proportion of protein kinase C (PKC)isozymes I and II that are associated with hindbrain membranes. In the cerebellum, the oppositeeffect has been noted (Brooks & Storey, 1993b). This change in PKC distribution has beensuggested to occur in associated with changes in membrane organisation during anoxia.From the mammalian literature, a number of studies have noted that in rat brain, Na andK (Numann et a!., 1991), and in heart, Na and Ca2 (Schubert et a!., 1989), flux throughthe respective ion channels can be reduced by either direct administration of PKC or additionof phorbol esters to activate PKC. In liver cells, this has been developed further where transientopening of Na, K and Ca2 channels occurs by increases in [Ca2], and deactivation by PKC(Fitz et a!., 1994). In this model, extracellular ATP mediated the increase in [Ca2J1througha 5’nucleotide receptor. Similarly, PKA and cGMP dependent protein kinases appear capableof modulating ion channel flux and G-proteins have been implicated in the control of ionchannels through direct association with Na and Ca2 channel proteins (Schubert et a!., 1989).Given the fact that changes in channel flux rates occur in response to progressive hypoxia,the recent finding that an oxygen-sensing mechanism modulates K channel activity in brain isparticularly interesting (Jiang & Haddad, 1994). In this study, progressive hypoxia deactivatedK channels in a manner that suggested the direct involvement of a metal binding protein thatwas not aFe2-heme protein. The implications of this study are significant since it demonstratesa direct associate between ion flux, channel opening and oxygen concentration.Alteration of pump-ATPase activity also occurs as the result of phosphorylation. Na/KATPase is inhibited through a protein kinase which is appears to be PKC (Bertorello et a!.,1991). However, Na/K ATPase activity has also been demonstrated to fall with reduced[Na]0or removal of available ATP. At this time it remains unclear if NafK ATPase activity28is suppressed allosterically or by effective substrate limitation during metabolic arrest.An overlooked, but potentially important regulator of ion channel and pump activity is themembrane itself. Since channel and pump proteins are membrane-bound they are subject toregulation by changes in membrane fluidity and composition. In comparison with mammaliancell membranes, reptilian cell membranes are markedly less leaky with lower rates of K andNa exchange (Else & Hubert, 1987). This has been correlated with a markedly differentmembrane composition in reptiles. In the liver, phospholipid concentration is 35% greater inrat than lizard with a significantly greater proportion (16.2%) of rat fatty acids aspolyunsaturates. Total protein content of rat liver is 54% greater than lizard liver and this isreflected in a higher protein:lipid ratio in rat liver mitochondria (Hubert & Else, 1989).Although the proportion of cholesterol was not measured in these studies, the lower proportionof polyunsaturated fatty acids, and the lower protein:lipid ratio of organelle membranes pointsto an inherently lower fluidity quotient in reptilian membranes (higher microviscosity).Possessing a less leaky membrane does not present any metabolic saving during anoxia in itselfsince standard metabolic rate is also 4-5 times lower in reptiles than in mammals. However,since membrane-bound proteins can be activated and de-activated depending on the lipidenvironment (Sweet & Schroeder, 1988), a predisposition to fluidity-related changes in channeland pump conformation in reptiles could be argued.The plasma membrane is a dynamic 2-dimensional fluid environment in which lipidmolecules and proteins continually diffuse, forming regional domains with locally variedcomposition and biophysical properties according to the relative concentrations of cholesterol,phospholipid and glycosphingolipid (Yechiel & Edidin, 1987; Curtain et a!., 1988). Throughzonal changes in membrane microviscosity a large number of membrane protein-dependent29processes are affected. From the channel arrest perspective it is significant that diffusion ratesof Na and K ions decrease with lowered membrane fluidity (Cooper et a!., 1975), as doactivities of Na/K ATPase (Kimelberg, 1975), Ca2ATP se (Warren et a!., 1975) and ionflux rates through the acetyicholine receptor channel (AchR; McNamee & Tong, 1988). Fromstudies such as these, it appears that functional membrane proteins perform most effectivelyover an optimal viscosity range, an in association with specific phospholipids, which supportsan active protein conformation (Sweet & Schroeder, 1988). The AchR channel has arequirement for cholesterol and negatively charged lipids which appear to interact with thesecondary and tertiary structure of the protein subunits. Altering either of these componentsresults in reversible channel deactivation and denaturation (Fong & McNamee, 1985; Sunshine& McNamee, 1994). Hypothetically, if anoxia caused regional changes in the fluidity of cellmembranes in turtle brain, the observed decrease in Na-channel density could reflect a changein Na-channel conformation which is not recognised by the ligand used to measure channeldensity[3H-brevatoxin (Pérez-Pinzón eta!., 1992c)]. Discovering whether the lipid compositionof cell membranes from facultative anaerobes predisposes ion channels and pumps to anoxiainduced, but reversible, denaturation constitutes an interesting area for future research.Anabolic Processes. The third conserved feature of metabolic suppression centres on therequirement to preserve functional viability during the period of metabolic suppression.Anabolic processes such as nucleic acid synthesis, protein synthesis, and membrane synthesisand assembly, are involved in the continuous renewal and replacement of existing cellstructures, enzyme pathways and so forth, and are vital in staving off entropic forces. Anabolicprocesses are energetically expensive, but demonstrate a wide functional scope depending onthe growth state of the tissue. For example, in static cultures of rat hepatocytes, protein30synthesis accounts for 13% of total ATP demand (Schneider et a!., 1990). However, in morerobust, growing cells such as the ascites tumour cell, protein synthesis alone accounts for nearly30% of total ATP demand (Muller et al., 1986). A cost for RNA synthesis of 4% and 11% hasalso been measured in these respective cell types bringing the anabolic cost of protein synthesisto 17% and 41% of total ATP demand in each case.To date, the only observations relating to anoxic anabolic responses during dormancy stemfrom research on the brine shrimp embryo (Anemia fransiscana). The encysted gastrula iscapable of suppressing metabolism by 99.6% after one day of entry into anoxia (Hand, 1990)and can persist in this state for up to 18 months when hydrated with little effect on recoverysuccess (Clegg, 1992). During this time, processes involved in sub-cellular differentiationincluding protein synthesis, mitochondrial maturation and liberation of lipid from yolk plateletsare completely inhibited (Kwast & Hand, 1994; Hoffman & Hand, 1990). A major differencebetween this system and vertebrate anoxia tolerant systems is that the brine shrimp gastrulaexhibits relatively poor intracellular buffering capacity. Therefore changes in metabolic,anabolic and catabolic rate processes are governed much more closely by intracellular pH in thissystem.The response and control of anabolic cell processes, and in particular protein turnover,during metabolic suppression has received very little attention to date in vertebrate anoxiatolerant systems. Yet, as a cell process, it is vital for ensuring stability within the protein pooland amongst cell structures and is also an important means of implementing phenotypic andmetabolic changes within tissues. The next section discusses the response of protein turnoverand gene expression to hypoxia in anoxia intolerant systems and uses these examples to drawup a hypothetical picture of the control of these processes in anoxia tolerant systems. This also31brings the literature review to the principal thrust of this thesis: the response, energetic cost,and significance of protein turnover and protein expression in metabolic suppression.Protein Turnover and Gene Expression in Anoxia Intolerant Systems.Kinetics ofProtein Induction. Protein turnover is a multifaceted cellular process comprisedof gene activation, expression of mRNA, translation and protein degradation. Alteration of thepattern of protein expression in response to an external stimulus (anoxia, for instance) requiresa balanced interaction between these processes in order to remove inappropriate proteins andsynthesise new ones.Figure 2 (upper panel) demonstrates the relationship between the turnover of the mRNApool and the protein pooi. From this interaction, Hargrove and Schmidt (1989) have developeda two compartment model that describes the relationship between the stability of a mRNA andthe subsequent synthesis of its translate. In this model, rate of formation of both mRNAs andproteins are independent of their own concentrations and as such, exhibit zero order kinetics.Rates of degradation are exponentially linked to the concentration of a mRNA or protein andexhibit first order kinetics. Since rates of degradation are the only concentration-dependentvariable, the rate of change in concentration for a given mRNA or protein is governed by itsrate of degradation often expressed as the half-life. A further important feature of the twocompartment model is that rates of protein synthesis are dependent on the stability andconcentration of mRNA. Therefore the induction of a protein to a new steady-stateconcentration can be described as the following solution (Hargrove & Schmidt, 1989):RSS.lcp(Equ. 1)kdpFIGURE 2. Two compartment model of Protein ExpressionSignalTiming of PEPCK Expression.+12MatureProtein32+ Transcription(Ks-i)GeneTranslation(Ks-2)mRNAPool(R)ProteinPool(P)RNaseGene H mRNADeg.(Kd-i) ProteaseGene HNucleotidesProteolysis(Kd-2)AminoAcids2-3 mmcAMPGene —MaturemRNA10mm2-5 mm4NascentProteinI NascentL mRNAtl/2=0.5-2.5hNucleotidest 1/2=5hAminoAcidsCompiled from Hargrove & Schmidt, 1989.33where P8 denotes new steady state protein concentration, R,, denotes new steady state mRNAconcentration, k8p denotes the new rate constant for protein synthesis and kdp is the new proteindegradation rate constant.This model presents overall protein turnover as a function of mRNA and protein half-life.Extending protein half-life, as predicted during metabolic suppression, therefore extends thelength of time a protein is present irrespective of its rate of synthesis. Because of the strongdependence of protein synthesis on mRNA concentration, changes in protein concentration toa new steady state can occur either by an alteration in transcription rates or a change in mRNAstability. This dominates over rates of protein degradation since mRNA half-lives for individualproteins are generally much shorter than their active protein translates, and the averagetranslational yield from one mRNA is 10,000 protein translates representing a largeamplification effect (Hargrove et a!. 1991). As a result of the apparent dependence of proteinsynthesis rates on mRNA concentrations, Hargrove and Schmidt suggest that the mostenergetically favourable method of suppressing protein synthesis rates is to effectively decreasethe concentration of translatable mRNA. Conversely, an increase in protein synthesis rates isbest achieved by increasing the rate of translation of existing mRNAs.A further point of note demonstrated in figure 2 (lower panel) is the timing of gene andprotein induction, demonstrated for phosphoenolpyruvate carboxykinase (PEPCK). Signalling,gene induction, transcription, mRNA maturation and translation only require about 2.5% of thetime required to produce a fully active, mature protein. Given that the half-life of the matureprotein is 5h, and post-translational processing of the nascent protein requires 12h, it appearsthat further control of rates of protein appearance and removal could be exerted at this ratelimiting step.34Control of translational rates via phosphorylation of initiation and elongation factors appearsto be a major mechanism for acute modulation of protein synthesis. About 20 proteins involvedin initiation and elongation are known to undergo phosphorylation by protein kinases. Since thesynthesis of all proteins begins with the binding of Met-tRNA to the 40S ribosomal subunit, thisstep represents the rate limiting step for the initiation of protein synthesis and as such, initiationis under tight control. The key enzyme in the binding of Met-tRNA to the 40S ribosomalsubunit is eukaryotic initiation factor-2 (eIF-2) whose activity can be suppressed in vivo byphosphorylation of a serine residue on the a-subunit (Davies et al., 1989; Kaufmann, et a!.1989). Phosphorylation also occurs amongst several proteins involved in elongation, mostsignificantly, elongation factor-2 (eEF-2) which catalyses the translocation of peptidyl-tRNAfrom the A to P sites on the ribosome. Phosphorylation by a Ca2-dependent protein kinaseleads to its inactivation (Ryazanov et a!., 1988).The GTP/GDP ratio represents a further level of control in protein synthesis. IncreasedGDP concentrations results in the competitive inhibition of GTP binding to the eIF-2 initiationcomplex, together with a number of elongation factors possessing GTPase activity (Walton &Gill, 1976).Control of protein degradation is much less well understood. Cellular protein degradationis an amalgam of several different pathways, some compartmentalised (eg in the lysosome) andothers cytosolic (eg ubiquitin-dependent pathways). Most protein degradation pathways havea requirement for ATP hydrolysis or binding either in their regulation, or the maintenance ofa favourable proteolytic environment. For instance, the ubiquitin-dependent pathway requiresthe binding of ATP in the activation of co-factors involved in binding ubiquitin to proteins,prior to their degradation (Hershko, 1988). In the lysosome, ATP is hydrolysed in the action35of an ATP-dependent proton pump that maintains a low pH within (Schneider, 1981). As such,proteolytic activity ceases if ATP concentrations dissipate (Gronostaj ski et al., 1985; Plomp eta!., 1987)Phosphorylation appears to play a role in altering the susceptibility of proteins todegradation. For instance, phosphorylation of neurofilaments decreases their susceptibility todegradation by calmodulin (Greenwood et a!., 1993) whereasCa2-dependent phosphorylationof lipocortins leads to selective degradation by an undescribed membrane-bound protease (Chuah& Pallen, 1989).Overall then, protein turnover in a steady state system is regulated at numerous points.Gross-level control of the concentration of a protein is manifested through the relativeconcentration of its mRNA versus its rate of degradation. Generally, acute modulation andfine-tuning of protein synthesis rates occurs via phosphorylation which modulates the catalyticefficiency of initiation and elongation factors. Control of protein degradation is likely muchmore specific to individual proteins, depending on the pathway employed and mode of substraterecognition. How then, does this framework of control fit with current research on the effectsof hypoxia on protein turnover?Effect ofHypoxia on Protein Turnover. Hypoxia has the general effect of reducing proteinturnover rates in mammalian systems. Short-term hypoxia (10% 02 for 6h) in rats for example,leads to a 20-35% decrease in protein synthesis rates in various tissues with the morecharacteristically energy demanding tissues (such as liver, kidney and brain) demonstrating thelargest inhibition of 25, 29 and 32% respectively (Preedy et a!., 1989). Studies on rathepatocytes have attempted to find the intracellular basis for this inhibition: l2Omin of anoxicincubation (OmmHg) results in a cessation of protein synthesis within 7 minutes of anoxia whilst36ATP concentrations remain at 80% of normoxic values. Progressively severe hypoxia (80 and5OmmHg) also shows a near complete cessation of protein synthesis rates despite highconcentrations of cellular ATP, continued rates of amino acid transport and no apparent increasein rates of protein degradation (Lefebvre et al., 1993). This indicates that the hypoxicinhibition of protein synthesis is rapid and apparently independent of changes in ATPconcentrations.What of the RNA pool? Hypoxia appears to lead to a dissociation of polyribosomalcomplexes from mRNA resulting in an effective change in translatable mRNA concentration(Surks & Berkowitz, 1970) and in rat hepatocytes, results in a slow fail in transcription rates(Lefebvre et a!., 1993). However, as with metabolic changes, neither of these processes occuron the same time-scale as the inhibition of protein synthesis.The implication from these studies is clear: acute and global changes in protein synthesisrates occur on a time-scale that is different from metabolic changes incurred by hypoxia(evidenced by ATP and AMP concentrations) or changes in RNA concentrations. Proteindegradation rates and rates of amino acid incorporation also appear to be negligible in slowingapparent protein synthesis rates (Lefebvre et a!., 1993). However, the fall does occur directlyand proportionately with the loss of 02, strongly implying a role for molecular oxygen itself inthe regulation of protein synthesis.Oxygen Sensing Processes in Hypoxic Gene Expression. In mammalian systems, hypoxiais associated with a tissue-specific increase in the expression of numerous gene products. Theoverall response is geared towards improving the capacity for metabolising glucose, increasingthe capability of the vascular system to carry and deliver oxygen and substrate, and protectingagainst protein damage caused by defective post-translational processes. For instance, cultured37skeletal muscle cells demonstrate a 3-5 fold increase in transcription rates for aldolase,triosephosphate isomerase, pyruvate kinase and lactate dehydrogenase over normoxic valuesover 3 days of incubation at 3% 02 (Webster, 1987). On a more rapid time-scale, liver andkidney show a several fold increase in the erythropoietic stimulating hormone, erythropoietin(Epo), and endothelial cells lining blood vessels and capillaries express a number of factorsincluding autocoids, vascular epithelial growth factor (VEGF), platelet derived growth factor-j3(PDGF-j3) and endothelin-1, all involved in stimulating capillary growth or modulating vascularsmooth muscle contractility.Particularly significant is the manner in which some of these genes are activated andtranscribed. Studies on the hypoxic induction of the Epo gene product in rat hepatocytes showa close relationship between the amount of Epo mRNA and protein translate expressed, and theseverity of hypoxia (Eckhardt et al., 1993). Expression of this gene can be manipulated byfactors such as carbon monoxide, cobalt and nickel that affect the oxygen-dependentconformation of heme proteins. Epo gene induction is also blocked by inhibitors of ferrochetalase activity in protoprophyrin ring synthesis. This kind of evidence has established thatEpo gene induction is linked directly to changes in tissue oxygen content by a widely distributed(tissue and species) membrane-bound heme protein (Goldberg et a!., 1988; Eckhardt et a!.,1993; Maxwell et a!., 1993). With increasing hypoxia, the heme group within this proteinchanges from oxy- to deoxy-conformations with a resulting increase in the expression of oneor more Epo transcription factors which subsequently activate the Epo gene (Tsuchiya et a!.,1993). The signalling mechanism that transduces the change in heme conformation into agenetic response has not yet been characterised but studies with agonists and second messengersthat classically activate protein kinase C did not result in Epo gene expression (Eekhardt et a!.,381993).This kind of02-sensing mechanism has also been implicated in the modulation of hormonalcontrol over phosphyoenolpyruvate carboxykinase (PEPCK) activity in the liver. Theperiportal-perivenous (afferent-efferent) zones of the liver acinus demonstrate a large oxygengradient from 65mmHg to 35mmHg on the venous side (Jungermann & Katz, 1989). Thisdefines both functional and metabolic characteristics of hepatocytes distributed along thisgradient such that periportal cells express higher activities of gluconeogenic enzymes (glucose-6-phosphatase, PEPCK for e.g), and perivenous cells appear preferentially glycolytic (expressinghigher activities of glucokinase and pyruvate kinase) (Jungermann and Katz, 1989).Administration of glucagon induces expression of the PEPCK gene via cAMP and increases theamount of active protein present but the magnitude of this response diminishes along theperiportal-perivenous oxygen gradient (Keitzmann et a!., 1992). Studies using homogenouspopulations of isolated hepatocytes demonstrate that glucagon-induced synthesis of PEPCK ismodulated by oxygen, and the oxygen effect can be diminished with cobalt and stimulated underhypoxia with carbon monoxide (Keitzmann et at., 1993). This strongly suggests that a hemeprotein oxygen sensing mechanism is involved in the oxygen-dependent modulation of PEPCKexpression and possibly accounts for the differential expression of enzymes along the periportalperivenous 02 gradient (Jungermann and Katz, 1993). Whether the oxygen-sensitive expressionof glycolytic enzymes is also connected with a heme protein oxygen sensing mechanism is notclear at present.Oxygen-dependent modulation of gene expression appears to be a widespread andevolutionarily conserved phenomenon. In mammals, endothelial cells express vascular epithelialgrowth factor, interleukin-2, platelet derived growth factor-a, angiotensin converting factor,39endothelin-1, and carotid body tyrosine hydroxylase, in an oxygen-dependent manner (seeFanburg et a!., 1990). The involvement of a heme-protein based mechanism in transducingchanges in 02 concentrations also appears to be widespread. The heme-protein mechanismdescribed for Epo expression is present in liver and kidney cell cultures derived from speciesthroughout the class mammalia (Maxwell et a!., 1993). Furthermore, the evidence that a hemeprotein is involved in hepatic PEPCK expression suggests that this kind of02-sensingmechanism is not unique to a single gene system.Numerous classes of oxygen-sensing genes are also well described in the prokaryotes. InRhizobium for example, the expression of proteins involved in nitrogen fixation is controlledby two proteins, FixL (a heme-protein) and FixJ which are increasingly expressed withdecreasing oxygen 02 tensions. E. coli is capable of modulating the expression of high and lowaffinity 02 binding cytochrome genes, fumarate reductase and nitrate reductase through the arc(aerobic respiration control) and fnr (fumarate and nitrate reductases) 02-sensing systemsrespectively (Tuchi et a!., 1990). The mechanism for02-sensing in the arc system appears todepend on the redox state of a component in the electron transport system whereas the fursystem demonstrated some of the characteristics of a heme-sensing mechanism.Implications for Anoxia-tolerant Systems. The preceding discussion has established thatthe turnover of a protein in mammalian systems is regulated by the concentration of its mRNAand the protein’s half-life. This in turn is fine-tuned by “acute regulators” of both translationand the various pathways of protein degradation. In anoxia-intolerant systems, progressivehypoxia results in a rapid inhibition of protein synthesis rates that