Regulation of fatty acid oxidation in mammalian liver

227

Biochirrticaet BiophysicsActu, 1167 (1993) 227-241 0 1993 Elsevier Science Publishers BV. Ah rights reserved ~5-27~/93/$~.~

Review

BBALIP 54149

Regulation of fatty acid oxidation in mammalian liver

’

Manuel GuzmiirP and Math J&i. Geefen” aDepartmentufBiochemistryand MolecularBiology4 Faculty ofChemistry,Compl’utenseUniversity,Madrid (Spain1 and b Laboratoryof VeterinaryBiochemistry,Vtrecht Vniuersity,Utrecht(The Netherlands) (Received 15 October 1992)

Key words: Fatty acid oxidation; Membrane, outer; Ketogenesiq ~it~hondrion; Carnitine p~mitoyltransferase; 3-~yd~~-3-methylgluta~l-~oen~me A synthase; (Mammalian liver]

Contents ...............................

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I.

Intr~~ctio~

II.

Uptake and activation of fatty acids ..............

III.

Translocation of fatty acids into mitochondria

IV.

~-O~datian

V.

Ketogenesis

VI.

Tricarboxyhc acid cycte activity ..................

VII.

Peroxisomal fatty acid oxidation .................

VIII.Futurep~~ects.. Ac~owl~dgements References

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of fatty acids. .....................

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I. Introduction Depending on the ~hysio~athologi~al status of the animal, the liver is a tissue that can express either high

Correspondence to: M.J.H. Geelen, Laboratory of Veterinary Biochemistry, Utrecht University, P.O. Box 80.176, 3508 I’D Utrecht, The Netherlands. t Dedicated to Dr. S.G. van den Bergh at the occasion of his retirement as Professor of Veterinary Bi~hemist~. Abbreviations: CoA, coenzyme A; CPT, carnitine palmitoyltransferase; CPTi ( = CPT-II, CPT-B), CPT, ( = CPT-I, CPT-Al and CPT,, CPT activity located in the mit~hondriai inner membrane, mitochondrial outer membrane and peroxisomes, respectively; FABP, fatty acid-binding protein; HMG-CoA, 3-hydro~-3-methylgIuta~ICoA, HSL, hormone-sensitive Iipase; PMA, 4&phorbol 12@myristate lfa-acetate.

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228 230 233 233 236 237

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rates of lipogenesis or high rates of fatty acid oxidation. It is thus very important that these two processes should be inversely controlled. Hence a number of regulatory mechanisms exist which ensure that fatty acid synthesis is blunted when fatty acid oxidation is activated, and vice versa. Wepatic fatty acid oxidation plays an essential role as a source of energy not only for the liver, but also for extrahepatic tissues; thus, under different catabolic situations the liver supplies e~rahepati~ tissues (including the brain) with ketone bodies as a giu~se-replating fuel (Fig. 1X Availabili~ of the fatty acid substrate is not the only factor determining the capacity of the liver to oxidize fatty acids. In addition, and more im~rtantly, it involves a number of fine-tuning mech~isms that control the flux through putative regulatory steps, such as the reactions catalyzed by the mitochondrial outer membrane carnitine

228 palmitoyltransferase (= CPT,, CPT-I, CPT-A) or by the mitochondrial 3-hydroxy-3-methylglutaryl-CoA (HMG-CoA) synthase. Two excellent reviews on hepatic fatty acid oxidation were published at the beginning of the 1980’s [1,2]. Moreover, other thorough reviews have dealt with partial aspects of the fatty acid-oxidative pathway (see below). Hence the present review is not intended to be comprehensive. Instead, it focuses on recent findings regarding the molecular processes that govern the control of fatty acid oxidation in liver and on the physiological (hormonal, nutritional) mechanisms underlying the coordinate control of several regulatory enzymes involved in this process. II. Uptake and activation

of fatty acids

The liver metabolizes fatty acids from both endogenous and exogenous origin. Endogenous fatty acids include those synthesized de novo from acetyl-CoA and those released upon lipolysis of hepatocellular triacylglycerol stores. Exogenous sources of fatty acids are the fatty acid-albumin complexes and the lipid components of circulating lipoproteins. Fatty acid synthesis de novo is achieved by the consecutive action of acetyl-CoA carboxylase and fatty acid synthase. The former is generally considered to catalyze the rate-limiting step of the fatty acid-synthe-

sizing process in both liver and most extrahepatic tissues (reviewed in Refs. 3,4). As will be discussed below, the product of the reaction catalyzed by acetyl-CoA carboxylase - i.e., malonyl-CoA - is precisely the physiological inhibitor of CPT,, one of the key regulatory enzymes of long-chain fatty acid oxidation. This has traditionally offered a feasible explanation for the coordinate control of fatty acid synthesis and oxidation in the liver [1,2]. Nevertheless, it is currently agreed that malonyl-CoA concentration within the hepatocytc is not the only factor involved in the control of the fatty acid-oxidative process (see below). Very little is known about the mobilization of hepatic triacylglycerols. Rat liver contains a lysosomal triacylglycerol lipase which is responsible for the hydrolysis of intracellular triacylglycerol stores (reviewed in Ref. 5). This lipase is immunologically different from the well-characterized hormone-sensitive lipase (HSL) from adipose tissue, heart and skeletal muscle [6]. No regulatory mechanism of hepatic triacylglycerol lipase has been unequivocally established. It has been suggested that phagocytosis of cytosolic fat droplets by lysosomes (autophagocytosis) is the key point in the control of the overall process of hepatic lipolysis [5]. Previous observations indicated that glucagon rapidly induces the formation of hepatic autophagic vacuoles, whereas insulin has the opposite effect [7]. However, Seglen and co-workers demonstrated more recently

VLDL

BLOODSTREAM

4 I

I

TRIACYLGLYCEROLS

PEROXISOME

LIPOPROTEINS

FATTY ACYL-Co’4 MITOCHONDRION

FATTY ACIDS

MALONYL-CoA *

KETONE BODIES

ACETY L-&A 7

Fig. 1. Overview of the fatty acid-oxidative process in the liver. After activation to their corresponding coenzyme A thioesters, fatty acids from both endogenous and exogenous sources may undergo p-oxidation upon translocation into the mitochondrial matrix. In addition, peroxisomes may make a complementary contribution to fatty acid p-oxidation. Inside mitochondria, the acetyl moieties produced by these two P-oxidizing systems may subsequently be converted to ketone bodies via the ketogenesis pathway. Alternatively, acetyl-CoA may be further oxidized to CO, by the action of the tricarboxylic acid cycle. (0) Major regulatory enzymes of fatty acid oxidation: 1, CPT,; 2, HMG-CoA synthase. (0) Enzymes which might play an additional role in the regulation of fatty acid oxidation: 3, CPT,; 4, acyl-CoA dehydrogenase; 5, p-3-hydroxybutyrate dehydrogenase; 6, isocitrate dehydrogenase; 7, 2-oxoglutarate dehydrogenase. (e) Enzymes not directly involved in fatty acid oxidation but which might indirectly play a regulatory role in fatty acid oxidation: 8, acetyl-CoA carboxylase; 9, phosphatidate phosphohydrolase; 10, diacylglycerol acyltransferase. HLPL, hepatic lipoprotein lipase; VLDL, very low density lipoprotein; AcAc, acetoacetate; OHBut. 3-hydroxybutyrate.

229 that hepatocytic autophagy is inhibited by CAMP analogues, phosphodiesterase inhibitors and phosphatase inhibitors [S-lo]. Clearly, further research is required to elucidate how the hydrolysis of hepatocellular triacylglycerols is controlled by hormones [S-13]. Ketotic states are usually characterized by an increase in circulating non-esterified long-chain fatty acids complexed with albumin [2]. Adipose tissue triacylglycerol is quantitatively the most important energy store in the organism, and thus plasma free fatty acid levels are mostly determined by the activity of adipose tissue HSL (reviewed in Ref. 14). It is well-established that this lipase is subject to regulation by phosphorylation-dephosphorylation; thus, incubation of adipocytes with lipolytic agents such as norepinephrine increases the phosphorylation state and the activity of HSL, whereas insulin treatment is associated with a decrease in the phosphorylation state and the activity of HSL [14]. Fatty acids are taken up very efficiently by the liver. Like in other tissues, fatty acid uptake by the liver is greatly dependent on the concentration of fatty acids in the extracellular medium. However, the molecular basis of the translocation process across the plasma membrane is still a matter of debate. It has been traditionally assumed that fatty acids penetrate the plasma membrane by a simple diffusion mechanism that is non-saturable [l&161. During the last few years, however, certain evidence has accumulated supporting the existence of a carrier-mediated, saturable uptake system (reviewed in Ref. 17). Thus, Stremmel and co-workers identified a 40 kDa membrane fatty acidbinding protein (FABP) which was proposed to function as a Na+/long-chain fatty acid cotranspo~er [18201. In fact, hepatic fatty acid uptake was depressed either by antibodies raised against that membrane protein [19-211 or by supression of the transmembrane Na+ gradient [19,221. In contrast with the well-established notion that fatty acid uptake is tightly coupled to intracellular fatty acid metabolism, it has been suggested that hepatic influx of long-chain fatty acids reflects membrane transport rather than intracellular metabolism or binding [23]. Nevertheless, experiments performed by Zakim’s group fully support the classical notion that the uptake of long-chain fatty acids by the liver is due to a series of spontaneous, uncatalyzed reactions [24-271. As recently determined by Ferraresi-Filho et al. [281, rates of influx of oleate across the hepatocyte plasma membrane do not fit with the experimental evidence obtained by Stremmel’s group. Hormonal control of fatty acid uptake has been shown to occur in isolated adipocytes. In these cells, epinephrine causes a stimulation of uptake which is counteracted by insulin [29]. Such a mechanism of control in liver tissue is currently under study (Stremmel, W., personal communication).

