Utilization of dietary lipids requires that they first be absorbed through the intestine. As these molecules are oils they would be essentially insoluble in the aqueous intestinal environment. Solubilization (emulsification) of dietary lipid is accomplished initially via the agitation action as food passes through the stomach and then continues within the intestine via bile salts that are synthesized in the liver and secreted from the gallbladder.
The emulsified fats can then be degraded by salivary, gastric and pancreatic lipases. The lipases found in the gastrointestinal tract include lingual lipase (secreted by the serous glands of the tongue), gastric lipase (secreted by chief cells of the stomach), pancreatic lipase, and pancreatic lipase-related proteins 1 and 2. These enzymes generate free fatty acids and a mixture of mono- and diacylglycerols from dietary triacylglycerols. Pancreatic lipase degrades triacylglycerols at the 1 and 3 positions sequentially to generate 1,2-diacylglycerols and 2-acylglycerols. Phospholipids are degraded at the 2 position by pancreatic phospholipase A2 releasing a free fatty acid and the lysophospholipid.
Following absorption of the products of pancreatic lipase by the intestinal mucosal cells, the resynthesis of triacylglycerides occurs. The triacylglycerides are then solubilized in lipoprotein complexes (complexes of lipid and protein) called chylomicrons. A chylomicron contains lipid droplets surrounded by the more polar lipids and finally a layer of proteins. Triacylglycerides synthesized in the liver are packaged into VLDLs and released into the blood directly. Chylomicrons from the intestine are then released into the blood via the lymph system for delivery to the various tissues for storage or production of energy through oxidation.
The triacylglyceride components of VLDLs and chylomicrons are hydrolyzed to free fatty acids and glycerol in the capillaries of tissues such as liver, adipose tissue and skeletal muscle by the actions of lipoprotein lipase (LPL) and hepatic triglyceride lipase (HTGL). The free fatty acids are then absorbed by the cells and the glycerol is returned via the blood to the liver (principal site) and kidneys. The glycerol can then converted to the glycolytic intermediate dihydroxyacetone phosphate DHAP or phosphorylated by glycerol kinase to glycerol-3-phosphate for reuse in triglyceride synthesis.
The classification of blood lipids is distinguished based upon the density of the different lipoproteins. As lipid is less dense than protein, the lower the density of lipoprotein the less protein there is.
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The primary sources of fatty acids for oxidation are dietary and mobilization from cellular stores. Fatty acids from the diet are absorbed from the gut, packaged into lipoprotein particles called chylomicrons within intestinal enterocytes and then delivered to cells of the body via transport in the blood. Fatty acids are stored in the form of triacylglycerols (triacylglycerides: TAGs or TGs) within all cell but predominantly within adipose tissue. In response to energy demands, the fatty acids of stored TGs can be mobilized for use by peripheral tissues. The release of metabolic energy, in the form of fatty acids, is controlled by a complex series of interrelated cascades that result in the activation of triglyceride hydrolysis. The primary intracellular lipases are adipose triglyceride lipase (ATGL, also called desnutrin), hormone-sensitive lipase (HSL), and lysosomal acid lipase (LAL). LAL is the most important lipase involved in lysosomal lipid metabolism as evidenced by the fact that LAL deficiency results in the significant accumulation of cholesteryl esters in tissues such as the spleen and liver. LAL deficiency is commonly called Wolman disease.
ATGL/desnutrin belongs to the family of patatin domain-containing proteins that consists of nine human members. The patatin domain was originally discovered in lipid hydrolases of certain plants and named after the most abundant protein of the potato tuber, patatin. Because some members of the family act as phospholipases, the proteins were originally called patatin-like phospholipase domain-containing protein A1 to A9 (PNPLA1–PNPLA9). ATGL (designated PNPLA2 in the patatin domain nomenclature) preferentially hydrolyzes TGs. The human ATGL gene is located on chromosome 11p15.5 and is composed of 9 exons encoding a 504 amino acids protein. In analogy with patatin, the active site of ATGL contains an unusual catalytic dyad (S47 and D166) within the patatin domain in the N-terminal half of the enzyme. The C-terminal half of the enzyme comprises the regulatory functions and also contains a predicted hydrophobic region involved in lipid droplet (LD) binding.
ATGL expression and enzyme activity are both under complex regulation. Expression of ATGL is induced by peroxisome proliferator-activated receptor (PPAR) agonists, glucocorticoids, and fasting. In addition, activation of FoxO1 by SIRT1-mediated deacetylation activates lipolysis by increasing ATGL expression. Conversely, silencing of SIRT1 has the opposite effect. Increased insulin release and food intake both result in decreased expression of ATGL. Reductions in ATGL expression have also been shown to be associated with mTOR complex 1 (mTORC1)-dependent signaling. The level of lipase activity of ATGL (as well as HSL) does not always correlate to the level of expression of the gene. For example, TNFα activity reduces both ATGL and HSL expression at the level of the gene but result in increased lipase activity resulting in fatty acid and glycerol release from TGs. Similarly, the use of non-selective beta blockers (e.g. isoproterenol) results in the same type of gene inhibition with enzyme activation. The discrepancy between ATGL and HSL mRNA levels and enzyme activities is the result of extensive post-translational regulation of both ATGL and HSL.
There are two known serine residues in ATGL that are subject to phosphorylation. Unlike phosphorylation of HSL (described below), ATGL phosphorylation is not PKA-dependent. In the mouse, it has been shown that AMPK phosphorylates ATGL resulting in increased lipase activity. However, it is important to point out that there remains a level of controversy as to the role of AMPK in the regulation of the activity of ATGL since induction, inhibition, and no effects results have been published.
In addition to phosphorylation regulating ATGL activity, the enzyme requires a coactivator protein for full lipase activity. This coactivator gene is known as comparative gene identification-58 (CGI-58). The term comparative gene identification relates to the use of computational methods to identify protein sequences highly conserved across various species and CGI-58 was originally discovered in a screen comparing the proteomes of humans and C. elegans. The official nomenclature for CGI-58 is α/β hydrolase domain-containing protein-5 (ABHD5), owing to the presence of an α/β hydrolase domain commonly found in esterases, thioesterases, and lipases. It is unlikely that CGI-58 possesses hydrolase activity because an asparagine residue is found in the catalytic domain where other hydrolases have a nucleophilic serine residue that is required enzymatic activity. In addition to functioning as an ATGL coactivator CGI-58 has also been shown to possess enzymatic activity as an acyl-CoA-dependent acylglycerol-3-phosphate acyltransferase (AGPAT). The physiological significance of this activity is currently unknown but it may be involved in the regulation of phosphatidic acid or lysophosphatidic acid signaling.
Additional proteins associated with lipid droplets (LD) in adipocytes participate in the CGI-58-mediated regulation of ATGL. In resting adipocytes of both WAT and BAT, the LD protein perilipin-1 interacts with CGI-58, preventing its binding to and, induction of ATGL. Following β-adrenergic stimulation of WAT, PKA phosphorylates perilipin-1 at multiple sites resulting in the release of CGI-58 which in turn, binds and stimulates ATGL. This demonstrates that β-adrenergic stimulation of PKA induces ATGL activity, not by direct phosphorylation of ATGL itself, but through phosphorylation of perilipin-1. This model of ATGL regulation is evident from frameshift mutants that have been identified in human perilipin-1. These mutations are identified as V398fs and L404fs indicating that the frame shift occurs at valine 398 and leucine 404, respectively. Each of these mutant ATGL proteins fail to bind CGI-58, resulting in unrestrained lipolysis, partial lipodystrophy, hypertriglyceridemia, and insulin resistance.
