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Ketogenesis in chick embryo isolated hepatocytes
Comp. Biochem. Physiol. Vol. 78B, No. 2, pp. 437~142, 1984
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KETOGENESIS IN CHICK EMBRYO ISOLATED HEPATOCYTES ALISON J. BATE* and ALAN J. DICKSON
Department of Biochemistry, The Medical School, University of Manchester, Manchester M 13 9PT, UK (Tel: 061-273-8241) (Received 21 November 1983)
Abstract--1. Isolated hepatocytes from 17-day old chick embryos exhibit high endogenous rates of ketogenesis. 2. The addition of long-chain fatty acids stimulated ketogenesis with potency ordered as follows: palmitate > oleate > stearate. 3. Octanoate produced a slight stimulation of ketogenesis when added at low concentrations (less than 0.25 mM). At higher concentrations the effect of octanoate was inhibitory. 4. The addition of glucose to incubations failed to lessen endogenous ketogenesis whereas propionate, pyruvate and lactate produced inhibition. 5. Ketogenesis from both endogenous sources and added fatty acids was not altered by the addition of glucagon, insulin, adrenalin or vasopresin.
and organs for different classes of nutrient. As a preliminary stage in investigations of tissuedependent variations in nutrient utilization, we describe in this paper the role of the liver in the production of ketone bodies (D-3-hydroxybutyrate and acetoacetate) from fatty acids. Under conditions of diabetes and prolonged starvation in mammals (nutritional situations analogous to those during chick embryo development) ketones, produced by the liver during conditions of excessive fatty acid availability, can be utilised by several tissues (McGarry and Foster, 1972). Ketone utilization reduces the requirement for glucose and plays an important role in the adaption to decreased glucose availability. Relatively little is known of ketogenesis and its control in avian species. Available data is confined to measurement of ketogenesis with isolated liver preparations from 2-6 week old chickens and pigeons (Bailey and Horne, 1972; Srling et al., 1975; Ogato et al., 1982; Sugano et al., 1982; Zammit, 1983). Measurement of the activity of D-3-hydroxybutyrate dehydrogenase in chick embryo brain suggests that hydroxybutyrate may be an important energy source for this tissue (Nehlig et al., 1980; Nehlig and Lehr, 1982). There is, however, no data available on ketone production by chick embryo liver. In this paper we describe the use of hepatocyte suspensions isolated from embryos 4 days prior to hatching to investigate the control of ketone body synthesis from a range of fatty acids by nutrients and hormones.
INTRODUCTION Growth and development of the embryonic chick is dependent on the mobilization and consequent utilization of nutrients from stores within egg white and yolk. Carbohydrate availability during embryonic development is severely limited. Less than 1% of egg nutrients are carbohydrate and these are mostly utilised within the first 7 days of the 21 day period of incubation (Hazelwood, 1972). Protein from egg white and yolk is degraded to its component amino acids by yolk sac and extra embryonic membranes (Hazelwood, 1972). Following their uptake into embryonic blood, while amino acids are available to promote protein synthesis and growth, certain amino acids may be catabolised for energy provision or enter the gluconeogenic pathway of liver, kidney and muscle (Yarnell et al., 1966; Dickson, 1983). Although no data are available on issue amino acid catabolism, considerable information exists on the role of amino acids in gluconeogenesis. Conversion of amino acids to glucose or glycogen results in the elevations in plasma glucose concentration and, in liver and muscle, tissue glycogen contents during the later stages of embryonic development (Freeman, 1969; Langslow, 1975; Wittman and Weiss, 1981). Studies with isolated tissue preparations from embryonic chicken liver have shown the importance of the gluconeogenic pathway in that tissue (Yarnell et al., 1966; Dickson, 1983). Stored lipid within the egg makes up 50~ of available nutrients (Romanoff, 1967). Data on respiratory quotients indicate that, especially during the final 7 days of embryonic development, fatty acids are the major oxidised substrate for tissue metabolism (Hazelwood, 1972). Although fatty acids are a major fuel for the embryo as a whole, there has been no definition of the requirements of individual tissues *Correspondence to be addressed to: Alison J. Bate.
MATERIALS AND
METHODS
Sources of animals and materials Fertilised White Leghorn (Gallus domesticus) eggs were
obtained from the University of Manchester Medical School and used after 17 days of incubation (4 days prior to hatching). Adrenalin, firefly lantern extract (FLE-50), glucagon, insulin, L-lactic acid, n-octanoic acid, palmitic acid, stearic acid and vasopressin were all obtained from Sigma, Poole, Dorset, UK. ATP, collagenase, dibutyryl cyclic AMP, D-3-hydroxybutyrate dehydrogenase, NAD +,
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ALISON J. BATE a n d ALAN J. DICKSON
NADH and pyruvate were purchased from Boehringer Corporation Ltd., Lewes, Sussex, UK. All other biochemicals (Analar grade) were supplied by B.D.H. Chemicals Ltd., Poole, Dorset, UK. Bovine serum albumin (BSA) was purchased from Park Scientific Ltd., Northampton, UK and was defatted and dialysed prior to use (Chen, 1967). Hepatocvte isolation and incubation Chick embryo hepatocytes were isolated by the collagenase digestion method of Picardo and Dickson (1982). Portions (0.5 ml) of the final cell suspension which contain on average 1.5 2.0 mg dry wt of cells were incubated in stoppered 5 ml polypropylene vials. The Krebs Henseleit bicarbonate buffer used for incubations contained BSA at a final concentration of 11!;. The vials were flushed with O2:CO 2 (19: 1) gas mixture and incubated with shaking at 37C. Reactions were terminated by the addition of perchloric acid (final concn, 0.2 M) to the entire ceil suspension. After centrifugation (2500g, 5min), portions of the supernatant were analysed for acetoacetate, D-3hydroxybutyrate and ATP. Analytical procedures Perchloric acid extracts were neutralised with 2 M KOH containing 0.5 M triethanolamine prior to analysis. ATP was measured by a modification of the luminescent assay of Stanley and Williams (1969). D-3-Hydroxybutyrate and acetoacetate were determined by a modification of enzymatic procedure of Williamson et al. (1962). For both metabolites, 0.1 ml of neutralised cell extracts were analysed in a final cuvette vol of 0.5ml. The assay medium for D-3-hydroxybutyrate contained 1.9 M Tris, 0.32 M hydrazine sulphate, 8 mM MgSO4, 8 mM EDTA and 2.4 mM NAD + (final pH 9.8). For acetoacetate the assay medium consisted of 80mM triethanolamine, pH 7.4 containing 0.14 mM NADH. Expression qf results Portions of cell suspensions were dried to constant weight at 120'C and values were used to calculate results in terms of dry weight.