The liver is also able to use fatty acids from the lipid components (mostly triacylglycerols) of plasma lipoproteins. Membrane receptors are present in the liver for intermediate-density lipoproteins and low-density lipoproteins (ape B/E receptor), for chylomicron remnants and apo E-rich high-densi~ lipoproteins (ape E receptor, also known as ‘LDL-receptor related protein’), and for the subfractions 2 and 3 of high-density lipoproteins (ape A-I/A-II receptor). Hepatic lipoprotein lipase - located on the surface of endothelial cells - as well as the aforementioned hepatic lysosomal triacylglycerol lipase - located inside parenchymal cells - take part in the process by which lipoprotein-triacylglycerol is hydrolysed to give intracellular fatty acids, although the relative importance of the two lipases depends on the nutritional state of the animal [30]. The interaction of the different li~protein fractions with hepatocytic receptors and the role of hepatic lipoprotein lipase in the uptake of lipoprotein lipids are outside the scope of this review. The reader is hence referred to a number of excellent overviews of this field [31-341. Intracellular traffic of long-chain fatty acids seems to be mediated in a number of tissues by cytosolic FABP (reviewed in Ref. 35). As a matter of fact, FABP has been shown to facilitate the diffusion of oleate in a model cytosol system 136-381. The liver contains a specific form of FABP, the so called GFABP, which is more abundant in the periportal than in the perivenous zone of liver. This L-FABP may be involved in processes such as peroxisomal long-chain fatty acid oxidation [39,401 and cholesterol metabolism [34]. The expression of L-FABP increases on the long term in response to enhanced hepatocyte fatty acid flux [34,35]. No data are available on the possibility that the binding activity of L-FABP is controlled by cellular agonists. Activation of fatty acids to their CoA thioesters is a prerequisite for their utilization by the cell. This reaction is catalyzed by a family of acyl-CoA synthetases which differ in their chain-length specificity and subcellular location [2,41-431. Thus, the liver contains shortchain, medium-chain, long-chain and very-long-chain acyl-CoA synthetases. Short-chain acyl-CoA synthetase is present in liver cytosol and mito~hondrial matrix, whereas medium-chain acyl-CoA synthetase is especially abundant in mitochondrial matrix. Although some octanoate can be converted to its CoA ester outside the mitochondrial matrix and then undergo p-oxidation by a carnitine-dependent carrying system 1441,the presence of short- and medium-chain acyl-CoA synthetases in the mitochondrial matrix may obviate the need for carnitine in most of the transport of shortand medium-chain fatty acids into mitochondrial oxidative metabolism (see below). Long-chain acyl-CoA synthetase is a well-characterized enzyme [45] which is

230 present in the outer mitochondrial membrane, microsomes and peroxisomes. Although it is tempting to speculate that the mito~hondrial and the peroxisomal enzymes synthesize acyl-CoA for oxidative metabolism, whereas the microsomal enzyme synthesizes acyl-CoA for complex lipid synthesis, there is no direct evidence that this is the case in the liver cell. Chemical and immunological studies have shown identical properties of long-chain acyl-CoA synthetases associated with the mitochondrial outer membrane, microsomes and peroxisomes [42,43]. Finally, a very-long-chain acyl-CoA synthetase has been detected in liver peroxisomes (461. This enzyme could therefore be involved in the peroxisomal oxidation of vex-long-chain fatty acids 146,471. It is generally agreed that the acyl-CoA synthetases have a minor regulatory influence on fatty acid metabolism [2,421. Although rat liver long-chain acylCoA synthetase may be controlled by the nutritional status of the animal [45], no evidence has been presented thus far for the hormonal control of acyl-CoA synthetases. However, since ATP is a substrate for the enzyme, it has been suggested that changes in the intramitochondrial content of ATP might result in concomitant changes in the rates of activation and oxidation of short- and medium-chain fatty acids [2]. It has also been put forward that variations in the intramitochondrial [acetyl-CoA]/[CoA] ratio may be transferred to the cytosol via changes in the [acetylcarnitine]/[carnitine] ratio, which might in turn control fatty acid activation by long-chain acyl-CoA synthetase 1481.Reg-

MOM

CYTOSOl

1

CoASH R

r

I

I

II MALONYL-CoA -

0-

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I

ulation of these processes by cellular stimuli needs to be proved. The percentage of fatty acyi-CoA diverted to the oxidation or the esterification pathways in the liver changes under different alterations of the metabolic status of the animal [1,2]. This is supposed to be due to variations in the flux through a number of key regulatory points involved in the coordinate control of hepatic fatty acid oxidation (see below) and esterification (reviewed in Refs. 49, 50). In addition, the availability of glycerol 3-phosphate - the precursor for the glycerol backbone of glycerolipids - was originally suggested to play a regulatory role in the contro1 of triacylgfycerol formation and hence in the distribution of fatty acids between oxidation and esterification I.5l-.56]. However, it is currently believed that the relation between gfycera1 3-phosphate content and triacylglycerol synthesis as observed in isofated hepatocytes may be due to the peculiar situation which occurs when liver cells are incubated in vitro (i.e., glycogen depletion and absence of glycerol) and which most likely does not occur in vivo under conditions of increased fat-store mobilization in the adipose tissue [2,57-601. III. T~ns~ocation

Short- and medium-chain fatty acids can readily penetrate mitochondria independently of carnitine and be subsequently activated to their CoA esters in the mitochondrial matrix by short- and medium-chain acyl-

ACYLCARNITINE

\

1

* .Iv cq I ’

CAANITINE

MITOCHONRR~AL MATRIX

CoASH

I T

BS 1 CPT, ‘4 4 I

ACY L-COA

MM

INTERMEMBRANE SPACE

f

of fatty acids into mitochond~a

I

L ACYL-CoA

WXIDATION

Fig. 2. Carnitine-dependent translocation of long-chain fatty acyl-CoA into the mitochondrial matrix. CPT, catalyzes the synthesis of a long-chain acylcarnitine complex from acyl-CoA and carnitine in the inner leaflet of the mitochondrial outer membrane (MOM). The acylcarnitine complex then diffuses across the mitochondrial intermembrane space up till the mitochondrial inner membrane (MIM), in which the consecutive action of the carnitine: acylcarnitine transiocase (T) and CPT, enables the long-chain fatty acyl-CoA moiety to enter the mitochondrial matrix for oxidative metabolism. CPT, is inhibited by malonyi-CoA at a site in the outer leaflet of the MOM. The existence of a malonyl-CoA binding site (BS) as a physically separated entity is still a matter of debate.

231 CoA synthethases to undergo P-oxidation 161-631. Long-chain acyl-CoA are able to cross the mitochondrial outer membrane; however, long-chain acyl-CoA as such cannot penetrate the mitochondrial inner membrane. Translocation of long-chain fatty acyl-CoA into mitochondrial matrix is a rather complex, carnitine-dependent process which involves the coordinate action of CPT,, a carnitine : acylcarnitine translocase and the mitochondrial inner membrane carnitine palmitoyltransferase ( = CPTi, CPT-II, CPT-B) (Fig. 2) [61-631. It is generally accepted that CPT, catalyzes the rate-limiting step of long-chain acyl-CoA translocation into mitochondria and represents a key regulatory point in the overall process of long-chain fatty acid oxidation by the liver. A number of excellent reviews highlight the importance of CPT, in the regulation of long-chain fatty acid oxidation in both liver and extrahepatic tissues [61-661. However, research on the molecular mechanisms underlying the regulation of CPT, has been hampered by problems related to (i) the molecular characterization of the enzyme, mostly due to the wrong classical belief that the enzyme was located in the mitochondrial inner membrane [671, (ii) the presence of peroxisomes in conventional mitochondrial preparations and subsequent cross-contamination with the easily-solubilized peroxisomal CPT [68,69], and (iii) solubilizing the protein from mitochondrial membranes without loss of enzyme activity and/or sensitivity to inhibition by malonyl-CoA [70-721. During the last few years, however, considerable efforts by a number of laboratories have helped to clarify this scenario. Most of this new information has arisen from experiments performed with a series of inhibitors of CPT, such as tetradecylglycidyl-CoA, 2-bromopalmitoyl-CoA or etomoxir-CJoA [63,66,72,73]. It is generally accepted nowadays that in the liver of a number of species CPT, is a protein of about 90 kDa, whereas heart and muscle isoenzymes are slightly smaller in size (about 86 kDa) [72,74-781. Nevertheless, the idea that the catalytic activity of CPT,, resides in a smaller polypeptide - very similar or even identical to CPT, - is still supported by others [79-821. The question of whether the malonylCoA-binding site resides in the same polypeptide as the catalytic center [71,72,74,75,771 or is a physically separated entity [76,79-821 is also a matter of debate. In the case of CPT,, cloning of its cDNA in both human [83] and rat liver [841 has shown a molecular mass of 74 kDa for the preprotein and 71 kDa for the mature, membrane-inserted enzyme. Using octyl glucoside extracts of heart mitochondria, Kerner and Bieber 1791 have put forward the hypothesis that CPT, and CPTi are associated with a complex of P-oxidation enzymes. This possibility is actually interesting since it would allow in vivo the channeling of the fatty acid substrate between the cytosol and the mitochondrial

matrix at points of contact of the two mitochondrial membranes, thus avoiding diffusion of intermediates space to the on the way from the intermembrane mitochondrial matrix. The discovery of malonyl-CoA inhibition of CPT, [1,85,86] attracted an enormous attention to the study of the regulation of this enzyme. As mentioned in Section II, malonyl-CoA is precisely the product of the reaction catalyzed by acetyl-CoA carboxylase, the ratelimiting step of fatty acid synthesis [3,4], so that coordinate control of synthesis and oxidation of fatty acids is achieved [1,21. Changes in hepatic malonyl-CoA levels appear upon alterations in the nutritional and hormonal status of the animal [1,2]. Malonyl-CoA itself not only inhibits CPT, activity, but could also play a role in determining the sensitivity of CPT, to inhibition by malonyl-CoA. Thus, either preincubation of intact hepatocytes with agents which increase intracellular malonyl-CoA levels 1871 or exposure of isolated rat liver mitochondria to physiological concentrations of malonyl-CoA [88] render CPT,, more sensitive to malonyl-CoA in a subsequent assay. Furthermore, a close correlation has been observed between the sensitivity of hepatic CPT, to inhibition by malonyl-CoA and the concentration of malonyl-CoA at which the enzyme was exposed in vivo before isolation of mitochondria for enzyme assay [891. However, Cook and Cox [901 have been unable to detect any sensitization of CPT,, after exposure of the enzyme to malonyl-CoA. Instead, these authors observed a lag phase in the assay of CPT, activity which was more pronounced when malonyl-CoA was present in the medium. This phenomenon they called hysteretic behaviour of WI’, [903. The sensitivity of CPT, to inhibition by malonyl-CoA also depends on other factors such as temperature of the assay, pH and ionic strength of the medium (reviewed in Ref. 64). Malonyl-CoA is thus a very important effector which controls the entry of long-chain fatty acyl moieties into the mitochondrial oxidative metabolism. However, it is currently well-established that malonyl-CoA concentration is not the only factor controlling the process of acyl-CoA transport into mitochondria [2,64,66]. Thus, diet- and hormone-induced changes in the flux through the fatty-acid-oxidative pathway are usually accompanied by parallel variations in the specific activity and sensitivity to malonyl-CoA of rat liver CPT,,. For example, under ketotic states such as starvation [91], diabetes 192,931 and hyperthyroidism [941 the specific activity of CPT, increases whereas enzyme sensitivity to inhibition by malonyl-CoA decreases. The opposite occurs in lipogenic states such as hypothyroidism [95], refeeding after starvation 1961or chronic ethanol feeding 1971.The importance of malonyl-CoA sensitivity of CPT,, in the control of long-chain fatty acid oxidation has been unequivocally demonstrated by Prip-Buus et