In non-adipose tissues with high rates of TG hydrolysis, such as skeletal muscle and liver, regulation of ATGL activity occurs via a mechanism distinct from that in adipose tissues. In these tissues, perilipin-1 is replaced by perilipin-5. During fasting, perilipin-5 recruits both ATGL and CGI-58 to LDs by direct binding of the enzyme and its coactivator. Data indicates that perilipin-5 is involved in the interaction of LDs with mitochondria and thereby inhibits ATGL-mediated TG hydrolysis. Other perilipins exist in cells including perilipin-2, -3, and -4 but it is unclear if these proteins are also involved in regulating the association of ATGL with LDs. In hepatocyte cell lines it has been shown that overexpression of perilipin-2 inhibits ATGL activity by restricting its physical access to LDs.
Recently, a specific peptide inhibitor for ATGL was isolated from white blood cells, specifically mononuclear cells. This peptide was originally identifed as being involved in the regulation of the G0 to G1 transition of the cell cycle. This peptide was, therefore, called G0G1 switch protein 2 (G0S2). The protein is found in numerous tissues, with highest concentrations in adipose tissue and liver. In adipose tissue G0S2 expression is very low during fasting but increases after feeding. Conversely, fasting or PPARα-agonists increase hepatic G0S2 expression. The protein has been shown to localize to LDs, cytoplasm, ER, and mitochondria. These different subcellular localizations likely relate to multiple functions for G0S2 in regulating lipolysis, the cell cycle, and, possibly, apoptosis via its ability to interact with the mitochondrial antiapoptotic factor Bcl-2. With respect to ATGL regulation, the binding of the enzyme to LDs and subsequent is dependent on a physical interaction between the N-terminal region of G0S2 and the patatin domain of ATGL.
The delivery of ATGL to LDs requires functional vesicular transport. When essential protein components of the transport machinery are defective or missing, such as ADP-ribosylation factor 1 (ARF1), small GTP-binding protein 1 (SAR1), the guanine-nucleotide exchange factor Golgi-Brefeldin A resistance factor (GBF1), or the coatamer protein coat-complex I (COPI) and COPII, ATGL translocation to LDs is blocked and the enzyme remains associated with the ER.
A landmark study published in 1964 demonstrated that a lipolytic activity present in adipose tissue was induced by hormonal stimulation. This work described the isolation and characterization of both HSL and monoacylglyceride lipase (MGL). This original study demonstrated that HSL had a higher level of activity as a DG hydrolase than as a TG hydrolase. Nevertheless, it became dogma that HSL was rate-limiting for the catabolism of fat stores in adipose and many non-adipose tissues. However, when HSL-deficient mice were produced and shown to efficiently hydrolyze TGs the model began to emerge demonstrating ATGL, and not HSL, to be rate-limiting for adipose tissue TG hydrolysis. HSL-deficient mice do not accumulate TGs in either adipose or non-adipose tissues, but they do accumulate large amounts of DGs in many tissues. This indicated for the first time that HSL was more important as a DG hydrolase than a TG hydrolase. It is now accepted that ATGL is responsible for the initial step of lipolysis in human adipocytes, and that HSL is rate-limiting for the catabolism of DGs. HSL not only hydrolyzes DGs but is also active at hydrolyzing ester bonds of many other lipids including TGs, MGs, cholesteryl esters, retinyl esters, and short-chain carbonic acid esters.
The HSL gene is located on chromosome 19q13.2. Alternative exon useage results in tissue-specific differences in mRNA and protein size. In adipose tissue the HSL protein is composed of 775 amino acids, whereas the testicular form is composed of 1,076 amino acids. The expression profile of HSL essentially mirrors that of ATGL. Highest mRNA and protein concentrations are found in WAT and BAT with low levels of expression found in muscle, testis, steroidogenic tissues, and pancreatic islets as well as several other tissues. Functional studies on the enzyme have identified an N-terminal lipid-binding region, the α/β hydrolase fold domain including the catalytic triad, and the regulatory module containing all known phosphorylation sites important for regulation of enzyme activity.
Model for the activation of hormone-sensitive lipase by epinephrine. Epinephrine binds its receptor and leads to the activation of adenylate cyclase. The resultant increase in cAMP activates PKA which then phosphorylates and activates hormone-sensitive lipase. Hormone-sensitive lipase hydrolyzes fatty acids from diacylglycerols that result from the action of ATGL/desnutrin. The final fatty acid is released from monoacylglycerols through the action of monoacylglycerol lipase (MGL), an enzyme that is also active in the absence of hormonal stimulation.
HSL and ATGL share many regulatory similarities yet the mechanisms of the regulatory processes differ markedly between the two enzymes. In adipose tissue, HSL enzyme activity is strongly induced by β-adrenergic stimulation, conversely insulin has a strong inhibitory effect. While β-adrenergic stimulation regulates ATGL primarily via recruitment of the coactivator CGI-58), HSL is a major target for PKA-mediated phosphorylation. Additional kinases, including AMPK, extracellular signal-regulated kinase (ERK), glycogen synthase kinase-4 (GSK-4), and Ca2+/calmodulin-dependent kinase 1 (CAMK1), also phosphorylate HSL to modulate the activity of the enzyme. HSL has at least five potential phosphorylation sites, of which S660 and S663 appear to be particularly important for hydrolytic activity. Enzyme phosphorylation affects enzyme activity only moderately resulting in an approximate 2-fold increase in hydrolytic activity. For full activation, HSL must gain access to LDs, which, in adipose tissue, is mediated by perilipin-1. In addition to phosphorylating HSL, PKA also phosphorylates perilipin-1 on six consensus serine residues. The result of these phosphorylations is the binding of HSL to the N-terminal region of perilipin-1. This protein-protein interaction is the means by which HSL gains access to LDs. The net effect, of HSL-phosphorylation and enzyme translocation to LDs, coupled with ATGL activation by CGI-58, leads to a more than 100-fold increase in TG hydrolysis in adipocytes.
Additional factors modulate the activation of HSL and ATGL. One such factor is receptor-interacting protein 140 (RIP-140) which induces lipolysis by binding to perilipin-1, increasing HSL translocation to LDs, and activating ATGL via CGI-58 dissociation from perilipin-1. In non-adipose tissues, such as skeletal muscle, HSL is activated by phosphorylation in response to epinephrine (β-adrenergic receptor-mediated activation of PKA) and muscle contraction (calcium release from sarcoplasmic reticulum). Since skeletal muscles lack perilipin-1 it has not yet been determined which alternative mechanisms regulate HSL access to LDs.
Insulin-mediated deactivation of lipolysis is associated with transcriptional downregulation of ATGL and HSL expression. Additionally, insulin signaling results in phosphorylation and activation of various phosphodiesterase (PDE) isoforms by PKB/Akt leading to PDE-catalyzed hydrolysis of cAMP which in turn results in reduced activation of PKA. These actions turn off lipolysis by preventing phosphorylation of both HSL and perilipin-1, activation and translocation of HSL, and activation of ATGL by CGI-58. In addition to its peripheral action, insulin also functions within the sympathetic nervous system to inhibit lipolysis in WAT. Increased insulin levels in the brain inhibit HSL and perilipin phosphorylation which results in reduced HSL and ATGL activities.
MGL is considered to be the rate-limiting enzyme for the breakdown of MGs that are the result of both extracellular and intracellular lipolysis pathways. The extracellular generation of MGs is the result of the action of endothelial cell lipoprotein lipase (LPL) on lipoprotein particle-associated TGs. Intracellular hydrolysis of TGs by ATGL and HSL, as well as intracellular phospholipid hydrolysis by phospholipase C (PLC) and membrane-associated DG lipase α and β results in the generation of MGL substrates.
The MGL gene is located on chromosome 3q21.3 and is composed of 7 exons encoding a protein of 313 amino acids. MGL has been shown to localize to LDs, cell membranes,and the cytosol. The enzyme is ubiquitously expressed with highest levels of expression in adipose tissue. MGL shares homology with esterases, lysophospholipases, and haloperoxidases. The enzyme contains a consensus GXSXG motif within a catalytic triad that is typical of lipases and esterases.