chain fatty acids (Fig. 1: Table 1). ]'he percentage stimulation caused by each agent was similar at all time points. For the experiments reported in Fig. I and Table 1, palmitate (0.5raM) and octanoate (0.125 mM) increased the hydroxybutyrate:acetate ratio by 14.9°,0 and 29.57~; respectively. This effect was observed at all time points studied. Oleate (0.5 mM), stearate ( l . 0 m M ) and butyrate (4.0raM) each decreased the ratio by 4(~507;; at all incubation times. The concentration dependence of ketogenesis on exogenous fatty acid varied between fatty acids (Fig. 2). Although palmitate, oleate and stearale gave similar degrees of stimulation of ketogenesis, the concentration required to attain this maximal sensitivity could be ordered: palmitate < oleate < stearate. Butyrate stimulation of ketogenesis was slight and the maximal effect produced was only 60!!1; of that attained with palmitate, oleate and stearate. Octanoate at concentrations below 0.25 mM produced a slight stimulation of ketogenesis. Further increases in octanoate concentration inhibited ketogenesis. Addition of the ketogenic precursors cited above produced minor effects on hepatocyte viability. Only octanoate reduced cell ATP content (by 5 15";, at incubation periods of up to 1 hr). To investigate the interaction of other possible plasma constituents with ketogenesis from endogenous and exogenous sources, experiments were performed over 60 min with or without additions of metabolites and hormones. Glucose (at concentrations up to 20 mM) had no effect on ketone body formation whereas fructose (10 mM) caused a 40",; inhibition. Addition of propionate, lactate and pyruvate produced significant inhibition at concentrations from 0.25-10 mM (Table 2). The effects of lactate and
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RESULTS The cell preparations from which results on ketogenesis were obtained had initial A T P contents in the range of 7.89 _+ 0.45 (12) nmoles/mg dry wt. This value changed by less than 2.3'I o over a 1 hr period of incubation. On this basis, the preparations used show close to physiological viability (Picardo and Dickson, 1982) and metabolic processes such as ketogenesis are unlikely to be impaired as a result of hepatocyte isolation. Hepatocyte suspensions maintained a constant rate of endogenous ketogenesis for incubations of up to 2 hr (results not shown). The major ketone produced was D-3-hydroxybutyrate. The ratio of hydroxybutyrate:acetoacetate in the extract progressively increased through incubation. After 15, 30, 60 and 120rain of incubation the ratio was 3.2_+0.4, 5.2 _+ 0.8, 17.8 _+ 4.3 and 53.2 + 8.9 respectively (from 4 experiments). The amount of acetoacetate produced was small and progressively of minor importance to the total sum of ketones synthesised during incubations. Except where stated, results are expressed only as total ketones produced by hepatocyte suspensions. Subsequent experiments were restricted to 60 min duration. The rapid endogenous ketogenesis was accelerated by the addition of long- and medium-
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Fig. 1. Time course of endogenous (O) and oleatestimulated (11) ketogenesis. Hepatocytes were prepared from 17-day-old chick embryo livers as described in Materials and Methods. Results are expressed as total ketones (hydroxybutyrate plus acetoacetate) formed in incubations. Final oleate concentration in the incubations was 0.5 mM. Values are means _+SEM for three separate cell experiments.
Ketogenesis in chick embryo isolated hepatocytes
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Table 1, Stimulation of endogenous ketogenesis by fatty acid addition* % of endogenous ketogenesis incubation time (min)
Added fatty acid
30 187.2 + 13.5 213.5 + 13.1 197.4 _+ 8.8 109. I + 3.8 152.4 _+ 8.0
15
Oleate (0.5 mM) Palmitate (0.5 mM) Stearate (1.0 mM) Octanoate (0.125 mM) Butyrate (4.0 raM)
171.3 + 16.9 N.D. 182.9 +_ 8.6 124.2 + 2.8 125.0 + 7.3
60 195.2 + 10.9 209.4 _+ 8.6 201.7 +- 4.3 125.3 + 4.7 167.4 + 4.5
*The final concentrations of added fatty acids are indicated in parentheses. Ketogenesis is expressed as the sum of hydroxybutyrate and acetoacetate. Methods for the isolation and incubation of hepatocytes and the determination of ketones is given in Materials and Methods. In each case values are the means+_SEM for thlee separate cell preparation (N.D. = not determined).
pyruvate were observed both when the metabolites were added singly or in combination (in a ratio of 9: 1 respectively). Inhibition occurred both for ketogenesis from endogenous sources and from added palmitate (0.25 raM). Ketogenesis was unaltered by the addition of glucagon, dibutyryl cyclic AMP, insulin, adrenalin or vasopressin (Table 3). Ineffectiveness of glucagon, dibutyryl cyclic AMP and adrenalin on ketogenesis was observed although each hormone stimulated glycogenolysis in parallel incubations (Picardo and Dickson, 1982).
the presence of fatty acids is equivalent to that found for fed or starved perfused rat and pigeon livers (S61ing et al., 1975). Hydroxybutyrate is the major product of ketogenesis from exogenous and endogenous fatty acid catabolism. Although the precise value for the hydroxybutyrate/acetoacetate ratio varied with the incubation time, the ratio was consistently high (from 275
250
DISCUSSION
The results presented in this paper are the first comprehensive report of hepatic ketogenesis and its control during chick embryonic life. A previous report described the production of ketones from [1J4C]palmitate by liver slices from 10 day old chick embryos (Koerker and Fritz, 1970). This report was brief and the use of liver siices is a technique of questionable value today. Our results indicate that ketogenesis is a major metabolic pathway in embryonic liver metabolism. High rates of ketogenesis (91.9 _+ 3.2 nmoles/hr/mg dry wt) were observed in the absence of any added fatty acids. Hepatocytes from embryonic chicks contain large lipid droplets within their cytosolic structure and have a high fat content as indicated by a 50% fat-free dry weight estimate determined for such a tissue (Goodridge, 1973a). The anoxia and stress incurred throughout hepatocyte isolation stimulates the mobilization of fatty acids from hepatic cytosolic stores and permits their entry into the processes of fatty acid oxidation. Hepatocyte stores of fatty acids sustain a constant rate of ketogenesis for incubation periods of over 1 hr (Fig. 1). The rate observed over this period, in the absence of added fatty acids, is close to 50% of that obtained for rat and pigeon isolated liver preparations in the presence of exogenous fatty acids (S61ing et al., 1975; Demaugre et aL, 1983). Addition of fatty acids to chick embryo hepatocytes accelerated ketone body synthesis by approximately two-fold (Table 1; Fig. 1). Once maximal ketogenesis was attained, further addition of fatty acids failed to further stimulate conversion to ketones. Limitations in mitochondrial transport will restrict fatty acid activation at the initial stages of ketone formation. At maximum rate, ketogenesis in
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2
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4
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Fig. 2. Modulation of hepatocyte ketogenesis by palmitate (O), stearate ([]), oleate (O), butyrate (A) and octanoate (11). Hepatocytes were incubated for 60 min and the effects of additions on total ketone body formation are expressed as % of the endogenous rate. Each addition represents a single hepatocyte preparation and the individual points are the means + SD for 3 separate determinations,
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ALISON J. BATE a n d ALAN J. DICKSON
Table 2. Inhibition of endogenous ketogenesis by propionate, lactate and pyruvate additions* Final concentration of addition (raM) 0.25 0.5 1.0 2.5 5.(1 10.0
'?i, of endogenous ketogenesis Lactate/pyruvate Propionate mixture 75.2 + 3.8 70.0 + 4.1 66.3 2 4 . 4 50.9 ~ 2.5 43.2 _~ 3.5 33.6_+ 2.4
74.5 -+ 5.4 52.9 ± 1.9 31.6_+ 5.7 15.2 ± 3.7 18.4 _+ 2.1 36.5 + 3.1
*The mixture of lactate and pyruvate was m a d e up to give the final concentration indicated with components in a ratio o[" 9:1 respectively. Ketogenesis was determined as the total sum of hydroxybutyrate and acetoacetate produced over a 60 min incubation period. Values are the means + SEM for 3 separate cell preparations in each case, Results are expressed as % of the endogenous rate (endogenous ketogenesis was 82.8 _+ 7.0 and 1 0 1 . 0 + 6 . 2 n m o l e s ketones produced/hr/mg dry cells in the experiments for propionate and lactate/pyruvate mixture respectively).