232 al. [981 in hepatocytes isolated from fetal and newborn rabbits. Moreover, control of hepatic long-chain fatty acid oxidation independent of malonyl-CoA concentration has been shown to occur in several pathophysiological situations, e.g. during neonatal development [981, at weaning [99], on feeding high-fat diets containing medium-chain triacylglycerols [loo] or after chronic ethanol feeding [loll and upon treatment of hepatocytes with a number of agonists such as vanadate [102] or the amino acids alanine and asparagine [103]. Most of the studies on rat liver CPT enzymes have focussed on the long-term control of kinetic and regulatory properties of CPT,,. Short-term adaptive changes of hepatic CPT, activity, on the other hand, have thus far not received as much attention. This may be due to the fact that short-term modulation of CPT, activity is difficult to preserve during the procedure of cell disruption and subsequent isolation of mitochondria for enzyme assay. This problem may be circumvented by assaying CPT, activity in a permeabilized-cell system [104,105]. The use of this procedure has shown that hepatic CPT, is controlled on the short term by different types of agonists. For example, insulin, vasopressin and the phorbol ester 12@-myristate 13cy-acetate (PMA) inhibit CPT, activity, whereas vanadate and CAMP-raising agents have the opposite effect [105]. In all these studies a qualitative relationship was observed between changes in CPT, activity and changes in the rate of mitochondrial palmitate oxidation, indicating that CPT,, is a key regulatory enzyme in the control of long-chain fatty acid oxidation by liver mitochondria, at least in the experimental system employed therein (Table 1). However, it was also pointed out that additional points of control of the fatty-acid-oxidative process seem to exist [105] (see below). The effects of the aforementioned cellular agonists on CPT, activity might be mediated via changes in the intracellular concentration of malonyl-CoA. For example, hepatocyte malonyl-CoA levels are decreased by glucagon and increased by insulin [106]. However, in order to measure enzyme activity the plasma membrane is permeabilized and this causes the cytosol to leak out from the cells, leading to a large dilution of cytosolic components including malonyl-CoA [105]. Therefore, the modulation of CPT, activity by cellular effecters should involve a persistent modification as well. Further support for this notion comes from experiments with okadaic acid. This protein phosphatase inhibitor activates palmitate oxidation 11071, and this stimulation occurs in concert with profound changes in the kinetic and regulatory properties of CPT,, namely increased specific activity, decreased sensitivity to inhibition by malonyl-CoA and reduced affinity for the palmitoyl-CoA substrate [107,1081. The exhaustive molecular characterization of CPT, has allowed the study of the mechanisms involved in

TABLE

I

Effects of short-term modulators of hepatic metabolism on CPI;, actkity in digitonin-permeabilized hepatocytes and on palmitate oxidation in intact hepatocytes After an incubation period of 30 min in the absence or in the presence of the substances indicated, aliquots of the hepatocyte suspensions were removed to measure CPT, activity in digitonin-permeabilized hepatocytes and the rate of palmitate oxidation in intact hepatocytes. 100% values were 4.05 +0.37 nmol palmitoylcarnitine formed/min per mg protein for CPT,, activity (as measured with 50 PM palmitoyl-CoA and 0.5% bovine serum albumin) and 43.7k5.Y nmol palmitate oxidized/h per mg protein for palmitate oxidation (as measured with 0.4 mM palmitate and 1% bovine serum albumin). see Ref. 108. Significantly different from incubations with no additions: ” P < 0.05; ’ P < 0.01, Cell incubation

No additions 10 nM glucagon 50 PM dibutyryl-CAMP 50 PM forskolin 85 nM insulin 100 nM EGF 100 nM vasopressin 1pMPMA 2 mM vanadate

CPT,, activity (%)

Palmitate oxidation

100 127+ 7” 131+ 3h 138* 5h 84i 8“ 86+ 4” 82+ 7” 67; 7h 145+16 h

100 136k 137* 140* 85 + X6& 8x+ 52; 142+

(%)

4” 3” 5” 1“ 3” 3 iI 9” 1I ”

the long-term regulation of this enzyme protein. Thus, Brady and co-workers have shown that the amount of CPTi mRNA and/or immunoreactive protein increase in rat liver in a number of situations which induce the ketogenic capacity of the liver [109-1131. However, the molecular processes underlying the alterations in the kinetic and regulatory properties of hepatic CPT,, are still unknown [63,64,66]. Very recently both monoclonal (Pegorier, J.P. and Prip-Buus, C., personal communication) and polyclonal antibodies [78] against rat liver CPT, have been obtained, and the levels of immunoreactive CPT, protein have been shown to increase several-fold under different ketotic states [78]. However, the question of how the regulatory properties of the enzyme change in these situations remains unsolved. It has been suggested that post-transcriptional modifications of the existing enzyme molecules could be involved in the alterations of the regulatory properties of CPT, induced by fasting and diabetes [92]. Kashfi and Cook have proposed that increased proteolysis of the CPT, protein under ketotic states might change the regulatory properties of the enzyme [114]. It is noteworthy that the modifications induced by okadaic acid (presumably increased enzyme phosphorylation) on hepatic CPT, [107,108] are precisely those appearing in ketotic states such as fasting, diabetes and hyperthyroidism [2,64,66], in which the phosphorylation degree of target regulatory enzymes is enhanced. Hence it is tempting to speculate that increased phosphorylation of CPT,, may be a key mecha-

233 nism to render the enzyme more active, less sensitive to inhibition by malonyl-CoA and less saturable by the acyl-CoA substrate. In summary, it is likely that the regulation of hepatic CPT, involves a series of complex and coordinate mechanisms, including at least (i) changes in the intracellular content of malonyl-CoA, (ii) changes in the kinetic and regulatory properties of the pre-existing enzyme molecules (as determined by enzyme phosphorylation and/or proteolysis?), and (iii) changes in the amount of enzyme molecules. It is also worth noting that the properties of the mitochondrial membrane environment determine (at least in vitro) the activity and sensitivity to malonyl-CoA of rat liver CPT, [115-1181, and this could be another physiological mechanism exerting control on the CPT, enzyme. IV. /3-Oxidation of fatty acids The knowledge of the P-oxidation of fatty acids has increased immensely during the last decade (reviewed in Ref. 42). The most important advances in this field include (i) the identification of a second mammalian P-oxidative system located in peroxisomes, together with the subsequent characterization of a trifunctional enzyme which is exclusive for peroxisomal P-oxidation and which possesses trans-2-enoyl-CoA hydratase, L3-hydroxyacyl-CoA dehydrogenase and A3-&-A*truns-enoyl-CoA isomerase activities [119-1221; (ii> the characterization of the enzymes involved in the poxidation of unsaturated fatty acids, i.e., 2,4-dienoylCoA reductase and A3-cis-A*-truns-enoyl-CoA isomerase [123-1251; (iii) the recognition of a number of inherited human defects of fatty acid P-oxidation [41,126]; and (iu> the detection of substrate channeling in mitochondrial P-oxidation, in line with the suggestion that mitochondrial P-oxidative enzymes exist as a multienzyme complex (‘metabolon’) which interacts with the complexes of the respiratory chain on the inner face of the mitochondrial inner membrane [127]. Measurements of acyl-CoA intermediates of P-oxidation in rat-liver mitochondria have also provided evidence for some organization of the enzymes of poxidation [128]. Other enzymes involved in fatty acid P-oxidation which have been recently characterized include a peroxisomal branched-chain fatty acid oxidase [129,1301, a mitochondrial very-long-chain acylCoA dehydrogenase [1311 and a mitochondrial WWS2-enoyl-CoA hydratase/L-3-hydroxyacyl-CoA dehydrogenase/3-ketoacyl-CoA thiolase trifunctional protein associated with the mitochondrial inner membrane which only appears to metabolize long-chain acyl-CoA esters [132]. Although the information on the concentration of P-oxidation enzymes in the mitochondrial and peroxisomal matrices is rather limited, Sumegi et al. 11271 have pointed out that some acyl-CoA esters

occur at lower concentrations than some of the poxidation enzymes. In spite of this all, very little is known about the control of the enzymes involved in the P-oxidation of fatty acids. The pathway has two oxidative steps (i.e., the reactions catalyzed by acyl-CoA dehydrogenase and L-3-hydroxyacyl-CoA dehydrogenase), and one possible regulatory factor could be then the intramitochondrial NADH/NAD+ concentration ratio [133, 1341. In fact, elevations of the intramitochondrial NADH/NAD+ ratio in isolated hepatocytes depress P-oxidation of fatty acids in parallel 11331. Additional effecters of the p-oxidation route may include acetoacetyl-CoA, which in vitro is an inhibitor of acyl-CoA dehydrogenase in liver 11351, and acetyl-CoA, which exerts feedback inhibition at least in heart on 3-ketoacyl-CoA-thiolase [ 136,137], and long-chain L-3-hydroxyacyl-CoA, which strongly inhibits enoyl-CoA hydratase [138]. Although it is likely that these potential regulatory steps will respond to hormone-induced changes in the aforementioned parameters, as far as we know there is no direct evidence that the rate of hepatic P-oxidation of intramitochondrial acyl-CoA is controlled by cellular effecters. In addition, there does not appear to be real evidence for allosteric effecters or covalent modification of p-oxidation enzymes. Fatty acids may also undergo w-oxidation, which will result in the formation of medium- and long-chain dicarboxylates with no need for CoA [1391. By this accessory pathway, monocarboxylic acids are firstly converted into w-hydroxymonocarboxylic acids by the microsomal mixed function oxidase system [140]. The w-hydroxylated product can be either P-oxidized in mitochondria and peroxisomes or further w-oxidized to w-oxomonocarboxylic and dicarboxylic acids by the action of cytosolic alcohol- and aldehyde dehydrogenase, respectively [141]. The latter may be activated to their CoA esters by a microsomal dicarboxyl-CoA synthetase and subsequently P-oxidized mainly in peroxisomes to medium-chain dicarboxylic acids, which are water soluble and can be excreted in the urine [139,142]. Although w-oxidation is induced in ketotic states or upon clofibrate feeding to rats, it actually makes only a marginal contribution to total hepatocellular fatty acid oxidation [143]. V. Ketogenesis Formation of ketone bodies is the major metabolic fate of acetyl-CoA produced by the P-oxidation spiral in liver. It is commonly agreed that the contribution of the tricarbcxylic acid cycle to the disposal of acetyl-CoA is not very high in situations of sufficient glucose supply and almost nil in catabolic states such as starvation and diabetes, in which practically all fatty acids oxidized by the liver are diverted into ketone body

