Hepatic Lipid Metabolism
Jiansheng Huang, Jayme Borensztajn, and Janardan K. Reddy
Introduction
The liver is a major regulator of lipid metabolism in the body. It plays a central role in the synthesis and degradation (oxidation) of fatty acids. Fatty acids serve as an important source of energy as well as energy storage for many organ-isms and are also pivotal for a variety of biological processes, including the synthesis of cellular membrane lipids and gen-eration of lipid-containing messengers involved in signal transduction [1]. Fatty acids can generally be stored effi-ciently as non-toxic triglycerides (triacylglycerols/fat), which generate more than twice as much energy, for the same mass, as do carbohydrates or proteins. Accordingly, liver is a key player in energy homeostasis: first, as it converts excess dietary glucose into fatty acids that are then exported to other tissues for storage as triglycerides as lipid droplets [2]; second, under conditions of increase in synthesis and decreased oxidation of fatty acids the liver contributes to the progressive accumulation of excess unspent energy in the form of energy-dense triglycerides in adipocytes of adipose tissue, which provide virtually limitless capacity to store energy and finally, under chronic energy over-load situations the liver may serve as a surrogate reservoir for storing consid-erable quantities of excess fat, leading to the development of hepatic steatosis and steatohepatitis [3]. For molecular patho-genesis of fatty liver, please see Chap. 29. Independent chapters are also included on non-alcoholic fatty liver dis-ease (NAFLD) (Chap. 34) and alcoholic liver disease (Chap. 35). This ability of the liver to store lipids is viewed as a protective mechanism, neutralizing the potential toxicity of long-chain fatty acids [4].
In addition to synthesis, oxidation, and secretion of fatty acids for transport and storage in extrahepatic tissues, the liver also functions in the maintenance of plasma lipid levels, a) through its ability to assemble and secrete lipoproteins into
the circulation and b) through its central role in the removal of lipoproteins from circulation [5]. Accordingly, an understand-ing of fat metabolism in liver is important for delineating the pathophysiological implications of altered energy balance and in developing pharmacological strategies for preventive and therapeutic approaches. The emphasis of this chapter is on the sources and synthesis of fatty acids, very low density lipopro-tein assembly and secretion, and fatty acid oxidation.
Sources and Synthesis of Fatty Acids
Fatty acids utilized by the liver for energy generation, for storage as triglycerides, and incorporation into lipoproteins, are generally derived from: (a) hepatic uptake of plasma non-esterified fatty acids (NEFAs) transported in the circulation after release by adipose tissue, as well as after hydrolysis of circulating triglyceride-rich lipoproteins; (b) breakdown of triglycerides of chylomicron remnants taken up by hepato-cytes; (c) hepatic cytoplasmic triglyceride stores that mani-fest as lipid droplets, big or small, in all cell types; and (d) synthesis in situ (lipogenesis).
Hepatic Uptake of Plasma NEFAs Released by Adipose Tissue and Hydrolysis of Circulating Triglyceride-Rich Lipoproteins
Fatty acids deposited as triglycerides in white adipose tissue represent the primary energy store in animals. Under condi-tions of caloric deficit (e.g., starvation) or increased energy demand, triglycerides are hydrolyzed and free fatty acids are released into the circulation. Thus, in these situations, most of the circulating fatty acids taken up by the liver are mobi-lized from adipose tissue triglycerides. The hydrolysis of triglyceride is catalyzed by adipose tissue lipases in sequen-tial steps leading to the formation of NEFAs and glycerol. Adipose triglyceride lipase (ATGL) and hormone-sensitive
133
J.K. Reddy (*)
Department of Pathology, Northwestern University, Feinberg School of Medicine, Chicago, IL, USA e-mail: jkreddy@northwestern.edu
S.P.S. Monga (ed.), Molecular Pathology of Liver Diseases, Molecular Pathology Library 5, DOI 10.1007/978-1-4419-7107-4_10, © Springer Science+Business Media, LLC 2011
134J. Huang et al.
as a central site of regulation of lipolysis (see below). The mechanism by which HSL attacks the lipid droplets is not yet firmly established, but may be involved in the activation of a lipid binding domain on HSL and translocation from the cytosol by a carrier protein [11].
Although circulating fatty acids derived from adipose tis-sue stores are important during starvation as well as during increased energy demand states, plasma fatty acids taken up by the liver in the postprandial state are derived mainly from the catalytic action of lipoprotein lipase (LPL). This enzyme is present on the endothelial surface of capillaries of skeletal lipase (HSL) are the major triglyceride lipases involved in this process (Fig. 10.1). ATGL specifically performs the first step in lipolysis generating diacylglycerol and fatty acid [6], and subsequent action of HSL most efficiently hydrolyses diacylglycerol, further releasing fatty acids. Together, ATGL and HSL are responsible for more than 95% of the triglycer-ide hydrolase activity present in white adipose tissue [7]. Also involved in the mobilization of fatty acids from stored sites are members of perilipin family [8]. The initial hydroly-sis of triglyceride to diacylglycerol is known to be the rate-limiting step, since hydrolysis of diacylglycerol is tenfold faster. The released free fatty acids become bound to circu-lating albumin and are carried to the liver, as well as other tissues, for uptake and utilization. Fatty acid mobilization from fat is tightly controlled by hormones such as insulin, glucagon, epinephrine, and adrenocorticotropic hormone. Mice deficient in HSL secrete less insulin and are glucose intolerant and reveal adipocyte size increase and decrease plasma free fatty acids, mostly due to the attenuation of lipolysis [9, 10]. Of interest is that the surface of the lipid storage droplet coated with a variety of proteins has emerged
Fig. 10.1 Adipose tissue lipolysis during fasting, fatty acid influx into
liver and PPARa sensing. Under conditions of caloric deficit such as
fasting, or increased energy demand, trigylcerides (TG) in adipose tis-sue stores are hydrolyzed and free fatty acids (FFAs) are released into the circulation. The hydrolysis of TG is catalyzed by adipose triglyc-eride lipase (ATGL) and hormone sensitive lipase (HSL). Nuclear receptor PPARa in liver, senses the influx of FFAs and upregulates all three fatty acid oxidation systems, reduces fatty acid esterification to TGs and minimizes hepatic steatosis (see wild type mouse liver on the left). If PPARa sensing is deficient as in PPARa/ACOX1 double knockout mouse liver (right), fatty acid oxidation systems are not upregulated to burn the influxed FFAs. This results in increased TG synthesis contri-buting to the development of hepatic steatosis. Modified from Yu et al. [91] with permission
and cardiac muscles and in adipose tissue and is responsible for the hydrolysis of circulating triglyceride-rich lipoproteins synthesized and secreted by the intestine (chylomicrons) or liver (very low density lipoproteins/VLDL) [12]. The action of LPL-mediated lipolysis of chylomicrons requires apoC-II as a cofactor acquired from plasma high density lipoproteins (HDLs) [12]. Most of the fatty acids generated from the LPL-mediated hydrolysis of chylomicrons and VLDL trig-lycerides are taken up by the tissues where the enzyme is located to be either oxidized (muscles) or stored (adipose tis-sue) [13]. However, a considerable fraction of the generated unesterified fatty acids become bound to plasma albumin and, in this form, are transported to the liver where they are taken up [13, 14]. It has been estimated that >50% of long chain fatty acids bound to albumin can dissociate and bind to liver cells in single pass through the liver [15]. The six-mem-ber family of fatty acid transport proteins (FATP-1 through FATP-6) present as integral transmembrane proteins facili-tates the uptake of long chain and very long chain fatty acids into cells [14, 16]. These proteins exhibit fatty acid syn-thetase activity implying that fatty acids are rapidly con-verted to acyl-CoAs in the cell after translocation across the plasma membrane. The mechanism whereby the fatty acids enter the hepatocytes has not yet been fully elucidated, but it appears that FATP5 plays a major role in hepatocellular uptake of fatty acids [17]. Furthermore, hepatocytes con-tain cytoplasmic fatty acid binding protein (FABP1) and fatty acid translocase (FAT/CD36) are also known to be involved in the cellular uptake of fatty acids [18]. Whether fatty acids enter into hepatocytes by passive diffusion through the plasma membrane or the entry is facilitated
exclusively by fatty acid membrane protein transporters remains unclear [19].