occurs on a differentmagnitude and time-scale from metabolic changes implying a regulatory role for 02 itself, overand above other controlling factors. Finally,02-sensing via conformation changes in a heme40protein is a conserved phenomenon that regulates a number of02-sensitive gene products. Whatare the implications for anoxia-tolerant systems?Since protein turnover is an energetically demanding process it is reasonable to assumeprotein turnover rates are likely suppressed along with other cellular processes during metabolicsuppression. Anoxia-tolerant systems classically conserve ATP concentrations such that theputative fall in protein turnover rates likely occurs with little or no change in metabolicparameters (as observed above in rat hepatocytes). If this is the case then a regulatory role for02 at some level is a serious consideration.A fall in protein turnover rates necessarily means that protein half-lives are extended andthe requirement for mRNA transcription reduced. Although the inhibition of protein turnoverand transcription rates would result in a large energetic saving, prolonged exposure of proteinsto the cellular environment raises a number of other problems. The combined effects ofchemical and physical protein modification, coupled with slow rates of protein replacement andremoval would result in an accumulation of denatured and dysfunctional proteins over a periodof time. For this reason, rates of protein synthesis and degradation are unlikely reduced to zeroto avoid overt accumulation of damaged protein. Nevertheless, extended protein half-lives meanreduced rates of removal; if tissues are to regain function rapidly on re-oxygenation, amechanism for maintaining protein structure and function during anoxia is implied.The finding that02-sensing mechanisms govern the expression of certain hypoxia sensitivegenes, and is evolutionarily conserved, has obvious significance in anoxia-tolerant organisms.Depending on the nature of the02-sensing mechanism, “tracking” changes in 02 concentrationprovides the potential for the expression of a phenotype more suited to anaerobic conditions wellbefore 02 becomes limiting. The function of differential protein expression under anoxia is41unclear but there exists the possibility that the expression of metabolic enzymes, stress proteinsand free radical scavengers represents a part of this response that is associated with preservedcell function in anoxia and neutralising free-radical production on recovery.Consequences of Suppressing Metabolism: Remaining Viable.To this point, I have discussed mechanisms for increasing the efficiency of anaerobic ATPyield per mole of substrate, minimising and tolerance to metabolic end-product accumulation,and the coordinated changes required for suppressing metabolism. The final component thatis required for the successful survival of a long-term period of anoxia involves maintainingfunctional stability: how are the metabolic characteristics of tissues preserved so that onrecovery, active functions can resume unimpaired? The following discussion demonstrates thatanoxia imposes both acute and chronic effects on cellular viability and that in metabolicsuppression, these are dealt with in ways which are either novel, or appear as secondaryconsequences of other metabolic adjustments made in anoxia.Acute Mechanisms of Cell Death in Anoxia. To understand why remaining viable undermetabolic suppression is a problem, it is first important to understand how and why anoxiaintolerant cells die as a result of oxygen limitation. Figure 3 demonstrates the sequence ofevents that result in anoxia-mediated cell death and compares them to the known sequence ofevents on entry into anoxia in anoxia-tolerant organisms. In anoxia-intolerant systems anaerobicmetabolic demand is supported by a high rate of glycolysis which results in the progressiveacidification of the cell. As anaerobic glycolysis becomes progressively unable to sustain ATPdemand, a large and rapid fall in ATP concentrations occurs which is associated withaccumulations of ADP, AMP and the adenosine breakdown products, xanthine and42FIGURE 3. Pathway of acute cell damage (cell death cascade) in anoxia-intoleranttissues, and mechanism of avoidance in anoxia-tolerant tissues. (1) = anoxiatolerant pathways associated with avoidance of acute cell damage, (2) = pathwaysassociated with avoidance of chronic cell damage.PROGRESSIVE HYPOXIAATP SmmEsIsPasteur effect sustainsATP demand at nearaerobi rates.Exhaustion offermentable substrate43+Net inward Naand K’ effluxMito, ER and plasma membranesdepolariseCa2 entry into cytosol fromorganelles and cell exteriorATP DEMAND“Oxygen Sensor”? Adenosine?________PhosphorylationeventsATP synthesis Reduced ATP demand andre-partitioning of ATP turnoverto favour vital cell functions1______Channel arrest balancesreduced ion pwnp activities1ATP DEMAND[ATPJ [pHJpH below protein optimaTemporary stabilisation ofmembrane potentialLai. Chronic membrane damageand leakageProton leakage - pH1 increasesAl? SThTHESIS -.Low rate of substrate usageand end-product accumulationMechanismsfor stabilisingfunctionally importantproteins/Membrane Potentialstabilised(1)(2)Antioxidantdefenses(1 &‘‘(1)REOXYGENATION ANDSURVIVALPositive Feedback opensf2+4,Activation of phospholipasesand Ca dependent proteasesLoss of functional organisationand membrane rupture1CELL DEATh44hypoxanthine.Classically, cell death has been ascribed to a large increase in intracellular Ca2concentrations ([Ca2]1), resulting from the failure of membrane and mitochondrial potentialsmaintained by Na/K ATPase andCa2ATPases, as ATP concentrations fall. The subsequentincrease in Ca2 entry via the Na/Ca2 exchanger activates phospholipases disrupting theplasma membrane, endoplasmic reticulum and mitochondrial membranes, and activates cytosolicproteases leading to the appearance of dysfunctional proteins. This serves to further increasethe cycle of Ca2 entry and cellular disruption. Although this model is appealing, it hasrecently been established that diminished ATP concentratons ([ATP]) are not associated withincreases in [Ca2j1.Furthermore, increasing [Ca2J1does not result in the appearance of overtfeatures of cell disruption such as bleb formation (Lemasters et al., 1987; Nieminen et a!.,1990).The finding that a fall in intracellular pH (pH1) facilitates cell survival during hypoxia (Binget al., 1973) lead to a new theory of hypoxic cell injury where buffering of low pH towardsneutrality served as the initial trigger for the sequence of cell death, and maintenance of a lowpH1 actually abrogated events in the cell death cascade (Lemasters et a!., 1993). This modelof events during anoxic cell death has subsequently been termed the “pH Paradox” and centreson the observation that falling pH1 during anoxia results in a crude inhibition of functionalproteins as pH1 falls below their various optima. This inhibition is thought to include a numberof processes that regulate Ca2 homeostasis and also some of the processes that play a role incell disruption. In muscle fibres for example, the chief routes for Ca2 entry [the Na/Ca2exchanger (Kim et al., 1987); Ca2 flux through slow Ca2 channels (Chenais et a!., 1975);Ca2 release from the sarcoplasmic reticulum (Nakamaru & Schwartz, 1972)1 and phospholipase45A2 activity (Harrison et al., 1991) are all progressively inhibited as pH4 falls below 6.9. ThepH paradox has been well characterised in liver tissue where a fall in pH, from 7.4 to 6.9 andlower, during periods of anoxia-induced ATP depletion, results in significant increases in cellsurvival times during both anoxia and normoxic recovery (Currin et al., 1991).As demonstrated in figure 3, the “pH paradox” model suggests that the onset of cell deathis function of pH1. Slow increases in plasma membrane permeability over time results in thegradual efflux of W and a subsequent increase in pH1 towards neutrality. The speed at whichpH approaches neutrality increases as the plasma membrane depolarises. Increased rates of Naentry result in rapid proton entry as Cl ions are extruded in exchange for Na and CO32 ionsvia the Na-dependent Cl/HCO; antiporter. At the point of neutrality, and as the plasmamembrane fully depolarises, Ca2-medi ted cell death occurs through the classical pathwaysdescribed aboveFigure 3 also demonstrates how acute cell death is avoided in turtle tissues. Steady ATPconcentrations coupled with channel and ion pump arrest avoids plasma and mitochondrialmembrane depolarisation and the consequent entry of Ca2 into the cell (cf Nieminen et al.,1994). Furthermore, the proton buffering capacity of turtle tissues results in a gradual fall inpH1 in anoxia as opposed to the rapid and severe fall found in anoxia-intolerant tissues. In theturtle liver for example, pH1 falls linearly from 7.5 to 6.8 over a period of 2h (Wasser et al.,1991) and in the isolated perfused heart, pH, is maintained at 7.2 during four hours of anoxia(Wasser et a!., 1990). Clearly, although the “pH paradox” extends anoxic survival of anoxiaintolerant species, other mechanisms come into play in facultative anaerobes which avoid thenecessity for pH-inhibition of processes for acute cell death.From this analysis it seems that acute effects on cell viability are avoided largely by46metabolic adjustments (coordinated suppression of ATP supply and demand) which also includea reduction in membrane permeability (channel arrest) and a stable membrane potential.However, metabolic suppression as a mechanism for avoiding acute cell death creates a new setof conditions that affect cell survival in anoxia. Periods of suppressed metabolism can last frommonths to several years in some cases opening the way for long-term deleterious effects on cellviability. The next section discusses the potential for chronic, accumulative cell damage duringmetabolic suppression.Chronic Accumulation of Cell Damage. Long-term effects of anoxia and metabolicsuppression on cellular viability are poorly characterised in anoxia-tolerant species. However,it is reasonable to assume that if protein turnover is slowed during metabolic suppression, andprotein half-lives increase accordingly, then the potential exists for accumulative damage to theprotein pool, and subsequent failure of cell function in the long-term.Extensive increases in protein half-lives for cytochrome c oxidase have been noted duringanoxic dormancy in the brine shrimp embryo (Artemia fransiscana). Anoxic incubation ofAnemia gastrulae results in a large depression in protein turnover rates that are associated witha reduced number of polysomes and a static mRNA pooi (Hoffman & Hand, 1992). Studieson the anoxic turnover of cytochrome-c oxidase demonstrate a 77 fold increase in half-life forthis enzyme from 1.3 to 101 days (Anchordoguy et al., 1993). The significance of increasedcellular protein half-life has been noted in cellular pathology studies and in particular, the fieldof cellular aging. Aging in tissues is associated with a chronic increase in the half-lives of totalcell proteins. By extending the time a protein is present within the cell, the probability of post-translational modification resulting in conformational changes increases. With time, andreduced rates of protein removal, this leads to an accumulation of dysfunctional cellular protein47that can have a profound effect on the capability of tissues to remain functional (reviewed byReff, 1985). If during metabolic suppression, protein turnover was substantially suppressed,the infinite increase in protein half-life could lead to a lethal accumulation of dysfunctionalproteins. This clearly does not occur to lethal extents; anoxic incubation of Artemia embryosfor 1½ years results in minimal reduction in hatch success rates (Clegg, 1992). Similarly inturtles after 6 months of anoxic submergence at 3°C, survivorship was 40%, with a number ofdeaths associated with fungal disease rather than overt signs of metabolic failure (Ultsch &Jackson, 1982). Given the potential for protein damage during long-term metabolicsuppression, and the clear ability of anoxia-tolerant animals to regain full aerobic tissuefunction on recovery, it seems that the implication exists for a mechanism that preferentiallystabiises proteins. By examining rates of protein synthesis and degradation in turtlehepatocytes, the potential for protein damage during metabolic suppression is assessed in thefollowing research chapters and further discussed in chapter 5.Cell Damage on Recovery From Anoxia. Recovery from anoxia has the potential to bemore lethal than the period of anoxia itself due to re-exposure to 02. The risk of oxygensuperanion (02j production during hypoxia or recovery from anoxia is high, especially ifassociated with the catabolism of purines to xanthine as is often the case after short periods ofhypoxia in anoxia-intolerant systems. Under aerobic conditions xanthine is catabolized byxanthine dehydrogenase (XDH) to uric acid and NADH. However, under hypoxic conditions,depletion of ATP results in the influx of Ca2 ions into the cell activating a Ca2 protease. Thisprotease catalyses the conversion of XDH to its oxidase form, xanthine oxidase (XO), with theresulting catabolism of xanthine to uric acid and 02 (McKelvey et a!., 1988). Once again,anoxia-tolerant organisms avoid this cascade by maintaining ATP concentrations high and48constant during the period of metabolic suppression thus avoiding changes in [Ca2]1and theloss of adenine purine nucleotides beyond IMP (Land et al., 1993). Therefore under steady-state conditions, superoxide formation by this route is likely negligible, with the greatest riskoccurring during metabolic adjustment from one condition to another, such as entry intometabolic suppression or recovery. During these times, rapid increases in O2 concentrationsare likely contained by superoxide dismutases, catalases and peroxidases, as has beendemonstrated during anoxia in the garter snake (Hermes-Lima & Storey, 1992).Overall, this discussion demonstrates that the successful survival of a long-term period ofanoxia requires a coordinated metabolic response to negate rapid changes in cell membranepotential and substrate stores. The subsequent hypometabolism creates the potential for a newset of conditions that could foster accumulative damage to cellular macromolecules. Above all,tissues must regain function during the recovery phase and at the same time, reduce the risk ofoxygen free radical production and resulting tissue damage.This literature review has discussed the current views on the mechanisms of metabolicsuppression and the proposed significance of protein turnover and protein stability to tissuesurvival under anoxia. Although the magnitude and fine-tuning of suppressed metabolism aretissue-specific, it is clear that the overall control of events occurs at the cellular level, withoutexogenous input. To successfully probe the molecular and biochemical events that occur inmetabolically suppressed tissues, a model cell culture system is required that demonstrates theability to survive anoxia by metabolic suppression.49The Turtle Hepatocyte as a Model System for the Study of Anoxia Tolerance.Over the past 25 years, hepatocyte preparations from numerous vertebrate classes have beenwidely used to investigate aspects of cellular metabolism and its control. The reason for thislarge body of interest centres on the ease of hepatocyte preparation from numerous species andthe wide metabolic and functional scope of liver tissue in general (Berry et a!. 1991).Survival of anoxia clearly requires a coordinated response between organ systems withinthe animal. Whereas whole animal studies generate highly valuable data regarding thecoordination of metabolic changes between organ systems, these preparations are of limited usein examining the cellular energetics and control of anoxic metabolism in facultative anaerobes.Therefore a hepatocyte preparation was developed from the western painted turtle, Chrysemyspicta bellii and used to further investigate the sub-cellular characteristics and control ofmetabolism in anoxia (Buck et al., 1993b). Table 1 demonstrates the metabolic features of thiscell preparation under normoxia and anoxia, without exogenous substrate. The principal pointof note is that these isolated cells demonstrate all of the overt features of anoxia-tolerancediscussed in this chapter, vis: extended viability in anoxia, high concentrations of fermentablesubstrate (glycogen), metabolic suppression, constant [ATPJ and energy charge, reduced ion-pump activity with a constant membrane potential, and an ability to recover from extendedperiods of anoxia. The metabolic response to anoxia in rat hepatocytes is very different.Whole liver glycogen concentrations in rat are about 200mM, but much of this is lost duringisolation, such that freshly isolated rat hepatocytes possess in the region of 1-10mM glycogen,and are heavily dependent on exogenous glucose (Senglen, 1976; Berry et a!., 1991). As aresult, a 90% loss of [ATP] occurs during the first 30mm of anoxia and a complete loss ofmembrane potential and viability occurs after a period of 2h, coinciding with dissipation of the50TABLE 1. Metabolic Characteristics of Isolated Turtle Hepatocytes.Character Normoxia Anoxia(lOh) (lOh)Metabolic Rate 68.4mol ATP/g/h 6.5Heat Flow 1.O8mW/g 0.26[ATP] 2.21cmolIg 2.01Energy Charge 0.95 0.90Energy Charge(10mM lodoacetate) 000 0.18Glycogen 720jimolIg 630Glucose ReleaseRate 22.6tmoJIg/h 7.5Lactate ProductionRate n.m. 4.2Gluconeogemc Rate 1.95imol glucose/glh 0.00Na/K1-ATPase 19.1mo1 ATP/g/h 4.8Membrane Potential -31.3mV-30.6Data Compiled from: Buck, Land & Hochachka, 1993b; Buck et a!., 1993a;Buck & Hochachka, 1993; n.m.- not measurable.51mitochondrial membrane potential and exhausted glycogen stores (Andersson et al., 1987; Aw& Jones, 1989).Based on this data it appears that the turtle hepatocyte preparation represents an excellentmodel system for studying cellular control and energetics of metabolic suppression. Althoughthe cells are isolated and not subject to systemic signals, they still display the same responseto anoxia as whole liver in vivo: metabolic suppression, membrane stabilisation, glycogenolysisand glucose release. Also, since turtle hepatocytes demonstrate an ability to recovery from longperiods of anoxia they must also conform to the principal of preservation of cell function.Aims of the Research and Thesis Overview.Metabolic suppression requires the coordinated suppression of multiple energy consumingcellular pathways. Each pathway is important to a greater or lesser extent in the continuedsurvival of the cell but none is more important than protein synthesis and protein degradationin governing the phenotypic characteristics of the cell. Clearly, at the most basic level, thebalance of protein turnover and gene expression governs cellular function before, during andafter acute and chronic periods of dormancy. The aims of this research were therefore:(1) To calculate the energetic costs of protein synthesis and protein degradation duringnormoxia and anoxic metabolic suppression (Chapters two and three).(2) To determine the ATP-dependence of proteolysis and relative change in the stabilityof short and long lived protein pools as a result of reduced protein turnover (Chapter three).(3) To examine the expression and suppression of anoxia-specific proteins, and inparticular, whether an oxygen sensing mechanism is important in the control of this response52(Chapter four).(4) To determine the relative energetic costs and significance of protein turnover andgene expression relative to other cellular processes during metabolic suppression (Chapter five)Within these goals, a picture is presented of how a multifaceted cellular process such asprotein turnover (grossly comprised of gene activation, transcription, translation, stabilisationand numerous proteolytic pathways) is suppressed and controlled as part of a larger and highlycoordinated molecular, metabolic and physiological response to oxygen lack.53Chapter 2.Protein Synthesis During Anoxia and Recovery in Anoxia TolerantHepatocytes from the Western Painted Turtle Chrysemys picta bellii.Preface.This chapter appears as a paper published by S.C. Land, L.T. Buck and P.W. Hochachkain the American Journal of Physiology 265(R34): R41-R48, 1993. L.T. Buck developed themethod for isolating the hepatocytes. All aspects of the research were performed by myself.Introduction.The biosynthesis of proteins is a costly process requiring the hydrolysis of 4 ATPequivalents per peptide bond. In addition, a further cost of 1 ATP has been estimated to be theincidental cost of amino acid transport into the cell (Reeds eta!., 1985). Protein synthesis aloneaccounts for 18-26% of total energy production in various organisms and cell types (reviewedby Hawkins, 1991) and clearly consumes significantly more when the costs of mRNA synthesisand protein degradation are taken into account. In certain differentiating cell types, forexample, protein synthesis and degradation (i.e. protein turnover) account for 35% to 41 % ofthe total ATP utilization rate [Ehrlich ascites tumour cells and rabbit reticulocytes (Muller eta!., 1986 and Siems et a!., 1984 respectively)] with RNA synthesis accounting for a further11 % in the former case. For animals that depress metabolism in order to survive periods ofchronic anoxia, sustaining rates of protein turnover would be an expensive process. It wastherefore hypothesized that suppression of protein synthesis rates would account for a significant54proportion of the depression in metabolic rate under anoxia.Tolerance to hypoxia is well developed in the western painted turtle (Chiysemys pictabellii). The ability of this animal to withstand anoxic submergence is well characterised withforced dive survival times of up to 2 weeks at 18°C and 4-5 months at 3°C (Robin et a!., 1964;Ultsch & Jackson, 1982). A major adaptive strategy enabling survival of such a chronic periodof anoxia is the ability to coordinate the suppression of ATP utilizing processes, with slowedATP synthesis rates sustained by anaerobic glycolysis. The result is that overall ATP turnoveris suppressed while ATP concentrations remain steady (Hochachka, 1986; Kelly & Storey,1988). Hepatocytes isolated from this species show a similar response when exposed to anoxiaand have been demonstrated to maintain viability and energy charge for at least 10 hours underanoxia and cyanide administration at 25°C (Bucket al., 1993b). By comparison, rat hepatocytesunder cyanide lose up to 90% of the intracellular ATP pool after 30 mins of anoxia and displaya complete loss in viability after 2 hours (Anderson et al., 1987; Aw & Jones, 1989). As such,hepatocytes isolated from C. picta provide an excellent model for studies in anoxia toleranceand metabolic suppression.In order to assess the role of protein synthesis in this metabolic suppression, fractional ratesof protein synthesis during normoxia, anoxia and recovery were measured in painted turtlehepatocytes. From this an energy cost for protein synthesis was calculated under normoxic andanoxic conditions. Since protein synthesis rates are closely correlated with energy state throughalterations in the GTP:GDP ratio (Walton & Gill, 1976; Garcia-Esteller et a!., 1983), theresponse of the purine nucleotide pooi to anoxia was also investigated. Finally, rates of ureaproduction were measured to provide a window into the costs of amino acid and nucleotidecatabolism under normoxia and anoxia.55Materials and Methods.Chemicals. L-[2,6HJphenylalanine (specific activity (SA)= 56.OCi.mmol1), L[U’4C]serine (SA= l5OmCi.mmol’) and