5 20). In mammalian blood the ratio of ketones would be between 2-3 in both the fed, starved and severely diabetic conditions (Page et al., 1971: Sarkar, 1972; Blackshear et al., 1975). Avian species have consistently high plasma hydroxybutyrate:acetoacetate ratios. Ratios between 6 and 20 are normal for fed and starved chickens and pigeons after hatching (Bailey et al,, 1971: Bailey and Horne, 1972: Sarkar, 1972; Davison and Langslow, 1975; Brady et al., 1978; Zammit, 1983). Plasma from 17 day old chick en embryos contained 2.3 2.5 mM total ketones in a ratio of24:1 (hydroxybutyrate:acetoacetate) (A. J. Bate and A. J. Dickson, unpublished results). Thus the ratio of ketones produced by isolated chick embryo hepatocytes is an accurate reflection of the situation in ovo. High ratios of hydroxybutyrate:acetoacetate may be a consequence of limitations in the transfer of reducing equivalents from mitochondria to cytosol in chicken liver, an event of importance in the control of gluconeogenesis from lactate and pyruvate (Dickson and Langslow, 1978; Bannister and O'Neill, 1981: Ogato et al., 1982; Sugano et al., 1982). Within embryonic liver, the major fatty acids available are palmitate, stearate and oleate (34, 34 and 30°o of total fatty acids respectively) (Goodridge, 1973b). The potency of these fatty acids upon ketogenesis when added exogenously to hepatocytes underlines that they will play an important physiological role in ketone body formation hi ovo. Utilization of fatty acids produced one of two effects on
hydroxybutyrate:acetoacetate ratios: (a) an increase of 20°J~, (with palmitate or octanoate); (b) a decrease of 40~500~, (with oleate, stearate or butyrate). Rapid oxidation of fatty acids should produce a high N A D H : N A D ÷ ratio within mitochondria. This should in turn depress the hydroxybutyrate:acetoacetate ratio. The observed depression in the ketone ratio is likely to be a result of high rates of fatty acid oxidation. Increased ratios observed with palmitate and octanoate cannot be explained from the available background material. Octanoate exerts a dual effect on ketogenesis (Fig. 2). At concentrations below 0.25 mM, this medium chain fatty acid produced a small stimulation of ketogenesis. At higher concentrations a progressive inhibition occurred. The transport of medium chainlength fatty acids (e.g. octanoate) into mitochondria is not subject to limitations at the level of membrane permeation (Fritz, 1959: McGarry and Foster, 1974). Increases in octanoate concentration will result in high intramitochondrial concentrations of this fatty acid and the mitochondrial pool of coenzyme A will be sequestered in the form of an octanoyl-coenzyme A complex. When sufficiently high concentrations of coenzyme A are complexed, inadequate amounts will be available for acetyl-coenzyme A t~rmation fiom fatty acid catabolism. This mechanism would reduce the rate of ketogenesis as a consequence of inadequate precursor (acetyl coenzyme A) supply. Data on the effects of propionate, lactate and pyruvate (see later sections) support the concept of ketogenesis limitations as a result of coenzyme A availability. Although there are reports on the use of octanoate (1 mM, final concentration) as a ketogenic precursor in isolated chicken liver preparations (Ogato eta/.. 1982; Sugano et al., 1982). such reports are for birds after hatching and represent a situation with low basal rates of ketogenesis. Competition between octanoate and endogenous fatty acids is liable to be much reduced under such conditions and inhibition of ketogenesis may only be apparent at high concentrations of octanoate (i.e. from above 1.0 raM). The use of octanoate in mammalian liver preparations (e.g. S61ing et al., 1975) may be possible at concentrations of 1 2 m M for the same reason. Interactions between the metabolism of likely plasma constituents and ketogenesis in chick embryos were investigated by addition of glucose, lactate and pyruvate to liver cell suspensions (Table 2). Glucose, which wilt be present in plasma oi" 17-day-old embryos at a concentration of 7 8raM (Langslox~,
Table 3. Effects of hormone addition on ketogenesis from endogenous and exogenous fauy acids* ?i, of control ketogcnesis From endogenous In the presence of fatty acids added palmitate Glucagon (2.87 x 10 r M ) Dibutyryl cyclic A M P (5.0 x 10 5 M) Insulin (1.74 x 10 7 M) Insulin + glucagon Adrenalin (1.1 x 10 5 M) Vasopressin (2.47 x l0 ' M)
102.9+2.8 99.1 4_-4 6 106.3 _+ 4.1 106.6 + 3.6 102.6_+ 8.6 110.6 + 5.0
101.5~ 1.0 N.D. 99.4 2 2.2 100.3 ± 2.2 105.3 -+ 3.6 102.4 + 6.5
*Incubations were for 60 min. Ketogenesis was determined as the sum of hydroxybutyrate and acetoacetate production. The control rate describes the extent of ketone production from endogenous sources or palmitate (at a final concn of 0,25 raM) alone and the response to hormones expressed as a percentage of the control value in each case. Values are means + SEM fl~r 3 separate cell preparations in each case.
Ketogenesis in chick embryo isolated hepatocytes 1975), had no effect on ketogenesis, As hepatic glycolytic enzymes are of low activity until after hatching (Wallace and Newsholme 1967) and lipogenesis from glucose is absent prior to hatching (Goodridge, 1968, 1970), the ineffectiveness of glucose probably results from a lack of glucose catabolism by embryonic liver. The effect of lactate and pyruvate upon ketogenesis was observed with mixtures or individual solutions of the components (Table 2). As each component was individually inhibitory on ketogenesis, the mode of action is not related to alterations in redox state produced in cellular metabolism. The effects would appear to be linked to the subsequent metabolism of the carbon skeleton of lactate and pyruvate. That propionate gives rise to the same maximal inhibition of ketogenesis as lactate and pyruvate (Table 2) suggests a common mechanism of action. It appears likely that the common mechanism is the utilisation of coenzyme A in entrance of propionate, pyruvate and lactate into the tricarboxylic acid cycle. Reduction of coenzyme A levels in mitochondria will impair the activation of fatty acids and result in reduced rates of ketogenesis. Inhibition of ketogenesis by addition of propionate, lactate and pyruvate has been observed with isolated hepatocytes from other species (Lomax et al., 1983). Although the precise mechanism for inhibition remains uncertain, the data presented here for chick embryo hepatocytes favour the possibility of regulation through competition for coenzyme A utilization. Whatever the mechanism, inhibition of ketogenesis by lactate and pyruvate will be important for the regulation of ketogenesis in ovo. As plasma lactate concentrations in chick embryos during the last few days of incubation are in the range of 0.6-0.7 mM (Dickson, 1983) maximal possible rates of ketogenesis will be reduced by up to 50~o. Although glucagon and insulin are known to play important roles in the acute regulation of fatty acid esterification and oxidation in hepatocytes from chicks after hatching (Goodridge, 1973a; Mooney and Lane, 1982), with concomitant effects on ketogenesis, there was no effect of these hormones on ketone body formation in 17-day-old chick embryo hepatocytes (Table 3). This occurred at a time when glucagon increased glycogenolysis (Picardo and Dickson, 1982) and produced dramatic increases in hepatocyte cyclic A M P concentrations (unpublished work). The lack of effect with dibutyryl cyclic A M P further emphasises that unresponsiveness is unrelated to receptor numbers of sensitivity but can be restricted to intracellular reception of hormonal stimulation. Previous work with hepatocytes (Picardo and Dickson, 1982) and adipocytes (Langslow, 1972) have shown similar diminished responses to glucagon at this stage of chick embryonic life. Adrenalin, which acts through a cyclic AMP-mediated mechanism in chick embryos (Picardo and Dickson, 1982) and the calcium-mobilising hormone, vasopressin, were also ineffective. The data presented in this paper indicates that ketogenesis is a major metabolic pathway in chick embryo liver. The provision of ketones as an alternative fuel for several tissues may be an essential part of the metabolic homeostasis in the development of chick embryos. Although the nature of the tissues