MGL is critically important for efficient degradation of MGs since it has been shown in mouse models that lack of MGL impairs lipolysis and is associated with increased MG levels in adipose and non-adipose tissues alike. MGL has received particular attention in recent years due to the discovery that the enzyme is responsible for the inactivation of 2-arachidonoylglycerol (2-AG) which is an endogenous cannabinoid monoglyceride (endocannabinoid).
For more information on the activities of ATGL, HSL and other lipases regulating triacylglycerol levels in adipocytes visit the Adipose Tissue page.
In contrast to the hormonal activation of adenylate cyclase and (subsequently) hormone-sensitive lipase in adipocytes, the mobilization of fat from adipose tissue is inhibited by numerous stimuli. The most significant inhibition is that exerted upon adenylate cyclase by insulin. When an individual is in the well fed state, insulin released from the pancreas prevents the inappropriate mobilization of stored fat. Instead, any excess fat and carbohydrate are incorporated into the triacylglycerol pool within adipose tissue.back to the top
When fatty acids are released from adipose tissue stores they enter the circulation as free fatty acids (FFAs) and are bound to albumin for transport to peripheral tissues. When the fatty acid–albumin complexes interact with cell surfaces the dissociation of the fatty acid from albumin represents the first step of the cellular uptake process. Uptake of fatty acids by cells involves membrane proteins with high affinity for fatty acids. There are several members of the fatty acid receptor family including fatty acid translocase (FAT/CD36), plasma membrane-associated fatty acid-binding protein (FABPpm), and fatty acid transport proteins (FATPs). The FATPs are a family of at least six fatty acid transport proteins (FATP1–FATP6) that are also members of the solute carrier family of transporters. The FATPs facilitate the uptake of very long-chain (VLCFA) and long-chain fatty acids (LCFA).
|FAT/CD36||fatty acid translocase; FAT is also known as CD36 which is a member of the scavenger receptor class (class B scavenger receptors) of receptors that bind lipids and lipoproteins of the LDL family; located on chromosome 7q11.2, composed of 15 exons spanning 32 kb|
|FABPpm||plasma membrane-associated fatty acid-binding protein|
|FATP1||FATP1 is SLC27A1; FATP1 is also known as acyl-CoA synthetase very long-chain family, member 4 (ACSVL4); highest levels of expression in adipose tissue, skeletal and heart muscle; located on chromosome 19p13.11 spanning 13 kb, composed of 12 exons encoding a 646 amino acid protein|
|FATP2||FATP2 is SLC27A2; FATP2 is also known as acyl-CoA synthetase very long-chain family, member 1 (ACSVL1) as well as very long-chain acyl-CoA synthetase (VLCS); highest levels of expression in liver and kidney; present in peroxisome and microsomal membranes; located on chromosome 15q21.2 composed of 10 exons encoding a 620 amino acid protein|
|FATP3||FATP3 is SLC27A3; FATP3 is also known as acyl-CoA synthetase very long-chain family, member 3 (ACSVL3); located on chromosome 1q21.3 composed of 10 exons encoding a 730 amino acid protein|
|FATP4||FATP4 is SLC27A4; FATP4 is also known as acyl-CoA synthetase very long-chain family, member 5 (ACSVL5); is the major intestinal long-chain fatty acid transporter; located on chromosome 9q34.11 spanning 17 kb, composed of 13 exons encoding a 643 amino acid protein|
|FATP5||FATP5 is SLC27A5; FATP5 is also known as acyl-CoA synthetase very long-chain family, member 6 (ACSVL6), very long-chain acyl-CoA synthetase-related protein (VLACSR), or very long-chain acyl-CoA synthetase homolog 2 (VLCSH2); highest levels of expression in the liver; capable of activating 24- and 26-carbon VLCFAs; located on chromosome 19q13.43 composed of 10 exons encodong a 690 amino acid protein|
|FATP6||FATP6 is SLC27A6; FATP6 is also known as acyl-CoA synthetase very long-chain family, member 2 (ACSVL2), very long-chain acyl-CoA synthetase homolog 1 (VLCSH1); expressed at highest levels in the heart; protein only detected in heart and testis; exhibits a preference for the transport of palmitic acid and linoleic acid, does not transport fatty acids less than 10 carbons long; located on chromosome 5q23.3 spanning 67 kb and composed of 11 exons encoding a 619 amino acid protein|
The result of the interaction of fatty acids with plasma membrane receptors/binding proteins is a transmembrane concentration gradient. At the plasma membrane the apparent pKa of the fatty acid shifts from about 4.5 in aqueous solutions to about 7.6. This pKa change is independent of fatty acid type. As a consequence, about half of the fatty acids are present in the un-ionized form. This local environment effect promotes a transfer (flip-flop) of uncharged fatty acids from the outer leaflet across the phospholipid bilayer. At the cytosolic surface of the plasma membrane, fatty acids can associate with the cytosolic fatty acid binding protein (FABPc) or with caveolin-1. Caveolin-1 is a constituent of caveolae (Latin for little caves) which are specialized "lipid rafts" present in flask-shaped indentations in the plasma membranes of many cells types that perform a number of signaling functions by serving as lipid delivery vehicles for subcellular organelles. In order that the fatty acids that are thus taken up to be directed to the various metabolic pathways (e.g. oxidation or triglyceride synthesis) they must be activated to acyl-CoA. Members of the atty acid transport protein (FATP) family have been shown to possess acyl-CoA synthetase (ACS) activity. Activation of fatty acids by FATPs occurs at the highly conserved cytosolic AMP-binding site of these proteins. The overall process of cellular fatty acid uptake and subsequent intracellular utilization represents a continuum of dissociation from albumin by interaction with the membrane-associated transport proteins, binding to FABPc and caveolin-1 at the cytosolic plasma membrane, activation to acyl-CoA (in many cases via FATP action) followed by intracellular trafficking via FABPc and/or caveolae to sites of metabolic disposition.back to the top
Oxidation of fatty acids occurs in the mitochondria and the peroxisomes (see below). Fatty acids of between 4–8 and between 6–12 carbon atoms in length, referred to as short- and medium-chain fatty acids (SCFAs and MCFAs, respectively), are oxidized exclusively in the mitochondria. Long-chain fatty acids (LCFAs: 10–16 carbons long) are oxidized in both the mitochondria and the peroxisomes with the peroxisomes exhibiting preference for 14-carbon and longer LCFAs. Very-long-chain fatty acids (VLCFAs: C17–C26) are preferentially oxidized in the peroxisomes.
Fatty acids must be activated in the cytoplasm before being oxidized in the mitochondria. Activation is catalyzed by fatty acyl-CoA synthetases (also called acyl-CoA ligases or thiokinases). The net result of this activation process is the consumption of 2 molar equivalents of ATP.
The transport of fatty acyl-CoA into the mitochondria is accomplished via an acyl-carnitine intermediate, which itself is generated by the action of carnitine palmitoyltransferase 1 (CPT-1 or CPT-I) an enzyme that resides in the outer mitochondrial membrane. There are three CPT-1 genes in humans identified as CPT-1A, CPT-1B, and CPT-1C. Expression of CPT-1A predominates in the liver and is thus, referred to as the liver isoform. CPT-1B expression predominates in skeletal muscle and is thus, referred to as the muscle isoform. CPT-1C expression is exclusive to the brain and testes. The CPT-1A gene (symbol = CPT1A) is located on chromosome 11q13.3 and consists of 20 exons spanning 60 kb encoding a 773 amino acid protein. The CPT-1B gene (symbol = CPT1B) is located on chromosome 22q13.33 and consists of 21 exons spanning 10 kb. The CPT-1C gene (symbol = CPT1C) is located on chromosome 19q13.3 and consists of 20 exons spanning 23 kb. The activity of CPT-1C is distinct from those of CPT-1A and CPT-1B in that it does not act on the same types of fatty acyl-CoAs that are substrates for the latter two enzymes. However, CPT-1C does exhibit high-affinity malonyl-CoA binding.