L-[U14Cjleucine (SA= 147Ci.mmol)were purchasedfrom Amersham Canada Ltd (Oakville, ON). HPLC gradeK2HPO4was from BDH Chemicals(Vancouver, BC). Minimum essential medium (MEM) amino acids were purchased from GibcoCanada (Burlington, ON). Electrophoresis reagents were purchased from Bio-Rad(Mississaugua, ON). All other chemical were purchased from Sigma Chemical Co (St Louis,MO).Animals. Adult painted turtles were obtained from Lemberger Co., Inc. (Oshkosh WI) andhoused in an outdoor tank equipped with a basking platform and heat lamps. Water temperaturewas maintained at an average of 22°C. Animals were fed chopped beef heart and liver ad liband exhibited excess visceral fat and stomach contents prior to hepatocyte isolation. All animalsused were females in the early stages of seasonal sexual maturation and were sampled over afive week period in mid summer.Solutions. Isolation media were as previously described (Buck et a!., 1993b). Briefly theywere as follows (in mM): Solution A: 78.5 NaC1, 34.5 NaHCO3,2.2 KC1, 0.9 Na2HPO4and10 Na-HEPES, pH7.5. Solution B: Solution A with 5.8 CaCl2, 3.8 MgCl2, 0.1 MgSO4 and2% BSA. Solution C: Solution A with 2U/ml Sigma protease XXIV. All isolation solutionswere saturated with 95%02:5%C.Primary culturing media were: i) Storage Medium: Solution B with 2% BSA (dialysedagainst water), MEM amino acids and 200U/ml penicillin. MEM amino acids werereconstituted according to the manufacturers recommendations and diluted 100 fold into thestorage medium. ii) Incubation Medium: Solution B with 2% dialysed BSA, MEM amino56acids (without phenylalanine), plus 0.6mM phenylalanine (phe) and 200U/ml penicillin. iii)Labelling medium: As for incubation medium but with 0.6mM non-radioactive phe and0.6Ci.mg cells-1 L[2,6HJ phe added separately, during the experiment. Each primaryculturing medium was saturated with 95%air:5%C02unless otherwise indicated. All culturingmedia were sterile filtered and treated aseptically.Hepatocyte Preparation. Hepatocytes were isolated by protease disaggregation aspreviously described (Buck et al., 1993b). Immediately after isolation, cells were placed intostorage medium, plated and kept overnight at 5°C. Contamination by non parenchymal cellswas always <2%. Adenylate energy charge (EC) was 0.9 lhr post isolation and increased to0.94 overnight. Prior to the experiment, cells were warmed to 25°C for lhr before beingplaced in the appropriate experimental incubation medium. Cell viability throughout theexperiment was assessed in terms of the adenylate energy charge as determined from parallelexperiments.Experimental Design. The experiments were conducted in two parts. In part A., eachhepatocyte preparation was divided into three distinct pools for the parallel assessment ofprotein synthesis parameters, rates of oxygen consumption and purine nucleotidephosphateprofiles. Cells from the protein synthesis pooi were further subdivided into two groups fortechnique validation and the measurement of fractional rates of protein synthesis. Part B.describes separate experiments, conducted to investigate rates of urea production and labellingof proteins during recovery from anoxia for 1 dimensional SDS polyacrylamide gelelectrophoresis (l-D SDS-PAGE). All determinations were made at 25°C.Pail A. - Rate of Protein Synthesis.Validation. To validate the conditions used to measure fractional rates of protein57synthesis, cells were incubated in normoxic incubation medium with the following alterations:i) 0.6mM non-radioactive phe + 30Ci:1mlL[2,6-3H]phe. ii) 2 Ci:’ml L-[U-14C]serine (ser),iii) 2Ci.ml1L-[U-’4C]ser + 1.2mM non-radioactive phe. Incorporation of amino acids intoprotein was assessed over the course of an hour by removing 5Ojl samples at 5, 20, 40 and 60minutes onto Whatman #3 filter discs. These were washed in l0%(w/v) TCA (5m1/filter) for20 minutes before being placed in fresh 5% TCA and heated to 100°C for 5mm. Finally, filterswere washed in 10% TCA at 20°C, then 95% ethanol and air dried. The radioactivity of theintracellular free pooi for condition i) was determined by centrifuging a 100ILl aliquot of cellsthrough a mix of dimethyl phthalate and dioctyl phthalate (1:5 v:v) into 10% TCA. Theintracellular protein was allowed to precipitate on ice for 30mm before being removed bycentrifugation. The resulting TCA soluble radioactivity was measured to give intracellular phefree pool activity. [‘4C]-ser and[3H1-phe counts were determined on a LKB Rack-beta 1214scintillation counter using ACSII (Amersham) scintillation cocktail with a counting efficiencyof 43.5% for[3H] and 95% efficiency for[14CJ. Condition i) above, represents the incorporationbehaviour of cells under the labelling conditions used to measure fractional rates of proteinsynthesis (see below) and groups ii) and iii) represent the effects of double the dosage of nonradiolabelled phe on rates of ser incorporation into protein.Incorporation of Isotope into Protein. Cells apportioned to the measurement of fractionalrates of protein synthesis were divided into the following experimental groups: Normoxic(NOR, 12h aerobic), anoxic (ANX, 12h anoxic), cycloheximide (CYC, 12h aerobic; 10molcycloheximide.mg packed cells1, administered 2hr prior to, and during, addition of isotope),1st recovery period (R-1, l2hr anoxic, lhr normoxic), 2nd recovery period (R-2, l2hr anoxic,2hr normoxic). For each experimental group, imi of cells at 5Omg.ml1,in duplicate, were58suspended in either air (95%air;5%C02 -aerobic) or nitrogen (95%N2;5%C0 - anoxic)saturated incubation media as appropriate and placed in sterile culture tubes. Each culture tubewas set up to maintain a pressure head of the appropriate, humidified gas mixture for theduration of the experiment as previously described (Buck et a!., 1993b). Cells were keptcontinuously suspended by rotation (— 1.5 cycles.sec1)using an IKA-Vibrax-VXR rotator(Janke & Kunkel Co., W. Germany).30mm before the end of the l2hr experimental incubation period, cells were removed fromincubation medium by gentle centrifugation (3Oxg for 2 minutes) and resuspended in the samevolume of labelling medium without isotope, saturated with the appropriate gas mixture.Labelling medium for the CYC group contained the same dosage of cycloheximide as duringthe incubation. At this time, recovery groups were also resuspended in normoxic labellingmedium. Cells were then allowed to recover from handling for 30mm before the addition ofisotope.At l2hr, 0.6 Ci,L-[2,6 3H] phe.mg cells1 was administered by injection though thechamber cap. Cells were allowed to incorporate isotope into protein under the appropriateexperimental condition for lhr. For the R-2 group, label was administered lhr post anoxicrecovery. Incorporation was stopped by centrifuging cells out of labelling medium (3Oxg, 2mm)and washing the cell pellet 3 times with BSA-free solution B to remove all extracellular labeland BSA (1-D SDS-PAGE confirmed that 3 washes in BSA-free solution B were required toremove all of the BSA from the cells). A relatively high dose of isotope was used in order tomaximise the amount of label incorporated into cycloheximide and anoxia treated cells. Anexcess of non-radioactive phe in the labelling medium was used to give linear rates of radiolabeluptake with a constant intracellular free phe pooi activity (fig 4).59Total cellular protein was precipitated by adding lml 5% perchioric acid (PCA) to cellpellets. Precipitates were allowed to form on ice for 30mm before centrifugation and removalof acid soluble radioactivity (intracellular free pool). The protein pellet was washed a further3 times in 2m1 5% PCA, then resuspended in 2.5m1 0.5M NaOH and incubated for lhr at 37°Cto re-solubilise the protein and hydrolyse RNA. The resulting solution was used to determineprotein content (BioRad protein assay kit) and total acid insoluble radioactivity. Alldeterminations of radioactivity were carried out by liquid scintillation spectrophotometry asdescribed above.In order to determine total RNA content and the specific activity of protein-incorporated[3H]-phe, protein dissolved in the remaining NaOH solution was re-precipitated with 0.444m120% PCA and pelleted by centrifugation. The supernatant was removed to measure RNAcontent using the dual wavelength procedure of Munro & Fleck (1969). The extinctioncoefficient for RNA was A2=32.9 with contaminating protein corrected as A232 =6.11 (Ashford& Pain, 1986). The remaining protein pellet was washed a further 3x with 2m1 5% PCA andthen hydrolysed in 6m1 6M HC1 for 48hr at 110°C. The HC1 was removed by evaporation andthe resulting amino acids were washed with water and again evaporated to dryness. Thehydrolysate was then dissolved in 2m1 0.5M Na-citrate (pH6.3).Determination of the specific activity of [3Hj-phe in the intracellular free pool andhydrolysate fractions was measured by the enzymatic conversion of phe to 13-phenylethylamine(13-PEA) as previously described (Garlick et al., 1980). We found that the formed 13-PEAproduct was light sensitive and as a result this step and all ensuing steps (including organicextraction) were performed in the dark. To imi sample, 3ml 3M NaOH and 8ml nheptane:chloroform (3: lv/v) were added and the sample mixed thoroughly. The NaOH was60removed by freezing and the 13-PEA was extracted from the organic layer by mixing thoroughlywith 0.O1MH2S04.After extraction, 0. imi of hydrolysates and lml of free pools were assayedfor [13-PEA] using the method of Suzuki & Yagi (1976). Briefly, this involved incubation ofthe samples with 10mM ninhydrin, 0.2mM L-leucyl-L-alanine and 0.5M potassium phosphate(pH8.0) at 60°C for 1 hour. After incubation, samples were cooled to room temperature andfluorescence measured at 495nm (excitation 390nm) on a Farrand Ratio Fluorometer-2 (FarrandOptical Co, NY). Since the light from the fluorometer caused a constant rate of decay of thefluorescent product, readings were standardized to lOs after exposing the sample to the lightsource.The fractional rate of protein synthesis (ks, % .hr’) was calculated as follows:k8 Bx.Q xlOO (Equ.2)SA twhere SB is the protein bound specific activity of phenylalanine (disintegrations.mirf’(DPM)nmol phe’), SA is the radioactivity of the intracellular free pooi (DPM.nmol phe1) andt is the length of time the cells were incubated with label (in minutes).Absolute rates of protein synthesis (mg protein synthesized.g cells’.hour1),used to computeATP turnover rates, were calculated as the fractional rate of protein synthesis multiplied by thetotal cellular protein content. Finally, the RNA:protein ratio (gRNA.mg proteint)was usedto express rates of protein synthesis relative to the tissue RNA content (kA, gProtein.mgRNA1.hour’) as follows:kA = lOk,. (Equ. 3)RNA:protein(Millward et al., 1973; Preedy et a!., 1988).Oxygen Consumption. Rates of oxygen consumption were measured polarographically61using Clark type electrodes (Yellow Springs, Ohio) in a 2ml thermostatted Gilson oxycell at25°C (Middleton, WI). Electrodes were calibrated using air equilibrated super-pure H20 at25°C, assuming an 02 solubility of 258.3mol02.dm3at 1 atmosphere (Forstner & Gnaiger,1983). Cultures in storage medium were equilibrated to 25°C and continually gassed with 95%air;5 % CO2 for 2hr prior to being placed in the oxycell at approximately 6mg. ml-’. To assessthe proportion of cycloheximide inhibitable oxygen consumption, 1Oj.mol cycloheximide.mg’cells was administered at a point after a linear rate of oxygen consumption had been obtainedunder control conditions. The concentration of cycloheximide used was based on the dosage atwhich a sustained maximal effect could be obtained immediately. All experiments were carriedout in duplicate.Extraction and Measurement of Purine Nucleotide Phosphates. Purine nucleotidephosphate profiles were measured in normoxic, anoxic and cycloheximide treated cultures fromexperiments conducted parallel to the protein synthesis group. The gassing and cycloheximideadministration procedure was the same with the exception that the cells were incubated in 25m1erlenmeyer flasks at an initial volume of 4m1 at 5Omg.m1’. PCA extracts were taken at 0,5 andlOhr for normoxic and anoxic cultures and at 2hr post administration for the cycloheximidetreated culture; each culture was sampled in duplicate. The procedure used to extract for purinenucleotides was as described (Buck et a!., 1993b). After extraction, samples were immediatelyfrozen in liquid N2 and stored at -80°C until required.ATP, ADP, AMP and IMP were measured using an LKB 2152 HPLC with a 7jLmAquapore AX300 ion exchange column (Brownlee Columns, Applied Biosystems mc, CA) aspreviously described (Schulte eta!., 1992). GTP,GDP and GMP were measured simultaneouslywith the adenylates by extending the run time to 36mm allowing the elution of GTP.62Patt B. - Urea Production. PCA extracts were taken from experiments conductedseparately from the studies in part A. Hepatocytes suspended in solution B. were incubated in25m1 erlenmeyer flasks under normoxia and anoxia as previously described (Buck et a!.,1993b). PCA extracts of the cell suspension were taken in duplicate at 0 2.5,5,7.5 and lOhrof incubation. Urea was measured in K2C03 neutralised extracts using the glutamatedehydrogenase linked assay (Bergmeyer, 1974). A rate of urea production was calculated fromthe steady state rate of urea accumulation over 5, 7.5 and lOhr under normoxic and anoxicconditions.In Vivo Labelling ofProteins During Recoveryfor Electrophoresis andAutoradiography.A separate series of experiments was conducted to investigate the 1 dimensional electrophoreticpattern of total cellular proteins exposed to l2hr normoxia and l-3hr recovery from 12h anoxia.Incubation procedures, prior to labelling, were as described above for the protein synthesis ratestudy with the exception that a third hour recovery group was also included. After l2hrincubation under the appropriate gassing regime, the normoxic and first hour recovery groupswere resuspended in normoxic storage medium without leucine (leu) but with 50Ci L[U14C]leu.ml’). At this time, second and third hour recovery groups were resuspended in non-radioactive normoxic storage medium. At 2 and 3hr into the recovery period, these groups weresuspended in medium containing L-[U14CJleu as described above. Each culture was allowed toincorporate label for lhr. At the end of the labelling period, each experimental group waswashed in BSA-free solution C x3 and the final cell pellet lysed by boiling for 10 minutes in2x loading buffer (100mM Tris base (pH6.8), 200mM dithiothreitol, 4% SDS, 0.2%bromophenol blue, 20% glycerol and 1mM phenylmethyonylsulfonyl fluoride (PMSF)). Theresulting slurry was sonicated for 2x 15s and centrifuged at l0000xg for 10 minutes. L63[U14C]leu was used as the labelling amino acid as it labelled the protein pool with the highestspecific activity when compared to [‘4C]-ser,[3H1-phe and[35S]-methionine.[14C]-leu labelled proteins were electrophoresed on a one dimensional gel using thediscontinuous buffer system of Laemmi (1970). Briefly, this employed a 5% acrylamidestacking gel and a 11.5% resolving gel. The running buffer consisted of 26mM Tris base (pHto 8.8 using HC1), 200mM glycine and 0.1% SDS. lx i0 trichioroacetic acid insoluble countsper minute, per sample, were loaded and the gel run overnight at 4°C using a constant amperageof lOmA. At the end of the run, gels were soaked for 30mm in Amplify fluor (Amersham),rinsed in distilled water and dried using a Bio-Rad gel drier. Flourographs were exposed for72hr on pre-flashed Kodak OMat X-AR film at -70°C. Molecular weights were determinedusing Bio-Rad low molecular weight range standards.Statistical Analysis. All data are presented as mean ± standard error with the number ofobservations indicated in each case. The value of n is defined as the number of independenthepatocyte preparations used in each series of experiments. Student’s t-test was used tocompare control and experimental groups with multiple groups compared using ANOVA withTukey’s HSD. Statistical significance is at p <0.05.Results.Validation for the Measurement of Fractional Rates of Protein Synthesis. In order toinvestigate whether the conditions used to measure fractional rates of protein synthesis reflectedan accurate rate of amino acid incorporation into cellular protein, a series of validationexperiments were carried out. For this to be an accurate technique, the intracellular free poolactivity should remain high and constant with rates of phe incorporation into protein linear over64the sampling period. In addition, administration of a flooding dose of phe should notartefactually affect rates of protein synthesis. Figure 4 demonstrates that the rate of L-[U‘4C]ser incorporation into protein was not affected by a two fold excess of non-radioactive phe.This suggests that the use of a high, flooding dose of phe did not affect the overall rate ofamino acid incorporation into protein. Under the conditions used to measure fractional ratesof protein synthesis, the rate of incorporation of L-[2,6-3HJphe into cellular protein remainedlinear over the lhr sampling period with the activity of the intracellular free pool remaininghigh and constant. Therefore, the conditions employed provide an accurate indication of therates of protein synthesis in turtle hepatocytes.Rates ofProtein Synthesis. Anoxia induced a fall in the fractional rate of protein synthesis(the percentage of the protein pooi that turns over per hour) by 92% of control values whichwas indistinguishable from the rate obtained with the inhibitor, cycloheximide (Fig. 5A).During normoxic recovery, k increased to 160% of control values and returned to normal after2hr. These changes in protein synthesis rates were not accompanied by any change in theRNA:protein ratio such that kA reflected the same pattern as the lç results from normoxia toanoxia and under cycloheximide administration (Figs. 5B and C). However, first hour recoverykA values were statistically insignificant from control groups.Specific ATP Requirement for Protein Synthesis. In order to investigate rates of ATPturnover for protein synthesis under normoxia and anoxia, it is necessary to calculate a specificATP requirement for the synthesis of each peptide bond. Reeds et a!. (1985) have suggestedthat the minimum theoretical cost for protein synthesis is 5 ATP equivalents per peptide bond.From this, a theoretical value of 46mmol ATP.g protein synthesized’ is arrived at, assumingan average amino acid molecular weight of 1 lOda. Table 2 demonstrates the route taken to65I Intracellular TCA Soluble Radioactivity4x1 + .: 2000 —.9 _;41500—Time (mm)FIGURE 4. Validation of experimental conditions used to measure fractional ratesof protein synthesis. 0.6mM non-radioactive phe + 30Ci:’m1 L[2,6-3H]phe(Closed circles) and the resulting intracellular TCA soluble radioactivity (closedsquares, error bars within symbols), 2 jCi:1ml L-[U-’4C]ser (open circles),2Ci. ml ‘L-[U-‘4C]ser + 1.2mM non-radioactive phe (open triangles). N = 3.Abbreviations: DPM, Disintegrations per minute; TCA, Trichioroacetic acid.0.15-• A0.09- i.2.E 0.06c) —.c 0 ‘0.03••O.OOr ‘7’.7.50.3. 0.20.1• *•0.0 I I I___NOR ANX CYC R-1 R-2FIGURE 5. Response of translation to anoxia and recovery. (A.) Fractional ratesof protein synthesis, (B.) RNA:Protein ratio and (C.) KA for l2hrs normoxia(NOR), l2hr anoxia (ANX), l2hr normoxic- final 2hr with 1Omol.mg’ cellscycloheximide (CYC), lhr post anoxic recovery (R-1) and 2hr post anoxicrecovery (R-2). N=5. **)<().(fl, *p.(O.05.67empirically determine the specific ATP requirement for protein synthesis from the present data.By using cycloheximide inhibitable 02 consumption to calculate a cellular rate of ATP turnoverfor protein synthesis and dividing this by the change in absolute rates of protein synthesis withcycloheximide, we were able to calculate a specific ATP requirement for protein synthesis of47.6 ± 6.8mmols ATP.g Protein1 Assuming an average amino acid molecular weight ofilOda, we arrived at 5.2 ± 0.9 ATP per peptide bond, close to the predicted theoretical value.Proportion of Metabolism Utilized by Protein Synthesis. Using the specific ATPrequirement for protein synthesis, we calculated protein synthesis ATP turnover rates fromabsolute rates of protein synthesis measured under normoxia and anoxia (Table 3). These wereexpressed as percentages of total ATP turnover, calculated from the rate of 02 consumption(normoxic total rate) and from the rate of lactate production [anoxic total rate; data from Bucket a!., (1993b)]. In addition, the extent of cycloheximide inhibitable 02 consumption was usedto provide an independent measurement of ATP turnover required for protein synthesis undernormoxic conditions. ATP turnover rates for protein synthesis fell 92% under anoxia, asexpected from the k values presented above. Despite the reduction in ATP turnover, theproportion of total metabolism required by protein synthesis remained statistically insignificantbetween normoxia and anoxia. The normoxic rate of ATP turnover calculated fromcycloheximide inhibitable 02 consumption was similar to that from the absolute protein synthesisrate (19.6 and 24.4molATP.g cells4.hr’, respectively). Under normoxic conditions, theproportion of ATP turnover for protein synthesis calculated from these routes was 28.1 to36.2%.Proportion ofMetabolism Utilized by Urea Production. Rates of urea production fell 72%from 0.51 ± 0.13 to 0.16 ± 0.O9mol urea.g cells’.hr’ under anoxia. Assuming a cost of 468TABLE 2. Calculation of a Specific ATP Requirement for Protein Synthesis fromCycloheximide Inhibitable Oxygen Consumption and Absolute Rates of ProteinSynthesis.Change in ATP turnover with cycloheximide.Control rate of 02 consumption 12.0 ± 1.3mol02.gcells’.hrtCycloheximide rate of 02 consumption 8.7 ± 1.2mol0.gcells1.hrtz 02 consumption 3.3 ± 0.3cmo102.gcells’.hr’Assuming a P.0 ratio of 3, then.ATP turnover withcycloheximide 19.6 ± 1. 7mo1ATP.g’ . hr1Change in absolute rates of protein synthesis with cycloheximide.Normoxic rate 0.53 ± 0.O8mgP.g cells4.hr’Cycloheximide rate 0.06 ± 0.O4mgP.g ce1ls.hr’Absolute rate of proteinsynthesis 0.47 ± 0.O8mgP.g cells1.hrtATP Requirement = 47.6 ± 6.8 mmolATP.g Protein1Assuming average amino acid molecular weight as liOda then:Specific ATP Requirement = 5.2 ± 0.9 ATP.Peptide Bond’Values are mean ± S.E., n=5. All determinations made under normoxic conditions.t ATP Requirement = LATP turnover with cycloheximideAbso1ute rate of protein synthesis with cycloheximide69TABLE 3. Percentage of Normoxic and Anoxic ATP Turnover Utilized byProtein Synthesis.Conditions Total ATP Absolute Rate of Protein Synthesis % of TotalTurnover Protein Synthesis ATP Turnover(mol.g cells1.hr’) (mgP.g cells’.hr1) (mol.g cells’.hrt)Normoxia 67.0 ± 0.53 ± 0.08 24.4 ± 3.9 36.2 ± 5.8(19.6 ± 1.7) (28.1 ± 2.1)Anoxia 6.3 ± 0.4* 0.03 ± 0.01 1.6 ± 0.5” 25.0 ± 7.3Values are mean ± S.E., n=5. *p<()() relative to Normoxic group.Calculated from rate of oxygen consumption assuming P:0 ratio of 3. Numbers inparentheses for the normoxic group indicate protein synthesis ATP turnover calculated fromcycloheximide inhibition of 02 consumption (Control and inhibited 02 consumption valuesgiven in Table 2).* Calculated from rate of lactate accumulation; data from Buck et al. ,(1993b).