441
which may utilise ketones has not been determined, data from chick embryo brain indicates high concentrations of hydroxybutyrate dehydrogenase (Nehlig et al., 1980; Nehlig and Lehr, 1982). In the development of chicks from embryos to neonates plasma fatty acid concentrations decrease from 1.2 to 0.3mM (Freeman and Manning, 1971; Langslow, 1975). In parallel with this, plasma ketone concentrations drop from 2.4mM in 17-day-old embryos (this paper) to 0.6 mM in neonates (Sarkar, 1972). Thus ketones provide large amounts of alternative nutrients for the chick embryo at a time of high plasma fatty acid concentration. REFERENCES
Bailey E. and Home J. A. (1972) Formation and utilization of acetoacetate and o-3-hydroxybutyrate by various tissues of the adult pigeon. Comp. Biochem. Physiol. 42B, 659-667. Bailey E. J., Horne J. A., lzatt M. E. G. and Hill L. (1971) Concentrations of acetoacetate and 3-hydroxybutyrate in pigeon blood and desert desert locust haemolymph. Life Sci. 10, 1415-1415. Bannister D. W. and O'Neill I. E. (1981) Control of gluconeogenesis in chick (Gallus domesticus) isolated hepatocytes: effect of redox state and phosphoenolpyruvate carboxykinase [EC 4.1.1.32] location. Int. J. Biochem. 13, 437-444. Blackshear P. J., Holloway P. A. H. and Alberti K. G. M. M. (1975) Metabolic interactions of dichloroacetate and insulin in experimental diabetic ketoacidosis. Biochem. J. 146, 447-456. Brady L. J., Romsos D. R., Brady P. S., Bergen W. G. and Leveille G. A. (1978) The effects of fasting on body composition, glucose turnover, enzymes and metabolites in the chicken. J. Nutr. 108, 648-657. Chen R. F. (1967) Removal of fatty acids from serum albumin by charcoal treatment. J. biol. Chem. 242, 173-181. Davison T. F. and Langslow D. R. (1975) Changes in plasma glucose and liver glycogen following the administration of gluconeogenic precursors to the starving fowl. Comp. Biochem. Physiol. 52A, 645-649. Demaugre F., Buc H. A., Cepanec C., Monacion A. and Leroux J-P. (1983) Comparison of the effects of 2-chloropropionate and dichloroacetate on ketogenesis and lipogenesis in isolated rat hepatocytes. Biochem. Pharmac. 32, 1881-1885. Dickson A. J. (1983) Gluconeogenesis in chick embryo isolated hepatocytes. Int. J. Biochem. 15, 861-865. Dickson A. J. and Langslow D. R. (1978) Hepatic gluconeogenesis in chickens. Molec. Cell. Biochem. 22, 167-181. Freeman B. M. (1969) The mobilization of hepatic glycogen in Gallus domesticus at the end of incubation. Comp. Biochem. Physiol. 28, 1169-1176. Freeman B. M. and Manning A. C. C. (1971) Glycogenolysis and lipolysis in Gallus domesticus during the perinatal period. Comp. Gen. Pharmac. 2, 198-204. Fritz I. B. (1959) Action of carnitine on long chain fatty acid oxidation by liver. Am. J. Physiol. 197, 297-304. Goodridge A. G. (1968) Conversion of [U-~4C]glucoseinto carbon dioxide, glycogen, cholesterol and fatty acids in liver slices from embryonic and growing chicks. Biochem. J. 108, 655-661. Goodridge A. G. (1970) Regulation of lipogenesis. Stimulation of fatty acid synthesis in vivo and in vitro in the liver of the newly hatched chick. Biochem. J. 118, 259 263. Goodridge A. G. (1973a) Regulation of fatty acid synthesis in isolated hepatocytes prepared from the livers of neonatal chicks. J. biol. Chem. 248, 1924-1931.
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Goodridge A. G. (1973b) Regulation of fatty acid synthesis in the liver of prenatal and early postnatal chicks. Hepatic concentrations of individual free fatty acids and other metabolites. J. biol, Chem. 248, 1939 1945. Hazelwood R. L. (1972) In Avian Biology (Edited by Farner D. S. and King J. R.), pp. 471-526. Academic Press, New York. Koerker D. J. and Fritz I. B. (1970) Studies on the control of fatty acid oxidation in liver preparations from chick embryos. Can. J. Biochem. 48, 418 424. Langslow D. R. (1972) The development of lipolytic sensitivity in isolated fat cells from Gallus domesticus during the foetal and neonatal period. Comp. Biochem. Physiol. 43B, 689-701. Langslow D. R. (1975) The pancreatic insulin content and its relationship to plasma glucose and free fatty acid concentrations in the embryo and neonatal chick. Br. Poult. Sci. 16, 329-333. Lomax M. A., Donaldson I. A. and Pogson C. I. (1983) The control of fatty acid metabolism in liver cells from fed and starved sheep. Biochem. J. 214, 554-560. McGarry J. D. and Foster D. W. (1972) Regulation of ketogenesis and clinical aspects of the ketotic state. Metabolism 21,471~,89. McGarry J. D. and Foster D. W. (1974) The metabolism of (-)-octanoylcarnitine in perfused livers from fed and fasted rats. J. biol. Chem. 249, 7984~7990. Mooney R. A. and Lane M. D. (1982) Control of ketogenesis and fatty acid synthesis at the mitochondrial branch-point for acetyl-CoA in the chick liver cell: effect of adenosine 3',5'-monophosphate. Eur. J. Biochem. 121, 281-287. Nehlig A., Crone M-C. and Lehr P. R. t1980) Variations of 3-hydroxybutyrate dehydrogenase activity in brain and liver mitochondria of the developing chick. Biochim. biophys. Acta 633, 22 32. Nehlig A. and Lehr P. R. (1982) Activity o1 acetoacetylCoA thiolase and regulation ot" ketone body metabolism in the brain of the developing chick. Brain Reds. 241, 291 297. Ogato K., Watford M., Brady L. J. and Hanson R. W. (1982) Mitochondrial phosphoenolpyruvate carboxy-
kinase (GTP) and the regulation of gluconeogenesis and ketogenesis in avian liver. J. biol Chem. 257, 5385-5391. Page M. A., Krebs H. A. and Williamson D. H. (1971) Activities of enzymes of ketone body utilization in brain and other tissues of suckling rats. Biochem. J. 121, 49 53. Picardo M. and Dickson A. J. (1982) Hormonal regulation of glycogen metabolism in hepatocyte suspensions isolated from chicken embryos. Comp. Biochem. Physiol. 71B, 689 693. Romanoff A. L. (1967) In Biochemist O, ~/ the Avian Embryo. A Quantitatit,e Analysis of Prenatal Development. John Wiley, New York. Sarkar N. K. (1972) Ketone body metabolism in chickens. hit. J. Biochem. 3, Ill 116. S61ing H. D., Mewes W., Unger G. and Kuhn A. (1975) Different regulation of hepatic ketogenesis during starvation in rats and pigeons, hit. J. Biochem. 6, 867 870. Stanley P. E. and Williams S. G. (1969) Use of the liquid scintillation spectrometer for determining adenosine triphosphate by the luciferase enzyme. Analyt. Biochem. 29, 381 392. Sugano T., Shiota M., Khono H. and Shimada M. (1982) Intracellular redox state and control of gluconeogenesis in perfused chicken liver. J. Biochem. 91, t917-1929. Wallace J. C. and Newsholme E. A. (1967) A comparison of the properties of fructose 1,6-diphosphatase and the activities of other key enzymes of carbohydrate metabolism in the livers of embryoic and adult rat, sheep and domestic fowl. Bioehem. J. 104, 378 384. Williamson D. H., Mellanby J. and Krebs H. A. (1962) Enzymatic determination of D( - )-fi-hydroxybutyric acid and acetoacetic acid in blood. Biochem. J. 82, 90-96. Wittmann J. and Weiss A. (1981) Studies on the metabolism of glycogen and adenine nucleotides in embryonic chick liver at the end of incubation. Comp. Biochem. Physiol. 69C, I 6. Yarnell G. R., Nelson P. A. and Wagle S. R. (1966) Biochemical studies of the developing embryo. 1. Metabolism of glyconeogenic precursors. Archs Biochem. Biophys. 114, 539 542. Zammit V. A. (1983) Regulation of hepatic fatty acid oxidation and ketogenesis. Proc. Nutr. Soc. 42, 289-302.