Following carnitine acyl-carnitine-mediated transfer of the CPT-1-generated fatty acyl-carnitines across the inner mitochondrial membrane, the fatty acyl-carnitine molecules are acted on by the inner mitochondria membrane carnitine palmitoyltransferase 2 (CPT-2 or CPT-II) regenerating the fatty acyl-CoA molecules. The CPT-2 gene (symbol = CPT2) is located on chromosome 1p32.3 and consists of 5 exons that span 20 kb.
Transport of fatty acids from the cytoplasm to the inner mitochondrial space for oxidation. Following activation to a fatty-CoA, the CoA is exchanged for carnitine by CPT-1. The fatty-carnitine is then transported to the inside of the mitochondrion where a reversal exchange takes place through the action of CPT-2. Once inside the mitochondrion the fatty-CoA is a substrate for the β-oxidation machinery.
The process of mitochondrial fatty acid oxidation is termed β-oxidation since it occurs through the sequential removal of 2-carbon units by oxidation at the β-carbon position of the fatty acyl-CoA molecule. The oxidation of fatty acids and lipids in the peroxisomes (see below) also occurs via a process of β-oxidation. Each round of β-oxidation involves four steps that, in order, are oxidation, hydration, oxidation, and cleavage.
The first oxidation step in mitochondrial β-oxidation involves a family of FAD-dependent acyl-CoA dehydrogenases. Each of these dehydrogenases has a range of substrate specificity determined by the length of the fatty acid. Short-chain acyl-CoA dehydrogenase (SCAD, also called butyryl-CoA dehydrogenase) prefers fats of 4–6 carbons in length; medium-chain acyl-CoA dehydrogenase (MCAD) prefers fats of 4–16 carbons in length with maximal activity for C10 acyl-CoAs; long-chain acyl-CoA dehydrogenase (LCAD) prefers fats of 6–16 carbons in length with maximal activity for C12 acyl-CoAs.
The next three steps in mitochondrial β-oxidation involve a hydration step, another oxidation step, and finally a hydrolytic reaction that requires CoA and releases acetyl-CoA and an acy-CoA two carbon atoms shorter than the initial substrate. The water addition is catalyzed by an enoyl-CoA hydratase activity, the second oxidation step is catalyzed by an NAD-dependent long-chain hydroxacyl-CoA dehydrogenase activity (3-hydroxyacyl-CoA dehydrogenase activity), and finally the cleavage into an acyl-CoA and an acetyl-CoA is catalyzed by a thiolase activity. These three activities are encoded in a multifunctional enzyme called the mitochondrial trifunctional protein, MTP. MTP is composed of eight protein subunits, four α-subunits encoded by the HADHA gene and four β-subunits encoded by the HADHB gene. The α-subunits contain the enoyl-CoA hydratase and long-chain hydroxyacyl-CoA dehydrogenase activities, while the β-subunits possess the 3-ketoacyl-CoA thiolase (β-ketothiolase or just thiolase) activity. The mammalian genome actually encodes five distinct enzymes with thiolase activity.
|Thiolase Gene Symbol||Comments|
|ACAA1||acetyl-CoA acyltransferase 1; also called peroxisomal 3-oxoacyl-CoA thiolase; involved in peroxisomal fatty acid β-oxidation; located on chromosome 3p22.2 spanning 11 kb composed of 12 exons encoding a 424 amino acid protein|
|ACAA2||acetyl-CoA acyltransferase 2; also called mitochondrial 3-oxoacyl-CoA thiolase; catalyzes the terminal reaction of mitochnodrial fatty acid β-oxidation in addition to that catalyzed by HADHB of the MTP; located on chromosome 18q21.1 encoding a 397 amino acid protein|
|ACAT1||acetyl-CoA acetyltransferase 1; also called mitochondrial acetoacetyl-CoA thiolase; involved in ketone body synthesis (see below) in the liver; located on chromosome 11q22.3 spanning 27 kb composed of 12 exons encoding a 427 amino acid protein|
|ACAT2||acetyl-CoA acetyltransferase 2; also called cytosolic acetoacetyl-CoA thiolase; involved in cholesterol biosynthesis and in the utilization of ketone bodies by the brain; located on chromosome 6q25.3|
|HADHB||hydroxyacyl-CoA dehydrogenase/3-ketoacyl-CoA thiolase/enoyl-CoA hydratase, beta subunit; 3-ketoacyl-CoA thiolase; β-ketothiolase; HADHB encodes the β-subunit of mitochondrial trifunctional protein (MTP); located on chromosome 2p23.3 composed of 16 exons|
Each round of β-oxidation produces one mole of FADH2, one mole of NADH, and one mole of acetyl-CoA. The acetyl-CoA, the end product of each round of β-oxidation, then enters the TCA cycle, where it is further oxidized to CO2 with the concomitant generation of three moles of NADH, one mole of FADH2 and one mole of ATP. The NADH and FADH2 generated during the fat oxidation and acetyl-CoA oxidation in the TCA cycle then can enter the respiratory pathway for the production of ATP via oxidative phosphorylation.
The oxidation of fatty acids yields significantly more energy per carbon atom than does the oxidation of carbohydrates. The net result of the oxidation of one mole of oleic acid (an 18-carbon fatty acid) will be 146 moles of ATP (2 mole equivalents are used during the activation of the fatty acid), as compared with 114 moles from an equivalent number of glucose carbon atoms.back to the top
The majority of natural lipids contain an even number of carbon atoms. A small proportion of plant derived lipids contain odd numbers and upon complete β-oxidation these yield acetyl-CoA units plus a single mole of propionyl-CoA. The propionyl-CoA is converted, in an ATP-dependent pathway, to succinyl-CoA. The succinyl-CoA can then enter the TCA cycle for further oxidation.
The oxidation of unsaturated fatty acids is essentially the same process as for saturated fats, except when a double bond is encountered. In such a case, the bond is isomerized by a specific enoyl-CoA isomerase and oxidation continues. In the case of linoleate, the presence of the Δ12 unsaturation results in the formation of a dienoyl-CoA during oxidation. This molecule is the substrate for an additional oxidizing enzyme, the NADPH requiring 2,4-dienoyl-CoA reductase.back to the top
In addition to mitochondrial oxidation of fatty acids, the peroxisomes also play an important role in overall fatty acid metabolism. Very-long-chain fatty acids (VLCFAs: C17–C26) are preferentially oxidized in the peroxisomes with cerotic acid (a 26:0 fatty acid) being solely oxidized in this organelle. The peroxisomes also metabolize di– and trihydroxycholestanoic acids (bile acid intermediates); long-chain dicarboxylic acids that are produced by ω-oxidation of long-chain monocarboxylic acids; pristanic acid via the α-oxidation pathway (see below); certain polyunsaturated fatty acids (PUFAs) such as tetracosahexaenoic acid (24:6), which by β-oxidation yields the important PUFA docosahexaenoic acid (DHA); and certain prostaglandins and leukotrienes.
The enzymatic processes of peroxisomal β-oxidation are very similar to those of mitochondrial β-oxidation with one major difference. During mitochondrial oxidation the first oxidation step, catalzyed by various acyl-CoA dehydrogenases, results in the reduced electron carrier FADH2 that then delivers its' electrons directly to the electron transport chain for synthesis of ATP. In the peroxisome the first oxidation step is catalyzed by acyl-CoA oxidases which is coupled to the reduction of O2 to hydrogen peroxide (H2O2). Thus, the reaction is not coupled to energy production but instead yields a significant reactive oxygen species (ROS). Peroxisomes contain the enzyme catalase that degrades the hydrogen peroxide back to O2.