234 synthesis [1,2,144,145]. Thus, long-chain fatty acid oxidation by hepatocytes inhibits glucose phosphorylation, glycolytic flux through phosphofructokinase-1 and pyruvate oxidation, and shifts pyruvate into gluconeogenesis, thus reducing the amount of acetyl-CoA produced by glucose breakdown (reviewed in Ref. 144). Furthermore, some studies put forward the notion of substrate channeling between the enzymes of the poxidative spiral and the ketogenic route as well as between pyruvate oxidation and the tricarboxylic acid cycle, i.e., acetyl-CoA produced by P-oxidation of fatty acids is preferentially diverted into the ketogenic pathway, whereas that formed by pyruvate dehydrogenase is mostly employed by the tricarboxylic acid cycle [1461481. It is well established that hepatic ketogenesis is controlled on the short- and the long term by the nutritional status of the animal as well as by hormones (reviewed in Refs. 1,2). Among the latter, the polypeptide hormones insulin and glucagon display opposite effects on hepatic ketogenesis, the former depressing and the latter stimulating ketone body formation [55]. The ketogenic effect of glucagon is supposed to be mediated, at least in part, via the classical CAMP route (type 2 glucagon receptor), since it is mimicked by CAMP analogues as well as by other agonists which either increase intracellular CAMP levels (forskolin) or inhibit hepatocellular protein phosphatases (okadaic acid) [107]. Whether Ca*+ mobilization triggered by the putative type 1 glucagon receptor is also involved in glucagon action in hepatocytes is still unknown [14911511. In this respect, Halestrap [152] has shown that P-oxidation of fatty acids in isolated liver mitochondria is very sensitive to osmolarity compared with the oxidation of other substrates. Since Ca*+ causes mitochondrial swelling and increases the rate of fatty acid oxidation and matrix swelling and increased ketogenesis are observed in glucagon-treated hepatocytes [152], it may be assumed, therefore, that this may be a mechanism leading to the stimulation of ketogenesis by glucagon. Nevertheless, Ca*+ -mobilizing hormones such as vasopressin rapidly inhibit ketogenesis [105,153-1561, whereas the effect of c-u,-agonists such as noradrenaline has been variously reported ranging from stimulation [157,158] to no effect [159] or inhibition [160]. The effect of vasopressin on hepatic long-chain fatty acid oxidation seems to be rather complex. Addition of this hormone to hepatocyte incubations causes a depression of CPT,, activity which is accompanied by a dual effect on fatty acid oxidation: ketone body formation is decreased while CO, production is enhanced [105]. Thus, vasopressin may reduce the entry of fatty acids into mitochondria and diverts mitochondrial acylCoA into the tricarboxylic acid cycle at the expense of the ketogenic pathway, apparently uncoupling the putative channeling between fatty acid oxidation and ke-

togenesis (see above). This indicates that apart from CPT, other factors exert control over the fatty-acidoxidizing system. Vasopressin is known to trigger the receptor-mediated breakdown of phosphatidylinositol 4,5_bisphosphate to produce diacylglycerol and inositol 1,4,5-trisphosphate as second messengers. It has been suggested that the diacylglycerol component of the hormonal response is solely responsible for the regulation of acetyl-CoA carboxylase and fatty acid synthesis de novo by vasopressin [161]. Interestingly, the phorbol ester PMA and vasopressin exert a similar inhibition of hepatic CPT, activity [105] but a dissimilar effect on ketone body and CO, production by isolated hepatocytes [105,1621. Therefore, it is possible that activation of protein kinase C mediates the short-term regulation of acetyl-CoA carboxylase and CPT,, by vasopressin, whereas the Ca*+-mobilizing limb of vasopressin action may control intramitochondrial metabolism of acyl-CoA [105,161,162]. As stated above, there is good evidence that the step catalyzed by CPT, is a key regulatory point in the hormonal control of hepatic long-chain fatty acid oxidation (and so of ketogenesis). However, as suggested for example by experiments concerning vasopressin action, it seems that additional factors exert control on hepatic ketogenesis at the intramitochondrial level. A similar conclusion was inferred by Brady and Brady on the basis of inhibition by tetradecylglycidyl-CoA of palmitoyl-CoA oxidation by isolated liver mitochondria 11631.In addition, the acute reversal of diabetic ketosis [93] or the rapid depression of the ketogenic capacity of the liver on refeeding starved rats [96] are not accompanied by short-term changes in the kinetic and regulatory properties of hepatic CPT,, (see also Ref. 164). The first commited step of ketone body formation is the condensation of two acetyl-CoA molecules to yield acetoacetyl-CoA in a reaction catalyzed by acetyl-CoA acetyltransferase ( = acetoacetyl-CoA thiolase) (liver and extrahepatic tissues contain at least another mitochondrial thiolase isoenzyme which takes part in the last step of fatty acid p-oxidation, i.e., thiolysis of 3-ketoacyl-CoA, as well as an immunologically different cytosolic isoenzyme which catalyzes acetoacetylCoA cleavage in the process of ketone body utilization). Acetyltransferase-catalyzed formation of acetoacetylCoA is inhibited by the free CoA product, which decreases enzyme affinity for its substrate, acetyl-CoA [2]. This may represent a mechanism for the regulation of ketogenesis by changes in the intramitochondrial [acetyl-CoA]/[CoA] ratio [137]. Acetoacetyl-CoA is an important metabolite in the control of fatty acid metabolism in liver mitochondria. It has been observed in vitro that acetoacetyl-CoA inhibits acyl-CoA dehydrogenase, the first enzyme commited to fatty acid P-oxidation (see above) and exerts product inhibition

235

Acyl-CoA dehydrogenase

3- ketoacyl-CoA thiolase

’

I

ACETYL-CoA

-

-

w

\ \

\ \

\

Acetyi-CoA ace~t~nsferase

\

\ I / /

\\ c ACETOACETYL-CoA 3-oxoacid-CoA transferase

1

/

/ \e

EB

Y

I

\

HMG-CoA synthase

HIWG-CoA

TRlCARBOXYLtC

I

AC/O CYCLE

I

HMG-CoA lyase

ACETOACETATE

NADH

NAD+

4

D-3-HYDROXYBUTYRATE Fig. 3. Control of the intramitochondrial reactions of ketogenesis by CoA and its derivatives. Both free CoA and the CoA thioesters acetyl-CoA, acetoacetyl-CoA and succinyl-CoA may take part in the metabolite control of ketogenesis from acetyl-CoA in liver mitochondria. Dashed lines indicate negative (- 1 or positive control (+I of enzyme activity. Regulation of pyrwate carboxylase, pyrwate dehydrogenase and tricarboxylic acid-cycle enzymes by CoA thioesters is not represented in the figure.

on acetyl-CoA acetyltransferase and substrate inhibition on HMG-CoA synthase [165], the putative regulatory enzyme of hepatic ketogenesis (Fig. 3). However, it is unlikely that this happens in viva, First, because the acetyltransferase-catalyzed reaction may be relatively close to equilibrium and the level of acetoacetylCoA may be low enough under the prevailing conditions to impair, at least in part, the rate of ketogenesis

[145]. Second, if ch~neling occurs, as mentioned above, a~toace~l-boa may not have access to the acyl-CoA dehydrogenases. The liver contains two HMG-CoA synthase isoenzymes, one cytosolic [166] and one mitochondrial [167]. The former catalyzes the synthesis of HMG-CoA for cholesterol biosynthesis, whereas mitochondrial HMGCoA synthase catalyzes the synthesis of HMG-CoA for ketone body formation and seems to represent the major rate-limiting enzyme in ketogenesis from acetylCoA. The activity of this enzyme is controlled in vivo by two mechanisms which operate in concert: (i) short-term modification of pre-e~sting enzyme molecules by covalent modification (su~inylation/desuccinylation); (ii) long-term changes in the amount of enzyme mRNA and immunoreactive protein. Succinyl-CoA is the most important effector of HMG-CoA synthase. In the presence of physiological concentrations of succinyl-CoA, the enzyme rapidly inactivates itself by catalyzing self-su~inylation at the active .site [168]. The succinylated enzyme can be desuccinylated and reactivated in the presence of acetyl-CoA, which accelerates desuccinylation and prevents resuccinylation 11691. Quant et al. [170,171] have shown that glucagon rapidly lowers hepatic succinylCoA content in vivo and increases in parallel hepatic HMG-CoA synthase activity by lowering the succinylation extent of the enzyme. It is also worth noting that the effects of glucagon on both the activity and the succinylation extent of hepatic HMG-CoA synthase can be perfectly reproduced by m~ipulating mitochondrial succinyl-CoA content [170,171]. In addition, lowering plasma insuiin concentration by mannoheptulose injection produces the same effects as glucagon on the HMG-CoA synthase enzyme [170,171]. All these rapid alterations of HMG-CoA synthase activity are not accompanied by changes in the amount of enzyme molecules (P.A. Quant, personal communication). The activity of rat liver mitochondrial HMG-CoA synthase increases on the long term in ketotic situations such as starvation, fat feeeding, diabetes and lactation. In all these situations the amount of immunoreactive enzyme protein increases, whereas the extent of succinylation of the enzyme molecules decreases [ 172, P.A. Quant, personal communication]. The cDNA for mit~hondrial HMG-CoA synthase has been cloned by Hegardt’s group 11671. These authors have recently shown that the nutritional and the hormonal status of the animal exert control at the level of enzyme gene expression. Thus, in rat liver the levels of mitochondrial HMG-CoA synthase mRNA are strongly increased by starvation, fat feeding, diabetes and CAMP, whereas they decrease upon refeeding starved animals and by insulin injection (1731. In conclusion, changes in CPT, and mitochondrial HMG-CoA synthase activities operate in the same