0305-0491/84 $3.00 + 0.00 © 1984 Pergamon Press Ltd
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KETOGENESIS IN CHICK EMBRYO ISOLATED HEPATOCYTES ALISON J. BATE* and ALAN J. DICKSON
Department of Biochemistry, The Medical School, University of Manchester, Manchester M 13 9PT, UK (Tel: 061-273-8241) (Received 21 November 1983)
Abstract--1. Isolated hepatocytes from 17-day old chick embryos exhibit high endogenous rates of ketogenesis. 2. The addition of long-chain fatty acids stimulated ketogenesis with potency ordered as follows: palmitate > oleate > stearate. 3. Octanoate produced a slight stimulation of ketogenesis when added at low concentrations (less than 0.25 mM). At higher concentrations the effect of octanoate was inhibitory. 4. The addition of glucose to incubations failed to lessen endogenous ketogenesis whereas propionate, pyruvate and lactate produced inhibition. 5. Ketogenesis from both endogenous sources and added fatty acids was not altered by the addition of glucagon, insulin, adrenalin or vasopresin.
and organs for different classes of nutrient. As a preliminary stage in investigations of tissuedependent variations in nutrient utilization, we describe in this paper the role of the liver in the production of ketone bodies (D-3-hydroxybutyrate and acetoacetate) from fatty acids. Under conditions of diabetes and prolonged starvation in mammals (nutritional situations analogous to those during chick embryo development) ketones, produced by the liver during conditions of excessive fatty acid availability, can be utilised by several tissues (McGarry and Foster, 1972). Ketone utilization reduces the requirement for glucose and plays an important role in the adaption to decreased glucose availability. Relatively little is known of ketogenesis and its control in avian species. Available data is confined to measurement of ketogenesis with isolated liver preparations from 2-6 week old chickens and pigeons (Bailey and Horne, 1972; Srling et al., 1975; Ogato et al., 1982; Sugano et al., 1982; Zammit, 1983). Measurement of the activity of D-3-hydroxybutyrate dehydrogenase in chick embryo brain suggests that hydroxybutyrate may be an important energy source for this tissue (Nehlig et al., 1980; Nehlig and Lehr, 1982). There is, however, no data available on ketone production by chick embryo liver. In this paper we describe the use of hepatocyte suspensions isolated from embryos 4 days prior to hatching to investigate the control of ketone body synthesis from a range of fatty acids by nutrients and hormones.
INTRODUCTION Growth and development of the embryonic chick is dependent on the mobilization and consequent utilization of nutrients from stores within egg white and yolk. Carbohydrate availability during embryonic development is severely limited. Less than 1% of egg nutrients are carbohydrate and these are mostly utilised within the first 7 days of the 21 day period of incubation (Hazelwood, 1972). Protein from egg white and yolk is degraded to its component amino acids by yolk sac and extra embryonic membranes (Hazelwood, 1972). Following their uptake into embryonic blood, while amino acids are available to promote protein synthesis and growth, certain amino acids may be catabolised for energy provision or enter the gluconeogenic pathway of liver, kidney and muscle (Yarnell et al., 1966; Dickson, 1983). Although no data are available on issue amino acid catabolism, considerable information exists on the role of amino acids in gluconeogenesis. Conversion of amino acids to glucose or glycogen results in the elevations in plasma glucose concentration and, in liver and muscle, tissue glycogen contents during the later stages of embryonic development (Freeman, 1969; Langslow, 1975; Wittman and Weiss, 1981). Studies with isolated tissue preparations from embryonic chicken liver have shown the importance of the gluconeogenic pathway in that tissue (Yarnell et al., 1966; Dickson, 1983). Stored lipid within the egg makes up 50~ of available nutrients (Romanoff, 1967). Data on respiratory quotients indicate that, especially during the final 7 days of embryonic development, fatty acids are the major oxidised substrate for tissue metabolism (Hazelwood, 1972). Although fatty acids are a major fuel for the embryo as a whole, there has been no definition of the requirements of individual tissues *Correspondence to be addressed to: Alison J. Bate.
MATERIALS AND
METHODS
Sources of animals and materials Fertilised White Leghorn (Gallus domesticus) eggs were
obtained from the University of Manchester Medical School and used after 17 days of incubation (4 days prior to hatching). Adrenalin, firefly lantern extract (FLE-50), glucagon, insulin, L-lactic acid, n-octanoic acid, palmitic acid, stearic acid and vasopressin were all obtained from Sigma, Poole, Dorset, UK. ATP, collagenase, dibutyryl cyclic AMP, D-3-hydroxybutyrate dehydrogenase, NAD +,
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ALISON J. BATE a n d ALAN J. DICKSON
NADH and pyruvate were purchased from Boehringer Corporation Ltd., Lewes, Sussex, UK. All other biochemicals (Analar grade) were supplied by B.D.H. Chemicals Ltd., Poole, Dorset, UK. Bovine serum albumin (BSA) was purchased from Park Scientific Ltd., Northampton, UK and was defatted and dialysed prior to use (Chen, 1967). Hepatocvte isolation and incubation Chick embryo hepatocytes were isolated by the collagenase digestion method of Picardo and Dickson (1982). Portions (0.5 ml) of the final cell suspension which contain on average 1.5 2.0 mg dry wt of cells were incubated in stoppered 5 ml polypropylene vials. The Krebs Henseleit bicarbonate buffer used for incubations contained BSA at a final concentration of 11!;. The vials were flushed with O2:CO 2 (19: 1) gas mixture and incubated with shaking at 37C. Reactions were terminated by the addition of perchloric acid (final concn, 0.2 M) to the entire ceil suspension. After centrifugation (2500g, 5min), portions of the supernatant were analysed for acetoacetate, D-3hydroxybutyrate and ATP. Analytical procedures Perchloric acid extracts were neutralised with 2 M KOH containing 0.5 M triethanolamine prior to analysis. ATP was measured by a modification of the luminescent assay of Stanley and Williams (1969). D-3-Hydroxybutyrate and acetoacetate were determined by a modification of enzymatic procedure of Williamson et al. (1962). For both metabolites, 0.1 ml of neutralised cell extracts were analysed in a final cuvette vol of 0.5ml. The assay medium for D-3-hydroxybutyrate contained 1.9 M Tris, 0.32 M hydrazine sulphate, 8 mM MgSO4, 8 mM EDTA and 2.4 mM NAD + (final pH 9.8). For acetoacetate the assay medium consisted of 80mM triethanolamine, pH 7.4 containing 0.14 mM NADH. Expression qf results Portions of cell suspensions were dried to constant weight at 120'C and values were used to calculate results in terms of dry weight.