Chylomicron Remnants
As indicated above, a major source of fatty acids for the liver in the postprandial state are triglycerides associated with chylomicrons. These lipoproteins are synthesized in the intestine and are responsible for the transport of most of the
10 Hepatic Lipid Metabolism135
absorbed dietary fatty acids and cholesterol. However, the chylomicrons themselves do not contribute to the hepatic fatty acid pool because they are not cleared from circulation by the liver. Instead, chylomicrons, as well the triglyceride-rich VLDL, are acted upon by LPL in the peripheral vascular bed, which hydrolyzes the triglycerides in the core of the particles generating unesterified fatty acids, some of which bind to plasma albumin for transport to the liver [12]. As a result of LPL action, chylomicrons are converted into rem-nant particles, which are smaller than their parent chylomi-crons but retain most of their original cholesterol content as well as a considerable amount of triglycerides. Unlike the perilpin-2 (adipocyte differentiation-related protein/ADRP/adipophilin), perilipin-3 (tail-interacting protein of 47 kDa/TIP47), perilpin-4 (S3–12), and perilipin-5 (oxidative tissue-enriched/OXPAT) [27]. These five members share sequence similarity and are differentially expressed in different cell types [28]. These proteins are amphiphilic proteins that are associated with the phospholipid monolayer surrounding lipid droplets and participate in lipid droplet maturation and metabolism [29]. Perilipin-2 and -3 are expressed in most cell types. Although perilipin-4 (S3–12) is expressed mostly in adipocytes and to a lesser extent in heart and skeletal muscles, it can be induced in liver cells in response chylomicrons, remnants are readily cleared from circulation by the liver where they are disassembled, thus providing fatty acids to hepatocytes. The efficient uptake of chylomi-cron remnants – but not intact chylomicrons – by the liver is the result of a complex and as yet not a fully elucidated pro-cess that involves the interaction of apoprotein E on the sur-face of the particle first with glycosaminoglycans in the space of Disse, followed by its binding to the low density lipoprotein receptor (LDLR) on the surface of hepatocytes and finally endocytosis of the particle [20].
Hepatic Cytoplasmic Lipid Droplet Stores
All eukaryotes, from yeast to humans, synthesize triglycer-ides and store this excess energy in the form of cytoplasmic lipid droplets for use when needed [21, 22]. Lipid droplets, once considered inert or static, are emerging as metaboli-cally dynamic structures. During times of energy scarcity, this stored energy from lipid droplets is retrieved by the action of lipases [23]. Although the lipid droplet-rich adi-pocytes of adipose tissue are the principal sites of energy storage and retrieval, all other cell types in animals, includ-ing hepatocytes, can accommodate under normal physio-logical conditions, limited quantities of energy in the form of smaller, less conspicuous, lipid droplets. These smaller lipid droplets provide immediate energy source for hepato-cytes and other non-adipocyte cells, and serve as fatty acid source for utilization in intracellular signaling cascades [24]. However, in steatotic states, hepatocytes can accumu-late massive amounts of energy-rich macro- or micro-vesic-ular lipid droplets and lead to the development of fatty liver disease [25].
The entry and sequestration of lipid in lipid droplets as well as lipolysis to release the packaged fatty acids are physiologi-cal processes regulated by evolutionarily conserved families of lipid-droplet surface proteins, including the members of the perilipin (perilipin amino-terminal/PAT proteins) and Cide (cell-death-inducing DFFA-like effector) families [26]. The perilipin family includes five members: Perilpin-1 (perilipin),
to PPARg overexpression reflecting its function in hepatic adiposis [30]. Perilipin-5 is expressed in tissues with high fatty acid oxidation capability including the liver [31]. Fasting and PPARa activation are known to induce per-ilipin-5 expression in liver, consistent with its role in fatty acid oxidation [32–35] (Fig. 10.1). It appears that per-ilipin-4 and perilipin-5 are reciprocal in function, in that the former is associated with lipid storage and the latter with fatty acid oxidation [31, 36]. Perilipins decorate lipid droplets either constitutively (perilipins 1 and 2), or in response to lipolytic challenge (perilipins 3–5) [37]. Because of this differential presence, lipid droplets may be heterogeneous in a given cell with reference to perilipin composition [38]. Evidence suggests that perilipin-1 and perilpin-2 serve as physical barriers to lipolytic enzymes under basal conditions, but in response to lipolytic stimula-tion, these two proteins can also facilitate interactions with lipases [39]. Perilipin-1 is more effective at attenuating lipid droplet lipolysis than perilipin-2 [40]. Therefore, absence of perilipin-1 or perilipin-2 reduces the amount of triglyceride in adipose tissue and liver, possibly due to increased lipolysis resulting from the removal of barrier to lipase action [41]. The relative contributions of various members of perilipin family in lipid droplet composition, assembly, and hydrolysis of triglycerides in the progression of fatty liver disease remain to be clarified [42].