* Calculated from absolute rate of protein synthesis values assuming 46mmol ATP.gprotein’.70ATP per urea produced (Atkinson, 1992), an ATP turnover rate for urea synthesis wascalculated (Table 4). Under normoxic conditions, the proportion of metabolism utilized by ureasynthesis was 3 % of the total ATP turnover rate. In anoxia, this proportion remainedstatistically insignificant from the normoxic rate.Purine Nucleotide Phosphate Profiles. In order to investigate the relationship between therate of protein synthesis and energy state, adenine and guanine nucleotide phosphate profileswere measured during anoxia and under cycloheximide administration (Table 5). Neither [ATP]nor [GTP] changed significantly, however both [ADP] and [GDP] increased 2 fold up to 5hranoxia and remained constant to lOhr. As a result of the rise in [GDPJ the GTP:GDP ratiofell 3 fold by 5hr anoxia and remained constant to lOhr. The control nucleotidediphosphokinase mass action ratio (MAR) was calculated at 1.5 and this did not changesignificantly between normoxia and anoxia.To investigate whether there was any loss of purine nucleotides by deamination, inosinemonophosphate (IMP) concentrations were measured. [IMP] increased 4 fold over controls by5hr of anoxia, decreasing to 2 fold by lOhr. The elevation in [IMP] was not large enough toreflect a loss of adenylates or guanylates however; despite the [IMP] change the total purinenucleotide phosphate pooi remained statistically similar in all groups.Administration of cycloheximide did not alter purine nucleotide phosphate concentrationswith the exception of AMP and IMP, both being significantly elevated relative to control.Patterns of Protein Synthesis During Recovery. Figure 6 demonstrates the pattern ofradiolabel incorporation into normoxic and first, second and third hour recovery proteinsseparated by 1 dimensional SDS-PAGE. Two minor protein bands were detected duringrecovery at 44.5 and 36.3 kDa. However, expression of these protein bands alone could not71TABLE 4. Percentage of Normoxic and Anoxic ATP Turnover Utilized by UreaSynthesis.Conditions. Total ATP Turnover Urea Synthesis % of TotalATP Turnover(jmol. g cells-’. hr’) (mol. g cells-’. hr’)Normoxic 67.0 ± 4.3 2.0 ± 0.5 3.0 ± 0.7Anoxic 6.3 ± 0.4 0.6 ± 0.3* 10.3 ± 5.5Values are mean ± SE, n=6. *P <0.05 relative to normoxic group. ATP- turnover for ratesof urea synthesis were calculated assuming a cost of 4 moles of ATP per mole of urea(Atkinson, 1992). Rates were calculated from slopes between 5, 7.5 and lOhr time points.Total ATP turnover rates are as defined in Table 3.TABLE5.Purinenucleotidephosphateprofilesduringnormoxia,anoxiaandcycloheximideadministration.ConditionATPADPAMPGTPGDPGMPIMPNDPKaseGTP:GDPMARControl.0hr2.208±0.0310.268±0.0130.036±0.0100.237±0.0090.020±0.0030.015±0.0070.016±0.0031.51±0.2218.57±2.59Normoxia.5hr2.250±0.1340.235±0.0770.040±0.0130.193±0.0280.015±0.0030.012±0.0030.017±0.0031.68±0.5017.52±3.46102.314±0.1640.261±0.0520.034±0.0100.246±0.0230.018±0.0040.014±0.0020.016±0.0042.16±0.8320.05±2.23Anoxia.5hr1.740±0.0780.475±0.040*0.096±0.0460.193±0.0260.041±0.006*0.015±0.0020.067±0.007*1.63±0.416.33±1.79*102.008±0.3040.517±0.052*0.065±0.0230.281±0.0240.045±0.012*0.010±0.0020.034±0.002’4.01±1.746.62±1.86’Cycloheximide.2.136±0.0320.214±0.0320.117±0.032*0.289±0.0380.013±0.0010.012±0.0020.027±0.0062.45±0.8016.72±1.71Valuesaremeans±SEinmo1.g’cellwetweight.Valueofnineachcaseis7.*1) <0.05relativetoOhrcontrolpoint.All’,ADPandAMPdataareexpandedfromresultspreviouslypublishedinBucketal.,(7).Abbreviations:GTP,Guanosine5’triphosphate;GDP,Guanosine 5’diphosphate;GMP,Guanosine 5’monophosphate;IMP,Inosine 5’monophosphate;NDPKase,Nucleotidediphosphatekinase;MAR,massactionratio.NDPKaseMAR=LATPI.LGDPI/[ADPI.LGTPI.73FIGURE 6. Protein profile of recovery from 12h anoxia in whole turtle hepatocyte lysates.l-D SDS-PAGE autoradiogram of proteins labelled with [‘4C]-leu under normoxia (NOR) andfirst (R-l), second (R-2) and third (R-3) hour aerobic recovery from anoxia. Positions andweights of the Bio-Rad molecular weight markers are shown on the right.44.5—Nor R—1 R2 R—374Mr36.3—75account for the full extent of the recovery elevation in protein synthesis rates. It is thereforelikely that this is a generalised increase in protein expression of the total protein pool response.Discussion.A previous study reported a fall of 90% in turtle hepatocyte metabolic rate on entry intoanoxia (Buck et al., 1993b) which is similar to the 83% reduction in metabolism reported forthe whole animal during an anoxic dive (Herbert & Jackson, 1985). In this study, proteinsynthesis and urea production were investigated as single aspects of metabolism and it wasfound that the extent of anoxic depression (92 and 72%, respectively) was similar in magnitudeto the fall in total cellular metabolism. Furthermore, energy requirements for both proteinsynthesis and urea production remained at constant proportions of total ATP turnover fromnormoxia to anoxia. From this, it is concluded that anoxia results in a suppression of thesepathways that is coordinated with the fall in total cellular metabolism.The question remains as to how the depression in protein synthesis might be controlled.A number of studies have noted a correlation between basal k and the RNA:protein ratio(Houlihan et a!., 1990 and refs. therein) and this has been explained in terms of a highproportion of the RNA pooi being ribosomal (Young, 1970). If this is the case, theRNA:protein ratio serves as a gross measure of tissue ribosomal content. In this study, theobserved changes in k occurred independently of any alteration in the RNA:protein ratiosuggesting that acute, environmentally induced changes in k8 are not the result of changingrelative concentrations of the macromolecules required for protein synthesis. However, thisdoes not preclude a redistribution of the protein synthesis components. Previous studies haveindicated that hypoxia induces a breakdown of polysomes in rat liver (Surks & Berkowitz, 1971)76and a recent study on dormant Anemia cysts suggests that anoxia induces polysomaldisaggregation there also (Hoffman & Hand, 1992).A regulator of the state of polysomal aggregation is the GTP:GDP ratio. GDP is acompetitive inhibitor of GTP binding to several protein synthesis factors and has beendemonstrated to inhibit formation of the Met-tRNAf.GTP.eIF-2 initiation complex (Walton &Gill, 1976) and several elongation factors possessing GTPase activity (reviewed by Pall, 1985).The process of protein synthesis is therefore under sensitive feedback control with a fall in theGTP:GDP ratio resulting in an inhibition of polyribosome formation and peptide elongation inexisting polyribosomes. Furthermore, the guanylate pool is in equilibrium with the adenylatepool through the enzyme nucleoside diphosphate kinase (NDPKase). In this study, the fall inrates of protein synthesis under anoxia was accompanied by a 3 fold decrease in the GTP:GDPratio despite the maintenance of the ATP and GTP pools. The rise in nucleotide diphosphatesled to the speculation that changes in this ratio may be linked to inhibition of NDPKase asprevious studies in Anemia have suggested that rates of protein synthesis might be altered underanoxia by acidotic inhibition of NDPKase (Hoffman & Hand, 1992). However, this does notappear likely for turtle hepatocytes since i) the MAR for NDPKase (1.5) did not changesignificantly between normoxic, anoxic or cycloheximide groups, ii) NDPKase from rat liverhas a pH optimum between 6.5 and 9.5 (Kimura & Shimada, 1988) suggesting that this enzymeis active over a wide range of pH values and iii) pH is reported to fall gradually from 7.5 to6.8 at 20°C in whole turtle liver over 2hr of anoxic submergence (Wasser et a!., 1991).To explain the NDP accumulation within the cell, it is tentatively suggested that ATP andGTP are hydrolysed to new, lower steady state values during the first 5hr of anoxia. AlthoughNTP concentrations appear to remain steady during the anoxic incubation the standard deviation77is large enough to mask the small perturbation in [NTP] that would be required to account forthe observed increase in [NDP]. This may be coupled with an inhibition of the pathway forpurine catabolism. The total purine nucleotide pooi remains constant with accumulating IMPsuggesting that purine catabolism is inhibited past 5’-nucleotidase. This is supported by theobserved decrease in rates of urea production, the final end-product of amino acid and purinecatabolism, which is largely inhibited by anoxia (Table 3).During recovery from anoxia, fractional rates of protein synthesis rose to 160% ofnormoxic values before returning to normal after 2hr. Figure 6 shows that, other than thesynthesis of two minor protein bands, there were no major differences in normoxic andrecovery 1-dimensional SDS-PAGE patterns suggesting that this increase is not attributable tothe synthesis of specific sets of proteins but is a generalised response of total cell protein.Hypoxic recovery treatment in mammalian cell lines induces the synthesis of a set of stressproteins known as oxygen regulated proteins (ORP) or hypoxia associated proteins (HAP)(Heacock & Sutherland, 1986; Zimmermann et al., 1991). Synthesis of these proteins isthought to confer tolerance to repeated exposure to hypoxia and recovery in these cell types.In turtle hepatocytes there was no apparent up-regulation of proteins of similar molecular weighton recovery from anoxia suggesting that either these cells may constitutively expressORP/HAP’s or are not subject to the same stresses as mammalian cells during recovery fromanoxia.Exactly what leads to the increase in protein synthesis rates during recovery is unclear,however it is possible that this is part of an elevation in total cellular metabolism. Such anevent has been characterised during recovery from anoxia in marine intertidal organisms wheretotal oxygen consumption and metabolic heat production are elevated relative to pre-anoxic78levels [“oxygen debt repayment” (Shick et a!., 1988)]. Although this response is partiallyexplained in terms of the removal of anaerobic end-products, it is possible that there could bean anabolic component to this also.Empirical determinations of the number of ATP’s required per peptide bond tend to rangewidely in the literature with the closest determination to the theoretical estimate (5 ATP perpeptide bond) put at 7.5 ATP per peptide bond (68mmol ATP.gProtein synthesized1)(Aoyagiet a!., 1988). In this study it was assumed that cycloheximide does not affect aspects of energymetabolism aside from the inhibition of protein synthesis, that attenuation of protein synthesisis not accompanied by an increased rate of protein degradation and that the mitochondrialgenome contributes negligibly to whole cell protein synthesis (not inhibited by cycloheximide).Calculation of energy charge values from table 4 reveals that, with the exception of increased[AMP], cycloheximide administration did not perturb [ATP], [ADP] or the GTP:GDP ratio.Furthermore, based on low state 3 rates of mitochondrial oxidation and from qualitative electronmicroscopy, it appears that mitochondrial volume density is low in turtle liver tissue (AlamediaVal and Buck, unpubi. obs). An ATP requirement of 5.2 ± 0.9 ATP per peptide bond wascalculated which is close to the theoretical value of 5. From this, it was determined that proteinsynthesis constitutes 28-36% of the normoxic and 25% of the anoxic ATP utilization rate in thistype of tissue.The explanation as to why protein synthesis should account for such a high proportion ofATP turnover may lie in the state of sexual maturation of the animals from which thehepatocyte preparations are obtained. During seasonal gonadal development in oviparousvertebrates, the liver becomes increasingly involved in the de novo synthesis of vitellogenin.In fish, the result is that hepatocyte protein synthesis rates double over the course of79gonadogenesis (Haschmeyer & Mathews, 1983). In turtles therefore, it is likely that sexualmaturation in females is also accompanied by elevated hepatic protein synthesis rates.To conclude, this chapter has shown that metabolic suppression in turtle hepatocytes isassociated with the coordinated down-regulation of the pathways for both protein synthesis andurea production. The reduction in the GTP:GDP ratio, an event that is likely independent ofchanges in pH1, may play a role in the inhibition of protein synthesis through inhibiting theformation of polyribosomal complexes. Recovery from anoxia is associated with a transientincreased protein synthesis rates and this does not appear to be related to major changes inexpressed protein profiles. Such post-anoxic increases in protein synthesis rates may be aimedat clearing and replacing dysfunctional proteins formed during the anoxic bout or on recovery.80Chapter 3.Protein Turnover During Metabolic Suppression:Role and Energy Dependence of Proteolysis.Preface.The data presented in this chapter were published in a paper by S.C. Land and P.W.Hochachka (American Journal of Physiology (Cell Physiology 35): C1028-C1036, 1994. Allaspects of this work were performed by myself.Introduction.Entrance into a hypometabolic state in response to adverse environmental conditionsrequires two key conditions to be met: I) the reduction in rates of ATP synthesis must bebalanced and coordinated with reduced ATP demand such that ATP concentrations remainconstant (Hochachka, 1986), and ii) the functional capability of tissues must be maintainedduring the period of metabolic arrest so that on recovery, the tissue may resume activemetabolism unimpaired.Although there has been much interest placed on the mechanisms behind the coordinatedreduction in ATP turnover during metabolic arrest, little is known about the status of cellularfunctions that are responsible for preserving the functional stability of tissues. Of particularimportance to this question is the stability of the protein pool. As noted in cellular agingstudies, suppression of protein turnover rates increases the exposure of proteins to chemicalpost-translational modifications resulting in conformational changes and inactivation (reviewedby Reff, 1985). With reduced rates of removal by proteolysis, damaged proteins accumulate81eventually leading to the death of the tissue. Since dormancies can last from months to yearsin some cases, avoiding the occurrence or accumulation of damaged proteins is vital tosustaining tissue functional stability during chronic metabolic arrest. Therefore it washypothesised that there would be a significant component of remaining ATP turnover concernedwith the continued replacement and renewal of cellular protein.The experiments presented in chapter two demonstrated that during such a metabolicsuppression, turtle hepatocyte protein synthesis, and the ATP-turnover specific to this process,was reduced by 92% under anoxia. This occurred in a manner that was proportional to, andcoordinated with, the suppression of whole cell metabolic rate. The arrest of protein synthesiswas not complete, however, with remaining protein synthesis rates accounting for the sameproportion of total anoxic metabolism as under normoxic conditions (about 30%).If protein synthesis is sustained under anoxia, despite a large down regulation fromnormoxic rates, what is the role and cost of sustaining proteolysis during this time? Althoughpeptide bond hydrolysis is a thermodynamically favoured process, most proteolytic pathwayshave a functional requirement for metabolic energy either for maintenance of the proteolyticenvironment [eg, lysosomal ATP-dependent proton pump, (Schneider, 1981)] or activation ofco-factors [as in ATP-dependent ubiquitin conjugating enzymes, (Hershko, 1988)]. It is difficultto assess the relative costs incurred by each proteolytic pathway in vivo; however, previousstudies indicate that the total energy dependence of proteolysis may account for about 1-14%of total ATP consumption depending on the synthetic status of the tissue (Siems et a!., 1984;Muller et a!., 1986). When considered together with protein synthesis, the costs for turningover total cell protein can amount to as much as 40% of total normoxic ATP turnover rates inthe various differentiating cell types studied so far (Siems et a!., 1984; MUller et a!., 1986;82Schneider et al., 1990).Because of the costs involved in sustaining protein turnover, a balance must be reachedbetween the conflicting demands of energy conservation and the renewal of cellular protein.To investigate the role and energy dependence of protein degradation during anoxic metabolicsuppression, this study aims to 1) measure changes in protein degradation rates during exposureto anoxia and recovery for fast- and slow-turnover proteins, 2) calculate an energy budget forproteolysis and total protein turnover under normoxic and anoxic conditions and 3) measuredifferences in the proportion of coupling between labile and stable protein pools and ATPturnover.Materials and Methods.Chemicals. L[U-14Cjphenylalanine ([14Cjphe; specific activity (SA) = 479mCi. mmol’),L[2,6-3H]phenylalanine ([3H]phe; SA= 53Ci.mmol’) and L[4,5-3H]leucine ([3H]leu;SA= 147Ci.mmol’) were purchased from Amersham (Oakville, Ontario, Canada). Minimumessential medium (MEM) amino acids were from GIBCO (Burlington, Ontario, Canada). a2-Macroglobulin was purchased from Boehringer Mannheim (Laval, Quebec, Canada).Cycloheximide, emetine dihydrochioride, leupeptin, pepstatin-A, penicillin-streptomycin,iodoacetate, antimycin-A and all other bench chemicals were purchased from Sigma ChemicalCo, (St. Louis, MO).Primary Culture. Adult female western painted turtles weighing 300-500g were obtainedfrom Lemberger (Oshkosh, WI). Hepatocytes were isolated from females in the early stagesof sexual maturation as described in chapter two. After several cleaning stages, cells weresuspended in storage medium containing 4% bovine serum albumin (BSA) and 100U/ml83penicillin: lOJLg/ml streptomycin as modified from Land et al. (1993). 3m1 of cells, at a densityof 35mg.ml1, were then plated in sterile 35xlOmm Coming tissue culture dishes and thenmaintained at 25°C in a 5% CO2 humidified incubator. Under these conditions cells had firmlyadhered to the base of the culture dish within 12h of plating and LDH leakage remained lessthan 4% to 72h post-plating. Procedures for measuring cell viability through lactatedehydrogenase (LDH) leakage, glycogen content and adenylate concentrations between 0 and55h post-plating are previously described (Buck et al., 1993b).Measurement of Protein Degradation.Protein degradation rates were measured by following the rate of liberation of radioactivephenylalanine from prelabelled proteins in plated, static cultures of hepatocytes as described byBradley (1977). The method required prelabelling incubations with two radiolabels ([14C]pheand[3H]phe) to discern if there existed differences between protein degradation rates in culturespre-labelled to favour slow or rapidly turning over proteins (Vandenburgh & Kaufmann, 1980).Assessment of Conditionsfor Measuring Protein Degradation. Appropriate conditions forthe conduct of the protein degradation study were determined in the following pilot experiments.The minimum concentration of non-radioactive phe required to minimize isotope reincorporation into protein during the study was assessed in cultures labelled with 2Ci[14C]phefor 22h at 25°C. After washing cultures free of excess label, the release of[14C]phe into themedium was measured at 6h post-labelling in the presence of various concentrations of nonradioactive phe. From this, 1. 1mM non-radioactive phe was determined as the optimalminimum concentration required to prevent the reincorporation of isotope into cellular protein.To ensure that the use of 1. 1mM phe did not affect rates of cellular protein degradation,cultures were pre-labelled for 22h with 5JLCi[3H]leu and 2Ci [‘4C]phe. The rates of liberation84for each isotope were followed in the presence of 0.1 and 1. 1mM non-radioactive phe.Intracellular TCA-soluble radioactivity for [‘4C]phe labelled cultures was also followed in thepresence of 1. 1mM phe to confirm that there was no accumulation of radioactivity within cells.To ensure that there was no appreciable loss of[14C]phe label to[14C]02,oxidation rates weremeasured as described by French et al. (1981). Briefly, cells were incubated at 25°C for 1-3hin the presence of 1Ci [14C]phe. [‘4CJ02 that evolved during the incubation period wascollected on glass-fibre filters impregnated with 1M benzethonium hydroxide (hyaminehydroxide). The resulting radioactivity was quantified by liquid scintillation spectroscopy asdescribed below. Finally, radioactivity eluting with phenylalanine was assessed bychromatographic separation of amino acids on a Bio-Rad Rosil C-18 HL column (250 x 4.6mm,5m particle size) using a Waters 625 LC system (Toronto, Ontario, Canada) equipped witha Waters 995 photodiode array detector. Data were analyzed using Millennium vi. 1 software(Waters).Pre-labelling of Proteins. 3ml storage medium without phe but with 2Ci.ml1[‘4C]phewas added to each culture at 20h post-plating to label slower turning over proteins. Cells wereallowed to incorporate label for a further 22h at which point the [‘4C]phe labelling medium wasremoved and each culture washed 5 times with 3ml, non-radioactive storage medium containing1.1mM excess phe (degradation medium).[14C]phe was removed from the more labile proteinpool by incubating the cultures for 2h in degradation medium at 25°C after which, each culturewas again washed as described above. At this point, rapidly turning over proteins were labelledby incubating cultures in 3ml storage medium without phe, containing 10Ci.mlt[3H]phe for30mm at 25°C. Cultures were then rinsed 5 times with 3ml degradation medium reducing [‘4C]and [3H1 radioactivity in the supernatant to a constant minimum number of counts.85Experimental Procedure. After the proteins had been pre-labelled, independent sets ofexperiments were conducted with cultures maintained under the following conditions: 1) Fourexperimental groups incubated under normoxia, anoxia, anoxia+ 1mM NaCN, 5h anoxia+5hnormoxic recovery. 