chain fatty acids (Fig. 1: Table 1). ]'he percentage stimulation caused by each agent was similar at all time points. For the experiments reported in Fig. I and Table 1, palmitate (0.5raM) and octanoate (0.125 mM) increased the hydroxybutyrate:acetate ratio by 14.9°,0 and 29.57~; respectively. This effect was observed at all time points studied. Oleate (0.5 mM), stearate ( l . 0 m M ) and butyrate (4.0raM) each decreased the ratio by 4(~507;; at all incubation times. The concentration dependence of ketogenesis on exogenous fatty acid varied between fatty acids (Fig. 2). Although palmitate, oleate and stearale gave similar degrees of stimulation of ketogenesis, the concentration required to attain this maximal sensitivity could be ordered: palmitate < oleate < stearate. Butyrate stimulation of ketogenesis was slight and the maximal effect produced was only 60!!1; of that attained with palmitate, oleate and stearate. Octanoate at concentrations below 0.25 mM produced a slight stimulation of ketogenesis. Further increases in octanoate concentration inhibited ketogenesis. Addition of the ketogenic precursors cited above produced minor effects on hepatocyte viability. Only octanoate reduced cell ATP content (by 5 15";, at incubation periods of up to 1 hr). To investigate the interaction of other possible plasma constituents with ketogenesis from endogenous and exogenous sources, experiments were performed over 60 min with or without additions of metabolites and hormones. Glucose (at concentrations up to 20 mM) had no effect on ketone body formation whereas fructose (10 mM) caused a 40",; inhibition. Addition of propionate, lactate and pyruvate produced significant inhibition at concentrations from 0.25-10 mM (Table 2). The effects of lactate and
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RESULTS The cell preparations from which results on ketogenesis were obtained had initial A T P contents in the range of 7.89 _+ 0.45 (12) nmoles/mg dry wt. This value changed by less than 2.3'I o over a 1 hr period of incubation. On this basis, the preparations used show close to physiological viability (Picardo and Dickson, 1982) and metabolic processes such as ketogenesis are unlikely to be impaired as a result of hepatocyte isolation. Hepatocyte suspensions maintained a constant rate of endogenous ketogenesis for incubations of up to 2 hr (results not shown). The major ketone produced was D-3-hydroxybutyrate. The ratio of hydroxybutyrate:acetoacetate in the extract progressively increased through incubation. After 15, 30, 60 and 120rain of incubation the ratio was 3.2_+0.4, 5.2 _+ 0.8, 17.8 _+ 4.3 and 53.2 + 8.9 respectively (from 4 experiments). The amount of acetoacetate produced was small and progressively of minor importance to the total sum of ketones synthesised during incubations. Except where stated, results are expressed only as total ketones produced by hepatocyte suspensions. Subsequent experiments were restricted to 60 min duration. The rapid endogenous ketogenesis was accelerated by the addition of long- and medium-
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40
50
60
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Fig. 1. Time course of endogenous (O) and oleatestimulated (11) ketogenesis. Hepatocytes were prepared from 17-day-old chick embryo livers as described in Materials and Methods. Results are expressed as total ketones (hydroxybutyrate plus acetoacetate) formed in incubations. Final oleate concentration in the incubations was 0.5 mM. Values are means _+SEM for three separate cell experiments.
Ketogenesis in chick embryo isolated hepatocytes
439
Table 1, Stimulation of endogenous ketogenesis by fatty acid addition* % of endogenous ketogenesis incubation time (min)
Added fatty acid
30 187.2 + 13.5 213.5 + 13.1 197.4 _+ 8.8 109. I + 3.8 152.4 _+ 8.0
15
Oleate (0.5 mM) Palmitate (0.5 mM) Stearate (1.0 mM) Octanoate (0.125 mM) Butyrate (4.0 raM)
171.3 + 16.9 N.D. 182.9 +_ 8.6 124.2 + 2.8 125.0 + 7.3
60 195.2 + 10.9 209.4 _+ 8.6 201.7 +- 4.3 125.3 + 4.7 167.4 + 4.5
*The final concentrations of added fatty acids are indicated in parentheses. Ketogenesis is expressed as the sum of hydroxybutyrate and acetoacetate. Methods for the isolation and incubation of hepatocytes and the determination of ketones is given in Materials and Methods. In each case values are the means+_SEM for thlee separate cell preparation (N.D. = not determined).
pyruvate were observed both when the metabolites were added singly or in combination (in a ratio of 9: 1 respectively). Inhibition occurred both for ketogenesis from endogenous sources and from added palmitate (0.25 raM). Ketogenesis was unaltered by the addition of glucagon, dibutyryl cyclic AMP, insulin, adrenalin or vasopressin (Table 3). Ineffectiveness of glucagon, dibutyryl cyclic AMP and adrenalin on ketogenesis was observed although each hormone stimulated glycogenolysis in parallel incubations (Picardo and Dickson, 1982).
the presence of fatty acids is equivalent to that found for fed or starved perfused rat and pigeon livers (S61ing et al., 1975). Hydroxybutyrate is the major product of ketogenesis from exogenous and endogenous fatty acid catabolism. Although the precise value for the hydroxybutyrate/acetoacetate ratio varied with the incubation time, the ratio was consistently high (from 275
250
DISCUSSION
The results presented in this paper are the first comprehensive report of hepatic ketogenesis and its control during chick embryonic life. A previous report described the production of ketones from [1J4C]palmitate by liver slices from 10 day old chick embryos (Koerker and Fritz, 1970). This report was brief and the use of liver siices is a technique of questionable value today. Our results indicate that ketogenesis is a major metabolic pathway in embryonic liver metabolism. High rates of ketogenesis (91.9 _+ 3.2 nmoles/hr/mg dry wt) were observed in the absence of any added fatty acids. Hepatocytes from embryonic chicks contain large lipid droplets within their cytosolic structure and have a high fat content as indicated by a 50% fat-free dry weight estimate determined for such a tissue (Goodridge, 1973a). The anoxia and stress incurred throughout hepatocyte isolation stimulates the mobilization of fatty acids from hepatic cytosolic stores and permits their entry into the processes of fatty acid oxidation. Hepatocyte stores of fatty acids sustain a constant rate of ketogenesis for incubation periods of over 1 hr (Fig. 1). The rate observed over this period, in the absence of added fatty acids, is close to 50% of that obtained for rat and pigeon isolated liver preparations in the presence of exogenous fatty acids (S61ing et al., 1975; Demaugre et aL, 1983). Addition of fatty acids to chick embryo hepatocytes accelerated ketone body synthesis by approximately two-fold (Table 1; Fig. 1). Once maximal ketogenesis was attained, further addition of fatty acids failed to further stimulate conversion to ketones. Limitations in mitochondrial transport will restrict fatty acid activation at the initial stages of ketone formation. At maximum rate, ketogenesis in
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o
175
2 150
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OI Final
02
03
04
05
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2
3
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4
5
(mMI
Fig. 2. Modulation of hepatocyte ketogenesis by palmitate (O), stearate ([]), oleate (O), butyrate (A) and octanoate (11). Hepatocytes were incubated for 60 min and the effects of additions on total ketone body formation are expressed as % of the endogenous rate. Each addition represents a single hepatocyte preparation and the individual points are the means + SD for 3 separate determinations,
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ALISON J. BATE a n d ALAN J. DICKSON
Table 2. Inhibition of endogenous ketogenesis by propionate, lactate and pyruvate additions* Final concentration of addition (raM) 0.25 0.5 1.0 2.5 5.(1 10.0
'?i, of endogenous ketogenesis Lactate/pyruvate Propionate mixture 75.2 + 3.8 70.0 + 4.1 66.3 2 4 . 4 50.9 ~ 2.5 43.2 _~ 3.5 33.6_+ 2.4
74.5 -+ 5.4 52.9 ± 1.9 31.6_+ 5.7 15.2 ± 3.7 18.4 _+ 2.1 36.5 + 3.1
*The mixture of lactate and pyruvate was m a d e up to give the final concentration indicated with components in a ratio o[" 9:1 respectively. Ketogenesis was determined as the total sum of hydroxybutyrate and acetoacetate produced over a 60 min incubation period. Values are the means + SEM for 3 separate cell preparations in each case, Results are expressed as % of the endogenous rate (endogenous ketogenesis was 82.8 _+ 7.0 and 1 0 1 . 0 + 6 . 2 n m o l e s ketones produced/hr/mg dry cells in the experiments for propionate and lactate/pyruvate mixture respectively).