In recent years, the importance of the three members of the Cide family of lipid droplet proteins CideA, CideB, and CideC (also known as Fsp27), in lipid droplet metabolism is being increasingly recognized [43]. Similar to perilipins, Cide proteins are also associated physically with lipid drop-lets and appear to modulate droplet size and lipid metabo-lism in liver during lipid overload conditions [44]. The expression of perilipin-1, CideA, and CideC is markedly elevated in liver with severe steatosis [31, 45]. CideC expres-sion is elevated in the liver following PPARg overexpression [46]. Mice deficient in perilipin-2, CideB, or CideC do not develop fatty liver disease implying that these proteins nor-mally function to facilitate lipid storage in the cell [47]. Although the lipid droplet biology is gaining enhanced atten-tion, the state of knowledge is rudimentary since very little
136J. Huang et al.
reactions performed by enzymes located in the endoplasmic reticulum [62]. Palmitoyl-CoA is elongated by type III fatty acid synthetases, now known as elongases (elongation of very long chain fatty acids; ELOVLs) [63, ]. Seven ELOVL enzymes (ELOVL1–7) are known in mammals, and they reveal different fatty acid substrate preferences for catalyz-ing the elongation reaction []. ELOVL1 and ELOVL3 exhibit high activity toward all of the saturated C20- to C26-CoAs, while ELOVL2 elongates C20 and C22 PUFAs [65]. Fatty acids with chain lengths longer than C26 are elongated by ELOVL4 [65]. ELOVL5 has been shown to be responsi-De Novo Lipogenesisble for the elongation of C18 substrates and ELOVL6 par-ticipates in the elongation of C12–C16 fatty acids [65]. Lipogenesis encompasses fatty acid synthesis and their utili-ELOVL7 exhibits significant activities to C18-CoAs and zation for phospholipid and triglyceride generation. The less to C16:0-CoA. ELOVL1 and ELOVL7 are expressed human body is able to synthesize all fatty acids with the pos-in many tissues, suggesting that ELOVL7 elongates sible exception of two polyunsaturated fatty acids, namely C18:0-CoA to C20:0-CoA, which is then transferred to linoleic acid (C18:2) and a-linolenic acid (C18:3). Liver and ELOVL1 [63, ].adipose tissue are the major sites of fatty acid synthesis and mammary glands also generate fatty acids during lactation. Lipogenesis requires acetyl-CoA precursors that are gener-ated during certain metabolic processes as these precursors
Very Low Density Lipoprotein Assembly
provide all the carbon atoms necessary for fatty acid synthe-and Secretionsis [53]. Liver is the principal organ responsible for the con-version of excess carbohydrate (glucose), beyond organism's
energy needs, to fatty acids via a series of metabolic steps Fatty acids taken up by the liver and which do not undergo that are regulated by several factors, including nutritional, oxidation, are mainly esterified and collected in a common hormonal, and genetic elements [54]. Glucose is first con-cytosolic triglyceride pool [66]. Conditions in which the verted to pyruvate, which enters the Krebs cycle in the mito-delivery of fatty acids to the liver is increased may be associ-chondria to yield citrate [55]. Citrate is then transported into ated with increased hepatic triglyceride content. However, the cytosol and broken down by ATP citrate lyase to yield under normal conditions, triglycerides do not accumulate in acetyl-CoA and oxaloacetate. Acetyl-CoA is converted to the cytoplasm of hepatocytes but are mobilized and, together malonyl-CoA, the rate-limiting step in the lipogenesis path-with cholesterol and phospholipids, are assembled into way catalyzed mainly by acetyl-CoA carboxylases (ACC1 VLDL particles for secretion into the circulation. The secre-and ACC2) [56]. Successive molecules of malonyl-CoA, tion of VLDL into the blood is crucial in preventing the which serves as a two carbon donor, are added to the acetyl-accumulation of triglycerides by liver cells that may lead to CoA primer by a multifunctional enzyme complex, the fatty steatosis and its resulting pathologic consequences.
The synthesis of VLDL requires the availability of apoli-acid synthase (FAS) [57]. Palmitic acid (C16:0) is the pre-dominant fatty acid generated by FAS [58]. FAS is expressed poprotein B (apoB), a complex protein that serves as the in the liver and adipose tissue, but in the humans, the liver scaffolding upon which the VLDL is assembled. ApoB appears to be the major site for de novo lipogenesis [59]. occurs in two forms, apoB-100 and apoB-48 [67]. ApoB-100 Palmitic acid is desaturated by stearoyl-CoA desaturase-1 is essential for the assembly of VLDL in the liver, and apoB-(SCD-1) to palmitoleic acid or elongated to yield stearic acid 48 is essential for the assembly of chylomicrons in the intes-(C18:0). SCD-1 catalyzes the conversion of stearoyl-CoA to tine. Both apoBs are the products of a single gene. ApoB-100 oleoyl-CoA, which is a major substrate for triglyceride syn-is synthesized as a 4536-amino acid polypeptide and apoB-thesis [60]. Oleic acid (C18:1) is formed as a result of desatu-48 – a truncated form containing 48% of the protein from the ration of stearic acid and is regarded as the end product of N-terminus – is synthesized as a result of apoB mRNA de novo fatty acid synthesis. However, the saturated C16 fatty editing, a process that requires a multi-component enzyme acid, first synthesized during de novo lipogenesis, is the essen-complex containing an RNA-specific cytidine deaminase, tial precursor for almost all the newly synthesized fatty acids apobec-1, and an RNA-binding subunit, apobec-1 comple-mentation factor (ACF) [68]. In humans, the RNA editing including the formation of very long chain fatty acids [61].
Very long chain (>C22) saturated, monounsaturated, and occurs in the intestine but not in the liver. In rodents, apoB-48 PUFAs are synthesized by elongation and desaturation is also synthesized in the liver and secreted into the circulation
is known about the ~100 lipid droplet associated proteins [48]. It is noteworthy that the storage of lipid in the liver may also protect the cells against lipotoxicity [49–51]. The pack-aged lipids in droplets may also be important in transporta-tion of lipid to specific cellular destinations or for specific functions. Moreover, the lipid droplets may provide a shelter for some special proteins when they are at a higher level in the cells [22, 52].
10 Hepatic Lipid Metabolism肝脏在脂肪酸氧化产生能量和酮体合成基质在空腹的条件下被肝外组织利用中起着核心的作用137associated with VLDL. Thus, human VLDLs carry only apoB-100 whereas rodent VLDL particles carry either apo B-100 or apoB-48 [67].The synthesis of apoB in the liver for the assembly of VLDL apparently occurs at a constant rate. However, only a fraction of newly-synthesized apoB protein serves as the scaffold for VLDL assembly [69]. The assembly of VLDL has been proposed to occur in two sequential steps [70]. In the first step, apoB, during its co-translational translocation through a protein channel in the membrane of the rough endoplasmic reticulum, acquires a small quantity of triglyc-erides, phospholipids and cholesterol ester, forming a small, dense, VLDL precursor. At this stage, the apoB, if not properly folded or if it acquires an insufficient amount of lipids, is rapidly degraded and this process appears to be the principal determinant of the amount of apoB that is secreted into the circulation associated with the VLDL [71]. The acquisition of triglycerides by apoB is mediated by microsomal triglyceride transfer protein (MTP), which func-tions in the endoplasmic reticulum lumen as a chaperone shuttling triglycerides and phospholipids to the newly syn-thesized apoB [72]. In a second step, the small, dense VLDL precursor undergoes maturation by the further acquisition of triglycerides, a process that is still poorly understood, but which is thought to occur primarily by fusion between the newly lipidated particle with triglyceride droplets in the smooth endoplasmic reticulum [73]. This process does not appear to require the mediation of MTP [74, 75]. Under physiological conditions, the production of VLDL depends, primarily, on the availability of fatty acids that are taken up by the liver. However, overproduction of VLDL can occur resulting in hypertriglyceridemia. The mechanisms underly-ing VLDL overproduction are poorly understood although insulin appears to play an important role. High levels of this hormone increase levels of MTP expression; increase apoB availability, and induce the transcriptional regulation of hepatic lipogenic enzymes, all of which can lead to increased VLDL production and secretion [76].