236 direction. Control of hepatic ketogenesis may thus be exerted in a coordinate fashion at these two major regulatory sites. It is also likely that in situations in which the liver is flooded with medium-chain fatty acids, e.g. after administration of diets containing medium-chain triacylglycerols [174] or during the suckling period [173], the CPTO step is bypassed and HMGCoA synthase will exert pivotal control over the fatty acid-oxidative pathway in the liver. Interestingly, in Fao hepatoma cells oleate oxidation is limited by the hypersensitivity of CPT, to inhibition by malonyl-CoA (Pegorier, J.P. and Prip-Buus, C., personal communication). However, these celfs are able to oxidize octanoate to acetyl-CoA and CO, but they are unable to produce ketone bodies owing to a complete deficiency in mitochondrial HMG-CoA synthase (activity, protein and mRNA) (Pegorier, J.P. and Prip-Buus, C., personal communication). Completion of the ketogenic pathway is achieved by the action of HMG-CoA lyase and D-3-hydro~butyrate dehydrogenase. The former is not believed to constitute a regulatory step of hepatic ketogenesis [2], whereas the latter catalyzes the NADH-linked reduction of acetoacetate to o-3-hydroxybutyrate. It has been traditionally assumed that reoxidation of reducing equivalents by the action of D-3-hydroxybutyrate dehydrogenase would prevent the blockade of fatty acid /?-oxidation by elevated NADH levels, particularly at high rates of fatty acid oxidation [2]. In rat hepatocytes, however, the rate of ketogenesis from long-chain fatty acids is higher in periportal than in perivenous hepatocytes [175] but the ratio of production of 3-hydroxybutyrate: acetoacetate is lower in periportal than in perivenous hepatocytes [175,176]. Moreover, this ratio is decreased by treatment with glucagon in vitro 11751 and by diabetes in vivo (Agius, L., personal communication). The effects of glucagon treatment in vitro and of diabetes are not additive and presumably therefore share a similar mechanism of action (Agius, L., personal communication). Thus, low intramitochondrial ratios upon increased ketone body NADH,‘NAD+ production might be related to the higher gluconeogenic capacity of the liver in this situation. VI. Tricarboxylic acid cycle activity Complete oxidation of ace@-CoA to CO, is achieved in the mito~hondrial matrix by the enzymes of the tricarboxylic acid cycle. It has been suggested that the enzymes of the cycle are assembled in a functional multienzyme complex (“metabolon’) aimed to provide adequate coordination and efficiency (reviewed in Refs. 177, 178). Although the enzymes which catalyze the reactions of the cycle are well characterized, the mechanisms underlying the regulation of this complex enzyme system are still a matter of debate.

The activity of the tricarboxylic acid cycle in hepatic mitochondria is controlled by various hormones. Ca’+mobilizing hormones comprise the group of cellular effecters exerting the most prominent action on the metabolic flux through the tricarboxylic acid cycle (reviewed in Refs. 179, 180). Thus, vasopressin, angiotensin and a,-agonists such as phenylephrinc, adrenaline and noradrenaline markedly increase the turnover of the tricarboxylic acid cycie in rat liver [181,182]. This effect of Ca2’-mobilizing hormones is not detected in Cazf-depleted cells [151,1X3], supporting the notion that Ca2+ redistribution within the ceII plays a major role in the stimmation of the tricarboxylic acid cycle (see below). Although one report shows that glucagon has no effect on the rate of the cycle in the perfused rat liver [181], it is generally agreed that this hormone stimulates tricarboxylic-acid-cycle activity as well as mitochondrial respiration [179,180,182]. Furthermore. the observations that cu,-agonists (which decrease intracellular CAMP levels by inhibiting adenylate cyclase activity) have no effect on the flux through the cycle [181] and that the stimulation of respiration by glucagon depends on the presence of intracellular CaZt [151,1833, suggest that glucagon action on mitochondrial oxidative metabolism may be mediated (at least in part) by the putative type I glucagon receptor, i.e., by the CaZ+ connection (see above) [149,150,184,18.5]. A slight increase in the flux through the tricarboxylic acid cycle has been observed upon incubation of hcpatocytes with insulin [182]. It should be pointed out that citrate is not solely used for oxidation of acetyl-CoA molecules through the tricarboxylic acid cycle, but it also represents a means to shuttle C, units from the mitochondrial matrix to the cytosol for fatty acid synthesis de novo. Therefore, a certain flux through the step catalyzed by citrate synthase should be maintained in order to allow high lipogenic rates. Although the stimulatory effect of insulin on fatty acid synthesis de novo is majorly exerted at the acetyl-CoA carboxylase site [3], this hormone sustains the hepatocellular citrate content at an appropiate value to allow enough substrate for lipogenesis [106]. The molecular mechanism(s) by which hormonal stimuli control the activity of the tricarboxylic acid cycle are not clearly established. The competition between citrate synthase and acetoacetyI-CoA thioiase for the acetyl-CoA substrate, as well as the mitochondrial content of oxalacetate (the second substrate for citrate synthase), might represent a first regulatory mechanism of the cycle [2]. In case of substrate channeling between pyruvate oxidation and the tricarboxylic acid cycle, as well as between p-oxidation of fatty acids and ketogenesis (see above), it is very unlikely that this potential competition operates in the living cell. Instead, the specific activity of citrate synthase may be

237 controlled by a number of effecters such as ATP, citrate and succinyl-CoA [2]. Although ATP inhibits citrate synthase activity in vitro, studies on adenine nucleotide compartmentation in isolated hepatocytes indicate that it is not very likely that this mechanism takes place in vivo [2,179,180,186]. In contrast, inhibition of citrate synthase by succinyl-CoA could provide a means for the coordinated control of citrate synthase [187], 2-oxoglutarate dehydrogenase [188,189] and HMG-CoA synthase [1711. Feedback inhibition of citrate synthase by citrate might be important when isocitrate dehydrogenase and 2-oxoglutarate dehydrogenase are inhibited, e.g. in the absence of Ca*+ 11901. Evidence has accumulated during the last few years supporting the importance of the intramitochondrial Ca*+ concentration in regulating the tricarboxylic acid cycle (reviewed in refs 179, 180, 191). Thus, all the aforementioned hepatocellular effecters which raise the Ca*+ concentration in the cytosol also increase the Ca*+ concentration in the mitochondrial matrix, and this in turn stimulates pyruvate oxidation and tricarboxylic-acid-cycle activity via the three Ca*+-sensitive intramitochondrial dehydrogenases, namely pyruvate dehydrogenase, NAD+-isocitrate dehydrogenase and 2-oxoglutarate dehydrogenase 1179,180l. Ca*+-induced activation of pyruvate dehydrogenase is achieved through activation of pyruvate dehydrogenase phosphatase, which dephosphorylates and thus activates pyruvate dehydrogenase [192]. In contrast, the effect of Ca*+ on NAD+-isocitrate dehydrogenase and 2-0x0glutarate dehydrogenase occurs via direct binding of the Ca*+ ions to the enzymes, causing allosteric activation by strongly reducing the k, values for their respective substrates, isocitrate and 2-oxoglutarate [179, 180,183]. The catalytic activity of these three dehydrogenases is also subject to ‘classical’ feedback inhibition by increases in the NADH/NAD+ and ATP/ADP concentration ratios [183]. However, it is worth noting that treatment with Ca*+-mobilizing hormones rapidly increases pyruvate oxidation, tricarboxylic-acid-cycle activity and oxygen uptake by liver mitochondria, although NADH and ATP levels remain unchanged or even increased [179,180,186]. Therefore, as pointed out by McCormack and Denton [179,180,183], the use of Ca*+ as a second messenger within mitochondria would allow cells to maintain or even increase their ATP levels when the demand for ATP is enhanced. It has been suggested that Ca*+ sensitivity of 2-oxoglutarate dehydrogenase may be directly responsible for the stimulation of gluconeogenesis by Ca*+-mobilizing hormones via activation of the malate/aspartate shuttle [193]. Thus, redistribution of Ca*+ within the hepatocyte by hormonal action may play a critical role in the coordinate stimulation of mitochondrial oxidative metabolism (including fatty acid oxidation) and gluconeogenesis.

VII. Peroxisomal fatty acid oxidation Peroxisomes possess a particular enzymatic equipment to p-oxidize fatty acids, dicarboxylic acids, prostaglandins, hydroxylated SP-cholestanoic acids and various fatty acid analogues (reviewed in Ref. 122). Compared to mitochondrial P-oxidation, peroxisomal P-oxidation has a broader substrate specificity, being especially active towards very-long-chain fatty acids [ 1221. Unlike mitochondria, peroxisomes only chainshorten fatty acids [122,194,195]. In addition, part of the acetyl-CoA formed by incomplete P-oxidation in peroxisomes may be deacylated to free acetate [ 1961. In any case, peroxisomal oxidation products will subsequently be transferred to the cytosol and then to the mitochondrial matrix for further metabolism [63,122]. The contribution of peroxisomal P-oxidation to total hepatocellular long-chain fatty acid oxidation depends on both the concentration and the chain length of the fatty acid used [197,198]. In fed rats, it ranges from about 20 to 35% in the case of palmitate or oleate [108,197-1991. Peroxisomal P-oxidation may be readily induced by diets and upon long-term administration of a number of hypolipidemic agents and xenobiotics 11221. Although the mechanism of entry of fatty acid into peroxisomes remains unclear [122], it has been suggested that the transport of long-chain fatty acyl substrates into the peroxisomal matrix may occur by a carnitine-dependent mechanism [200,201]. Like the mitochondrial outer membrane, peroxisomes have a carnitine palmitoyltransferase (CPT,) which shares with CPT, a number of properties such as chain-length specificity for acyl-CoA substrate and pattern of sensitivity to inhibition by malonyl-CoA [202,203]. CPT, activity is also determined by the nutritional status of the animal. Thus, CPT, activity has been shown to be induced upon high-fat feeding [llO] and starvation 12041,although glucagon has no effect on the levels of CPT, mRNA 11101.In addition, the sensitivity of CPT, to inhibition by malonyl-CoA decreases in starvation 12041. The observation that CPT, is sensitive to inhibition by malonyl-CoA [202,2031 has prompted speculation about the possible control of peroxisomal fatty acid oxidation by changes in the intracellular content of malonyl-CoA. However, it has recently been shown that incubation of hepatocytes with the phosphatase inhibitor okadaic acid, which dramatically depresses acetyl-CoA carboxylase activity [205,206] and hepatocyte malonyl-CoA content, increases CPT, activity and mitochondrial palmitate oxidation (see above) but has no effect on CPT, activity and peroxisomal palmitate oxidation [1081. It has been suggested that the longchain acylcarnitines oxidized by peroxisomes are firstly synthesized by CPT, in the mitochondrial outer membrane and then converted to their corresponding CoA

238 esters by CPT, to undergo p-oxidation in the peroxisoma1 matrix [200,201]. If that was the case, increased CPT, activity upon okadaic acid treatment will most likely be accompanied by a stimulation of peroxisomal long-chain fatty acid oxidation. However, this is not what has been observed [108]. CPT, itself might thus be involved in the formation of long-chain acylcarnitines for peroxisomal oxidative metabolism.

intermediary metabolism should be determined in relation to the mechanisms of hormonal action on hepatic fatty acid oxidation [212,213]. In this respect, the specific effects of osmolarity on P-oxidation in isolated mitochondria have been mentioned above. In addition, we are currently testing the possibility that CPT,, is also controlled by changes in hepatocyte volume (see also Ref. 103).