5 20). In mammalian blood the ratio of ketones would be between 2-3 in both the fed, starved and severely diabetic conditions (Page et al., 1971: Sarkar, 1972; Blackshear et al., 1975). Avian species have consistently high plasma hydroxybutyrate:acetoacetate ratios. Ratios between 6 and 20 are normal for fed and starved chickens and pigeons after hatching (Bailey et al,, 1971: Bailey and Horne, 1972: Sarkar, 1972; Davison and Langslow, 1975; Brady et al., 1978; Zammit, 1983). Plasma from 17 day old chick en embryos contained 2.3 2.5 mM total ketones in a ratio of24:1 (hydroxybutyrate:acetoacetate) (A. J. Bate and A. J. Dickson, unpublished results). Thus the ratio of ketones produced by isolated chick embryo hepatocytes is an accurate reflection of the situation in ovo. High ratios of hydroxybutyrate:acetoacetate may be a consequence of limitations in the transfer of reducing equivalents from mitochondria to cytosol in chicken liver, an event of importance in the control of gluconeogenesis from lactate and pyruvate (Dickson and Langslow, 1978; Bannister and O'Neill, 1981: Ogato et al., 1982; Sugano et al., 1982). Within embryonic liver, the major fatty acids available are palmitate, stearate and oleate (34, 34 and 30°o of total fatty acids respectively) (Goodridge, 1973b). The potency of these fatty acids upon ketogenesis when added exogenously to hepatocytes underlines that they will play an important physiological role in ketone body formation hi ovo. Utilization of fatty acids produced one of two effects on
hydroxybutyrate:acetoacetate ratios: (a) an increase of 20°J~, (with palmitate or octanoate); (b) a decrease of 40~500~, (with oleate, stearate or butyrate). Rapid oxidation of fatty acids should produce a high N A D H : N A D ÷ ratio within mitochondria. This should in turn depress the hydroxybutyrate:acetoacetate ratio. The observed depression in the ketone ratio is likely to be a result of high rates of fatty acid oxidation. Increased ratios observed with palmitate and octanoate cannot be explained from the available background material. Octanoate exerts a dual effect on ketogenesis (Fig. 2). At concentrations below 0.25 mM, this medium chain fatty acid produced a small stimulation of ketogenesis. At higher concentrations a progressive inhibition occurred. The transport of medium chainlength fatty acids (e.g. octanoate) into mitochondria is not subject to limitations at the level of membrane permeation (Fritz, 1959: McGarry and Foster, 1974). Increases in octanoate concentration will result in high intramitochondrial concentrations of this fatty acid and the mitochondrial pool of coenzyme A will be sequestered in the form of an octanoyl-coenzyme A complex. When sufficiently high concentrations of coenzyme A are complexed, inadequate amounts will be available for acetyl-coenzyme A t~rmation fiom fatty acid catabolism. This mechanism would reduce the rate of ketogenesis as a consequence of inadequate precursor (acetyl coenzyme A) supply. Data on the effects of propionate, lactate and pyruvate (see later sections) support the concept of ketogenesis limitations as a result of coenzyme A availability. Although there are reports on the use of octanoate (1 mM, final concentration) as a ketogenic precursor in isolated chicken liver preparations (Ogato eta/.. 1982; Sugano et al., 1982). such reports are for birds after hatching and represent a situation with low basal rates of ketogenesis. Competition between octanoate and endogenous fatty acids is liable to be much reduced under such conditions and inhibition of ketogenesis may only be apparent at high concentrations of octanoate (i.e. from above 1.0 raM). The use of octanoate in mammalian liver preparations (e.g. S61ing et al., 1975) may be possible at concentrations of 1 2 m M for the same reason. Interactions between the metabolism of likely plasma constituents and ketogenesis in chick embryos were investigated by addition of glucose, lactate and pyruvate to liver cell suspensions (Table 2). Glucose, which wilt be present in plasma oi" 17-day-old embryos at a concentration of 7 8raM (Langslox~,
Table 3. Effects of hormone addition on ketogenesis from endogenous and exogenous fauy acids* ?i, of control ketogcnesis From endogenous In the presence of fatty acids added palmitate Glucagon (2.87 x 10 r M ) Dibutyryl cyclic A M P (5.0 x 10 5 M) Insulin (1.74 x 10 7 M) Insulin + glucagon Adrenalin (1.1 x 10 5 M) Vasopressin (2.47 x l0 ' M)
102.9+2.8 99.1 4_-4 6 106.3 _+ 4.1 106.6 + 3.6 102.6_+ 8.6 110.6 + 5.0
101.5~ 1.0 N.D. 99.4 2 2.2 100.3 ± 2.2 105.3 -+ 3.6 102.4 + 6.5
*Incubations were for 60 min. Ketogenesis was determined as the sum of hydroxybutyrate and acetoacetate production. The control rate describes the extent of ketone production from endogenous sources or palmitate (at a final concn of 0,25 raM) alone and the response to hormones expressed as a percentage of the control value in each case. Values are means + SEM fl~r 3 separate cell preparations in each case.
Ketogenesis in chick embryo isolated hepatocytes 1975), had no effect on ketogenesis, As hepatic glycolytic enzymes are of low activity until after hatching (Wallace and Newsholme 1967) and lipogenesis from glucose is absent prior to hatching (Goodridge, 1968, 1970), the ineffectiveness of glucose probably results from a lack of glucose catabolism by embryonic liver. The effect of lactate and pyruvate upon ketogenesis was observed with mixtures or individual solutions of the components (Table 2). As each component was individually inhibitory on ketogenesis, the mode of action is not related to alterations in redox state produced in cellular metabolism. The effects would appear to be linked to the subsequent metabolism of the carbon skeleton of lactate and pyruvate. That propionate gives rise to the same maximal inhibition of ketogenesis as lactate and pyruvate (Table 2) suggests a common mechanism of action. It appears likely that the common mechanism is the utilisation of coenzyme A in entrance of propionate, pyruvate and lactate into the tricarboxylic acid cycle. Reduction of coenzyme A levels in mitochondria will impair the activation of fatty acids and result in reduced rates of ketogenesis. Inhibition of ketogenesis by addition of propionate, lactate and pyruvate has been observed with isolated hepatocytes from other species (Lomax et al., 1983). Although the precise mechanism for inhibition remains uncertain, the data presented here for chick embryo hepatocytes favour the possibility of regulation through competition for coenzyme A utilization. Whatever the mechanism, inhibition of ketogenesis by lactate and pyruvate will be important for the regulation of ketogenesis in ovo. As plasma lactate concentrations in chick embryos during the last few days of incubation are in the range of 0.6-0.7 mM (Dickson, 1983) maximal possible rates of ketogenesis will be reduced by up to 50~o. Although glucagon and insulin are known to play important roles in the acute regulation of fatty acid esterification and oxidation in hepatocytes from chicks after hatching (Goodridge, 1973a; Mooney and Lane, 1982), with concomitant effects on ketogenesis, there was no effect of these hormones on ketone body formation in 17-day-old chick embryo hepatocytes (Table 3). This occurred at a time when glucagon increased glycogenolysis (Picardo and Dickson, 1982) and produced dramatic increases in hepatocyte cyclic A M P concentrations (unpublished work). The lack of effect with dibutyryl cyclic A M P further emphasises that unresponsiveness is unrelated to receptor numbers of sensitivity but can be restricted to intracellular reception of hormonal stimulation. Previous work with hepatocytes (Picardo and Dickson, 1982) and adipocytes (Langslow, 1972) have shown similar diminished responses to glucagon at this stage of chick embryonic life. Adrenalin, which acts through a cyclic AMP-mediated mechanism in chick embryos (Picardo and Dickson, 1982) and the calcium-mobilising hormone, vasopressin, were also ineffective. The data presented in this paper indicates that ketogenesis is a major metabolic pathway in chick embryo liver. The provision of ketones as an alternative fuel for several tissues may be an essential part of the metabolic homeostasis in the development of chick embryos. Although the nature of the tissues