但是应当指出的是，进入细胞后，该脂肪酸是通过酰基辅酶Ａ合成酶被激活为酰基辅酶Ａ酯并且靶向进入线粒体β氧化螺旋but fatty acids are considered the major source of energy for many cell types except the brain, which uses glucose and also ketone bodies for ATP generation [4, 77].Liver plays a central role in the fatty acid oxidation for energy generation and for the production of substrates for the synthesis of ketone bodies for use by extrahepatic tissues under fasting conditions for energy. In liver cells, fatty acids are oxidized in three cellular organelles, with b-oxidation confined to mitochondria and peroxisomes, and the CYP4A catalyzed w-oxidation taking place in the endoplasmic retic-ulum [4, 78]. The major pathway for the catabolism of fatty acids is mitochondrial fatty acid b-oxidation [4, 79]. The fol-lowing is a brief overview of the fatty acid oxidation pro-cesses in the liver.
Mitochondrial b-Oxidation
所得到的链缩短的酰基辅酶Ａ在线粒体中穿梭完成氧化Fatty Acid Oxidation
Breakdown of the major energy fuels namely, carbohydrates, amino acids, and fats, generates ATP, which is the universal cellular energy source. For ATP to be synthesized from these complex fuels, they first need to be broken down into their basic components. In general, carbohydrates are hydrolyzed into simple sugars, such as glucose and fructose, proteins to amino acids and fats (triglycerides) to fatty acids. Mitochondria use these energy-generating fuels and play a dominant role in ATP generation. The extent to which these fuels contribute to ATP production within an organism varies,
Mitochondrial b-oxidation is responsible for the degradation of the major portion of the short- ( Mitochondrial fatty acid b-oxidation is a complex process which is regulated at several levels, but mainly by carnitine palmitoyltransferase 1 (CPT1), the carnitine concentration, and malonyl-CoA, which inhibits CPT1. It should be noted that, after entry into the cell, the fatty acids are activated to acyl-CoA esters by acyl-CoA synthetases and targeted into the mitochondrial b-oxidation spiral [4]. Because the mito-chondrial inner membrane is impermeable to long-chain acyl-CoAs, they are transported across to the mitochondrial matrix by the so-called carnitine shuttle. This rate-controlling shuttle utilizes three proteins: CPT1, carnitine acylcarnitine translocase (CACT), and CPT2 [81]. CPT1 exchanges the CoA group of long-chain acyl-CoA for carnitine to form long-chain acylcarnitines, which are transported across the mitochondrial inner membrane by carnitine acylcarnitine translocase [82]. CPT2 located at the mitochondrial inner 138J. Huang et al. membrane releases the carnitine group from acylcarnitines in exchange for a CoA group and delivers the CoA esters to mitochondrial matrix for oxidation. The released carnitine shuttles back to the cytosol for reuse [81, 82]. In the mitochondria, fatty acyl-CoAs ( The mitochondrial b-oxidation has two distinct compo-nents. The first, which is mitochondrial inner membrane bound, is active with long-chain fatty acyl-CoAs (Cmedium-chain acyl-CoAs 12–C20) and after 2–3 cycles, generates for further oxidation in the mitochondrial matrix. The inner membrane-bound mitochondrial b-oxidation system involves four enzy-matic reactions performed by two membrane bound proteins. The first protein, a very long-chain acyl-CoA-dehydrogenase (VLCAD), catalyzes the a-b-dehydrogenation of the acyl-CoA ester, the initial, rate-limiting first step of mitochondrial fatty acid b-oxidation, to generate trans-2,3-enoyl-CoA [77]. The second membrane bound protein involved in long-chain acyl-CoA oxidation is a mitochondrial trifunctional protein (MFP, L-PBE), with long-chain enoyl-CoA hydratase/ long-chain 3-hydroxy acyl-CoA dehydrogenase/long-chain 3-ketoacyl-CoA thiolase activities responsible for performing steps 2–4 of b-oxidation. In the second step, enoyl-CoAs are hydrated by 2-enoyl-CoA hydratases to generate L-3-hydroxyacyl-CoAs, which in the third step are catalyzed by short chain L-3-hydroxyacyl-CoA dehydrogenase to produce 3-ketoacyl-CoAs. The final step is catalyzed by 3-ketoacyl-CoA thiolases [77, 85]. After 2–3 cycles using these mem-brane bound enzymes, the resulting medium acyl-CoAs ( •Mitochondria lack very long-chain fatty acyl-CoA syn-thetase, hence VLCFAs (>CPeroxisomal membrane 20) cannot enter these organ-elles. on the other hand has at least two acyl-CoA synthetases: a long chain acyl-CoA synthetase and a very long-chain acyl-CoA synthetase capable of activating LCFAs and VLCFAs, respectively [88]. While LCFAs can be oxidized both in the mitochon-dria and peroxisomes, the presence of very long-chain fatty acyl-CoA synthetase on peroxisomal membrane accounts for the exclusively streamlined b-oxidation of VLCFA within the peroxisomes. •The first oxidation step in the peroxisomal b-oxidation of fatty acids is catalyzed by fatty acyl-CoA oxidase 1 (ACOX1) in the classic inducible pathway, but unlike in mitochondria, the b-oxidation in peroxisomes is not cou-pled to ATP synthesis. Instead, the high-potential elec-trons are transferred to Oconverted into H2 to yield H2O2, which is further energy released during peroxisomal fatty acid oxidation is 2O and O2 by peroxisomal catalase. The dissipated as heat. •Unlike the mitochondrial system, peroxisomal b-oxida-tion does not go to completion, as the appropriately chain-shortened acyl-CoAs are exported to the mitochondria for the completion of b-oxidation. •Peroxisomal b-oxidation generated chain-shortened acyl-CoAs are shuttled to mitochondria, either as carnitine esters and/or as free fatty acid for the completion oxida-tion. Peroxisomes contain carnitine acetyltransferase and carnitine octanoyltransferase for conjugation and trans-port of short- and medium-chain acyl-CoAs respectively. Mitochondria on the other hand use carnitine shuttle with carnitine palmitoyl transferase-1 (CPT1) and CPT2 as major players.It is noteworthy that the peroxisomal b-oxidation is uniquely geared toward the metabolism of less abundant and relatively more toxic and biologically active VLCFAs (>C2-methyl-branched fatty acids, dicarboxylic acids, pros-20), tanoids, and C27 bile acid intermediates, among others []. VLCFAs are not completely b-oxidized in peroxisomes, but 10 Hepatic Lipid Metabolism139 this system serves to shorten the chain length for further completion of oxidation in mitochondria [77, 90]. Although this system normally functions in the shortening of VLCFA-CoA, it also breaks down LCFA-CoA when the mitochon-drial b-oxidation is decreased or overwhelmed. Long-chain dicarboxylic acids produced by the microsomal w-oxidation of LCFAs and VLCFAs (see below) are also metabolized by the peroxisomal b-oxidation system [91]. Dicarboxylic acids are generally considered more toxic than VLCFAs and are known to inhibit mitochondrial fatty acid oxidation system and thus may contribute to the development of hepatic ste-atosis. Peroxisomal