2) Six experimental groups incubated under normoxic conditions withadditions of 0.5mM leupeptin, 0.1mM pepstatin-A, 1U.ml a2macroglobulin,leupeptin +pepstatin-A+a2macroglobulin together (same concentrations), 0.1mM cycloheximideand 0.5mM emetine. Titers of proteolytic inhibitors, cycloheximide and emetine, weremaintained by replacing medium after each sampling point with degradation medium containingthe appropriate concentration of each inhibitor; 3) Two experimental groups incubated undernormoxia+ 1mM antimycin-A and anoxia + 10mM iodoacetate. All cultures to be incubatedunder normoxic (95% air; 5% C02) or anoxic (95% N2;5 % C02) conditions were placed insealable plexiglass chambers with a positive pressure of the appropriate atmosphere maintainedthroughout the experiment. All cultures were incubated at 25°C on a rotary shaker (50 rpm)for the duration of each experiment.At 0.5, 1, 1.5, 2, 3, 5, 7 and lOh into the experiment, the entire 3m! of degradationmedium was removed and replaced with 3m! freshly gassed, and temperature equilibrated,medium containing the appropriate concentration of proteolytic or metabolic inhibitor asrequired. Total radioactivity was measured directly in a 1O0d aliquot of the removedsupernatant. To 0.9m1 of the remaining supematant, 0. lml 100% (vol/vol) trichioroacetic acid(TCA) was added, the sample vortexed and the precipitate allowed to form for 20h at 4°C.TCA insoluble material was removed by centrifugation for 30mm at 10,000xg and the TCAsoluble radioactivity determined in a 100d aliquot. After sampling the medium at the final timepoint, each culture was washed three times in degradation medium before being resuspended86in 2ml 10% (vol/vol) TCA. Cells were then sonicated on ice for 2x15s and the resultingmixture was also incubated for 20h at 4°C. After centrifugation, an aliquot was removed fromthe supernatant for the determination of intracellular TCA soluble radioactivity. The remainingprotein pellet was then washed a further two times in 10% TCA and finally dissolved in lml1M NaOH by vigorous shaking on a IKA-Vibrax-VXR rotator (Janke & Kunkel, Bremen,Germany) at 25°C for 2h. Radioactivity in each sample was determined by scintillationcounting in ACSII aqueous scintillation cocktail (Amersham, ON) using an LKB Rack-beta 1214scintillation counter set to dual mode for [‘4C] and [3H] spectroscopy (efficiency = 95 and43.5%, respectively).Total initial radioactivity incorporated into protein was determined as the sum of the totalintracellular radioactivity and the total radioactivity released into the medium. Percentradioactivity remaining at each time point was calculated as:Total Initial Radioactivity - Released TCA Soluble Radioactivity x 100 (Equ. 4)Total Initial RadioactivityHalf-lives of[3H]phe- and[14C]phe-labelled protein pools were calculated as:T½ = ln2/k (Equ. 5)Where k= the logarithmic rate constant for the protein fraction degraded per hour (taken fromthe slopes between 0.5-3h for[3H]phe-labelled proteins and 5-lOh for [‘4C]phe-labelledproteins).Measurement of Normoxic and Anoxic Proteolytic ATP Turnover. The method used tomeasure ATP requirement for proteolysis was essentially as described by Seims et a!. (1984).Normoxic ATP requirement was assessed by the polarographic measurement of oxygen87consumption rates. Cells suspended in degradation medium at a density of 8mg.mt’ were placedinto 2m1 Gilson oxycell chambers (Middleton, WI) equipped with Clark-type electrodes andthermostatted to 25°C. Data were collected and analyzed using Datacan Acquisition Systemsoftware (Sable Systems, UT) and criteria for electrode calibration were as described in chapter2. After obtaining normoxic control rates of oxygen consumption, 0.1mM cycloheximide wasadministered to inhibit protein synthesis followed by 0.1mM cycloheximide with 0.5mMemetine-HC1 to inhibit both protein synthesis and proteolysis (Siems et a!., 1984). The changein oxygen consumption rates was used to calculate the ATP-turnover associated with proteinsynthesis and protein degradation by assuming a P:O ratio of 3.The anoxic ATP requirement for proteolysis was determined in cell suspensions from thesame preparations by incubation under a humidified atmosphere of 95 %N2;5% CO2 for 6h in thepresence of anoxia only, anoxia + cycloheximide and anoxia + cycloheximide + emetine-HC1in the same concentrations as above. Duplicate samples were removed at 0,2h,4h and 6h andextracted in a 7% (v/v) final concentration of perchioric acid (PCA) and finally neutralised aspreviously described (Buck et a!., 1993b). Lactate was measured on a Perkin-Elmer Lambda2 UV-vis spectrophotometer using the lactate dehydrogenase and glutamate-pyruvatetransaminase linked assay described by Bergmeyer (1974). Conversion of lactate productionrates to ATP turnover assumed P:lactate ratio of 1.5 (from glycogen).Data Handling and Statistics. Statistical manipulations were performed using Systatversion 5 (Evanston, IL) software. The value of n indicates the number of individualhepatocyte preparations used in each experiment and all percentage values were arcsintransformed. Confidence limits for significance are at 95 % with the statistical test for each dataset outlined in the figure or table legend. All data are expressed as mean ± standard error.88Results.Culture Conditions. Table 6 lists cell viability criteria up to 55h postplating in culturesheld under normoxic experimental conditions. LDH leakage remained at about 2% throughoutand remained less than 4% to 72h. Metabolic competency based on adenylate concentrationsfrom 0 to 55h was judged to be excellent and glycogen content remained high, although therewas some loss from glucose mobilization and release (Buck et a!., 1993b). Nevertheless,remaining glycogen at 55h would have been enough to fuel a further 17.8h of glycolyticallysupported metabolism at 25°C [assuming 2 lactates/glucosyl unit, a rate of lactate accumulationof 6.6mol.g’.hr’ (this study) and a rate of glucose release of 17. lmol.g.hr (Buck et a!.,1993b)].Validity of the Technique. Since the liver is a major site of amino acid catabolism, it wasof concern to investigate whether phe was metabolised significantly by turtle hepatocytes.[‘4C]02liberation rates from [‘4C]phe were 23.7nCi [‘4C102.g wet weight’.hr4 (n=3)representing approximately 1.8% of the rate of lactate oxidation in the same cell type (Buck etal., 1993). However, it should be noted that this method provides only a qualitative estimateof rates of decarboxylation and oxidation of phe in vivo due to randomisation of label. Whenanalyzed by HPLC, 80.3% of[14C] radioactivity eluted with phe with a further 3.4% and 6.1%appearing at elution times corresponding to tyrosine (tyr) and tryptophan (trp) respectively (Fig7A). Therefore, despite some loss of label to[14C]02and unidentified metabolic intermediates,about 90% of label could be accounted for as amino acids.Criteria for deciding the appropriate conditions to be used for measuring protein breakdownare shown in figures 7B and C. Figure 7B demonstrates that re-incorporation of released isotopeinto proteins was minimal at concentrations of non-radioactive phe in excess of 1mM in the89degradation medium. In each degradation experiment, an excess of non-radioactive phe to1. 1mM was therefore used to minimise isotope reutilisation. Figure 7C demonstrates that theuse of an excess of 1. 1mM phe did not alter the rate of liberation of[3H]leu indicating thatprotein degradation rates were not affected by the use of excess phe. Furthermore, there wasno intracellular accumulation of label indicating that label liberated from proteins equilibratedrapidly to the outside of the cell when in the presence of 1.1mM phe. Values for slopesregressed through each line are given in the figure legend.Protein Degradation Rates in13H]phe- and[‘4C]phe-labelled Proteins. The intermittentperfusion technique has previously been shown to be an effective method for the measurementof protein degradation in labile and stable protein pools from several cell types (Bradley, 1977;Vandenburgh & Kauffman, 1980) including rat hepatocytes (Hopgood, 1977). In this study,we were able to confirm the presence of both fast-turnover([3H]phe-labelled) and slow-turnover[14C]phe-labelled) protein pools. Figure 8 demonstrates the rate of[3HJphe and[14C]pheliberation from prelabelled proteins as a function of time for cells held under normoxicconditions. Slopes for the two curves from 0.5 to 3h were significantly different by analysisof covariance (p < 0.05, n = 9) at -2.519 and -1.582 % . h’ for[3H]phe and[14C]phe liberationrespectively. However, from 3-lOh,[3Hjphe liberation rates had slowed and matched[14CJpheliberation rates during this time. Rates of[14C]phe liberation did not differ significantlybetween 0.5-3h and 3-lOh.Effect of Anoxia and Recovery on Protein Half-hfe. By measuring the rate of isotopeliberation from pre-labelled protein (i.e. protein degradation rate) it was possible to calculateprotein half-lives for both [3H] and[‘4Cjphe-labelled proteins. The protein half-lives reportedreflect the time for 50% of protein to be degraded at the degradation rate measured over the90TABLE 6. Viability data for plated cells under primary culture conditions at 25°Cform 0 to 55 hours post-plating.Criterion Hours Post Plating0 55LDH Leakage’ 2.12 ± 0.61(5) 1.87 ± 0.54(5)(72h = 2.53 (3))Glycogen2 664.0 ± 57.7(6) 540.1 ± 44.6(6)ATP3 1.77 ± 0.17(5) 2.16 ± 0.23(5)ADP3 0.30 ± 0.02(5) 0.16 ± 0.007(5)AMP3 ND 0.055 ± 0.005(2)Values are mean ± SEM, with the value of n for each group given in parentheses.1 % of total cellular LDH activity in the extracellular medium.2 Values expressed as mol glucosyl units.g wet weight’.Values expressed as mol.g wet weight’.ND, not detectable.91FIGURE 7. Validation of experimental conditions used to measure proteolytic rates fromradiolabel release. A) Separation of radioactivity eluting with phe by HLPC. Broken lines:elution of [‘4C] with [‘4C]phe radioactive standard; Solid lines: elution of [14C1 radioactivityfrom deproteinised extracellular medium. Elution times for phe, tyrosine (Tyr) and tryptophan(trp) non-radioactive standards are shown. Results are the means of 3 samples. B) Release ofTCA soluble[14C]phe from prelabelled protein at 6h in the presence of increasing concentrationsof non-radioactive phe (means only, n=3). C) Release of TCA soluble radioactivity expressedas a percentage of total radioactivity. Closed triangles: cells pre-labelled with[14C]phe, releaseof[14C]phe followed in the presence of 0. 1mM phe (Slope=0.863±0.05,r2=0.964); Opentriangles: as above but in the presence of 1. 1mM phe (Slope =0.971 ±0.08,r2=0.927); Closedcircles, cells incubated in the presence of[3HJleu, release of[3H]leu followed in the presenceof 0.1mM phe (Slope=0.916±0.14, i9=0.761); Open circles: as above but release of labelfollowed in the presence of 1. 1mM phe (Slope=0.827±0. 10, i9=0.835); Open diamonds: cellspre-labelled with [‘4C]phe; intracellular TCA soluble radioactivity followed in the presence of1.1mM phe. Values are means ± SEM, n =4 and slopes were compared using analysis ofcovariance.3 QoOIsotopeLiberated(%ofTotalRadioactivity)Ui0UiIsotopeLiberatedIn6hr.(%ofTotalRadioactivity),-‘01.è.0%0II%TotalRadioactivitym L4 •600000C 10%93100-.o-.S. —4 6Time (Hrs)FIGURE 8. Isotope liberation from[3Hlphe-labelled (closed circles) and[‘4C]phe-labelled(open circles) protein pools expressed as percent of total radioactivity. Dotted transect linesindicate the O.5-3h sampling period used for[3HJphe-labelled proteins. Values are mean ±SEM, n=9. Data are plotted in an arithmetic relationship to emphasise differences betweenrates of fast- and slow-turnover protein pooi radiolabel washout.2 8 1094sampling period. This assumes that the rates of isotope liberation accurately reflect rates ofprotein degradation, in vivo which we deem reasonable based on the technique validation dataabove.Rate constants, half-lives and percentage suppression in response to anoxia and recoveryare shown in Table 7. In all treatments with the exception of recovery, the rate of isotopeliberation from[3H]phe-labelled protein was significantly different from the corresponding[‘4CJphe-labelled protein degradation rate (P <0.01 by analysis of covariance) confirming thepresence of labile and stable proteins under normoxia and anoxia. Anoxia and cyanideadministration resulted in an approximate doubling of protein half-lives for both the[3HJphe-and[‘4C]phe-labelled protein pools (Table 7). The percentage suppression of proteolytic rate(calculated from the difference between normoxic and anoxic rates of isotope release) was 30-39% for[3H]phe-labelled proteins but was more pronounced in[14C]phe-labelled proteins atvalues approaching 50%. The total mean suppression of proteolytic rate (the summedsuppression for both[3H]phe- and[14C]phe-labelled protein pools) was 35-41 %.To compare values for recovery half-lives in the[3HJphe-labelled protein pool, recovery[3HJphe release rates were compared to control release rates during the 5-lOh sampling period.[14C]phe-labelled protein recovery rates were compared as before. Protein half-lives in[3H]phe-labelled protein pooi were significantly extended relative to control with proteolytic ratessuppressed by 43%. However, in the[‘4CJphe-labelled protein pool, there were no significantchanges in half-life or proteolytic activity during recovery from anoxia.Proportion ofATP-dependent andATP-independentproteolysis. For the purposes of thisstudy, ATP-dependent proteolysis is defined as the part of protein degradation that requiresATP hydrolysis for continued activity. ATP-independent proteolysis is defined as that portionTAHLE7.Logarithmicrateconstants,half-livesandsuppressionof proteolyticactivityin[3HJpheand[‘4CIphe-labellJproteinsasinfluencedbyanoxiaandrecovery.ConditionI’HlPbeLabelledProteinsI’4ClPheLabelledProteinsTotalk’Half-Life%Suppression’Half-life%Suppression%Suppression’(xlO2)(hr)(1102)(hi)Normoxia3.07±05324.7±3.31.27±0.0855.6±3.5—Anoxia2.10±0.2034.4±3.T30.8±5.20.64±0.04109.6±7.444.2±2.235.7±3.8Anoxia1.92±0.4245.2±lI.838.6±4.70.59±0.10l31.7±22.747.5±3.241.0±4.1+NaCNRecovery1.80±0.304l.7±5.243.4±3.51.30±0.1658.1±8.7-10.3±8.739.8±4.8Valuesaremean±SEM, n=5ineachcase.SignificancewasassessedrelativetocontrolusingANOVAwithTukey’sHSD.P<0.05,_p<0.01,P<0.01asdeterminedbyStudent’st-testrelativetocontroldegradationslopesfromthe5-bItperiod.k=Logarithmicrateconstant(Log%.hr’)‘Calculatedasthe%differencecomparedtonormoxicdegradationslopes.‘Calculatedas%differencebetweensummedPHiphe-labelledand(‘4C)phe-Iabelledproteinslopesforeachgroupversusnormoxiccontrol.096of protein degradation that does not require the hydrolysis of ATP and demonstrates continuedactivity under administration of metabolic inhibitors. Figures 9A and B demonstrate the extentof proteolytic inhibition by administration of inhibitors of energy metabolism. For [Hjphelabelled proteins, antimycin-A and iodoacetate produced no statistical differences in rates ofisotope liberation from normoxia to anoxia. However, for[14C]phe-labelled proteins additionof antimycin-A under normoxic conditions resulted in a 51.6±5.6% inhibition of proteolysisrelative to the control slope. Administration of iodoacetate under anoxic conditions resulted ina further 24.0± 12.0% inhibition of proteolysis. From this it appears that about 82% ofproteolysis in turtle hepatocytes is ATP-dependent.Energetic Cost of Proteolysis. Figure 10 demonstrates lactate accumulation rates undercycloheximide alone, and cycloheximide with emetine-HCI administration over a 6 hour timecourse. Incubation with each inhibitor produced a statistically significant reduction in the rateof lactate production compared to that under anoxia alone. From this, anoxic ATP turnoverrates for protein synthesis, protein degradation and protein turnover were calculated and areshown, together with normoxic ATP turnover rates (calculated from rates of oxygenconsumption) in Table 8. Justification for the use of cycloheximide and emetine-HC1 to achievedifferential inhibition of protein synthesis and protein degradation are discussed below.Total Al? turnover was suppressed by 87% on exposure to anoxia with Al? turnoverspecific to protein synthesis (cycloheximide inhibitable metabolism) and proteolysis(cycloheximide + emetine inhibitable metabolism) falling by similar proportions of 88 and 93%respectively. Under both normoxic and anoxic conditions, Al? turnover for proteinsynthesis remained at the same proportion of total metabolic rate (— 33%), supporting theresults from chapter 2. However, the proportion of total ATP turnover utilised by Al?-100 97.1 II904;_______________Time (fir)Time (fir)FIGURE 9. Response of proteolysis to various metabolic inhibitors. A) [‘H]phe-labdlledprotein pool. Release of TCA soluble[3HJphe was followed in the presence of normoxia (closedcircles), normoxia + 1mM antimycin A (closed squares), anoxia + 10mM iodoacetate (opendiamonds). B)[‘4C]Phe-labelled protein pool. Release of TCA soluble [‘4C]phe was followedas detailed for figure 2A). Values are mean ± SEM, and are shown plotted as a semi-logarithmic relationship (n=5 in each case). Comparisons between slopes were by analysis ofcovariance.9850-.f4o.I 4Time (Hr)FIGURE 10. Rates of lactate production under anoxia (open circles), anoxia + 0.1mMcycloheximide (closed circles, P <0.01) and anoxia + 0.1mM cycloheximide + 0.5mMemetine-HC1 (closed squares, P <0.01). Values are mean ± SEM, n =5. Slopes ofexperimental plots compared to control (anoxia only) by analysis of covariance.99TABLE 8. Cost of protein synthesis and ATP-dependent proteolysis undernormoxic and anoxic conditions as determined by stepwise inhibition of proteinturnover using cycloheximide and emetine.Condition ATP Turnover % of TotalTotal Cyc Cyc + Eme Protein Proteolysis4 Protein ProteolysisInhibitable Synthesis3 SynthesisNormoxia’ 80.3±9.4 56.9±10.6’ 45.8±9.5 23.4±5.3 11.1±1.7 33.5±5.9 21.8±1.4Anoxia2 9.95±0.31 7.36±0.93 7.08±0.99 2.79±0.79 0.73±0.43 32.9±4.8 12.4±7.8% Reduction 87.6 88.1 93.4ATP turnover values are expressed as mol ATP.g’.hr1 and are means ± SEM, n=5.Significance was assessed relative to control using ANOVA with Tukey’s HSD, ‘P <0.05“P <0.01. 1 Calculated from cycloheximide and cycloheximide + emetine inhibitable oxygenconsumption assuming a P:O ratio of 32 Calculated from the rate of lactate accumulation (fig.4) under cycloheximide and cycloheximide + emetine administration assuming 1.5 moles ofATP per mole of lactate produced. Calculated as the difference between total ATP turnoverand cycloheximide inhibitable ATP turnover. “ Calculated as: Total ATP turnover -(cycloheximide ATP turnover- cycloheximide+ emetine ATP turnover).100dependent proteolysis was disproportionately reduced from 22 to 12% under anoxia.Effect of Cycloheximide, Emetine-HC1 and Protease Inhibitors. The translation inhibitors,cycloheximide and emetine-HC1, have been previously used to achieve a concentration-dependent inhibition of protein synthesis and protein turnover (Siems et al., 1984; MUller et al.,1986). It was of concern here to ensure the conditions employed resulted in a total inhibitionof protein synthesis rates and also ATP-dependent proteolysis to obtain protein turnover ATPrequirements. As demonstrated in Figure 11, cycloheximide had no significant effect on proteinhalf-lives in either protein pool at the concentrations employed (0. 1mM) but was found toproduce a total inhibition of protein synthesis rates (data not shown). Emetine-HC1 administeredat 0.5mM resulted in a maximal extension of protein half-lives for[14C]phe-labelled proteinswhich was similar in magnitude to the extension of half-life produced by the inhibitorsleupeptin, pepstatin and2macroglobulin summed together. No effect of emetine-HC1 wasevident on[3Hjphe-labelled proteins. Since most of the ATP-dependent proteolytic activityappears amongst[14C]phe-labelled proteins (Fig 9), we conclude that the administration of0.5mM emetine-HC1 was sufficient to inhibit all ATP-dependent proteolysis.The proteolysis inhibitors leupeptin and pepstatin-A have previously been shown to beeffective inhibitors of protein degradation rates in rat hepatocytes (Dean, 1975; Hopgood,1977). At the concentrations employed in this study it was found that these inhibitors produceda rapid and maximal, sustained inhibition of proteolysis for the duration of the experiment.Figure 11 demonstrates the effects of various proteolytic inhibitors on both rH]phe- and[‘4CJphe-labelled protein pools under normoxic conditions. The serine protease inhibitor,leupeptin, and acid protease inhibitor, pepstatin-A increased the turnover time for[14C]phe-labelled proteins significantly although leupeptin had little effect on the turnover of101___C-Labelled Proteins3H-Labelled Proteins150—• **125I.——I ©75.— *•— 50FIGURE 11. Effect of various protease inhibitors and protein synthesis inhibitors on H]pheand[‘4C]phe-labelled protein half-lives. Leu: 0. 1mM leupeptin, Pep: 0.1mM Pepstatin-A,a2Mg: 0.5U.m142Macroglobulin, LPM: Leupeptin+Pepstatin+a2Macroglobulin (sameconcentrations), Cyc: 0. 1mM Cycloheximide, Eme: 0.5mM Emetine hydrochloride. Values areMeans ± SEM, n=5. Values were compared using ANOVA with Tukey’s HSD. P<0.05,..P<0.001.102[3Hlphe-labelled proteins. The endoprotease,cr2macroglobulin, had no significant effect oneither pool. Addition of leupeptin, pepstatin-A and2macroglobulin together more than doubledthe turnover time for[‘4C]phe-labelled proteins, but turnover times for[3H]phe-labelled proteinswas affected no more than for pepstatin-A administration alone.Discussion.Anoxic submersion in Chrysemys picta is accompanied by a 83% suppression in metabolicrate (Herbert & Jackson, 1985). This reduction in metabolism has been well characterised inisolated hepatocytes where ATP turnover rates fall on the order of 88-90% during anoxia (Bucket at., 1993a&b, chapter 2 and this chapter) with coordinate and proportional reduction inturnover rates measured for Na/KATPase (Buck & Hochachka, 1993), protein synthesis andurea synthesis (Chapter 2). A significant finding from these parallel studies is thatNa4/KATP se activity accounts for the principal proportion of remaining metabolic rate undçranoxia (Buck & Hochachka, 1993).The current study set out to determine the energy requirements of protein degradation andfrom this, estimate the proportion of metabolism that is required to sustain remaining proteinturnover during anoxic metabolic arrest. From the present data it is clear that proteindegradation is an energetically expensive process that accounts for 22% and 12% of total ATPturnover under normoxic and anoxic conditions respectively. When the anoxic ATP turnoverrates for protein synthesis and protein degradation are summed together, we can account for45% of total ATP turnover supporting protein turnover during anoxic metabolic arrest. Giventhat protein turnover accounts for 45%, and Na/K ATPase, 74% (Buck & Hochachka, 1993),of total anoxic metabolic rate in turtle hepatocytes, it appears that we can account for over103100% of anoxic ATP metabolism. Some of this overestimation can be attributed to the use ofdifferent culture conditions and seasonal effects on protein turnover (Chapter 2). Nevertheless,these data indicate that during anoxic metabolic arrest, the primary energy expenditure isdirected towards the maintenance of a constant membrane potential with a substantial secondaryexpenditure concerned with the turnover of cellular protein.Normoxic protein half-lives for[3H]phe- and[‘4CJphe-labelled proteins were comparableto those previously measured for mammalian cell lines (Bradley, 1977; Vandenburgh &Kaufmann, 1980). However, there was a doubling of total protein half-life within the first lOhof anoxic metabolic arrest. This is in agreement with previous studies on the brine shrimpembryo (Artemia fransiscana) where chronic anoxic metabolic arrest resulted in a 77 foldextension of half-life for cytochrome c oxidase (Anchordoguy et al., 1993). In terms ofincreasing survival time, extending protein half-life appears paradoxical however. As noted incellular aging studies, the probability of protein structural damage by post-translational andchemical modifications increases significantly with time (reviewed by Reff, 1985). Therefore,if tissues are to retain overall functional stability during long-term metabolic arrest, the presenceof a stabilising mechanism is implied. As a means of achieving protein replacement andrenewal, the maintenance of reduced protein turnover during anoxic metabolic arrestdemonstrated in this study possibly represents an initial line of defense. It would be of interestto know if this is complemented by other innate mechanisms that stabilise proteins such asamino acid sequence features that do not predispose proteins to degradation (eg. Bachmair eta!., 1986; Rogers et a!., 1986) or the synthesis of stress proteins such as glucose regulatedprotein 78 (GRP78 or Binding Protein) that have been noted to stabilise nascent protein subunitsduring transport to the golgi body in other cell types (Hendershot et a!., 1988).104Although limited protein synthesis and degradation are maintained under anoxia, the currentdata suggest that they are not evenly matched. Here and in chapter 2, it was reported thatprotein synthesis rates were reduced by 88-92% during exposure to anoxia. Here, the totalmean proteolytic suppression for[3HJphe- and[‘4C]phe-labelled protein pools