Humans contain three peroxisomal acyl-CoA oxidases, ACOX1, ACOX2 and ACOX3. Human and rodent ACOX1 (also referred to as palmitoyl-CoA oxidase) is responsible for the oxidation of straight-chain mono- and dicarboxylic fatty acids, very long-chain fatty acids, prostaglandins, and xenobiotics. In humans, in contrast to rodents, 2-methyl branched fatty acids (primarily pristanoic acid) and the bile acid intermediates di- and tri-hydroxycoprostanic acids are desaturated in the peroxisomes by a single enzyme ACOX2 (also called branched-chain acyl-CoA oxidase). The human genome contains an ACOX3 gene but expression from the gene is detected in normal tissue only at extremely low levels. Rodent ACOX3 (also referred to as pristanoyl-CoA oxidase) is the oxidase responsible for oxidation of 2-methy branched chain fatty acids in these animals.
Pathway of peroxisomal β-oxidation. Fatty acids are taken into the peroxisome and esterified to CoA by the solute carrier family member proteins ABCD1, ABCD2, or ABCD3. ABCD1 is also called VLCFA-CoA synthetase. DBP is D-bifunctional protein. The peroxisomal thiolase indicated in the figure is ACAA1 (acetyl-CoA C-acyltransferase 1, also known as peroxisomal 3-oxoacyl-CoA thiolase). Acetyl-CoA generated by peroxisomal β-oxidation is transported out of the peroxisome after exchange of carnitine for the CoA. Peroxisomal and mitochondrial acetyl-carnitine is formed through the action of carnitine acetyltransferase (CAT). Acetyl-carnitine is transported out of the the peroxisomes and mitochondria via the action of carnitine-acylcarnitine translocase (CACT). Once in the cytosol acetyl-carnitine is converted to acetyl-CoA via the action of cytosolic CAT. These acetyl-CoA units can be used for cytosolic fatty acid synthesis or imported into the mitochondria for oxidation in the TCA cycle.
The hydration step and second oxidation step in peroxisomal β-oxidation is carried out by a single bifunctional enzyme as opposed to two separate enzymes as is the case in the mitochondria. There are two distinct bifunctional enzymes identified as L-bifunctional protein (LBP) and D-bifunctional protein (DBP). LBP is specific for L-3-hydroxyacyl-CoAs and DBP is specific for D-3-hydroxyacyl-CoAs. These bifunctional enzymes are also referred to as multifunctional proteins 1 and 2 (MFP-1 and -2) or L- and D-peroxisomal bifunctional enzymes (L-PBE and D-PBE). DBP is the primary, if not exclusive enzyme involved in the oxidation of VLCFAs, pristanic acid, and di- and trihydroxycholestanoic acids. The precise role of LBP in human peroxisomal lipid oxidation is unclear. Human peroxisomes contain the thiolase acetyl-CoA C-acyltransferase 1 (ACAA1) that catalyzes the terminal step in the peroxisoaml β-oxidation pathway.
The clinical significance of the activity of the acyl-CoA oxidases of peroxisomal β-oxidation is related to tissue specific oxidation processes. In the pancreatic β-cell there is little, if any, catalase expressed so that peroxisomal oxidation of VLCFAs results in an increased release of ROS that can damage the β-cell contributing to the progressive insulin deficiency seen in obesity.back to the top
The microsomal (endoplasmic reticulum, ER) pathway of fatty acid ω-oxidation represents a minor pathway of overall fatty acid oxidation. However, in certain pathophysiological states, such as diabetes, chronic alcohol consumption, and starvation, the ω-oxidation pathway may provide an effective means for the elimination of toxic levels of free fatty acids. The pathway refers to the fact that fatty acids first undergo a hydroxylation step at the terminal (omega, ω) carbon. Human ω-hydroxylases are all members of the cytochrome P450 family (CYP) of enzymes. These enzymes are abundant in the liver and kidneys. Specifically, it is members of the CYP4A and CYP4F families that preferentially hydroxylate the terminal methyl group of C10–C26 length fatty acids. CYP4A11 is the human homolog of the rat liver CYP4A1 gene whose encoded enzyme was the first ω-hydroxylase characterized. CYP4A11 utilizes NADPH and O2 to introduce an alcohol to ω-CH3– of several fatty acids including lauric (12:0), myristic (14:0), palmitic (16:0), oleic (18:1) and arachidonic acid (20:4). Following addition of the ω-hydroxyl the fatty acid is a substrate for alcohol dehydrogenase (ADH) which generates an oxo-fatty acid, followed by generation of the corresponding dicarboxylic acid via the action of aldehyde dehydrogenases (ALDH). Further metabolism then takes place via the β-oxidation pathway in peroxisomes.
Pathway of microsomal (omega) ω-oxidation as initiated by CYP4A11.
Another human CYP4A subfamily member has been identified and designated CYP4A22. This protein is highly homologous with CYP4A11 and has been shown to exhibit lauric acid ω-hydroxylase activity. Expression of CYP4A22 is low in all tissue in which it is found. The CYP4A subfamily is not the only CYP4 family of proteins that have been found to possess ω-hydroxylase activity. The CYP4F family enzyme CYP4F3A, which is expressed in leukocytes, is necessary for the ω-hydroxylation and subsequent degradation of leukotriene B4 (LTB4). LTB4 plays an important role in the modulation of inflammatory processes. The CYP4F3 gene is subject to alternative promoter usage and tissue-specific gene splicing, which results in two different proteins being produced. These two enzymes are designated CYP4F3A and CYP4F3B, with the latter enzyme being expressed in the liver. CYP4F3B has higher affinity for arachidonic acid.
Another CYP4F family member, identified as CYP4F2, has been identified that also has LTB4-hydroxylating activity. This CYP4F2 protein has a high degree of homology to the CYP4F3B protein and is expressed in the liver and kidneys. CYP4F2 has been shown to be the major arachidonic acid ω-hydroxylase in human liver and kidney. Indeed, the substrate specificity of CYP4F2 for arachidonic acid is much higher than that of CYP4A11 which was originally described as a signficant arachidonic acid β-hydroxylase. The formation of ω-hydroxylated arachidonic acid (20-hydroxyeicosatetraenoic acid, 20-HETE) by CYP4A11 plays an important role in the regulation of the cardiovascular system because 20-HETE is a known vasoconstrictor. Polymorphisms in the CYP4A11 gene are associated with hypertension in certain population, particular Asian populations. In addition to ω-hydroxylation of arachidonic acid and LTB4, CYP4F2 has been shown to be responsible for the ω-hydroxylation of the phytyl tail of the tocopherols and tocotrienols (collectively known as vitamin E). Metabolism of vitamin E requires an initial ω-hydroxylation step followed by subsequent β-oxidation.
Additional members of the CYP4F subfamily have been identified in humans. These genes are designated CYP4F8, CYP4F11, and CYP4F12. CYP4F8 is present in epithelial linings and catalyzes the (ω-1)-hydroxylation of prostaglandin H2 (PGH2). CYP4F11 is primarily expressed in liver, but also found in kidney, heart, brain and skeletal muscle. The primary endogenous substrates for CYP4F11 are long-chain 3-hydroxydicarboxylic acids (3-OHDCAs) and the enzyme is also very active at hydroxylating various xenobiotics. CYP4F12 is expressed liver, heart, gastrointestinal and urogenital epithelia and its primary substrates are eicosanoids and xenobiotics.back to the top
Phytanic acid is a fatty acid present in the tissues of ruminants and in dairy products and is, therefore, an important dietary component of fatty acid intake. Because phytanic acid is methylated, it cannot act as a substrate for the first enzyme of the mitochondrial β-oxidation pathway (acyl-CoA dehydrogenase). Phytanic acid is first converted to its CoA-ester and then phytanoyl-CoA serves as a substrate in an α-oxidation process. The α-oxidation reaction (as well as the remainder of the reactions of phytanic acid oxidation) occurs within the peroxisomes and requires a specific α-hydroxylase (specifically phytanoyl-CoA hydroxylase, PhyH), which adds a hydroxyl group to the α-carbon of phytanic acid generating the 19-carbon homologue, pristanic acid. Pristanic acid then serves as a substrate for the remainder of the normal process of β-oxidation. Because the first step in phytanic acid oxidation involves an α-oxidation step, the process is termed α-oxidation. For more details on peroxisome function see the Refsum disease page.