b-oxidation also acts in the synthesis and b-oxidation, especially when b-oxidation is defective. In the w-oxidation pathway, the first step involves the conversion within the endoplasmic reticulum of the w-methyl group of the fatty acid into a w-hydroxyl group by P450 enzymes belonging to the CYP4A/F subfamilies. In the human CYP4A11 of CYP4A family and CYP4F11 of the CYP4F family appear to be the predominant catalysts for fatty acid w-oxidation [96, 97]. The resulting w-hydroxy fatty acid is then dehydrogenated to a dicarboxylic acid in the cytosol. The dicarboxylic fatty acids generated by w-oxidation require b-oxidation in mitochondria and or peroxisomes to shorter chain dicarboxylic acids for excretion into the urine [98]. metabolism of docosahexanoic acid (DHA) and retroconver-sion of DHA to eicosapentaenoic acid. Similar to the mitochondrial b-oxidation system, the per-oxisomal b-oxidation spiral consists of four sequential steps with each metabolic conversion carried out by at least two different enzymes [90, 92]. These enzymes are separated into two pathways, inducible and non-inducible, with each pathway consisting of three separate enzymes. The first step of the peroxisomal b-oxidation in each pathway is catalyzed by a different ACOX, with an inducible classic ACOX1 exhibiting specificity for straight-chain VLCFA-CoA esters, dicarboxylic acids, and eicosanoids the non-inducible ACOX2 acting on CoA esters for 2-methyl branched-chain fatty acids [77, 93]. This first step converts acyl-CoA into enoyl-CoA. The second and third reactions, hydration and dehydrogena-tion of enoyl-CoA esters to 3-ketoacyl-CoA are catalyzed by one of two bi/multi-functional enzymes (PBE/MFP) with enoyl-CoA hydratase/L-3-hydroxyacyl-CoA dehydrogenase (L-PBE/MFP1) in the inducible pathway, or D-3-hydroxyacyl-CoA dehydratase/D-3-hydroxyacyl-CoA dehydrogenase (D-PBE/MFP2) in the non-inducible pathway. D-PBE/MFP2 of this pathway can substitute for the L-PBE/MFP1 function in the inducible [4, 90, 94]. The fourth step in the peroxisomal b-oxidation converts 3-ketoacyl-CoAs to acyl-CoA that is two carbon atoms shorter than the original molecule and acetyl-CoA. This function in the inducible pathway is performed by straight-chain 3-oxoacyl-CoA thiolase and in the noninduc-ible pathway by the sterol carrier protein x (SCPx), which possesses thiolase activity [94, 95]. The functional signifi-cance of peroxisomal b-oxidation system is to metabolize potentially toxic substrates such as VLCFAs and shuttle the chain-shortened metabolites to mitochondrial b-oxidation system for further degradation and to prevent hepatic steatosis. Microsomal w-OxidationLiver also utilizes the microsomal w-oxidation system to metabolize fatty acids (C10–C26) as an alternative pathway to Prior to entering b-oxidation spiral, dicarboxylic acids are converted into dicarboxylyl-CoAs by acyl-CoA synthase present in endoplasmic reticulum. Medium-chain dicarboxy-lyl-CoAs are oxidized in mitochondria, whereas long- and very long-chain dicarboxylyl-CoAs are metabolized exclu-sively by the classic inducible peroxisomal b-oxidation sys-tem in humans. Although w-oxidation is a minor catabolic pathway accounting for <10% of total fatty acid oxidation in the liver, this system constitutes a critical route in the elimination of potentially toxic levels of free fatty acids. First, w-oxidation is upregulated when b-oxidation is defec-tive, and this poses a problem as the dicarboxylic acids so generated cannot be adequately metabolized to avert accu-mulation of toxic levels of dicarboxylic acids. A defect in b-oxidation leads to dicarboxylic acid toxicity [77, 99]. Significant quantities of dicarboxylic acids are also generated under conditions of fatty acid overload in the liver, for example, in obesity and diabetes, and in conditions where mitochon-drial oxidation is inadequate to oxidize fatty acids. Fatty Acid a-Oxidation Plants synthesize fatty acids such as phytanic acid with odd numbers of carbon and these 3-methyl branched fatty acids cannot be b-oxidized, but use an alternate a-oxidation path-way present in peroxisomes [100]. Oxidation of phytanic acid proceeds as follows: phytanic acid is activated to phytanoyl-CoA by long-chain acyl-CoA synthetase, which is then hydroxylated to 2-hydroxyphytanoyl-CoA by phytanoyl-CoA 2-hydroxylase. Subsequently, 2-hydroxyphytanoyl-CoA is cleaved to yield formyl-CoA and pristanal. Pristanal is oxidized to pristanic acid and upon activation to pristanoyl-CoA, it enters peroxisomal b-oxidation spiral, eventually generating propionyl-CoA and acetyl-CoA. In essence, 3-methyl branched fatty acids in mammalian liver are sub-jected to a-oxidation in the initial steps and to b-oxidation process subsequently to generate acetyl-CoA and 2-methyl propionyl-CoA [100]. 140J. Huang et al. Ketogenesis Under conditions contributing to increased fatty acid uptake and oxidation in the liver, large amounts of acetyl-CoA are generated. When acetyl-CoA concentrations exceed the capacity of tricarboxylic acid cycle to metabolize, acetyl-CoAs are utilized to generate ketone bodies called acetoace-tate, b-hydroxybutyrate, and acetone. These products of ketogenesis, can be used as energy source by tissues such as brain, heart, skeletal muscle, liver, and kidneys [84, 101]. Accordingly, ketogenesis depends on the capacity of liver to fully oxidize fatty acids. It should be noted that the release of genes involved in lipogenesis, and peroxisome proliferator-activated receptor-a (PPARa), which controls the expression of genes of the fatty acid oxidation systems in liver [104–106]. Two other members of PPAR subfamily, namely, PPARb/d and PPARg, also play key roles in overall lipid metabolism of the organism [106]. Fatty acids and their metabolites control the expression of these factors. Retinoid X receptor a (RXRa), a heterodimerization partner of PPARs and several other nuclear receptors [105, 107], are also involved in lipid homeostasis. Evidence is also emerging about the role of microRNAs in the regulation of genes that participate in liver lipid metabolism [108]. glucose by the liver is essential to meet the energy needs of many cell types. During fasting conditions, after the hepatic glycogen reserves are exhausted, the organism relies on ketone bodies which become the main alternative fuel source. Ketone bodies released from liver provide up to 80% of the total energy needed. This glucose sparing effect of ketone bodies is extremely important in cerebral tissue in infants due to the high proportional glucose utilization [101]. Regulation of Hepatic Lipid Metabolism: Clues from Knockout Mouse ModelsOver the years, the importance of lipid metabolism in liver in the development of alcoholic steatohepatitis and chronic fatty liver disease is well recognized. Currently, lipid meta-bolic dysfunctions in liver, in particular steatosis