under anoxia was36-41% of control values with both protein pools exhibiting different individual magnitudes ofsuppression. The smaller suppression of total protein degradation rates indicates that underanoxia, a negative protein balance exists within the cell. Whether this negative protein balanceplays a part in setting the limits for maximum survival at 25°C remains unclear at this point.An imbalance between protein synthesis and degradation is also observed during recovery.In chapter 2, it was noted that normoxic recovery from anoxia resulted in a 160% increase inprotein synthesis rates compared to control values. However, proteolysis did not appear toshow the same response since proteolytic rates in the[3H]phe-labelled protein pool were lowerin recovery compared to controls by nearly 50%. The previously observed exaggeration inrecovery protein synthesis rates may partly be the result of inhibited proteolysis in the morelabile protein pool. Nevertheless, it appears that normoxic recovery results in a positive proteinbalance that may, in part, compensate for the negative protein balance that occurs upon entryinto anoxia.The basis for the observed imbalance in protein turnover during anoxia may be related tothe degree of coupling between proteolysis and ATP hydrolysis. A particularly strikingobservation from the data is the magnitude of the depression of ATP-dependent proteolysis(93%; Table 8) which closely matches the proportional suppression found for whole cellmetabolism and protein synthesis. Since total proteolytic suppression was on the order of 36-41 %, it appears that remaining proteolysis under anoxia is largely comprised of ATP105independent pathways which may be less responsive to alterations in cellular metabolism andtherefore could persist during the metabolic suppression. This implies that there is a shift inthe control of proteolysis away from ATP-coupled metabolism during the transition fromnormoxia to anoxia. With the majority of ATP-dependent proteolysis residing in the stable[14C]phe-labelled protein pool (fig 9), it appears that the negative protein balance under anoxiais largely attributable to loss of protein from more labile proteins, an important observationsince the majority of regulatory proteins possess short half-lives (Hargrove & Schmidt, 1989).Mechanisms resulting in the suppression of energy-dependent proteolysis are likely to bemany-fold depending on the dominant proteolytic pathway involved. However, a key featureof anoxic metabolic suppression in both turtle hepatocytes and the whole animal is that ATPdemand is coordinately down-regulated with ATP supply resulting in high and constantintracellular ATP concentrations (Chapter 2; Kelly & Storey, 1988; Buck et a!., 1993b; chapter2). Therefore, despite observations in rat hepatocytes that have demonstrated a direct positiverelationship between ATP concentrations and proteolytic rates (Gronostaj ski et al., 1985; Plompet a!., 1987) it appears that regulation by absolute changes in [ATP] alone is unlikely in thiscase. Other factors that may be involved in the suppression of energy-dependent proteolysismay include metaboiltes such as 2,3-bisphosphoglycerate [inhibits ATP dependent lysosomalproteolysis (Roche eta!., 1987)], increased expression of protease inhibitors, as has been shownfor mammalian hibernators (Srere et a!., 1992) and availability and successful targeting ofproteolytic substrates. Of particular significance to this final point is the observation thatphosphorylation-mediated changes in protein conformation alters protein susceptibility to Ca2-activated proteases (Chuah & Pallen, 1989; Greenwood et al., 1993). With the current evidencein turtle tissues for phosphorylation control of enzyme activities (reviewed by Storey & Storey,1061990) and the postulated role for phosphorylation in modulating ion channel and pump activities(Buck et al., 1993), control of protein stability or protease activity by such a mechanism shouldnot be ignored.In order to understand the nature of proteolytic suppression and activation it is importantto know which pathways are active and when. Although a number of studies have suggestedthat the bulk of cellular protein, including regulatory, denatured, short, and some long livedproteins are degraded by cytosolic ATP and ubiquitin dependent processes (Goldberg & Rock,1992), in liver, protein degradation by autophagic vacuole formation appears to be thepredominant proteolytic pathway (Mortimore & Kurana, 1990). The protease inhibitor pepstatinA inhibits lysosomal acid proteases such as cathepsin D with little or no effect on serine andthiol proteases. As is clear in figure 11, under pepstatin-A administration, protein half-livesfor both the[3Hjphe- and [‘4C]phe-labelled protein pools were extended by about 25%, arelatively small proportion of total cell proteolysis. Leupeptin, which inhibits a broad range ofthiol and serine proteases, resulted in a 50% increase in half-life of the[‘4C]phe-labelled proteinpool and, when summed together with pepstatin-A and2macroglobulin, doubled slow-turnovernormoxic proteolytic rates with minor, but significant effects on the[3H]phe-labelled proteinpool. This provides some initial evidence that serine, thiol and acid proteases constitute a largeproportion of total cell proteolytic activity in this type of cell.A final note concerns the fate of amino acids that are liberated by proteolysis duringanoxia. Previous studies have noted that hepatic phosphoenolpyruvate carboxykinase isexclusively mitochondrial in this, and other turtle species (Buck ec a!., 1993b; Land &Hochachka, 1993). It therefore appears unlikely that liberated amino acids play a major role inthe repletion of hepatic glycogen stores on recovery. Instead, it seems probable that the bulk107of liberated amino acids are either oxidised or reincorporated into protein as part of theobserved exaggeration in protein synthesis rates during re-oxygenation.In conclusion, evidence is presented that, despite an increase in total cellular protein half-life, a limited turnover of protein is maintained during anoxic metabolic arrest in painted turtlehepatocytes. The energetic costs to sustain this amount to approximately 45% of remainingATP turnover. The data presented on protein degradation support the coordinate suppressionof ATP-dependent proteolysis together with cellular energy metabolism but expose continuedenergy-independent proteolysis amongst the more labile[3H]phe-labelled proteins.108Chapter 4.A Heme-protein Based Oxygen Sensing Mechanism Controls theExpression and Suppression of Multiple Proteins inAnoxia-tolerant Turtle Hepatocytes.Preface.The data presented in this chapter are in a manuscript by S.C. Land and P.W. Hochachka.which is currently in submission stages to The Proceedings of the National Academy ofSciences, U.S.A.. All aspects of the work were performed by myself.Introduction.In the absence of oxygen, numerous species of facultative anaerobes suppress theirmetabolism to survive prolonged periods without oxygen. Intrinsic to this response is acoordinated re-organisation of ATP supply and demand that enables tissues to retain viabilityand function, in hand with slow rates of flux through all cell processes.The partitioning of energy demand and re-organisation of metabolism during metabolicsuppression has been examined in hepatocytes isolated from a successful vertebrate anaerobe,the western painted turtle [(Chrysenzys p1cm bellii (Buck et al., 1993b)]. On exposure toanoxia, these cells mount a coordinated physiological response, similar to that characterised inwhole liver, that is directed towards: 1) a 10-fold reduction in rates of ATP synthesis and ATPdemand, ii) conservation of the cellular membrane potential by concurrent reductions in ionchannel flux rates and ion pump ATPase activity, iii) reduced Al? demand by protein turnover,109urea synthesis and gluconeogenesis and, iv) maintenance of functional viability during the periodof metabolic suppression (Buck et at., 1993a&b, Buck & Hochachka, 1993, chapters 2 and 3).A notable feature of this metabolic re-organisation is that it occurs independently of changein ATP concentrations or cellular energy charge. Yet the changes in ATP demand for individualcellular processes are clearly coordinated with one-another and exhibit rate reductions inproportion to the depression of total ATP turnover (Buck et aL, 1993a&b, chapter 2). This isput further into perspective in both anoxia-tolerant and -intolerant systems, where numerousstudies report that the transition to anaerobic metabolism occurs in ranges where oxygen is stillsaturating at cytochrome c oxidase (cf figures 1 and 20). Taken together, these observationssuggest that during progressive hypoxia, alterations of cell function can occur independently ofmetabolic changes associated with oxygen limitation at the mitochondrion. Therefore, in theabsence of large-scale metabolite concentration changes and exogenous effectors, by process ofelimination, the oxygen molecule itself appears as the primary signaffing agent responsible forcoordinating metabolic changes in progressive hypoxia.The present study examines the relationship between the oxygen signal, its transductionmechanism, and changes in protein expression during anoxic metabolic suppression in turtlehepatocytes. Because protein expression is composed of multiple ATP-demanding pathways andexhibits control from gene induction to protein degradation, this cell process tests the ability ofthe oxygen signal to invoke and coordinate a hypoxic response between numerous inter-relatedpathways. Furthermore, protein turnover in turtle hepatocytes conforms well to an Al?independent pattern of control. During metabolic suppression protein synthesis and ATPdependent protein turnover are suppressed by 90% but these processes still account for about40% of remaining anaerobic ATP-turnover, suggesting that changes in protein expression are110important in metabolic suppression. However, the suppression of protein turnover occurswithout change in ATP concentrations or cellular energy charge, is synchronous with thesuppression of whole cell ATP supply and demand, and therefore appears independent ofoxygen limitation at the mitochondrion.Transduction of an oxygen signal over broad ranges of oxygen concentration requires amechanism that possesses appropriate sensitivity (a high K..) for oxygen. Particularlysignificant is the finding that in numerous cell types, hypoxia associated gene expression canbe regulated through oxygen-dependent confórmational changes in a ferro-heme protein. In theliver, this kind of mechanism appears to be at the basis for02-dependent expression oferythropoietin (Goldberg et a!., 1988) and modulates expression of phosphoenolpyruvatecarboxykinase (PEPCK) expression along the liver periportal-perivenous 02 gradient (Keitzmanneta!., 1992, 1993).Based on evidence for metabolism-independent control of protein turnover in turtlehepatocytes, and the emergence of control of hypoxia-associated gene expression via a hemeprotein oxygen receptor, the specific aims of this study were 1) to establish whether the controlof protein expression under normoxia and anoxia was specifically oxygen responsive in turtlehepatocytes, 2) to determine whether a conformational change in a heme-protein group waslikely involved as a mechanism for transducing oxygen concentration changes to a cellular-levelresponse and 3) to compare oxygen-responsive protein expression to the turtle hepatocyte heat-shock response.111Materials and Methods.Chemicals.L-[U-14C]Leucine ([‘4C]Leu, sp. act. 311 mCilmmol) was purchased from New EnglandNuclear, Dupont (Quebec, Canada) and[‘4C]-methylated protein molecular weight markers(high range) from Amersham (Oakville , Ontario, Canada). Minimum essential medium(MEM) amino acids were from GIBCO (Burlington, Ontario, Canada). All other chemicalswere purchased from Sigma Chemical (St. Louis, Mo).Culture Preparation.Adult female western painted turtles (300-500g) were purchased from Lemberger (Oshkosh,WI). Hepatocytes were prepared as previously described (chapter 2). Once the post-isolationcleaning stages were complete, cells were suspended in culture medium containing (in mM):78.5 NaCl, 34.5 NaHCO3, 10 Na-N-2-hydroxyethylpiperazine-N’-2-ethanesulfonic acid(HEPES, pH7.5), 5.8 CaCl2,3.8 MgCl2,2.2 KC1, 0.9Na2HPO4,0.1 MgSO4,4% bovine serumalbumin (BSA), 10% MEM amino acids and 100U/ml penicillin-10g/ml streptomycin andstored at 4°C until needed. Prior to each experiment, cells were warmed to the experimentaltemperature (25°C) for a minimum of 3h.Cell viability during the course of each experiment described below was assessed bymeasuring rates of lactate dehydrogenase (LDH) leakage (Buck et a!., 1993b). The expressionof a full molecular weight range of proteins in each experimental group was also taken as aconvenient post-hoc measure of overall cell competence.Erperimental Design.The experiments were conducted in three parts. In Part A, cells were divided into parallelgroups for the assessment of the oxygen-responsiveness of protein expression, and Part B was112designed to investigate whether the oxygen-responsive protein expression observed in Part A,could be manipulated through factors that affect the conformation of ferro-heme proteins.Previous studies, especially in the EPO field, have developed strategies for probing this system.Aerobic incubation with NaCN provides a “physiological mimic” of anoxic cell metabolismwhilst oxygen is still present. When compared to true anoxic incubation, this provides a meansfor determining oxygen-dependent effects on protein expression. The involvement of a hemeprotein may be determined by substitution of Co2 or Ni2 ions into the central Fe2 inprotoporphyrins, inducing the deoxygenated conformational state regardless of oxygenconcentrations. Conversely, an oxygenated conformation can be induced by incubation with CO,a potent ligand of ferro-heme proteins (Goldberg et at., 1988, Eckhardt et at., 1993).Finally, Part C examines whether the observed oxygen-responsive protein expression wasrelated to the heat shock response. Except for heat-shocked groups, all experiments wereconducted at 25°C.Part A: Oxygen-responsive Protein Expression. Hepatocyte suspensions were adjusted to adensity of 30mg/mi and split into two groups. Cells were then allowed to settle out ofsuspension, the supernatant removed, and replaced with an appropriate quantity of either air-equilibrated (95% air/S % C02,humidified) or nitrogen-equilibrated (95%N2/5 % C02,humidified)culture medium. After resuspension, cells were aliquotted into four experimental groups (3m1volume at 30 mg/mi) for the parallel determination of de novo protein expression (group 1) andcell viability criteria (group 2; 30mg/mi, lOmi volume) under the following conditions:normoxic (6h aerobic), normoxic+2mM NaCN (6h aerobic with 2mM NaCN added at 2hintervals), anoxia (6h anaerobic) and anoxia+2mM NaCN (6h anaerobic with 2mM NaCNadded as above). Because NaCN slowly oxidised in the normoxic-cyanide group, 2mM NaCN113was added at two-hourly intervals to ensure complete inhibition of oxidative phosphorylation.Rates of oxygen consumption were measured in cells from group 2. At two-hour intervals, 2m1of each cell suspension was removed and placed in 2m1 thermostatted oxycell chambers heldat 25°C. To reduce the risk of air diffusing into anoxic groups, samples were pipetted andanalyzed under a meniscus of paraffin oil. Oxygen consumption was then measuredpolarographically using Clarke-type electrodes as described in chapters 2 and 3.To determine the pattern of de novo protein synthesis under each condition, cells in group1 were washed free of culture medium 4h into the incubation and were resuspended in the samevolume of NaCN and gas-equilibrated culture medium lacking leucine, but with 12.5jCi[‘4C]LeuIml. After a 2h labelling period under each experimental condition, cells were washedthree times in BSA-free culture medium at 3°C by centrifuging at SOxg for 2mm. The finalwashed cell pellets were frozen in liquid N2 and stored at -80°C for later analysis.Part B: Heme-group conformation experiments. This series of experiments tested for theinvolvement of a heme-group in the regulation of oxygen-responsive protein expression. Toincorporate cobalt or nickel into the central iron position of heme proteins, 18h incubationswere constructed in appropriate experimental volumes held at a density of 3Omg/ml, as follows:200jM CoCl2, 30OM NiCl2, 20OM Co2+2mM 2,4-dioxoheptanoic acid (DHA) andremaining cells incubated as a control group. Throughout the incubation period, cells were heldunder a humidified, 95%airl5%C02atmosphere at 25°C. After incubating for 18-20h, cellswere washed once then aliquotted into groups 1 and 2 as indicated under Part A. Furtherseparation of groups 1 and 2 into the experimental treatments (same cell density in 3m1volumes) was as follows: normoxic (6h aerobic), Co2 (6h aerobic), Ni2 (6h aerobic),Co2+DHA (6h aerobic), Co2+95 %N2/5%C02 (6h anaerobic), 10% Carbon monoxide/S %114C02/85 % N2 (6h anaerobic), Co2+ 10% Carbon monoxide/5 % C02/85% N2 (6h anaerobic).Concentrations of Co2,Ni2 and DHA were held constant as above. Labelling of group 1 cellsfrom 4 to 6h into the incubation with [‘4C]Leu, and collection of final cell pellets, wereconducted as described in Part A.Part C. Heat Shock Response. A period of heat shock was used to induce stress proteinsgroups. The profile of stress protein expression under these conditions was then compared tothe oxygen-sensitive protein response observed in parts A and B. Control and heat-shock groupswere held at 25°C for 4h under normoxic conditions. At this time, cells apportioned to theheat-shock group were transferred to a waterbath pre-heated to 40°C and incubated with shakingfor lh. Because of the small volume of cells and media in each group (3ml at 3omgcells/ml)the incubation media equilibrated to 40°C in <5mm. Once the heat-shock period was over,controls and heat-shock groups were washed once and resuspended in leucine free-media with12.5Ci[14CJLeu/ml, previously equilibrated to 25°C, and incubated for a further lh. Cellswere then washed and stored as described above.Detection of Radiolabelled Proteins.[‘4C]Leu-labelled cells were lysed in 4 vols of lysis buffer [In mM: 100 NaCl, 10tris(hydroxymethyl)aminomethane-HC1 (Tris-HC1, pH7.6), 1 EDTA (pH 8.0), 1Phenylmethyonyl-sulfonyifluoride (PMSF) and 1g.ml aprotinin] and sonicated in an ice-saltslush bath twice for lOs. The crude lysate was then placed in 2vols, 2x Laemmi sample buffer[100mM Tris base (pH6.8), 200mM dithiothreitol, 4% sodium dodecyl sulfate (SDS), 0.2%bromophenol blue, 20% glycerol and 1mM PMSF] and heated to 100°C for 10mm. Sampleswere then sonicated again for 2xlOs and then centrifuged at l0,000xg for 20 mm. The115supernatants were saved and the amount of protein-bound radioactivity was determined bytrichioroacetic acid (TCA) precipitation. This was done by adding 3d of each supernate tol471Ll of distilled water in duplicate. A 50d aliquot of this mixture was then added to 5001il ofa solution containing 1M NaOH and 0.5% (vol) H20and incubated for 15mm at 40°C. Proteinwithin each sample was precipitated by the addition of 400JL1 50%(vol) TCA containing 0.02%(w:v) casein hydrolysate and stored on ice for a minimum of 30mm. Protein precipitates werecollected on Whatman GF/C filter disks, pre-soaked in 10% TCA, using a vacuum manifoldapparatus and then washed with 6xlOml 5% TCA containing 10mM Na2P4O7.The washed filterdisks were then dried at 40°C for 20mm and the radioactivity incorporated into proteindetermined by scintillation counting in ACSII scintillation cocktail (Amersham), using an LKBRack-beta 1214 scintillation counter set to an efficiency of 95% for [‘4C] activity.2.5x104TCA insoluble degradations per minute (DPM) per sample were loaded onto sodiumdodecylsulfate (SDS)-polyacrylamide gels consisting of 5% stacking and 12% resolving gels for1-dimensional SDS-polyacrylamide gel electrophoresis (PAGE). Proteins were electrophoresedat 3OmA for approximately 4h using a running buffer consisting of 26mM Tris base (pH8.8),200mM glycine and 0.1 %SDS. Once the dye front had reached the bottom of the gel, gels wereremoved and fixed for 3h with 3 washes of glacial acetic acid:methanol:water(0.Svol:0.lvol:lvol), rinsed in distilled water for 5mm and then washed in Amplify fluor(Amersham) for 30mm. Gels were then dried and the resulting fluorograph exposed to preflashed Kodak 0-Mat XARS autoradiography film for 24-48h at -80°C. Molecular weights weredetermined using high range (200-14.4 kDa) [‘4C]-methylated molecular weight markers(Amersham). Autoradiographs were quantified using a Molecular Dynamics laser scanningdensitometer operated with ImageQuant version 3.15 software (Molecular Dynamics, NY).116Data Presentation and Statistics.Statistical manipulations were performed using Systat version 5 (Evanston IL) software.Except where noted, significance was determined using analysis of variance with Dunnett’s ttest. Confidence limits for significance are at 95% and all data are expressed as means ±S.E.M. The value of ii indicates the number of individual hepatocyte preparations used in eachexperiment.Results.Cell Viability. Throughout each experiment, the percentage of extracellular LDH remainedless than 6% for all groups except carbon monoxide/anoxia where LDH leakage rose to10.4 ±3.8% over the last 2 hours. All groups demonstrated the incorporation of [‘4C]leu intoa wide molecular weight range of proteins (figs 12,14,16,18). Taken together, the low LDHleakage and pattern of protein expression suggests that the cell preparations remained healthythroughout the experimental period.Oxygen-sensitive Protein Expression. The experiments in Part A were designed to testwhether the presence of molecular 02 was required for expression of anoxia associated protein.Cyanide was used to mimic physiological anoxia under normoxic conditions and resulted in a89% inhibition of oxygen consumption over the experimental period (fig 12). Remaining oxygenconsumption is presumably comprised of oxygen requiring processes that are not involved inthe aerobic production of ATP (eg. peroxisome oxidation). Based on this, and previous studiesnoting that in turtle hepatocytes, aerobic incubation with cyanide does not appear to perturbcellular metabolism differently from anoxia (see discussion), the use of 2mM NaCN underaerobic conditions was considered to be an adequate mimic of physiological anoxia.117In response to anoxia, turtle hepatocytes express or suppress certain proteins in an oxygen-sensitive manner. This was judged on the basis of whether the presence of molecular oxygennegated the response despite mimicked physiological anoxia in the presence of 02, using NaCN.Figure 1 demonstrates the change in the distribution of protein expression from normoxia andnormoxic physiological anoxia, to complete anoxia. The expression of five proteins of 83, 70.4,42.5, 35.3, and 16.1 kDa was consistently increased in anoxia resulting in a rise in theproportional contribution of each protein to the total protein pooi (table 9). Similarly, expressionof a distinct set of five proteins of 63.7, 48.2, 36.9, 29.5 and 17.7 kDa were down-regulatedfrom normoxia to anoxia also demonstrating a decrease in the proportional contribution to thetotal protein pool (El’able 9). The mean molecular weights for expressed and suppressed proteinsin anoxia are given in table 9.Figure 13 demonstrates the densitometric change in protein-band peak absorbance area forthe changes in protein expression identified in figure 12. Panel A demonstrates the pattern ofoxygen-sensitivity for proteins that increase expression in anoxia. In each case, environmentalanoxia resulted in a significant increase in protein band peak absorbance area relative tonormoxic controls suggesting that the increased proportion of each protein in the anoxic proteinpool was not simply due to a change in the overall size of the protein pool. In addition, anoxicexpression of 83,42.5, 35.3 and 16.1 kDa proteins was significantly greater than undernormoxic treatment with cyanide suggesting that mimicked physiological anoxia was notsufficient to cause an increase in the expression of these proteins by itself. Since the principaldifference between environmental anoxia and normoxic physiological anoxia is the presence orabsence of oxygen (see discussion), we describe the expression of these proteins as beingoxygen-sensitive. Expression of the 70.4 kDa protein was distinct from this group in that it118FIGURE 12. Representative autoradiograph demonstrating the disthbution of anoxia-associatedprotein expression and suppression in the following conditions: normoxia (Nor),normoxia+2mM NaCN (Nor+NaCN), anoxia (Ax) and anoxia+2mM NaCN (Ax+NaCN).Molecular weights of protein up-regulated in anoxia are shown on the right side of the figureand those for down-regulated proteins, on the left. Positions of Amersham[14C]-methylatedprotein molecular weight markers (Mi) are shown on the right of the figure and all molecularweights are given in kilodaltons with positions. The rate of oxygen consumption (V02) undereach treatment is given at the base of each lane (n=5). Abbreviations: n.d.