VIII. Future prospects

Acknowledgements

As summarized in the present review, a series of mechanisms are very likely to take part in the control of hepatic fatty acid oxidation. However, a number of questions are still unanswered, especially with regard to the regulation of this complicated metabolic pathway in vivo. It is possible that the application of metabolic control analysis to hepatic fatty acid oxidation will help to elucidate the extent of metabolic control exerted by the different steps of this pathway. In this respect, P.A. Quant and co-workers (personal communication) are currently applying the top-down approach to metabolic control analysis of fatty acid oxidation and ketogenesis in isolated liver mitochondria and cells, the perfused liver and the whole animal in order to elucidate whether CPT,, HMG-CoA synthase or - most likely - both enzymes exert control over hepatic fatty acid oxidation. A similar approach has recently been used by Kunz [207], and it is expected that metabolic control analysis will become an important tool to understand how hepatic fatty acid oxidation is regulated. It is also to be expected that the fate of fatty acids metabolized by the liver may shortly be monitored in vivo through selective labeling of hepatic fatty acids, a procedure recently described [2081. This would allow quantitation, under different physiopathological conditions, of the partitioning of fatty acids between oxidation and esterification as well as of their utilization for the synthesis of triacylglycerols and phospholipids for secretion and/or retention by the liver. As a matter of fact, such studies have already produced data for the starved-to-fed transition [209] and for the insulin-deficient state (V.A. Zammit, personal communication). Another challenge will be to unravel the molecular mechanisms leading to the acute changes in CPT, activity as observed in isolated hepatocytes incubated in the presence of cellular agonists. On the other hand, all the studies carried out in isolated hepatocytes on the distribution of intermediary metabolism - including fatty acid oxidation - between the periportal and the perivenous zone of the liver should be extended to the whole organism, since this metabolic zonation may be dependent on the organization of the whole liver and its particular anatomical situation in vivo [210,211]. Last but not least, the newly emerging role of liver cell swelling in the control of

We are indebted to Drs. L. Agius, C. Prip-Buus, J.P. Pegorier, P.A. Quant, W. Stremmel and V.A. Zammit for providing access to unpublished results and to Dr. L.M.G. van Golde for critically reading the manuscript. Research in the authors’ laboratories was supported by the Netherlands Foundation for Chemical Research (SON) with financial aid from the Netherlands Organization for Scientific Research (NWO), as well as by the Research Foundation of the Spanish Ministry of Health (FISSS). References 1 M&arty, J.D. and Foster, D.W. (1980) Annu. Rev. Biochem. 49, 395-420. 2 Zammit, V.A. (1984) Prog. Lipid Res. 23, 39-67. 3 Geelen, M.J.H., Harris, R.A., Beynen, A.C. and McCune, S.A. (1980) Diabetes 29, 1006-1022. 4 Wakil, S.J., Stoops, J.K. and Joshi, V.C. (1983) Annu. Rev. Biochem. 52, 537-579. 5 Debeer, L.J., Beynen, A.C., Mannaerts, G.P. and Geelen, M.J.H. (1982) FEBS Lett. 140, 159-164. 6 Holm, C., Belfrage, P. and Fredrikson, G. (1987) Biochem. Biophys. Res. Commun. 148, 99-105. I Debeer, L.J., Thomas, J., De Schepper, P.J. and Mannaerts, G.P. (1979) J. Biol. Chem. 254, 8841-8846. 8 Seglen, P.O., Gordon, P.B., Holen, I. and Hoeyvik, H. (1991) Biomed. Biophys. Acta 50, 373-381. 9 Seglen, P.O. and Bohley, P. (1992) Experientia 48, 158-172. 10 Holen, I., Gordon, P.B. and Seglen, P.O. (1992) Biochem. J. 284, 633-636. 11 Duerden, J.M. and Gibbons, G.F. (1990) Biochem. J. 272, 583587. 12 Bjornsson, O.G., Duerden, J.M., Bartlett, SM., Sparks, J.D., Sparks, C.E. and Gibbons, G.F. (1992) Biochem. J. 281,381-386. 13 Wiggins, D. and Gibbons, G.F. (1992) Biochem. J. 284, 457-462. 14 Yeaman, S.J. (1990) Biochim. Biophys. Acta 1052, 128-132. A.A., Steinberg, D. and Tanaka, A. (1965) J. Biol. 1.5 Spector, Chem. 240, 1032-1041. 16 De Grella, R.F. and Light, R.J. (1980) J. Biol. Chem. 255. 9731-9738. W., Kleinert, H., Fischer, B.A., Gunawan, J., 17 Stremmel, Klaassen-Schhiter, C., Moller, K. and Wegener, M. (1992) Biochem. Sot. Trans. 20, 814-817. W., Strohmeyer, G., Borchard, F., Kochwa, S. and 18 Stremmel, Berk, P.D. (1985) Proc. Natl. Acad. Sci. USA 82, 4-8. G. and Berk, P.D. (1986) Proc. Natl. 19 Stremmel, W., Strohmeyer, Acad. Sci. USA 83, 3584-3588. L. (1986) Biochim. Biophys. Acta 20 Stremmel, W. and Theilmann, 877, 191-197. E., Gunawan, J., Manns, M. and 21 Diede, H.E., Rodilla-Sala, Stremmel, W. (1992) Biochim. Biophys. Acta 1125. 13-20.

239 22 Stremmel, W. (1987) J. Biol. Chem. 262, 6284-6289. 23 Stremmel, W. and Berk, P.D. (1986) Proc. Natl. Acad. Sci. USA 83,3086-3090. 24 Daniels, C., Noy, N. and Zakim, D. (1985) Biochemistry 24, 3286-3292. 25 Cooper, R.B., Noy, N. and Zakim, D. (1986) Hepatology 6,1187. 26 Cooper, R.B., C., Noy, N. and Zakim, D. (1987) Biochemistry 26,5890-5896. 27 Cooper, R.B., Noy, N. and Zakim, D. (1989) J. Lipid. Res. 30, 1719-1726. 28 Ferraresi-Filho, O., Ferraresi, M.L., Constantin, J., IshiiIwamoto, E.L., Schwab, A.J. and Bracht, A. (1992) Bichim. Biophys. Acta 1103, 239-249. 29 Abumrad, N.A., Harmon, CM., Barnela, U.S. and Whitesell, R.R. (1988) J. Biol. Chem. 263, 14678-14683. 30 Fu, B. and Hornick, C.A. (1992) Am. J. Physiol. 262, C1102Cl 108. 31 Have& R.S. and Hamilton, R.L. (1988) Hepatology 8, 1689-1704. 32 Kostner, G.M. (1989) Biochem. Sot. Trans. 17, 639-641. 33 Goldstein, J.L. and Brown, M.S. (1990) Nature 343, 425-430. 34 Johnson, W.J., Mahlberg, F.H., Rothblat, G.H. and Phillips, MC. (1991) Biochim. Biophys. Acta 1085, 273-298. 35 Veerkamp, J.H., Maatman, R.G.H. and Prinsen, C.F.M. (1992) B&hem. Sot. Trans. 20,801-805. 36 Stewart, J.M., Driedzic, W.R. and Berkelaar, J.A.M. (1991) B&hem. J. 275,569-573. 37 Vork, M.M., Glatz, J.F.C. and Van der Vusse, G.J. (1991) B&hem. J. 280,835. 38 Stewart, J.M. (1991) Biochem. J. 280, 835-836. 39 Brandes, R., Kaikaus, R.M., Lysenko, N., Ockner, R.K. and Bass, N.M. (1990) B&him. Biophys. Acta 1034,53-61. 40 Reubsaet, F.A.G., Veerkamp, J.H., Briickwilder, M.L.P., Trijbels, J.M.F. and Monnens, L.A.H. (1990) FEBS L&t. 267, 229230. 41 Groot, P.H.E., Scholte, H.R. and Hiilsmann, WC. (1976) In Advances in Lipid Research (Paoletti, R. and Kritchevsky, D., eds) Vol 14, pp. 75-126, Academic Press, New York. 42 Schulz, H. (1991) Biochim. Biophys. Acta 1081, 109-120. 43 Waku, K. (1992) B&him. Biophys. Acta 1124, 101-111. 44 Zammit, V.A. (1980) Biochem. J. 190, 293-300. 45 Suzuki, H., Kawarabayasi, Y., Kondo, J., Abe, T., Nishikawa, K., Kimura, S., Hashimoto, T. and Yamamoto, T. (1990) J. Biol. Chem. 265, 8681-8685. 46 Lazo, O., Contreras, M. and Singh, I. (1990) Biochemistry 29, 3981-3986. 47 Street, J.M., Singh, I. and Poulos, A. (1990) Biochem. J. 269, 671-677. 48 Oram, J.F., Wenger, J.I. and Neely, J.R. (1975) J. Biol. Chem. 250, 73-78. 49 Tijburg, L.B.M., Geelen, M.J.H. and Van Golde, L.M.G. (1989) B&him. Biophys. Acta 1004, 1-19. 50 Brindley, D.N. (1984) Prog. Lipid Res. 23, 115-133. 51 Fritz, I.B. (1961) Physiol. Rev. 41, 52-129. 52 Tzur, R., Tal, E. and Shapiro, B. (1964) Biochim. Biophys. Acta 84, 18-23. 53 Exton, J.H. and Park, C.R. (1967) J. Biol. Chem. 242,2622-2636. 54 Debeer, L.J., Declercq, P.E. and Mannaerts, G.P. (1981) FEBS Lett. 124, 31-34. 55 Beynen, A.C., Vaartjes, W.J. and Geelen, M.J.H. (1980) Horm. Metab. Res. 12, 425-430. 56 Declercq, P.E., Debeer, L.J. and Mannaerts, G.P. (1982) Biochem. J. 202, 803-806. 57 Zammit, V.A. (1981) B&hem. J. 198, 75-83. 58 Stals, H.K., Mannaerts, G.P. and Declercq, P.E. (1992) Biochem. J. 283, 719-725. 59 Dich, J., Bro, B., Grunnet, N., Jensen, F. and Kondrup, J. (1983) Biochem. J. 212, 617-623.