and steato-hepatitis, related to obesity and metabolic syndrome are receiving considerable attention. Accordingly, an under-standing of the regulatory factors influencing overall energy balance and attempts at pharmacological intervention are of paramount importance.Several genetic disorders affecting the mitochondrial and peroxisomal fatty acid oxidation system genes at different levels of the catabolic spiral have been discovered in humans [4, 80, 91]. These defects have provided fascinating insights into the role of fatty acid catabolism in maintaining energy homeostasis and causing dysfunction. In addition, during the past 25 years, several gene knockout mouse models have been generated to investigate the functional roles of many genes involved in various aspects of lipid metabolism.The following provides a brief overview of the regulatory molecules that influence lipogenesis, and fatty acid oxidation [102]. Abnormalities in lipogenesis, fatty acid oxidation, and in fatty acid intake and export, contribute to the development of hepatic steatosis and these functions in liver are influenced by many factors [103]. In this regard, attention is focused on two key transcription factors, the sterol regulatory element-binding protein 1-c (SREBP1-c) that controls the expression of Regulation of Hepatic LipogenesisAs de novo synthesis of fatty acids in liver is regulated by insulin and glucose, it is essential to understand their role in activating the membrane-bound lipogenic transcription fac-tor SREBP-1c, an important member of SREBP family of transcription factors [109]. Insulin is a key regulator of SREBP-1c, one of the three SREBP isoforms in humans and rodents [110]. SREBP-1c and SREBP-2 are the predominant forms in the liver. SREBP-1a and SREBP-1c activate hepatic fatty acid synthesis through regulation of lipogenic genes, such as ACC2, FAS, SCD-1 and glycerol phosphate acyl-transferase [111]. ACC2-generated malonyl-CoA inhibits fatty acid entry and oxidation in mitochondria, because malonyl-CoA inhibits CPT1 activity. As noted above, muta-tion of ACC2 results in increased fatty acid oxidation due to reduced malonyl-CoA production [112]. SREBP-1c induces lipogenic genes by binding to sterol response element (SRE) sites in their promoters, which is impaired in SREBP-1c−/− mice. SREBP-1c gene contains two response elements for lipogenic transcription factor liver X receptor (LXR), and the activation of LXR by oxysterol ligands induces the transcrip-tion of SREBP-1c [102]. Disruption of SREBP-1c gene expression in ob/ob mice improves hepatic steatosis, while overexpression of SREBP-1c in the liver leads to increases in hepatic glycogen and triglyceride contents [113]. Glucose regulates lipogenesis as glucose is converted to fatty acids. Glucose-mediated stimulation of lipogenesis is controlled by transcription factor carbohydrate response element-binding protein (ChREBP) [114]. ChREBP activates the expression of genes involved in the synthesis and uptake of fatty acids, including those encoding ACC, FAS, SCD-1, and ATP cit-rate lyase [102, 115]. Glucose exerts the ChREBP activation by regulating the entry of ChREBP from the cytosol into the nucleus thus influencing the DNA binding and transcrip-tional activity of this transcription factor [116].The transcriptional activities of ACC1, ACC2, and FAS in liver are regulated by both glucose and insulin [102, 117]. 10 Hepatic Lipid Metabolism141 Deficiency of ACC1 is embryonically lethal in mice, whereas ACC2 null mice are viable but exhibit enhanced metabolic rate, manifesting as continuously elevated fatty acid oxida-tion and reduced adiposity [118, 119]. In ACC2 null mice, CPT1 activity increases resulting in an increased rate of fatty acid oxidation [120]. It would appear that malonyl-CoA, generated by ACC1 and ACC2, work independently and that there is minimal, if any, overlap of their functions. FAS, which catalyzes the first committed step in fatty acid biosyn-thesis to generate mostly the saturated fatty acid palmitate, also plays an important role in fatty acid oxidation (see below). When FAS gene is conditionally inactivated in mouse humans include: deficiencies of CPT1, CPT2, CACT, VLCAD, MTP, MCAD, M/SCHAD, and among others [4, 77, 91]. Mouse models of gene disruption affecting mito-chondrial b-oxidation include: CPT1a, CPT1b, LCHAD/MTP, VLCAD, LCAD, MCAD, and SCAD among others [90, 91]. Most of these gene knockout mouse models exhibit severe or lethal phenotype and, in most part, mimic human disease with some exceptions [90, 128, 129]. In both humans, and mouse models, peroxisomal b- oxidation pathway disruptions have been studied in consid-erable detail [4, 50, , 91, 125, 127–129]. Three genetic disorders affecting peroxisomal b-oxidation spiral, namely liver, it causes decreased fatty acid oxidation suggesting that this enzyme is involved in generating ligands that activate the lipid sensing transcription factor, PPARa, a critical regu-lator of fatty acid oxidation [90, 121]. SCD-1 is also modu-lated by a number of dietary, physiological and hormonal factors including insulin and glucose, and is critical in main-taining intracellular lipid flux. SCD-1 has also been sug-gested in representing a key step in the partitioning of lipids between storage and oxidation [122]. Mice with global dele-tion of SCD-1 are resistant to high-fat diet induced obesity and SCD-1 deficiency also prevents the development of hepatic steatosis [123]. Significant increase of SCD-1 expres-sion and reduced energy expenditure in ob/ob mice result in marked hepatic steatosis, hyperlipidemia and increase of lipid secretion. Regulation of Fatty Acid Oxidation Oxidation of fatty acids in liver involves the participation of several enzymes that are affected by a multiplicity of factors including more prominently the nuclear receptor PPARa [90, 91]. Enzymes such as CPT1 and CPT2, and substrates (fatty acid metabolism intermediates), such as malonyl-CoA are known to influence mitochondrial fatty acid b-oxidation [82]. Likewise, several structurally diverse compounds called peroxisome proliferators [4, 90, 91], and certain metabolic pathways that generate or degrade biological (endogenous) PPARa ligands, also affect fatty acid catabolism, especially by activating PPARa, which is referred to as lipid and per-oxisome proliferator-sensor in liver [77]. PPARa transcrip-tionally regulates all key enzymes of peroxisomal and mitochondrial b-oxidation pathways, as well as microsomal w-oxidation system [4, 90, 91, 124–127]. Several genetic disorders in humans involving the enzymes of the mitochon-drial and peroxisomal fatty acid b-oxidation pathways have been identified that have provided considerable insights into the regulatory aspects of energy homeostasis and in particu-lar implications of disturbances in fatty acid metabolism. Genetic defects affecting