441
which may utilise ketones has not been determined, data from chick embryo brain indicates high concentrations of hydroxybutyrate dehydrogenase (Nehlig et al., 1980; Nehlig and Lehr, 1982). In the development of chicks from embryos to neonates plasma fatty acid concentrations decrease from 1.2 to 0.3mM (Freeman and Manning, 1971; Langslow, 1975). In parallel with this, plasma ketone concentrations drop from 2.4mM in 17-day-old embryos (this paper) to 0.6 mM in neonates (Sarkar, 1972). Thus ketones provide large amounts of alternative nutrients for the chick embryo at a time of high plasma fatty acid concentration. REFERENCES
Bailey E. and Home J. A. (1972) Formation and utilization of acetoacetate and o-3-hydroxybutyrate by various tissues of the adult pigeon. Comp. Biochem. Physiol. 42B, 659-667. Bailey E. J., Horne J. A., lzatt M. E. G. and Hill L. (1971) Concentrations of acetoacetate and 3-hydroxybutyrate in pigeon blood and desert desert locust haemolymph. Life Sci. 10, 1415-1415. Bannister D. W. and O'Neill I. E. (1981) Control of gluconeogenesis in chick (Gallus domesticus) isolated hepatocytes: effect of redox state and phosphoenolpyruvate carboxykinase [EC 4.1.1.32] location. Int. J. Biochem. 13, 437-444. Blackshear P. J., Holloway P. A. H. and Alberti K. G. M. M. (1975) Metabolic interactions of dichloroacetate and insulin in experimental diabetic ketoacidosis. Biochem. J. 146, 447-456. Brady L. J., Romsos D. R., Brady P. S., Bergen W. G. and Leveille G. A. (1978) The effects of fasting on body composition, glucose turnover, enzymes and metabolites in the chicken. J. Nutr. 108, 648-657. Chen R. F. (1967) Removal of fatty acids from serum albumin by charcoal treatment. J. biol. Chem. 242, 173-181. Davison T. F. and Langslow D. R. (1975) Changes in plasma glucose and liver glycogen following the administration of gluconeogenic precursors to the starving fowl. Comp. Biochem. Physiol. 52A, 645-649. Demaugre F., Buc H. A., Cepanec C., Monacion A. and Leroux J-P. (1983) Comparison of the effects of 2-chloropropionate and dichloroacetate on ketogenesis and lipogenesis in isolated rat hepatocytes. Biochem. Pharmac. 32, 1881-1885. Dickson A. J. (1983) Gluconeogenesis in chick embryo isolated hepatocytes. Int. J. Biochem. 15, 861-865. Dickson A. J. and Langslow D. R. (1978) Hepatic gluconeogenesis in chickens. Molec. Cell. Biochem. 22, 167-181. Freeman B. M. (1969) The mobilization of hepatic glycogen in Gallus domesticus at the end of incubation. Comp. Biochem. Physiol. 28, 1169-1176. Freeman B. M. and Manning A. C. C. (1971) Glycogenolysis and lipolysis in Gallus domesticus during the perinatal period. Comp. Gen. Pharmac. 2, 198-204. Fritz I. B. (1959) Action of carnitine on long chain fatty acid oxidation by liver. Am. J. Physiol. 197, 297-304. Goodridge A. G. (1968) Conversion of [U-~4C]glucoseinto carbon dioxide, glycogen, cholesterol and fatty acids in liver slices from embryonic and growing chicks. Biochem. J. 108, 655-661. Goodridge A. G. (1970) Regulation of lipogenesis. Stimulation of fatty acid synthesis in vivo and in vitro in the liver of the newly hatched chick. Biochem. J. 118, 259 263. Goodridge A. G. (1973a) Regulation of fatty acid synthesis in isolated hepatocytes prepared from the livers of neonatal chicks. J. biol. Chem. 248, 1924-1931.
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Goodridge A. G. (1973b) Regulation of fatty acid synthesis in the liver of prenatal and early postnatal chicks. Hepatic concentrations of individual free fatty acids and other metabolites. J. biol, Chem. 248, 1939 1945. Hazelwood R. L. (1972) In Avian Biology (Edited by Farner D. S. and King J. R.), pp. 471-526. Academic Press, New York. Koerker D. J. and Fritz I. B. (1970) Studies on the control of fatty acid oxidation in liver preparations from chick embryos. Can. J. Biochem. 48, 418 424. Langslow D. R. (1972) The development of lipolytic sensitivity in isolated fat cells from Gallus domesticus during the foetal and neonatal period. Comp. Biochem. Physiol. 43B, 689-701. Langslow D. R. (1975) The pancreatic insulin content and its relationship to plasma glucose and free fatty acid concentrations in the embryo and neonatal chick. Br. Poult. Sci. 16, 329-333. Lomax M. A., Donaldson I. A. and Pogson C. I. (1983) The control of fatty acid metabolism in liver cells from fed and starved sheep. Biochem. J. 214, 554-560. McGarry J. D. and Foster D. W. (1972) Regulation of ketogenesis and clinical aspects of the ketotic state. Metabolism 21,471~,89. McGarry J. D. and Foster D. W. (1974) The metabolism of (-)-octanoylcarnitine in perfused livers from fed and fasted rats. J. biol. Chem. 249, 7984~7990. Mooney R. A. and Lane M. D. (1982) Control of ketogenesis and fatty acid synthesis at the mitochondrial branch-point for acetyl-CoA in the chick liver cell: effect of adenosine 3',5'-monophosphate. Eur. J. Biochem. 121, 281-287. Nehlig A., Crone M-C. and Lehr P. R. t1980) Variations of 3-hydroxybutyrate dehydrogenase activity in brain and liver mitochondria of the developing chick. Biochim. biophys. Acta 633, 22 32. Nehlig A. and Lehr P. R. (1982) Activity o1 acetoacetylCoA thiolase and regulation ot" ketone body metabolism in the brain of the developing chick. Brain Reds. 241, 291 297. Ogato K., Watford M., Brady L. J. and Hanson R. W. (1982) Mitochondrial phosphoenolpyruvate carboxy-
kinase (GTP) and the regulation of gluconeogenesis and ketogenesis in avian liver. J. biol Chem. 257, 5385-5391. Page M. A., Krebs H. A. and Williamson D. H. (1971) Activities of enzymes of ketone body utilization in brain and other tissues of suckling rats. Biochem. J. 121, 49 53. Picardo M. and Dickson A. J. (1982) Hormonal regulation of glycogen metabolism in hepatocyte suspensions isolated from chicken embryos. Comp. Biochem. Physiol. 71B, 689 693. Romanoff A. L. (1967) In Biochemist O, ~/ the Avian Embryo. A Quantitatit,e Analysis of Prenatal Development. John Wiley, New York. Sarkar N. K. (1972) Ketone body metabolism in chickens. hit. J. Biochem. 3, Ill 116. S61ing H. D., Mewes W., Unger G. and Kuhn A. (1975) Different regulation of hepatic ketogenesis during starvation in rats and pigeons, hit. J. Biochem. 6, 867 870. Stanley P. E. and Williams S. G. (1969) Use of the liquid scintillation spectrometer for determining adenosine triphosphate by the luciferase enzyme. Analyt. Biochem. 29, 381 392. Sugano T., Shiota M., Khono H. and Shimada M. (1982) Intracellular redox state and control of gluconeogenesis in perfused chicken liver. J. Biochem. 91, t917-1929. Wallace J. C. and Newsholme E. A. (1967) A comparison of the properties of fructose 1,6-diphosphatase and the activities of other key enzymes of carbohydrate metabolism in the livers of embryoic and adult rat, sheep and domestic fowl. Bioehem. J. 104, 378 384. Williamson D. H., Mellanby J. and Krebs H. A. (1962) Enzymatic determination of D( - )-fi-hydroxybutyric acid and acetoacetic acid in blood. Biochem. J. 82, 90-96. Wittmann J. and Weiss A. (1981) Studies on the metabolism of glycogen and adenine nucleotides in embryonic chick liver at the end of incubation. Comp. Biochem. Physiol. 69C, I 6. Yarnell G. R., Nelson P. A. and Wagle S. R. (1966) Biochemical studies of the developing embryo. 1. Metabolism of glyconeogenic precursors. Archs Biochem. Biophys. 114, 539 542. Zammit V. A. (1983) Regulation of hepatic fatty acid oxidation and ketogenesis. Proc. Nutr. Soc. 42, 289-302.