In order to understand how the synthesis and degradation of fats needs to be exquisitely regulated, one must consider the energy requirements of the organism as a whole. The blood is the carrier of triglycerides in the form of VLDLs and chylomicrons, fatty acids bound to albumin, amino acids, lactate, ketone bodies and glucose. The pancreas is the primary organ involved in sensing the organism's dietary and energetic states by monitoring glucose concentrations in the blood. Low blood glucose stimulates the secretion of glucagon, whereas, elevated blood glucose calls for the secretion of insulin.
The metabolism of fat is regulated by two distinct mechanisms. One is short-term regulation, which can come about through events such as substrate availability, allosteric effectors and/or enzyme modification. The other mechanism, long-term regulation, is achieved by alteration of the rate of enzyme synthesis and turn-over.
ACC is the rate-limiting (committed) step in fatty acid synthesis. There are two major isoforms of ACC in mammalian tissues. These are identified as ACC1 and ACC2. ACC1 is strictly cytosolic and is enriched in liver, adipose tissue and lactating mammary tissue. ACC2 was originally discovered in rat heart but is also expressed in liver and skeletal muscle. ACC2 has an N-terminal extension that contains a mitochondrial targeting motif and is found associated with CPT-1 allowing for rapid regulation of CPT-1 by the malonyl-CoA produced by ACC. Both isoforms of ACC are allosterically activated by citrate and inhibited by palmitoyl-CoA and other short- and long-chain fatty acyl-CoAs. Citrate triggers the polymerization of ACC1 which leads to significant increases in its activity. Although ACC2 does not undergo significant polymerization (presumably due to its mitochondrial association) it is allosterically activated by citrate. Glutamate and other dicarboxylic acids can also allosterically activate both ACC isoforms.
ACC activity can also be affected by phosphorylation. Both ACC1 and ACC2 contain at least eight sites that undergo phosphorylation. The sites of phosphorylation in ACC2 have not been as extensively studied as those in ACC1. Phosphorylation of ACC1 at three serine residues (S79, S1200, and S1215) by AMPK leads to inhibition of the enzyme. Glucagon-stimulated increases in cAMP and subsequently to increased PKA activity also lead to phosphorylation of ACC. ACC2 is a better substrate for PKA than is ACC1. The activating effects of insulin on ACC are complex and not completely resolved. It is known that insulin leads to the dephosphorylation of the serines in ACC1 that are AMPK targets in the heart enzyme. This insulin-mediated effect has not been observed in hepatocytes or adipose tissues cells. At least a portion of the activating effects of insulin are related to changes in cAMP levels. Early evidence has shown that phosphorylation and activation of ACC occurs via the action of an insulin-activated kinase. However, contradicting evidence indicates that although there is insulin-mediated phosphorylation of ACC this does not result in activation of the enzyme. Activation of α-adrenergic receptors in liver and skeletal muscle cells inhibits ACC activity as a result of phosphorylation by an as yet undetermined kinase.
Insulin, a product of the well-fed state, stimulates ACC and FAS synthesis, whereas starvation leads to a decrease in the synthesis of these enzymes. Adipose tissue levels of lipoprotein lipase also are increased by insulin and decreased by starvation. However, the effects of insulin and starvation on lipoprotein lipase in the heart are just the inverse of those in adipose tissue. This sensitivity allows the heart to absorb any available fatty acids in the blood in order to oxidize them for energy production. Starvation also leads to increases in the levels of cardiac enzymes of fatty acid oxidation, and to decreases in FAS and related enzymes of synthesis.
Adipose tissue contains hormone-sensitive lipase (HSL), which is activated by PKA-dependent phosphorylation; this activation increases the release of fatty acids into the blood. This in turn leads to the increased oxidation of fatty acids in other tissues such as muscle and liver. In the liver, the net result (due to increased acetyl-CoA levels) is the production of ketone bodies (see below). This would occur under conditions in which the carbohydrate stores and gluconeogenic precursors available in the liver are not sufficient to allow increased glucose production. The increased levels of fatty acid that become available in response to glucagon or epinephrine are assured of being completely oxidized, because PKA also phosphorylates ACC; the synthesis of fatty acid is thereby inhibited.
The activity of HSL is also affected via phosphorylation by AMPK. In this case the phosphorylation inhibits the enzyme. Inhibition of HSL by AMPK may seem paradoxical since the release of fatty acids stored in triglycerides would seem necessary to promote the production of ATP via fatty acid oxidation and the major function of AMPK is to shift cells to ATP production from ATP consumption. This paradigm can be explained if one considers that if the fatty acids that are released from triglycerides are not consumed they will be recycled back into triglycerides at the expense of ATP consumption. Thus, it has been proposed that inhibition of HSL by AMPK mediated-phosphorylation is a mechanism to ensure that the rate of fatty acid release does not exceed the rate at which they are utilized either by export or oxidation.
Insulin has the opposite effect to glucagon and epinephrine: it increases the synthesis of triacylglycerols (and glycogen). One of the many effects of insulin is to lower cAMP levels, which leads to increased dephosphorylation through the enhanced activity of protein phosphatases such as PP-1. With respect to fatty acid metabolism, this yields dephosphorylated and inactive hormone-sensitive lipase. Insulin also stimulates certain phosphorylation events. This occurs through activation of several cAMP-independent kinases.
Fat metabolism can also be regulated by malonyl-CoA-mediated inhibition of CPT I. Such regulation serves to prevent de novo synthesized fatty acids from entering the mitochondria and being oxidized.back to the top
The glucose-fatty acid cycle describes interrelationships of glucose and fatty acid oxidation as defined by fuel flux and fuel selection by various organs. This cycle is not a metabolic cycle such as can be defined by the TCA cycle as an example, but defines the dynamic interactions between these two major energy substrate pools. The glucose-fatty acid cycle was first proposed by Philip Randle and co-workers in 1963 and is, therefore, sometimes referred to as the Randle cycle or Randle hypothesis. The cycle describes how nutrients in the diet can fine-tune metabolic processes on top of the more coarse control exerted by various peptide and steroid hormones. The underlying theme of the glucose-fatty acid cycle is that the utilization of one nutrient (e.g. glucose) directly inhibits the use of the other (in this case fatty acids) without hormonal mediation. The general interrelationships between glucose and fatty acid utilization in skeletal muscle and adipose tissue that constitutes the glucose-fatty acid cycle are diagrammed in the Figure below.