mitochondrial b-oxidation in ACOX1 deficiency, DPBE/MFP1 deficiency and SCPx defi-ciency have been identified and studied in detail in humans [4, 91, 127]. Gene knockouts in the mouse include disrup-tions of genes encoding ACOX1, L-PBE/MFP1, D-PBE/MFP2, and SCPx [4, , 91, 128, 129]. The following focuses, first, on the role of the transcrip-tion factor PPARa in regulating the fatty acid oxidation sys-tem genes, and second, as to how disruption of these genes contributes to high levels of unmetabolized substrates that function as PPARa ligands [128]. It is also worth noting that some of these substrates/ligands may be generated by a prox-imal enzyme in the oxidation pathway. It is becoming increasingly clear that some of the enzymes in fatty acid oxi-dation systems contribute to the formation or degradation of biological ligands of PPARa [4, 90, 92, 128]. First, all three members of the PPAR subfamily are known to function as sensors for fatty acids and fatty acid deriva-tives and thus control important metabolic pathways involved in lipid and energy metabolism [90, 91, 126]. Of these, PPARa is a key regulator of mitochondrial, peroxisomal, and microsomal fatty acid oxidation enzyme systems in liver [4, 90, 130–133]. Exposure to peroxisome proliferators, which are synthetic PPARa ligands, results in massive induc-tion of these fatty acid oxidation systems in liver culminating in increased energy combustion [4, 77]. This receptor is also activated by both saturated and poly-unsaturated fatty acids and their derivatives. PPARa also plays an important role in lipoprotein synthesis and inflammatory responses and in the development of liver cancer [90]. PPARa knockout mice have demonstrated unequivocally that this receptor is indeed the bona fide receptor for transcriptionally activating the fatty acid oxidation genes and inducing the peroxisome pro-liferator-mediated pleiotropic responses including liver tumors [130, 134]. PPARa null mice display normal basal levels of the inducible peroxisomal b-oxidation enzymes in liver but possess generally lower levels of mitochondrial b-oxidation enzyme activities [131]. Under fasting condi-tions, PPARa senses the fatty acid influx into the liver and up-regulates all three fatty acid oxidation systems to com-bust the energy and minimize hepatic steatosis [33–35]. Mice deficient in PPARa fail to upregulate fatty acid burning 142J. Huang et al. enzymes in liver and become grossly steatotic when fasted [33–35] (Fig. 10.1). When maintained on a diet deficient in methionine and choline, PPARa null mice develop severe steatohepatitis [135]. Second, increasing evidence suggests that PPARa senses certain endogenous lipid metabolic intermediates as ligands and participates in their metabolic breakdown by inducing downstream lipid metabolism genes [90, 128]. Studies with ACOX1 null mice revealed that disruption of this gene encoding the first and rate-limiting enzyme of the fatty acid b-oxidation system results in profound activation of PPARa in liver [94, 128] (Fig. 10.2). Animals with ACOX1 defi-ciency have high levels of VLCFAs and since VLCFA-CoAs are incapable of entering the fatty acid oxidation pathway due to ACOX1 deficiency, these unmetabolized substrates act as ligands to hyper activate PPARa in liver [4, 90, 128]. Recent studies with gene knockout mouse models suggest that enzymes such as ACOX1, L-PBE/MFP1, D-PBE/MFP2, and SCPx of peroxisomal b-oxidation spiral are necessary for the degradation of endogenously generated PPARa ligands [50, 78, 80, –91, 102, 104, 128, 129]. It is also becoming clear that other enzymes such as FAS, FACS, and certain lipoxygenases are required for PPARa ligand genera-tion in vivo [90, 104, 121]. For example, FAS deficient mice reveal a phenotype resembling PPARa deficiency. FAS generated 1-palmitoyl-2-oleoyl-sn-glycerol-3-phosphocholine (16:0/18:1-GPC), functions as a physiologically relevant PPARa ligand [104] (Fig. 10.2). Thus, absence of an enzyme such as FAS fails to generate a ligand, where as absence of an enzyme such as ACOX1 fails to degrade endogenous PPARa ligand(s). These observations establish that fatty acid metabolism in liver is intricately regulated to prevent to harmful accumulation of products that may lead to hepatic steatosis and steatohepatitis, or that fail to upregulate PPARa under conditions of stress such as that induced by excess fatty acid influx into liver during fasting or excess energy overload. MicroRNAs in the Regulation of Lipid Metabolism Recent studies suggest that microRNAs play a role in regu-lating lipid metabolism [136–139]. For example, ectopic expression of let-7 blocks 3T3 L1 cell growth and completely inhibits terminal differentiation [136]. miR-103(1), miR-103(2), and miR-107 within introns of the pantothenate Fig. 10.2 Biological ligands of PPARa. Lipid metabolism generates intermediary metabolites and if some of these, for example acyl-CoAs generated by fatty acyl-CoA synthetase (FACS) remain unmetabolized as in the absence of ACOX1, they can act as biological (endogenous) PPARa ligands in liver. Gene knockout mouse models have provided valuable insights regarding PPARa ligand degrading and ligand generating enzymes in liver. The model also shows peroxisome proliferator response element (PPRE) hexanucleotide direct repeat separated by one nucleotide (DR1) and the putative PPAR and RXR heteridimer binding half-sites. Also shown are the basal transcription machinery and the transcription coactivator PBP/MED1 as they appear critical for PPAR regulated tran-scriptional activity. From Pyper et al. [90] with permission 10 Hepatic Lipid Metabolism143 kinase gene appear to be important in the regulation of cellular acetyl-CoA synthesis [137]. miR-122 is a liver specific microRNA, expressed in rodents and appears to regulate cholesterol and hepatic lipid metabolism [138]. Inhibition of miR-122 in mice results in a significant improvement in liver steatosis, which may result from a reduction of several lipo-genic genes and stimulation of hepatic fatty-acid oxidation. Recent studies showed that PPARd and PPARa coactivator Smarcd1/Baf60a are the target genes of miR-122 [138]. Several microRNAs have been proposed to be associated and/or products generated by specific enzymatic processes appear to function as biological ligands for the transcription factors that control the expression of genes responsible for lipid metabolism in liver. Acknowledgment This work was supported by NIH Grant DK083163 (J.K.R). References with obesity. The expression of miR-335 in liver is elevated in obese mouse models, including ob/ob, db/db, and KKAy mice and its expression accounts for increase of body, liver size, and white adipose tissue stores [139]. miR-335 levels are closely correlated to the expression of PPARg, aP2, and FAS expression, the markers for adipocyte differentiation. Thus, the induction of