- not detectable).119MrDown//i//Z/i7/7‘7 ;/Up—83.O70.4—42.5.—35.363.7—’48.2—’36.9—29.5—b—97.4—69.0—46.0—30.0—14.417.7—b—161V02(nmol02/g/h)194.7 22.2 n.d. n.d.±23.1 ±5.2120TABLE 9. Oxygen-sensitive and Insensitive Protein Expression in Anoxia.Proteins ExDressed in Anoxia Proteins Sunoressed in Anoxia%ofTotal %o(TotalMolecular Weight Protein Pool3 Molecular Weight Protein Pool3(kDa) Normoxia Anoxia (kDa) Normoxia AnoxiaOxygen Sensitive1 Oxven Sensitive183.0± 0.8 3.01 ± 1.14 10.33 ± 2.89 63.7± 0.5 9.82 ± 2.42 4.96 ± 3.1342.5± 0.8 0.49 ± 0.49 7.12 ± 1.51 48.2± 0.5 9.00 ± 2.06 1.69 ± 0.7935.3 ± 0.2 0.27 ± 0.27 4.17 ± 1.40 36.9 ± 0.5 4.84 ± 1.30 2.06 ± 1.1216.1± 0.8 0.34 ± 0.26 2.03 ± 0.70 29.5± 0.2 4.85± 1.64 1.44 ±1.15’17.7± 0.3 1.91 ± 0.51 0.32 ± 0.24Oxygen Insensitive270.4 ± 0.2 7.12 ± 3.39 19.34 ± 2.30Values are means ± SE, n=5 in each case. P<0.05; P=0.08 (students paired t-test)1E)efil as a change in protein expression only in the absence of oxygen (response does not occurin normoxia + NaCN group).1Defined as a change in protein expression in all anoxic groups, including normoxia+NaCN.3Calculated as a percentage of total densitometric absorbance area.121FIGURE 13. Oxygen-sensitivity of protein expression during physiological anoxia (02 stillpresent in medium) and environmental anoxia. A) Anoxia-induced increases in specific protein-band peak absorbance area determined by densitometry; B) Anoxia induced decreases in specificprotein-band peak absorbance area. Error bars depict standard error and n=5 in each case;P <0.05 relative to normoxic group; ‘P <0.05 relative to normoxia+NaCN group.Abbreviations: normoxia (nor), normoxia+ 10mM NaCN (nor+NaCN), anoxia (ax) andanoxia+ 10mM NaCN (ax+NaCN).122AL. Mr(kDa)83.0IllDillllhI 70.4I 142.535.3_________ ____InMr (kDa)I 163.7Cl)*A***60-***:: pp 1JIIB.48.236.9I 1 29.5JAAA TT AI1J 17.7* II. A-ii IIh -r **0, ,z zz z+ +123exhibited an increase in expression in all anoxic groups regardless of the presence of oxygen(oxygen-insensitive).Panel 13B demonstrates the oxygen-sensitivity of proteins suppressed in anoxia. Allproteins demonstrated decreases in expression relative to normoxia and with the exception ofa protein at 63.7 kDa, this overall trend was also statistically significant for all protein bandsrelative to normoxia+cyanide.Oxygen-Sensing Mechanism. The substitution of cobalt or nickel into the central ironposition of protoporphyrin, changes the state of the heme group from oxy- to deoxyconformations (Goldberg et at., 1988). If a heme-protein oxygen receptor is active in regulatingthe expression of the oxygen-sensitive proteins observed in part A then normoxic incubationwith Co2 or Ni2 should result in increased expression of proteins described in fig. 13A(except 70.4 kDa), conversely decreased expression would be expected for proteins describedin fig. 13B. In addition, incubation of cells with DHA, an inhibitor of heme-protein synthesis,should abrogate the response with cobalt.Figure 14 shows the positions of the oxygen-sensitive proteins described above,demonstrating the predicted pattern of responsiveness to Co2, Ni2 and Co2+DHA. Thecompiled densitometric data for specific protein-band peak absorbance areas is shown in figure15. Panel A demonstrates statistically significant increases in the expression of 42.5, 35.3 and16.1 kDa with cobalt or nickel treatment. The oxygen-sensitive protein at 83kDa did not showresponsiveness to any treatment group. For each protein band, incubation with Co2+DHAresulted in a return of protein expression to normoxic values for 42.5 and 16. lkDa proteins butdid not abrogate the increase in expression with Co2 for the 35.3 kDa protein. Panel Bdemonstrates the same overall pattern for oxygen-sensitive proteins that were found to be¹124FIGURE 14. Response of oxygen-sensitive proteins to normoxia (nor) cobalt, nickel andcobalt+dioxoheptanoic acid (Co2+DHA) treatment. AU treatments were conducted undernormoxic conditions. Positions of oxygen sensitive protein bands and molecular weight markersare given as for figure 13.-c) 0 00 0r\) 0,126FIGURE 15. Changes in protein expression for oxygen sensitive protein bands duringtreatment with cobalt, nickel and cobalt+dioxoheptanoic acid. A) Increased protein expressionwith cobalt and nickel. B) Decreased protein expression with cobalt and nickel. Statisticalsignificance was determined as described in figure 14; P<O.05 relative to nor, ‘P<O.05relative to Co, n=4 in each case.127Mr (kDa)83.0- 42.560- T40-Tj* ***16.1.—Cl) ‘Cl) .20IIo liii ñr____*A+0C.)B. Mr (kDa)63748.236.960-29.5** 17.70• —Cl) 40. liii ***____20O”T Ti’÷0C.)128suppressed in anoxia in Part A. Incubation with cobalt or nickel resulted in clear differences inthe amount of protein present. Interestingly, the protein at 63.7kDa which did not show astatistical difference relative to mimicked physiological anoxia in figure 13B, demonstrated asignificant suppression with nickel treatment. This supports the possibility that this protein mayindeed be oxygen-sensitive. Incubation with Co2+DHA abrogated the cobalt or nickel inducedsuppression for 63.7, 48.2 and 17.7 kDa proteins but had no effect on the suppression of 36.9and 29.5 kDa proteins.To confirm that the responses observed with cobalt or nickel were indeed brought about bychanges in heme-protein conformation, carbon monoxide was used to alter heme proteinconformation to the “oxygenated state” whilst in the presence of anoxia and anoxia+cobalt.Figure 16 demonstrates the positions of oxygen and Co2/Ni-sensitive proteins on arepresentative autoradiograph and the change in protein band absorbance area is given in figure17. Panel A demonstrates that incubation with CO significantly reduced the anoxia, andCo2+anoxia-induced expression from normoxic control groups for 42.5, 35.3 and 16.1 kDaproteins. Panel B demonstrates the response of suppressed oxygen and Co/Ni-sensitiveproteins to CO. The most profound effect of CO was found in the 17.7 kDa protein band whichdemonstrated a complete abrogation of the anoxia-induced suppression by CO treatment.However, this did not occur if Co2 was included in the incubation. CO also lead to a modestreturn towards normoxia from a significantly suppressed point for 63.7 and 48.2 kDa proteins.However, CO treatment did not abrogate the anoxia and Co2/Ni suppression in the 36.9 and29.5 kDa protein bands.Heat-Shock Response. To determine if any of the oxygen-sensitive proteins expressed inanoxia were part of an overall stress response, hepatocytes were heat-shocked to 40°C for lh.129FIGURE 16. Response of oxygen and cobalt-sensitive protein expression to treatment with10% carbon monoxide +anoxia. Positions of oxygen and cobalt sensitive protein bands andmolecular weight markers are given as for figure 13.Down130/15’!I /oII 1.I0(CiUp M—97.4—69.0—46.0—30.063.7—p48.2—’36.9—’’29.517.7—‘—42.5—14.4131FIGURE 17. Changes in oxygen and cobalt-sensitive protein expression as a function of carbonmonoxide+anoxia treatment. A)Effect on proteins that increase expression with cobalt andanoxia. B) Effect on proteins that decrease expression during treatment with cobalt and anoxia.Significance was determined as described in fig 14; P <0.05 relative to nor, AP <0.05 relativeto Co2+Ax, P <0.05 relative to Co2, n =4 in each case.Expression(PeakAbsorbanceArea)Nor.CoCo+AxCo+Ax+COAx+CO.0000II______*___H*-I_____**H*.—1*—I*I*1**!IJI-u..!-ExpressionH(PeakAbsorbanceArea).0Nor CoCo+AxCo+Ax+COAx+CO****1*!!!C133FIGURE 18. Response of turtle hepatocyte protein expression to heat shock at 40°C for lh.Positions of heat shock proteins are given on the right of the figure and position of molecularweight markers on the left. Abbreviations: C, control; HS, heat shock; Hsp, heat shockproteins.134Hsp(kDa)—90.2—74.8—25.O[—15.5III__‘634135Figure 18 demonstrates the change in protein expression during the hour following the heat-shock episode. Protein bands of 90.2, 74.8, 63.4, 25 and 15.5 kDa were evident in the heatshocked group and were of a distinct molecular weight range compared to the increasedexpression of oxygen-sensitive proteins that appeared in anoxia. One possible exception is the16.1 kDa oxygen- andCo2/Ni-sensitive protein band which is within the margin of standarderror to the 15.5 kDa heat shock protein.Discussion.The major finding in this study was the presence of nine protein bands that exhibited adifferential change in expression directly in response to the presence or absence of oxygen. Inthe case of the 42.5, 35.3, 17.7 and 16.1 kDa protein bands, expression could be predictablymanipulated with cobalt, nickel, carbon monoxide and an inhibitor of heme synthesis. Whenconsidered together, this strongly suggests that a conformational change in a heme-group isinvolved in the transduction of this response. Not all of the described oxygen-sensitive proteinbands conformed to this pattern. Increased anoxic expression of the 83 kDa protein band wasoxygen-sensitive (fig 12), but was completely unresponsive to any kind of heme proteinmanipulation. Furthermore, with the exception of the 17.7 kDa protein band, all suppressedoxygen andCo2/Ni-sensitive proteins (panel B figs 13,15,17) were unresponsive to carbonmonoxide or an inhibitor of heme synthesis. Since heme-groups differ widely in their affinitiesfor carbon monoxide (Coburn, 1979), this observation may be suggestive of control by adifferent heme protein and raises the possibility of multiple oxygen-sensing mechanismsgoverning the expression or suppression of proteins in anoxia.This overall assessment is influenced by whether aerobic incubation with cyanide was a136realistic mimic of physiological anoxia. In ideal circumstances, the largest difference betweenaerobic cyanide treatment and anoxia would be the presence or absence of oxygen itself, allother physiological effects of both states being equal. This has been disputed in the mammalianhepatocyte literature where aerobic incubation with KCN results in a rapid loss of cell viability,diminished adenylate energy charge, mitochondrial swelling, increased intramitochondrialphosphate accumulation, and a 50% loss in the mitochondrial proton-motive force. In anoxiawithout cyanide treatment, these effects occurred on a longer time-scale suggesting that cyanidehas a more immediate and profound effect on cellular metabolism (Aw & Jones, 1989).However, anoxia-tolerant organisms demonstrate a markedly different metabolic response toreduced oxygen availability. Although in situ mitochondrial metabolism has not beeninvestigated in cyanide-treated turtle hepatocytes, aerobic incubation with 0.5mM KCN over lOhhas no effect on cellular viability or energy charge (Buck et al., 1993b), which agrees with theminimal change in LDH leakage under NaCN incubation noted in this study. Furthermore,turtle hepatocytes demonstrate a similar reduction in metabolic rate and microcalorimetric heatflux when under aerobic incubation with cyanide or under anoxia (Buck et al., 1993a) and inboth cases, the plasma membrane potential remains unperturbed from control aerobic conditions(Buck & Hochachka, 1993). Overall, the metabolic response of turtle hepatocytes to aerobicincubation with cyanide clearly represents a much closer mimic of true anoxia than in the caseof mammalian hepatocytes. The metabolic differences that might exist are probably minor andare unlikely to be sufficient to cause a differential protein expression in comparison with trueanoxia.The identity of the expressed and suppressed protein bands was not established in thepresent study. Previously it was noted that anoxic gene expression in turtle liver was associated137with de novo synthesis and increased expression of three mRNA transcripts that weresubsequently translated in vitro into proteins corresponding to 19.5 (de novo), 28.6 and 79.9kDa (Douglas et a!., submitted). It was suggested that these protein translates could be part ofa stress response that is active in stabilising cellular constituents both during anoxia and on reoxygenation. To determine whether the anoxic protein bands observed in this study were partof an overall stress response, we compared them to the turtle hepatocyte heat-shock response.Of the proteins expressed in anoxia, only the 16.1 kDa protein band demonstrated a molecularweight overlap with a heat-shock protein at 15.5 kDa, and all others bands were completelydistinct. It therefore appears that the increase in the expression of specific proteins in anoxiais not part of a stress response per Se. Whether the mRNA translates corresponded to any ofthe protein bands found in the present study is difficult to assess since in vitro translation doesnot produce a mature protein product. Nevertheless, the discrepancy between the number ofprotein bands found in this study, and the number of mRNA transcripts found by Douglas eta!. (submitted) could be suggestive of control at both transcription and translation. With regardto the latter, Lefebvre et a!. (1993) have demonstrated that a possibility exists for oxygen-sensitive control of translation in rat hepatocytes: progressive hypoxia induced rapid andcoordinate decreases in protein synthesis rates which occurred well before any change in cellularenergy charge, metabolite or mRNA concentration.A number of oxygen-sensitive genes that are controlled through conformational changes ina heme-protein have been characterised in liver tissue. Particularly significant is the observationthat hormonally regulated gluconeogenic and glycolytic enzyme expression along the periportalperivenous oxygen gradient, may be under modular control by a cobalt and carbon monoxidesensitive, oxygen-sensing mechanism. Under high periportal oxygen tensions, glucagon induces138the expression of PEPCK mRNA and protein, an effect which is diminished with progressivehypoxia. Incubation with cobalt relieves the oxygen effect and carbon monoxide reinstates it(Keitzmann et a!., 1992 & 1993). Although hormones, metabolites and autonomic input areall involved in defining basal expression of periportal-perivenous enzymes, it appears thatoxygen-sensing plays an adaptive role in altering enzyme profiles during periods of hypoxia(Jungermann & Katz, 1989). Under normoxia, incubation of isolated hepatocytes with eitherglucagon or insulin preferentially mimics periportal gluconeogenic, and perivenous glycolyticenzyme profiles respectively. However, reducing oxygen concentrations significantly lowersPEPCK activity in glucagon-treated cells and increases pyruvate kinase (PK) and glucokinaseactivity in both insulin and glucagon-treated cells (Nauck et a!., ,1981; Wölfle & Jungermann,1985). The control of this response appears to be different in the case of PEPCK and PK. Inwhole liver in vivo, PEPCK mRNA and specific activity is found predominantly in theperiportal region (Bartels et a!., 1989), whereas pyruvate kinase mRNA is found homogenously,but the active protein translate is only located in perivenous areas (Lames et a!., 1987). Thus,oxygen appears capable of modulating PEPCK enzyme concentrations through a heme-proteinoxygen-sensor at the transcriptional level and in the case of PK, through an unknownmechanism at the translational level.This agrees with the initial assessment that turtle liver also appears to demonstrate oxygen-sensitive control at transcription and translation, but whether the anoxic profile of proteinexpression observed in this study reflects an oxygen-sensing phenomenon similar to that foundin liver zonation remains unknown. Whereas the expression and suppression of protein bandsoccurred in the absence of added exogenous factors such as hormones, no single protein in thisstudy demonstrated absolute induction or suppression (from or to zero) in anoxia. Likewise,139incubation with metal ions, carbon monoxide or DHA did not result in an absolute loss or gainof the protein bands of interest. It seems clear that the background expression of these proteinsis governed by an oxygen-independent factor, which can be modulated by an oxygen-sensitiveheme protein based mechanism during anoxia.The transduction pathway of an oxygen signal, through a heme-protein has been examinedin studies on the regulation of the Epo gene in liver and kidney (Goldberg et al., 1988; Maxwellet al., 1993; Eckhardt et a!., 1993). In liver, the basal expression of this gene occurs almostentirely in parenchymal cells in the hypoxic perivenous region of the acinus and deepeninghypoxia increases its expression several fold (Koury et a!., 1991). The epo gene is flanked atboth ends by a series of positive and negative regulatory elements. Oxygen-sensitive inductionof this gene is believed to involve the expression of one or more oxygen regulated inducerelement(s) (Tsuchiya et al., 1993) which relieve the binding of a trans-acting repressor at a siteupstream from the epo gene. The inducer element(s) also remove inhibitory binding of aribonucleoprotein to an upstream promoter site, allowing the binding of a constitutive 47kDaprotein, and consequent expression of the epo gene (Imagawa et al., 1994). The intracellularsignalling pathway that transduces the change in heme conformation into a genetic response hasnot yet been characterised but there is some evidence to suggest that phosphorylation via proteinkinase C is not involved (Eckhardt et a!., 1993). The nature of this kind of oxygen-sensitivecontrol of gene expression remains unknown although there are numerous parallels in theprokaryotic literature (e.g Lois et a!., 1993). Nevertheless, studies on epo, and other oxygen-sensitive genes may provide good working models for future studies concerning oxygen sensingmechanisms in facultative anaerobes.In conclusion, evidence is provided for an oxygen-sensing mechanism that is based on140conformational change in a heme-protein and that is involved in the expression and suppressionof numerous proteins during anoxia in turtle hepatocytes. In addition, there is some evidenceto suggest that other oxygen-sensitive, non-heme based mechanisms may have been involvedin regulating an 83 kDa protein band. Because the oxygen-sensitive response is not absolute,the primary role for this kind of mechanism appears to be in the adaptive modulation of existingprotein concentrations in response to progressive hypoxia and anoxia. Given that oxygensensitive control of gene expression is phylogenetically conserved and widespread betweenorgan systems (Hochachka, 1994; Fanburg et al., 1992), there is a good probability that thiskind of oxygen-sensing mechanism may play a fundamental role in adjusting cellular phenotypeto the demands of anoxic survival.141Chapter 5.General Discussion.Partitioning of ATP Demand in Metabolically Suppressed Tissues.A schematic representation of the normoxic-anoxic transition in turtle hepaotcytes ispresented in figure 19. Although metabolic rate falls precipitously on entry into anoxia,remaining anoxic metabolism accounts for about 10% of the normoxic metabolic rate. This canbe extinguished by addition of the glycolytic inhibitor, iodoacetate which also results in therapid death of the cell. This demonstrates two important points: firstly, metabolism duringmetabolic suppression is maintained by a reduced rate of glycolysis and secondly, a slow, butsustained rate of metabolism is vital for the continued survival of tissues during metabolicsuppression.From the research presented in this thesis, together with other studies on turtle hepatocyteswe are now in a position to ascribe and rank energetic costs to the metabolic processes activeduring both normoxia and anoxic metabolic suppression (Figure 19). Under normoxia, rates ofA1’P turnover for Na/K ATPase, protein synthesis, protein degradation and urea synthesisoccupy a similar proportion of total metabolic rate as found in other cell types (eg. Ascitestumour cells, MUller et a!., 1986) with protein turnover (i.e. protein synthesis and degradation)accounting for a principal proportion of normoxic ATP turnover. Since we can account for89% of normoxic ATP turnover, processes not measured, such as RNA synthesis, Ca2 ATPaseand so forth, presumably account for the remainder. Normoxic gluconeogenesis rates suggesta gross ATP requirement of 1 1.4mol ATP.g cells1.hr’ but without knowledge of rates ofglycolysis under normoxia it is impossible to infer a net energy cost for this process.1420.9100•—0_N50•-..25 -[02]Suppression(%)94Total ATP Turnover 6.3 (100%) - 67.0 (100%)(jimol ATP!g/h)75NaIK ATPase 4.8 (74%) -4- 19.1 (28%)93Protein Synthesis 1.6 (25%) -- 24.4 (36%)ATP-dependent 94Protein Degradation 0.7 (12%) - 11.1 (22%)70Urea Synthesis 0.6 (10%) 2.0(3%)100Gluconeogenesis 0 (0%) -- 11.4 (16.7%)FIGURE 19. Partitioning of energy demand from normoxia to anoxia in turtlehepatocytes. Compiled from chapter 2, chapter 3, Buck et al., 1993b and Buck &Hochachka, 1993. Percentages are data means from each source.143Under anoxia, a particularly significant finding is that the suppression of whole cellmetabolism is matched by all individual ATP-dependent cellular processes investigated in thesecells so far. This supports the concept that entry into metabolic suppression is highly regulatedand coordinated. Of the remaining cellular metabolism, however, 76% is required by Na/KATPase followed by a further 35-45% required by protein turnover. Although this actuallyaccounts for more than 100% of anoxic ATP metabolism (differences in culture conditions andseasonal variation in the donor animals most likely account for this) it is clear that there is asignificant, primary energy expenditure for the maintenance of membrane potential and asecondary expenditure for the maintenance of protein turnover during anoxic metabolic arrest.Urea synthesis accounts for a diminishingly small proportion of anoxic metabolism and sinceATP concentrations remain steady, and glycogen content falls in proportion to the rate of lactateproduction and glucose mobilisation it is assumed that gluconeogenesis is not active underanoxia.Does the remaining ATP demand in anoxia represent a set-point beyond which