The glucose-fatty acid cycle represents the interactions between glucose uptake and metabolism and the consequent inhibition of fatty acid oxidation and the effects of fatty acid oxidation on the inhibition of glucose utilization. The reciprocal regulation is most prevalent in skeletal muscle and adipose tissue. When glucose levels are high it is taken into cells via the GLUT4 transporter and phosphorylated by hexokinase. The reactions of glycolysis drive the carbon atoms to pyruvate where they are oxidized to acetyl-CoA. The fate of the acetyl-CoA is complete oxidation in the TCA cycle or return to the cytosol via citrate for conversion back to acetyl-CoA via ATP-citrate lyase (ACL) and then into into malonyl-CoA and subsequent long-chain fatty acid (LCFA) synthesis. The synthesis of malonyl-CoA is catalyzed by acetyl-CoA carboxylase (ACC) and once produced will inhibit the import of long-chain fatty acyl-CoAs (LCFacyl-CoA) into the mitochondria via inhibition of carnitine palmitoyltransferase 1 (CPT-1). This effectively blocks the oxidation of fatty acids leading to increased triacylglyceride synthesis (TAG). The equilibrium between malonyl-CoA synthesis and breakdown back to acetyl-CoA is determined by the regulation of ACC and malonyl-CoA decarboxylase (MCD). As long as there is sufficient capacity to divert glucose carbons to TCA cycle oxidation and fatty acid synthesis there will be limited acetyl-CoA mediated inhibition of the pyruvate dehydrogenase complex (PDHc). On the other hand, when fatty acid levels are high they enter the cell via one of several fatty acid transporter complexes [fatty acid translocase (FAT)/CD36 is shown since this transporter has a preference for LCFAs], and are then transported into the mitochondria to be oxidized. The large increase in fatty acid oxidation subsequently inhibits the utilization of glucose. This is the result of increased cytosolic citrate production from acetyl-CoA and the inhibition of phosphofructokinase-1 (PFK1). The increased acetyl-CoA derived from fat oxidation will in turn further inhibit glucose utilization via activation of PDH kinases (PDKs) that will phosphorylate and inhibit the PDHc. Although not shown, PDKs are also activated by increased mitochondrial NADH/NAD+ ratios in response to increased fatty acid β-oxidation. Under conditions where fat oxidation is favored ACC will be inhibited and MCD will be activated ensuring that LCFA that enter the cell will be able to be transported into the mitochondria. PS is pyruvate symporter responsible for mitochondrial uptake of pyruvate. TCAT is tricarboxylic acid transporter.
How do the dynamics of the glucose-fatty acid cycle play out under various physiological conditions and changing fuel substrate pools? In the fasted state it is imperative that glucose be spared so that the brain can have adequate access to this vital fuel. Under these conditions, hormonal signals from the pancreas, in the form of glucagon, stimulate adipose tissue lipolysis releasing free fatty acids (FFAs) to the blood for use as a fuel by other peripheral tissues. When the released FFAs enter the liver they oxidized and also serve as substrates for ketogenesis. The oxidation of fatty acids inhibits glucose oxidation as outlined in the above figure. In addition to sparing glucose for the brain, fatty acid oxidation also preserves pyruvate and lactate which are important gluconeogenesis substrates. The effects of fatty acids on glucose utilization can also be observed in the well fed state after a high fat meal and during periods of exercise.
As outlined in the above Figure, the inhibition of glucose utilization by fatty acid oxidation is mediated by short-term effects on several steps of overall glycolysis that include glucose uptake, glucose phosphorylation and pyruvate oxidation. During fatty acid oxidation the resultant acetyl-CoA allosterically activates PDKs that phosphorylate and inhibit the PDHc. PDKs are also activated by increasing levels of NADH that will be the result of increased fatty acid oxidation. Thus, two products of fat oxidation result in inhibition of the PDHc. In addition, excess acetyl-CoA is transported to the cytosol either as citrate (as diagrammed) or as acetyl-carnitine. Mitochondrial acetyl-carnitine is formed through the action of carnitine acetyltransferase (CAT). Acetyl-carnitine is transported out of the the mitochondria via the action of carnitine-acylcarnitine translocase (CACT). Once in the cytosol acetyl-carnitine is converted to acetyl-CoA via the action of cytosolic CAT. In the cytosol, citrate serves as an allosteric inhibitor of PFK1 thus limiting entry of glucose into glycolysis. The increase in glucose-6-phosphate that results from inhibition of PFK1 leads to feed-back inhibition of hexokinase which in turn limits glucose uptake via GLUT4. Additional mechanisms of fatty acid metabolism that lead to interference in glucose uptake and utilization are the result of impaired insulin receptor signaling. These latter processes are discussed in detail in the Insulin Function page.
Mechanisms by which glucose utilization inhibits fatty acid oxidation are tissue specific due primarily to the differences in Km of hepatic glucokinase and skeletal muscle and adipose tissue hexokinase. In addition, hepatic CPT-1 is approximately 100-fold less sensitive to inhibition by malonyl-CoA than are the skeletal muscle and cardiac isoforms. When glucose is oxidized in glycolysis the resultant pyruvate enters the mitochondria via the pyruvate symporter. Increasing mitochondrial pyruvate inhibits the PDKs allowing for rapid decarboxylation of pyruvate by the PDHc ensuring continued entry of glucose into the glycolytic stream. Some of the acetyl-CoA derived from pyruvate oxidation will be diverted from the TCA cycle as citrate and transported to the cytosol by the tricarboxylic acid transporter (TCAT). The citrate is converted to acetyl-CoA and oxaloacetate by ATP-citrate lyase (ACL) and can now serve as a substrate for ACC. The resultant malonyl-CoA will inhibit CPT-1 thus, restricting mitochondrial uptake and oxidation of fatty acyl-CoAs. The inhibition of fatty acid oxidation in the liver re-routes LCFAs into triglycerides (TAGs). Long term effects of excess glucose are reflected in hepatic steatosis resulting from the diversion of fats into TAGs instead of being oxidized.
In addition to being regulated by intermediates of glucose and fat oxidation, several enzymes in these two pathways are regulated at the level of post-translational modification and/or gene expression. Most of these regulatory schemes have been covered in the above sections.back to the top
The majority of clinical problems related to fatty acid metabolism are associated with processes of oxidation. These disorders fall into four main groups:
1. Deficiencies in Carnitine: Deficiencies in carnitine lead to an inability to transport fatty acids into the mitochondria for oxidation. This can occur in newborns and particularly in pre-term infants. Carnitine deficiencies also are found in patients undergoing hemodialysis or exhibiting organic aciduria. Carnitine deficiencies may manifest systemic symptomology or may be limited to only muscles. Symptoms can range from mild occasional muscle cramping to severe weakness or even death. Treatment is by oral carnitine administration.
2. Carnitine palmitoyltransferase deficiencies: Deficiencies in CPT-1 are relatively rare and affect primarily the liver and lead to reduced fatty acid oxidation and ketogenesis. The most common symptom associated with CPT-1 deficiency is hypoketotic hypoglycemia. There is also an elevation in blood levels of carnitine. The liver involvement results in hepatomegaly and in muscles results in weakness. CPT-2 deficiencies can be classified into three main forms. The adult form affects primarily the skeletal muscles and is called the adult myopathic form. This form of the disease causes muscle pain and fatigue and myoglobinuria following exercise. The severe infantile multisystem form manifest in the first 6–24 months of life with most afflicted infants demonstrating significant involvement before 1 year. The primary symptom of this form of CPT-2 deficiency is hypoketotic hypoglycemia. Symptoms will progress to severe hepatomegaly and cardiomyopathy. Often times death from CPT-2 deficiency may be mis-diagnosed as sudden infant death syndrome, SIDS. The rarest form of CPT-2 deficiency is referred to as the neonatal lethal form. Symptoms of this form appear within hours to 4 days after birth and include respiratory failure, hepatomegaly, seizures, hypoglycemia, and cardiomegaly. The cardiomegaly will lead to fatal arrhythmias. Carnitine acyltransferases may also be inhibited by sulfonylurea drugs such as tolbutamide and glyburide.