60 Cronholm, T. and Curstedt, T. (1984) B&hem. J. 224,731-739. 61 Bremer, J. (1983) Physiol. Rev. 63, 1420-1468. 62 Bieber, L.L. and Fiol, C.J. (1986) Biochem. Sot. Trans. 14, 674-676. 63 Bieber, L.L. (1988) Annu. Rev. Biochem. 57, 261-283. 64 Zammit, V.A. (1986) Biochem. Sot. Trans. 14, 676-679. 65 Saggerson, E.D. (1986) B&hem. Sot. Trans. 14, 679-681. 66 McGarry, J.D., Woeltje, K.F., Kuwajima, M. and Foster, D.W. (1989) Diabetes Metab. Rev. 5, 271-284. 67 Murthy, M.S.R. and Pande, S.V. (1987) Proc. Natl. Acad. Sci. USA 84, 378-382. 68 Healy, M.J., Kerner, J. and Bieber, L.L (1988) Biochem. J. 249, 231-237. 69‘ Ramsay, R.R. (1988) Biochem. J. 249, 239-245. 70 Murthy, M.S.R. and Pande, S.V. (1987) Biochem. J. 248, 727733. 71 Woeltje, K.F., Kuwajima, M., Foster, D.W. and McGarry, J.D. (1987) J. Biol. Chem. 262, 9822-9827. 72 McGarry, J.D., Sen, A., Esser, V., Woeltje, K.F., Weis, B. and Foster, D.W. (1991) Biochimie 73, 77-84. 73 Foley, J.E. (1992) Diabetes Care 15, 773-784. 74 Woeltje, K.F., Esser, V., Weis, B.C., Cox, W.F., Schroeder, J.G., Liao, S.T., Foster, D.W. and McGarry, J.D. (1990) J. Biol. Chem. 265, 10714-10719. 75 Declercq, P.E., Falk, J.R., Kuwajima, M., Tyminski, H., Foster, D.W. and McGarry, J.D. (1987) J. Biol. Chem. 262, 9812-9821. 76 Zammit, V.A., Costorphine, C.G. and Kolodziej, M.P. (1989) Biochem. J. 263, 89-95. 77 Murthy, M.S.R. and Pande, S.V. (1990) Biochem. J. 268, 599604. 78 Kolodziej, M.P., Crilly, P.J., Costorphine, C.G. and Zammit, V.A. (1992) Biochem. J. 282,415-421. 79 Kerner, J. and Bieber, L. (1990) Biochemistry 29, 4326-4334. 80 Ghadiminejad, I. and Saggerson, E.D. (1990) FEBS Lett. 269, 406-408. 81 Ghadiminejad, I. and Saggerson, E.D. (1990) Biochem. J. 270, 787-794. 82 Woldegiorgis, G., Fibich, B., Contreras, L. and Shrago, E. (1992) Arch. Biochem. Biophys. 295, 348-351. 83 Finocchiaro, G., Taromi, F., Rocchi, M., Liras Martin, A., Colombo, I., Torri Tarelli, G. and Di Donato, S. (1991) Proc. Natl. Acad. Sci. USA 88, 661-665. 84 Woeltje, K.F., Esser, V., Weis, B.C., Sen, A., Cox, W.F., McPhaul, M.J., Slaughter, C.A., Foster, D.W. and McGarry, J.D. (1990) J. Biol. Chem. 265, 10720-10725. 85 McGarry, J.D., Mannaerts, G.P. and Foster, D.W. (1977) J. Clin. Invest. 60, 265-270. 86 McGarry, J.D., Leatherman, G.F. and Foster, D.W. (1978) J. Biol. Chem. 253, 41284136. 87 Guzman, M. and Castro, J. (1989) Biochim. Biophys. Acta 1002, 405-408. 88 Zammit, V.A. (1983) Biochem. J. 210, 953-956. 89 Robinson, I.N. and Zammit, V.A. (1982) Biochem. J. 206, 177179. 90 Cook, G.A. and Cox, K.A. (1986) Biochem. J. 236, 917-919. 91 Bremer, J. (1981) Biochim. Biophys. Acta 665, 628-631. 92 Cook, G.A. and Gamble, MS. (1987) J. Biol. Chem. 262, 20502055. 93 Grantham, B.D. and Zammit, V.A. (1988) Biochem. J. 249, 409-414. 94 Stakkestad, J.A. and Bremer, J. (1983) Biochim. Biophys. Acta 750, 244-252. 95 Saggerson, E.D. and Carpenter, C.A. (1986) B&hem. J. 236, 137-141. 96 Grantham, B.D. and Zammit, V.A. (1986) Biochem. J. 239, 485-488.

240 97 Guzmln, M., Castro, J. and Maquedano, A. (1987) Biochem. Biophys. Res. Commun. 149, 443-448. 98 Prip-Buus, C., P&gorier, J.P., D&e, P.H., Kohl, C. and Girard, J. (1990) Biochem. J. 269, 409-415. 99 Decaux, J.F., FerrC, P., Robin, D., Robin, P. and Girard, J. (1988) J. Biol. Chem. 263, 3284-3289. 100 Pegorier, J.P., D&e, P.H., Herbin, C., Laulan, P.-Y., Blade, C., Perft, J. and Girard, J. (1988) Biochem. J. 249, 801-806. 101 Guzman, M. and Castro, J. (1990) Alcohol. Clin. Exp. Res. 14, 472-477. 102 Guzman, M. and Castro, J. (1990) Arch. Biochem. Biophys. 283. 290-295. 103 Baquet, A., Lavoinne, A. and Hue, L. (1991) Biochem. J. 273. 57-62. 104 Stephens, T.W. and Harris, R.A. (1987) Biochem. J. 243, 405105 GuzmLn, M. and Geelen, M.J.H. (1988) Biochem. Biophys, Res. Commun.151, 781-787. 106 Beynen, A.C., Vaartjes, W.J. and Geelen, M.J.H. (1979) Diabetes 28, 828-835. 107 Guzman, M. and Castro, J. (1991) FEBS Lett. 291, 105-108. 108 Guzmln, M. and Geelen, M.J.H. (1992) Biochem. J. 287, 487492. 109 Brady, P.S. and Brady, L.J. (1987) Biochem. J. 246, 641-649. 110 Brady, P.S., Marine, K.A., Brady, L.J. and Ramsay, R.R. (1989) Biochem. J. 260, 93-100. 111 Brady, P.S. and Brady, L.J. (1989) Biochem. J. 258, 677-682. 112 Wang, L., Brady, P.S. and Brady, L.J. (1989) Biochem. J. 263, 703-708. 113 Brady, P.S., Park, E.A., Liu, J., Hanson, R.W. and Brady, L.J. (1992) Biochem. J. 286, 779-783. 114 Kashfi, K. and Cook, G.A. (1992) Biochem. J. 282, 909-914. 115 Brady, L.J., Silverstein, L.J., Hoppel, CL. and Brady, P.S. (1985) Biochem. J. 232, 445-450. 116 Brady, L.J., Hoppel, C.L. and Brady, P.S. (1986) Biochem. J. 233, 427-433. 117 Kolodziej, M.P. and Zammit, V.A. (1990) Biochem. J. 272, 421-425. 118 Castro, J., Cartes, J.P. and Guzmin, M. (1991) Biochem. Pharmacol. 41, 1987-1995. 119 Kunan, W.H., Biihner, S., Moreno de la Garza, M., Kionka, C., Metablowski, M., Schultz-Borchard, J. and Thieringer, R. (1988) Biochem. Sot. Trans. 16, 418-420. 120 Palosaari, P.M. and Hiltunen, J.K. (1990) J. Biol. Chem. 265, 2446-2449. 121 Palosaari, P.M., Vihinen, M., Mantslla, PI., Alexson, S.E.H., Pihlajaniemi, T. and Hiltunen, J.K. (1991) J. Biol. Chem. 266, 10750-10753. 122 Osmundsen, H., Bremer, J. and Pedersen, J.I. (1991) Biochim. Biophys. Acta 1085, 141-158. 123 Osmundsen, H. and Hovik, R. (1988) Biochem. Sot. Trans. 16, 420-422. 124 Wang, H.Y. and Schulz, H. (1989) Biochem. J. 264, 47-52. 125 Smeland, T.E., Nada, M., Cuebas, D. and Schulz, H. (1992) Proc. Natl. Acad. Sci. USA 89, 6673-6677. 126 Coates, P.M. and Tenaka, K. (1992) J. Lipid Res. 33, 1099-1110. 127 Sumegi, B., Porpaczy, Z. and Alconyi, I. (1991) Biochim. Biophys. Acta 1081, 121-128. 128 Watmough, N.J., Turnball, D.M., Sherratt, H.S.A. and Bartlett, K. (1989) Biochem. J. 262, 261-269. 129 Vanhove, G., Van Veldhoven, P.P., Vanhoutte, F., Parmentier, G., Eyssen, H.J. and Mannaerts, G.P. (1991) J. Biol. Chem. 266, 24670-24675. 130 Van Veldhoven, P.P., Vanhove, G., Vanhoutte, F., Dacremont, G., Parmentier, G., Eyssen, H.J. and Mannaerts, G.P. (1991) J. Biol. Chem. 266, 24676-24683.