metabolismcannot be suppressed without excessive tissue damage and possibly cell death? The answerwould appear to be dependent upon the effectiveness of membrane stabilisation by channel arrestand probably also the ability of cell proteins to remain stable. In endothermic tissues, this setpoint can be considered to be very much higher due to the porosity of cellular membranes andnaturally higher rates of protein turnover. Singer et a!. (1992) have proposed that the minimalmetabolic rate (MMR) of all mammals, including those that enter dormant states, can besuppressed down to a minimal level that is defined by the lowest metabolic limit for sustainedstructural integrity of tissues. Interestingly, this MMR set-point coincides with the basal mass-specific metabolic rate of the largest mammal, the blue whale. Suppression of metabolism144beyond this point (as would occur in hypothermia or ischemia) results in an inability to recovertissue function, also supporting the notion of vital cell processes in defining the set-point forMMR. In turtle hepatocytes, complete inhibition of ATP supply is clearly lethal. However,the absolute limit for sustaining viability with increasingly severe metabolic suppression has notbeen experimentally probed.Recoverable, near ametabolic states do exist in nature however, as in the case of dormancyin Artemia embryos (Hand, 1990; Clegg, 1992). The key to survival here appears to lie in therelatively low complexity of the system (cells are barely differentiated) and significantly, in theability to encyst. The cyst wall is impermeable to ions and metabolites (Clegg & Jackson, 1989)and in ensuring a constant internal environment, presumably stabilises cells against largechanges in external ion concentrations. However, the question of protein stability remains amajor area of interest during dormancy in this species (Anchordoguy & Hand, In Press).Importance of Protein Turnover in Metabolic Suppression.Protein synthesis presents an excellent example of how energetically expensive cellularprocesses are balanced against the requirement to conserve energy during metabolic suppression.In chapter 2, the specific cost for peptide bond synthesis in turtle hepatocytes was calculatedat 5.3 ATP equivalents per peptide bond. Theoretically, this would be distributed in translationas 1 ATP and 1 GTP in initiation and 2 GTP in elongation. In addition, the binding of anamino acid to tRNA involves the hydrolysis of two phosphate bonds in the potentiation of theaminoacyl-tRNA complex for peptide bond formation in elongation. Since most amino acids areco-transported with Na, 1 ATP is included as the incidental cost of amino acid transport byNa extrusion by Na/K ATPase. Theoretically then, 5 ATP equivalents are involved directly145and indirectly in the synthesis of a single peptide bond (amino acid transport, binding to tRNAand elongation), and a further 2 are involved in forming the initiation complex. Assuming thatthe hydrolysis of high-energy phosphate bonds yields a standard G° of -7.3kcallmol, then thecost of peptide bond synthesis possesses a G° of 36.6kcal/mol (assumes a negligible cost forinitiation which occurs once per peptide chain). The standard G° of the peptide bond isSkcallmol and therefore the energetic efficiency of protein synthesis is only about 13%. Clearly,the high degree of fidelity required by protein synthesis commands the vast proportion of energyexpenditure in protein synthesis.Once formed, peptide bonds are extremely stable. This results from resonance within ther-electron orbital between the carboxyl and amide groups along the length of the bond (Paulinget al., 1951). This resonance hybrid resists twisting in the plane of the bond and creates anextremely stable, low energy linkage between amino acids. Given the high cost of synthesisand high stability of peptide bonds, it would seem counter-productive to maintain proteinturnover during metabolic suppression. Nevertheless, in turtle hepatocytes, about 40% ofanoxic energy metabolism is invested in sustaining protein turnover; why so?Maintenance of reduced protein synthesis rates ensures that control of protein expressiondoes not shift too far towards protein degradation. Consider equation 1 (chapter 1) where thesteady state concentration of a protein was a function of its rate of transcription and translation,divided by its rate of degradation. Complete inhibition of protein synthesis would result in achange in the control of protein expression towards degradative processes alone. Therefore,maintaining protein synthesis rates at some level helps to defray the influence of proteindegradation on protein expression during metabolic suppression. The results presented inchapters 2 and 3 support this view since the control of protein expression did indeed shift146towards protein degradation in anoxia but remaining rates of protein synthesis were significant.However, protein degradation rates were also reduced in anoxia, resulting in a two-fold increasein protein half-lives for both stable and labile protein pools. Clearly proteins are generallypresent for longer during metabolic suppression before they are degraded and replaced. Takentogether, the shift in control of protein expression towards protein degradation and the overallincrease in protein half-life points to a cellular environment that favours the accumulation ofdamaged and denatured proteins.What are the principal sources of protein damage in these conditions? Post-translationalprotein modification and damage can occur by a variety of mechanisms. A particularlyinteresting discovery which may have some significance to protein conformation duringmetabolic arrest is the ability of asparagine and glutamine residues to undergo spontaneouschemical deamination to aspartic and glutamic acids (Robinson, 1979; Yuan et al., 1981). Theappearance of deaminated protein forms is common among proteins and these usually constitutea precursor to their catabolism. In the case of triosephosphate isomerase (TPI) for example,under conditions of normal protein turnover, spontaneous deamination of two specificasparagines results in the separation and unfolding of the TPI dimers leading to the appearanceof a more acidic enzyme prior to its subsequent degradation (Yuan et al., 1981). However, ifprotein turnover rates are reduced, as in the degenerative disease progeria, denatured TPIaccumulates, with potentially lethal consequences (Gracy et al., 1984).Another source of protein modification which may be particularly relevant to tissues thatstore and release concentrations of fermentable substrate results from non-enzymatic attachmentsof glucose to free amino groups (Vaissara et al., 1985). This association occurs in tissues withglucose concentrations in excess of 50mM and is characteristic amongst proteins with slow147turnover times. The resulting glycosylated proteins (lcnown as advanced glycosylation end-products - AGE) selectively bind to an AGE receptor and are degraded. In turtle liver, glucoseconcentrations reach 200mM during the first lOh of anoxia at 25°C (Buck et at., 1993b), a timewhen protein turnover is extended suggesting that AGE accumulation may be a very realproblem during metabolic suppression. Other potential sources of protein damage arise fromlimited proteolysis, and racemization. Proteolytic modification of functionally importantproteins may be particularly problematical given the relative significance of ATP-independentproteolysis in anoxia. Racemization (the substitution of L to D amino acid stereoisomers withina protein via racemase) occurs in proteins with extremely long turnover times and may be asignificant source of protein damage to slow-turnover structural proteins during dormancies thatare extremely chronic.Recovery from chronic anoxia represents a potentially critical period in terms of reinstatingaerobic cellular function. After 6h of forced anoxic submergence at 24°C, arterial P02 rapidlyrecovers to pre-dive within the first 30 mm of recovery. The decline in plasma lactate isimmediate in onset, but much less rapid, returning lactate concentrations to pre-dive levels over15h (Robin et a!., 1981). This suggests that after prolonged anoxia, aerobic conditions, suitablefor supporting oxidative metabolism are quickly reinstated. With the potential for oxygen freeradical formation during this time (chapter 1), compounded by anoxia-induced protein damage,the reinstatement of aerobic metabolism could be severely hampered. It seems likely thereforethat efficient animal anaerobes should possess mechanisms for stabilising proteins during anoxiaand dealing with oxygen free radical formation on recovery.A major mechanism of stabilising proteins against denaturation during many kinds ofmetabolic insult (heat, oxygen and glucose deprivation, metabolic toxins etc) involves the148synthesis of stress proteins. A key feature of stress protein synthesis is that repeated exposureto the given stress results in increased tolerance and a lower incidence of protein damage(Morimoto et a!., 1990). Although there are approximately 30 different kinds of stress proteinsynthesised under differing stresses (belonging to classes of heat shock proteins (HSPs) andglucose regulated proteins (GRPs), for example), the best characterised are those of the heatshock protein 70 (HSP 70) class. These proteins possess both constitutive and inducible formsand appear to play a role in stabilising nascent protein structures during formation of the tertiaryand quaternary structure. Other members of the HSP7O family (Munro & Pelham, 1986) andGRP78 (Hendershot et al., 1988) have been identified as similar to immunoglobulin bindingprotein (BiP), a protein involved in stabilising nascent protein chains prior to glycosylation orduring periods of low glucose exposure (Kim et a!., 1987). If the nascent protein chain remainsbound to BiP due to denaturation, BiP selectively identifies that protein chain for removal byproteases. Since protein folding is an ATP dependent process requiring the interplay ofnumerous stress protein classes and several bind-release stages, some workers have noted thatexpression of inducible forms of stress protein does not occur if ATP concentrations are highand constant (Benjamin et a!., 1992; Williams et a!., 1993). Once again, this typifies thesituation in tissues during metabolic arrest. In turtle hepatocytes, during recovery from anoxia,ATP concentrations have been found to remain high with no synthesis of stress proteins (Landeta!., 1993).There are numerous other mechanisms of protein stabilisation, all of which require furtherresearch under conditions of metabolic suppression. Receiving some current attention is theeffect of reduced metabolism on ubiquitinisation of proteins, a mechanism that ear-marksproteins for removal by ubiquitin-dependent cytoplasmic proteases (Hershko, 1988). During the- 149early stages of estivative dormancy in Artemia gastrulae, there is a 93% reduction in the numberof proteins conjugated to ubiquitin (Anchordoguy & Hand, 1994). The first step in ubiquitinprotein conjugation requires ATP and involves a protein whose action may be pH dependent.With the large and rapid fall in pH that occurs during the first stages of dormancy in thisspecies, it is possible that pH, combined with a change in adenylate ratios may be importantfactors in stabilising proteins against this kind of proteolytic degradation (Anchordoguy & Hand,1994). Another exciting possibility is whether proteins from facultative anaerobes possessinnate features of protein primary sequence that preferentially stabilise proteins againstproteolytic attack. For instance, certain amino acid sequences such as those rich in proline,glutamic acid, serine and threonine appear to destabiise proteins (the PEST hypothesis, Rogerset al., 1986) or features of amino terminal residues that appear to determine ease of degradationby the ubiquitin-proteasome pathway (The N-end Rule, Bachmair et al., 1986). In studies ofmetabolic suppression, this is a wide-open and important area for future investigation.If rates of protein synthesis and protein degradation are indeed imbalanced to favourliberation of amino acids, what is their fate or possible role? Clearly, in anoxia, they are notutilised as metabolic or gluconeogenic substrates although they likely fulfil this role duringrecovery where reinstatement of oxidative metabolism enables flux through gluconeogenic andoxidative pathways and also the re-instatement of aerobic protein synthesis rates. In the contextof cellular maintenance, a role for free amino acids in the stabilisation of cell structure has beensuggested. In hypoxia-intolerant cell-lines, a depolarisation resulting from diminished iongradients precedes cell death with the result that intracellular accumulations of amino acids arelost. However, it appears that if cells are provided with glycine or alanine in low mMconcentrations, viability is markedly extended. In mammalian kidney proximal tubules, the150protective effects of glycine and alanine have been characterised to include the preservation ofintracellular ATP concentrations, stabilisation of cell membrane structure, inhibition ofNa4/KATPase (although K leakage still occurs) and a delayed increase in intracellular free Ca2.The precise mechanism behind this effect is not yet known but it appears that protection isindependent of amino acid catabolism and there is a strict dependency for amino acids with aglycine-related structure (Venkatachalam & Weinberg, 1993).Implications of Oxygen Sensing in Metabolic Suppression.Changes in metabolism during progressive hypoxia are clearly linked, in some way, tochanges in oxygen. The question is, how coarse is this control and are there signallingmechanisms that sense changes in extracellular 02 over a wide range of ambient concentrations?Closer examination of individual biochemical pathways reveal apparent “set-points for hypoxia-associated rate changes. Figure 20 demonstrates the oxygen dependence for various cellularprocesses over the lower half of the curve presented in figure 1 in Chapter 1. The pointsindicated on the curve for each cellular process are given as the half-maximal (V05) responseto a decrease in oxygen concentration, measured at the cell surface. The most strikingobservation from this figure is the apparent change in rates of glycolysis and gluconeogenesiswhich occur at the same point, and in opposite directions, well before decreases in proteinsynthesis rates, and in turn, well before any fall in ATP concentrations. As with the fall inwhole tissue V02 (figure 1, Chapter 1) V0.5 for each of these processes occurs at points that areseveral-fold higher than the standard km of 02 at cytochrome c oxidase.This response can be interpreted in two ways: i) in intact hepatocytes, there is.a substantialgradient of 02 across the plasma membrane that results in diffusional limitation of in situA.151280>6040200 50 100 150[021 (?LM)B HYPOXIA INCREASES. EXPRESSION1008060HYPOXIADECREASES40 EXPRESSION2000 50 100 150 200[021 (pM)FIGURE 20. Control of anaerobic processes appears independent of mitochondrialoxygen consumption and ATP concentrations in rat hepatocytes. A) V05 pointsfor up- and down-regulated cell processes in anoxia. B) Induction and suppressionpoints for hypoxia sensitive gene and protein expression in anoxia. Compiled fromNauck eta!., 1981, Wölfle & Jungermann, 1985, Eckhardt eta!., 1993, Lefebvreet a!., 1993, Thurman et a!., 1993).INCREASE IN GLYCOLYSISFALL IN GLUCONEOGENESIS0200152mitochondrial metabolism even at high oxygen concentrations, and/or ii) cellular metabolismis regulated by a mechanism that is capable of sensing and transducing changes in extracellularoxygen concentrations (possesses a high Km for 02) into a coordinated cellular response. Joneset a!. (1990), has calculated that under normoxic steady-state conditions, the gradient of 02from the unstirred extracellular boundary layer to the mitochondrion is about 6M 02. Duringacute hypoxia, there is a dramatic increase in the relative resistance of oxygen diffusion overthis distance, due to functional anoxia around mitochondria (Jones et a!., 1990). Therefore,metabolism is functionally anaerobic before ambient 02 reaches zero. Nevertheless, themagnitude of the oxygen gradient is too small to account for the V0•5 of oxygen sensitive cellularprocesses demonstrated in figure 1.A further issue that is not settled by the suggestion that02-gradients may bring about thecoordinated change towards anaerobic metabolism, is the observation that cellular changes occurwhilst metabolic signals remain relatively unperturbed. Changes that do occur, cannot accountfor the global suppression of ATP synthesis and demand that is associated with numerousmetabolic processes during metabolic suppression. Clearly, something else is controlling fluxthrough metabolic pathways, that is sensitive to oxygen concentrations and is capable ofcoordinately modulating numerous cellular pathways.In an attempt to examine the relationship between signal and response, the studies inchapter 4 investigated oxygen-induced changes in protein expression and suppression. Theresults suggest that oxygen plays a significant role in modulating the expression and suppressionof proteins over their basal concentrations through conformational changes in a heme-proteinoxygen receptor. This kind of oxygen sensing mechanism is significant because it presents amechanism where, depending on the02-binding kinetics of the heme group, changes in oxygen153concentration can be detected early in hypoxia, and ensuing oxygen-sensitive changes titratedappropriately with increasingly severe hypoxia. Furthermore, the results suggested that oxygencontrol of protein expression could over-ride the primary factors controlling basal proteinexpression (eg hormones). This is therefore a system that operates independently of oxygensupply until a critical point is reached in hypoxia, at which time the system is capable ofinvoking oxygen sensitive-control and readjusting to functioning in a low 02 environment.Figure 20B demonstrates the points of induction for various oxygen sensitive gene products inmammalian liver, over the lower half of the curve presented in figure 1 (Chapter 1). As withthe pattern of protein induction in turtle hepatocytes, expression and suppression of proteinsoccurs in an oxygen-sensitive fashion, well before ATP concentrations fall, and long before 02is limiting at the mitochondrion. The advantage of modulating protein profiles in this way isclear: as progressive hypoxia becomes increasingly severe, so the profile of functional proteinscan be readjusted towards favoured function in anoxia well before oxygen and energy supplybecome limiting.Switching over control from the primary signals that govern cell functions, to an oxygen-dependent modulation of cell function requires an oxygen-sensing mechanism that can interactdirectly with normal modes of cellular signalling. Guanylate cyclases offer an excellent exampleof a possible bridge between hormone and oxygen based signalling mechanisms (fig 21).Receptor guanylate cyclases (GCases) are found in both insoluble (membrane-bound), andsoluble (cytosolic) forms and in each case, respond to the binding of agonists to produce cyclicGMP (cGMP). By activating protein kinases, cGMP modulates Na channel activity innumerous tissues and can abrogate cAMP-induced responses by activating cAMP-hydrolysingphosphodiesterases (Hille, 1991). In metabolic suppression, cGMP and protein kinase G’s have154been implicated in the inhibition of PK activity by phosphorylation in the anoxic whelk tissues(chapter 1). Although the membrane-bound GCase appears largely responsive to hormones, thesoluble form appears to be activated by other agonists. This protein possesses a ferro-hemesubunit which can change conformation to directly affect the rate of hydrolysis of GTP -‘cGMP+2Pi at the catalytic site and as such, allows alternate signals to be fed into the cGMPsignalling pathway. The principal agonist at this site appears to be nitric oxide which interactsdirectly with the central iron position to effect large changes in cGMP concentrations (Ignarro,1989). To my knowledge, no studies have been carried out to examine if changes in cytosolicoxygen can similarly activate GCase, or whether this is represents a physiologically importantsignalling route in hypoxia in vivo. However, the association of two signalling mechanismswithin one signal-transducing pathway is highly suggestive of the theoretical requirements foran oxygen-sensing process that operates to modulate existing cell functions (fig 21).How important are oxygen sensing mechanisms in the control of metabolic suppression?This is difficult to assess at this stage although the potential is enormous. The fact that proteinconcentrations change via an oxygen-sensing mechanism is indicative of fundamental phenotypicchange within cells during metabolic suppression. Further investigation may reveal subsequentmodulation of metabolic pathways or cellular processes as a result. Perhaps the most importantfeature of this mechanism is its demonstrated ability to bring about a coordinated change inprotein concentrations directly in response to a change in oxygen, as would be required forcellular responses during metabolic suppression.155PDE ActivationcAMP S’AMPHormone(eg ANP)VOther cell responsesincluding metabolicenzyme inhibitionIGTP02FIGURE 21. The oxygen-sensing potential of the cGMP signalling pathway:feeding two messages into one second-messenger system. Abbreviations: PDEphosphodiesterase, ANP-Atrial naturietic peptide, GCase-guanylate cyclase.Na/PProtein Kinase-G2Pi+cGMP 4?Protein Kinase-G IFe-j4 [Ca2j1156Perspective.The research presented in this thesis supports the concept of metabolic suppression as ahighly coordinated and regulated response to anoxia. The principal contribution made here isthat we can now assign energetic costs to individual cellular processes during normoxia andanoxia, and view changes in the partitioning of these costs across each environmental condition.In turtle hepatocytes, it was found that the maintenance of the membrane potential and proteinturnover account for the vast proportion of energy usage in anoxia. Is this observationtransferrable to other tissues during metabolic suppression? Almost certainly yes, in the caseof tissues that normally possess high metabolically activity such as heart and brain. Asestablished in chapter 1, membrane ion pumping accounts for the vast proportion of energyutilisation in the brain and despite changes in synaptic activity, the large number of ion channelspresent points to a good likelihood that ion pumping would be the most active component ofmetabolism in metabolic suppression. Metabolically active tissues, are usually also associatedwith higher rates of protein turnover where the consequences of accumulations of damagedprotein would otherwise be high in normoxia and anoxia.The control of this energy partitioning is particularly intriguing since in some cases relativeincreases in the overall proportion of energy utilised occur, during a time when ATP synthesis,together with ATP demand are in a balanced suppression. The coordination of these eventsremains a mystery, however this thesis also demonstrated that in the case of altering hepatocyteprotein profiles, oxygen itself was the key player. That oxygen can directly control numerousphysiological and biochemical processes through oxygen sensing mechanisms represents apotentially crucial and unexplored area of future research in facultative anaerobes. 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