3. Deficiencies in Acyl-CoA Dehydrogenases: A group of inherited diseases that impair β-oxidation result from deficiencies in acyl-CoA dehydrogenases. The enzymes affected may belong to one of three categories:
long-chain acyl-CoA dehydrogenase (LCAD)
medium-chain acyl-CoA dehydrogenase (MCAD)
short-chain acyl-CoA dehydrogenase (SCAD)
MCAD deficiency is the most common form of acyl-CoA dehydrogenase deficiency. In the first years of life this deficiency will become apparent following a prolonged fasting period. Symptoms include vomiting, lethargy and frequently coma. Excessive urinary excretion of medium-chain dicarboxylic acids as well as their glycine and carnitine esters is diagnostic of this condition. In the case of this enzyme deficiency taking care to avoid prolonged fasting is sufficient to prevent clinical problems.
4. Refsum Disease: Refsum disease is a rare inherited disorder in which patients harbor a defect in the peroxisomal α-oxidizing enzyme, phytanoyl-CoA hydroxylase (PhyH). Although mutations in PhyH are the primary cause of Refsum disease, the syndrome can also result from defects in the peroxisomal protein (PEX7) responsible for the import of PhyH into the peroxisome. Patients accumulate large quantities of phytanic acid in their tissues and serum. This leads to severe symptoms, including cerebellar ataxia, retinitis pigmentosa, nerve deafness and peripheral neuropathy. As expected, the restriction of dairy products and ruminant meat from the diet can ameliorate the symptoms of this disease. It should be noted that accumulation of phytanic acid is not solely the result of defects in PhyH. Phytanic acid accumulation is also seen when there are inherited defects in peroxisome function leading to Zellweger syndrome, neonatal adrenoleukodystrophy and infantile Refsum disease. In addition, rhizomelic chondrodysplasia punctata, type 1 (RCDP1) results in phytanic acid accumulation. Refsum disease due to deficiency in PhyH is properly referred to as classical Refsum disease to distinguish it from infantile Refsum due to peroxisome dysfunction.back to the top
During high rates of fatty acid oxidation, primarily in the liver, large amounts of acetyl-CoA are generated. These exceed the capacity of the TCA cycle, and one result is the synthesis of ketone bodies. The synthesis of the ketone bodies (ketogenesis) occurs in the mitochondria allowing this process to be intimately coupled to rate of hepatic fatty acid oxidation. Conversely, the utilization of the ketones (ketolysis) occurs in the cytosol. The ketone bodies are acetoacetate, β-hydroxybutyrate, and acetone.
The formation of acetoacetyl-CoA occurs by condensation of two moles of acetyl-CoA. This reaction is essentially a reversal of the thiolase (HADHB or ACAA2) catalyzed reaction of β-oxidation but is in fact catalyzed by the mitochondrial enzyme acetoacetyl-CoA thiolase (encoded by the ACAT1 gene). Acetoacetyl-CoA and an additional acetyl-CoA are converted to β-hydroxy-β-methylglutaryl-CoA (HMG-CoA) by mitochondrial HMG-CoA synthase (encoded by the HMGCS2 gene), an enzyme found in large amounts only in the liver. HMG-CoA in the mitochondria is converted to acetoacetate by the action of HMG-CoA lyase. Acetoacetate can undergo spontaneous decarboxylation to acetone, or be enzymatically converted to β-hydroxybutyrate through the action of β-hydroxybutyrate dehydrogenase. The ketone bodies freely diffuse out of the mitochondria and hepatocytes and enter the circulation where they can be taken up by non-hepatic tissues such as the brain, heart, and skeletal muscle.
When the level of glycogen in the liver is high the production of β-hydroxybutyrate increases. When carbohydrate utilization is low or deficient, the level of oxaloacetate will also be low, resulting in a reduced flux through the TCA cycle. This in turn leads to increased release of ketone bodies from the liver for use as fuel by other tissues. In early stages of starvation, when the last remnants of fat are oxidized, heart and skeletal muscle will consume primarily ketone bodies to preserve glucose for use by the brain. Acetoacetate and β-hydroxybutyrate, in particular, also serve as major substrates for the biosynthesis of neonatal cerebral lipids.
Ketone bodies are utilized by extrahepatic tissues via a series of cytosolic reactions that are essentially a reversal of ketone body synthesis. The initial steps involve the conversion of β-hydroxybutyrate to acetoacetate and of acetoacetate to acetoacetyl-CoA. The first step involves the reversal of the β-hydroxybutyrate dehydrogenase reaction. It is important to appreciate that under conditions where tissues are utilizing ketones for energy production their NAD+/NADH ratios are going to be relatively high, thus driving the β-hydroxybutyrate dehydrogenase catalyzed reaction in the direction of acetoacetate synthesis. The second reaction of ketolysis involves the action (shown below) of succinyl-CoA:3-oxoacid-CoA transferase (SCOT), also called 3-oxoacid-CoA transferase 1 (OXCT1). The latter enzyme is present at high levels in most tissues except the liver. Importantly, very low level of SCOT expression in the liver allows the liver to produce ketone bodies but not to utilize them. This ensures that extrahepatic tissues have access to ketone bodies as a fuel source during prolonged fasting and starvation.
The fate of the products of fatty acid metabolism is determined by an individual's dietary and physiological status. The overall rate of hepatic ketogenesis may by affected by several factors:
1. Control in the release of free fatty acids from adipose tissue directly affects the level of ketogenesis in the liver. This is, of course, substrate-level regulation. Fatty acid release from adipose tissue is controlled via the activity of hormone-sensitive lipase (HSL). When glucose levels fall, pancreatic glucagon secretion increases resulting in phosphorylation of adipose tissue HSL, thus resulting in increased hepatic ketogenesis due to increased substrate (free fatty acids) delivery from adipose tissue. Conversely, insulin, released in the well-fed state, inhibits ketogenesis via the triggering of dephosphorylation and inactivation of adipose tissue HSL.
2. Once fats enter the liver, they have two distinct fates. They may be activated to acyl-CoAs and oxidized, or esterified to glycerol in the production of triacylglycerols. If the liver has sufficient supplies of glycerol-3-phosphate, most of the fats will be turned to the production of triacylglycerols.
3. The acetyl-CoA generated by the oxidation of fats can be completely oxidized in the TCA cycle or it can be diverted into lipid biosynthesis. If the hepatic demand for ATP is high the fate of acetyl-CoA is likely to be further oxidation to CO2. This is especially true under conditions of hepatic stimulation by glucagon which results in increased gluconeogenesis and the energy for this process is derived primarily from the oxidation of fatty acids supplied from adipose tissue.
4. In addition, glucagon results in phosphorylation and inhibition of acetyl-CoA carboxylase (ACC), the rate limiting enzyme of de novo fatty acid synthesis. Conversely, under conditions of insulin release, hepatic ACC is activated and the excess acetyl-CoA will be converted into malonyl-CoA and then free fatty acids. The increased malonyl-CoA results in inhibition of fatty acid transport into the mitochondria resulting in reduced fat oxidation and reduced production of excess acetyl-CoA.back to the top
The production of ketone bodies occurs at a relatively low rate during normal feeding and under conditions of normal physiological status. Normal physiological responses to carbohydrate shortages cause the liver to increase the production of ketone bodies from the acetyl-CoA generated from fatty acid oxidation. This allows the heart and skeletal muscles primarily to use ketone bodies for energy, thereby preserving the limited glucose for use by the brain.
The most significant disruption in the level of ketosis, leading to profound clinical manifestations, occurs in untreated insulin-dependent diabetes mellitus. This physiological state, diabetic ketoacidosis (DKA) results from a reduced supply of glucose (due to a significant decline in circulating insulin) and a concomitant increase in fatty acid oxidation (due to a concomitant increase in circulating glucagon). The increased production of acetyl-CoA leads to ketone body production that exceeds the ability of peripheral tissues to oxidize them. Ketone bodies are relatively strong acids (pKa around 3.5), and their increase lowers the pH of the blood. This acidification of the blood is dangerous chiefly because it impairs the ability of hemoglobin to bind oxygen.back to the top