131 Izai, K., Uchida, Y., Orii, T., Yamamoto, S. and Hashimoto, T. (1992) J. Biol. Chem. 267, 1027-1033. 132 Uchida, Y., Izai, K., Orii, T. and Hashimoto. T. (1992) J. Biol. Chem. 267, 1034-1041. 133 Grunnet, N. and Kondrup, J. (1986) Alcohol. Clin. Exp. Res. IO. 64S-68s. 134 Latipiia, P.M., Karki, T.T., Hiltunen, J.K. and Hassinen, I.E. (1986) Biochim. Biophys. Acta 875, 293-300. 135 McKean, M.C., Frerman, F.E. and Mielke, D.M. (1979) J. Biol. Chem. 254, 2730-2735. 136 Olowe, Y. and Schulz, H. (1980) Eur. J. Biochem. 109, 425-429. 137 Wang, H.Y., Baxter, C.F. and Schulz, H. (1991) Arch. Biochem. Biophys. 289, 2744280. 138 He, X.Y., Yang, S.Y. and Schulz, H. (1992) Arch. Biochem. Biophys. 298, 527-53 1. 139 Van Hoof, F., Vameq, J., Draye, J.P. and Veitch, K. (1988) Biochem. Sot. Trans. 16, 423-424. 140 Pettersen, J.E. (1972) Clin. Chim. Acta 41, 231-237. 141 Mitz, M.A. and Heinrikson, R.L. (1961) Biochim. Biophys. Acta 46, 45-50. 142 Kolveraa, S. and Gregersen, N. (1986) Biochim. Biophys. Acta 876. 515-525. 143 Christensen, E., Gronn, M., Hagve, T.A. and Christophersen, B.O. (1991) Biochim. Biophys. Acta 1081, 167-173. 144 Sugden, M.C., Holness, M.J. and Palmer, T.N. (1989) Biochem. J. 263, 313-323. 145 Lopes-Cardozo, M., Mulder, I., Van Vught, F., Hermans. P.G.C. and Van den Bergh, S.G. (1975) Mol. Cell. Biochem. 9, 1555173. 146 Baranyai, J. and Blum, J.J. (1989) Biochem. J. 258, 121-140. 147 Norsten, C. and Cronholm, T. (1990) Biochem. J. 265. 569-574. 148 Des Rosiers, C., David, F., Garneau, M. and Brunengraber. H. (1991) J. Biol. Chem. 266, 1574-1578. 149 Corvera. S., Huerta-Bahem, J., Pelton, T.J., Hruby, V.J., Trivedi, D. and Garcia-Sanz, J.A. (1984) Biochim. Biophys. Acta 804, 4344441. I50 Wakelam, M.J.O., Murphy, G.J., Hruby, V.J. and Houslay, M.D. (1986) Nature 323, 68-71. I51 Kraus-Friedman, N. (1986) Trends Biochem. Sci. 1 I, 276-279. 152 Halestrap, A.P. (1989) Biochim. Biophys. Acta 973, 355-382. 153 Williamson, D.H., Illic, V., Tordoff, A.F.C. and Ellington. E.V. (1980) Biochem. J. 186, 621-624. 154 Almas, I., Singh, B. and Borrebaek, B. (1983) Arch. Biochem. Biophys. 222, 370-379. 155 Chihara, M., Nomura, T., Tachibana, M., Nomura, H., Nomura, Y. and Hagino, Y. (1989) Biochim. Biophys. Acta 1012, 5-9. 156 Harano, A., Hidaka, H., Kojima, H., Harano, Y. and Shigeta, Y. (1992) Horm. Metab. Res. 24, 5-9. 157 Kosugi, K., Harano, Y., Nakano. T.. Suzuki, M., Kashiwagi. A. and Sigeta, Y. (1983) Metabolism 32, 1081-1087. 158 Oberhaensli, R.D., Schwendimann, R. and Keller. U. (198.5) Diabetes 34. 374-379. 159 Sugden, M.C., Tordoff, A.F.C., illic, V. and Williamson, D.H. (1980) FEBS Lett. 120, 80-84. 160 Nomura, T., Nomura. Y., Tachibana, M., Nomura, H., Ukai, K.K., Yokohama, R. and Hagino, Y. (1991) Biochim. Biophys. Acta 1092, 94-100. 161 Vaartjes, W.J., Bijleveld, C.. Geelen, M.J.H. and Van den Bergh, S.G. (1986) Biochem. Biophys. Res. Commun. 139, 403409. 162 Vaartjes, W.J. and De Haas, C.G.M. (1985) Biochem. Biophys. Res. Commun. 129, 721-726. 163 Brady, P.S. and Brady, L.J. (1986) Biochem. J. 238, 801-809. 164 Penicaud, L., Robin, D., Robin, P., Kande, J., Picon, L., Girard. J. and Ferre, P. (1991) Metabolism 40, 873-876. 165 Lowe, D.M. and Tubbs, P.K. (1985) Biochem. J. 227. S91-599.

241 166 Gil, G., Goldstein, J.L., Slaughter, C.A. and Brown, MS. (1986) J. Biol. Chem. 261, 3710-3716. 167 Ayte, J., Gil-Gbmez, G., Haro, D., Marrero, P.F. and Hegardt, F.G. (1990) Proc. Natl. Acad. Sci. USA 87,3874-3878. 168 Lowe, D.M. and Tubbs, P.K. (1985) Biochem. J. 232, 37-42. 169 Lowe, D.M. and Tubbs, P.K. (1985) Biochem. J. 227, 601-607. 170 Quant, P.A., Tubbs, P.K. and Brand, M.D. (1989) Biochem. J. 262, 159-164. 171 Quant, P.A., Tubbs, P.K. and Brand, M.D. (1990) Eur. J. Biochem. 187, 169-174. 172 Quant, P.A., Robin, D., Robin, P., FerrC, P., Brand, M.D. and Girard, J. (1991) Eur. J. Biochem. 195,449-454. 173 Casals, N., Rota, N., Guerrero, M., Gil-Gbmez, G., Ayte, J., Ciudad, C.J. and Hegardt, F.G. (1992) Biochem. J. 283,261-264. 174 Pigorier, J.P., D&e, P.H., Herbin, C., Laulan, P.Y., Blade, C., Peret, J. and Girard, J. (1988) Biochem. J. 249, 801-806. 175 Tosh, D., Alberti, K.G.M. and Agius, L. (1988) Biochem. J. 256, 197-204. 176 Guzman, M. and Castro, J. (1989) B&hem. J. 264, 107-113. 177 Srere, P.A. (1987) Annu. Rev. B&hem. 56, 89-124. 178 Sumegi, B., Sherry, A.D., Mallor, C.R., Evans, C. and Srere, P.A. (1991) Biochem. Sot. Trans. 19, 1002-1005. 179 McCormack, J.G. and Denton, R.M. (1990) Annu. Rev. Physiol. 52,451-466. 180 McGxmack, J.G., Halestrap, A.P. and Denton, R.M. (1990) Physiol. Rev. 70, 391-425. 181 Pate], T.B. (1986) Eur. J. Biochem. 159, 17-22. 182 Mohan, C., Memon, R.A. and Bessman, S.P. (1991) Arch. B&hem. Biophys. 287, 18-23. 183 McCormack, J.G. and Denton, R.M. (1986) Trends B&hem. Sci. 11, 258-262. 184 Houslay, M.D., Wakelam, M.J.D., Murphy, G.J., Gawter, D.J. and Pyne, N.J. (1987) Biochem. Sot. Trans. 15,21-24. 185 Bataille, D. (1990) Ann. Endocrinol. (Paris) 51, 101-107. 186 Brocks, D.G., Siess, E.A. and Wieland, O.H. (1980) Eur. J. Biochem. 113, 39-43. 187 Smith, CM. and Williamson, J.R. (1971) FEBS Lett. 18, 35-38. 188 Yeaman, S.J. (1988) Biochem. J. 257, 625-632. 189 Reed, L.J. and Hackert, M.L. (1990) J. Biol. Chem. 265, 89718974.

190 Wan, B., LaNoue, K.F., Cheung, J.Y. and Scaduto, R.C. (1989) J. Biol. Chem. 264, 13430-13439. 191 Brown, G.C. (1992) Biochem. J. 284, 1-13. 192 Randle, P.J. (1986) B&hem. Sot. Trans. 14, 799-806. 193 Sterniczuk, A., Hreniuk, S., Scaduto, R.C. and LaNoue, K.F. (1991) Eur. J. Biochem. 196, 143-150. 194 Bartlett, K., Hovik, R., Eaton, S., Watmough, N.J. and Osmundsen, H. (1990) Biochem. J. 270, 175-180. 195 Pourfarzam, M. and Bartlett, K. (1992) Eur. J. Biochem. 208, 301-307. 196 Leighton, F., Bergseth, S., Rortveit, T., Christiansen, E.N. and Bremer, J. (1989) J. Biol. Chem. 264, 10347-10350. 197 Handler, J.A. and Thurman, R.G. (1988) Eur. J. Biochem. 176, 477-484. 198 Skorin, C., Necochea, C., Johow, V., Soto, V., Grau, A.M., Bremer, J. and Leighton, F. (1992) Biochem. J. 281, 561-567. 199 Rognstadt, R. (1991) Biochem. J. 279, 147-150. 200 Vamecq, J. (1987) Biochem. J. 241, 783-791. 201 Buechler, K.F. and Lowenstein, J.M. (1990) Arch. Biochem. Biophys. 281, 233-238. 202 Derrick, J.P. and Ramsay, R.R. (1989) Biochem. J. 262,801-806. 203 Nit A’bhaird, N. and Ramsay, R.R. (1992) Biochem. J. 286, 637-640. 204 Cook, G.A. and Weakley, L.J. (1990) Biochem. Sot. Trans. 18, 988. 205 Haystead, T.A.J., Slim, A.T.R., Carling, D., Honnor, R.C., Tsukitani, Y., Cohen, P. and Hardie, D.G. (1989) Nature 337,78-81. 206 Geelen, M.J.H., Papiez, J.S., Girgis, K. and Gibson, D.M. (1991) Biochem. Biophys. Res. Commun. 180,525-530. 207 Kunz, W.S. (1991) Biomed. B&him. Acta 50, 1143-1157. 208 Moir, A.B.M. and Zammit, V.A. (1992) Biochem. J. 283, 145149. 209 Moir, A.B.M. and Zammit, V.A. (1993) Biochem. J. 289,45-55. 210 Gebhart, R. (1992) Pharmacol. Ther. 53, 275-354. 211 Gardemann, A., Piischel, G.P. and Jungermann, K. (1992) Eur. J. Biochem. 207, 399-411. 212 Haiissinger, D. and Lang, F. (1991) Biochim. Biophys. Acta 1071,331-350. 213 Haussinger, D. and Lang, F. (1992) Trends Pharmacol. Sci. 13, 371-373.