miR-335 may contribute to the pathophysiology of obesity and to the development of hepatic steatosis [139]. Recent microarray data showed that miR-151, -192, -34a, -24, -10b, -132 are upregulated in hepatic steatosis [140]. The identification of target genes for these micro-RNAs and authentication of changes in their expres-sion require further attention to appreciate the regulatory implications of these pivotal molecules in hepatic lipid metabolism. Manipulating microRNA levels may be an another layer of gene regulation, and may present a potential for therapeutic targets. Summary Lipids constitute an important source of energy. Liver plays a central role in lipid metabolism as it is critical for lipogen-esis and fatty acid catabolism. Several enzymes participating in these energy balancing processes are affected by a variety of pathophysiological conditions and regulated by certain transcription factors. Evidence indicates that SREBPs and PPARa play prominent roles in controlling the hepatic expression of genes responsible for fatty acid synthesis and oxidation, respectively. Elucidation of the role of enzymes and of transcription factors that regulate the expression of these enzymes is critical for understanding the intricacies involved in the pathogenesis of hepatic steatosis and for the development of therapies. Several genetic disorders affecting the mitochondrial and peroxisomal fatty acid oxidation sys-tems have been identified, and in recent years, gene knock-out-mouse models have also been generated to investigate in detail the role of a given gene in modulating the metabolic pathways when their function is disrupted. These genetically altered mouse models are also providing considerable insights into the role of enzymes in generating and degrading endogenous ligands for transcription factors. The substrates 1. Eyster KM. The membrane and lipids as integral participants in signal transduction: lipid signal transduction for the non-lipid biochemist. Adv Physiol Educ. 2007;31:5–16. 2. Hostetler HA, Huang H, Kier AB, et al. Glucose directly links to lipid metabolism through high affinity interaction with per-oxisome proliferator-activated receptor alpha. J Biol Chem. 2008;283:2246–54. 3. Busetto L. Visceral obesity and the metabolic syndrome: effects of weight loss. Nutr Metab Cardiovasc Dis. 2001;11:195–204. 4. Reddy JK, Hashimoto T. Peroxisomal beta-oxidation and peroxi-some proliferator-activated receptor alpha: an adaptive metabolic system. Annu Rev Nutr. 2001;21:193–230. 5. Mensenkamp AR, Havekes LM, Romijn JA, et al. Hepatic steato-sis and very low density lipoprotein secretion: the involvement of apolipoprotein E. J Hepatol. 2001;35:816–22. 6. Zimmermann R, Lass A, Haemmerle G, et al. Fate of fat: the role of adipose triglyceride lipase in lipolysis. Biochim Biophys Acta. 2009;1791:494–500. 7. Schweiger M, Schreiber R, Haemmerle G, et al. Adipose triglyc-eride lipase and hormone-sensitive lipase are the major enzymes in adipose tissue triacylglycerol catabolism. J Biol Chem. 2006;281:40236–41. 8. Brasaemle DL. Thematic review series: adipocyte biology. The perilipin family of structural lipid droplet proteins: stabilization of lipid droplets and control of lipolysis. J Lipid Res. 2007;48: 2547–59. 9. Holm C. Molecular mechanisms regulating hormone-sensitive lipase and lipolysis. Biochem Soc Trans. 2003;31:1120–4. 10. Wang S, Soni KG, Semache M, et al. Lipolysis and the integrated physiology of lipid energy metabolism. Mol Genet Metab. 2008;95:117–26. WJ, Sridhar K, Bernlohr DA, et al. Interaction of rat hormone-sensitive lipase with adipocyte lipid-binding protein. Proc Natl Acad Sci U S A. 1999;96:5528–32. 12. Wang H, Eckel RH. Lipoprotein lipase: from gene to obesity. Am J Physiol Endocrinol Metab. 2009;297:E271–88. 13. Cianflone K, Paglialunga S, Roy C. Intestinally derived lipids: metabolic regulation and consequences – an overview. Atheroscler Suppl. 2008;9:63–8. 14. Stahl A. A current review of fatty acid transport proteins (SLC27). Pflugers Arch. 2004;447:722–7. 15. Guo W, Huang N, Cai J, et al. Fatty acid transport and metabolism in HepG2 cells. Am J Physiol Gastrointest Liver Physiol. 2006;290:G528–34. 16. Pohl J, Ring A, Hermann T, et al. Role of FATP in parenchymal cell fatty acid uptake. Biochim Biophys Acta. 2004;1686:1–6.17. Doege H, Baillie RA, Ortegon AM, et al. Targeted deletion of FATP5 reveals multiple functions in liver metabolism: alterations in hepatic lipid homeostasis. Gastroenterology. 2006;130:1245–58.18. Wu Q, Ortegon AM, Tsang B, et al. FATP1 is an insulin-sensitive fatty acid transporter involved in diet-induced obesity. Mol Cell Biol. 2006;26:3455–67. 11. Shen 144 19. Abumrad N, Coburn C, Ibrahimi A. Membrane proteins implicated in long-chain fatty acid uptake by mammalian cells: CD36, FATP and FABPm. Biochim Biophys Acta. 1999;1441:4–13. 20. Yu KC, Cooper AD. Postprandial lipoproteins and atherosclerosis. Front Biosci. 2001;6:D332–54. 21. Arrese EL, Soulages JL. Insect fat body: energy, metabolism, and regulation. Annu Rev Entomol. 2010;55:207–25. 22. Ducharme NA, Bickel PE. Lipid droplets in lipogenesis and lipol-ysis. Endocrinology. 2008;149:942–9. 23. Walther TC, Farese Jr RV. The life of lipid droplets. Biochim Biophys Acta. 2009;1791:459–66. 24. Murphy S, Martin S, Parton RG. Lipid droplet-organelle interac-tions; sharing the fats. Biochim Biophys Acta. 2009;1791:441–7. 25. Zehmer JK, Huang Y, Peng G, et al. A role for lipid droplets in J. Huang et al. and protein kinase A-stimulated lipolysis. J Biol Chem. 2003; 278:51535–42. 42. Fujii H, Ikura Y, Arimoto J, et al. Expression of perilipin and adi-pophilin in nonalcoholic fatty liver disease; relevance to oxidative injury and hepatocyte ballooning. J Atheroscler Thromb. 2009; 16:3–901. 43. Traini M, Jessup W. Lipid droplets and adipose metabolism: a novel role for FSP27/CIDEC. Curr Opin Lipidol. 2009;20:147–9. 44. Le Lay S, Dugail I. Connecting lipid droplet biology and the meta-bolic syndrome. Prog Lipid Res. 2009;48:191–5. 45. Meex RC, Schrauwen P, Hesselink MK. Modulation of myocellu-lar fat stores: lipid droplet dynamics in health and disease. Am J Physiol Regul Integr Comp Physiol. 2009;297:R913–24. 46. Kim JY, Liu K, Zhou S, et al. Assessment of fat-specific protein 27 inter-membrane lipid traffic. Proteomics. 2009;9:914–21. 26. Gong J, Sun Z, Li P. CIDE proteins and metabolic disorders. Curr Opin Lipidol. 2009;20:121–6. 27. Wolins NE, Brasaemle DL, Bickel PE. A proposed model of fat packaging by exchangeable lipid droplet proteins. FEBS Lett. 2006;580:5484–91. 28. Kimmel AR, Brasaemle DL, McAndrews-Hill M, et al. Adoption of PERILIPIN as a unifying nomenclature for the mammalian PAT-family of intracellular lipid storage droplet proteins. J Lipid Res. 2010;51:468–71. 29. Straub BK, Herpel E, Singer S, et al. Lipid droplet-associated PAT-proteins show frequent and differential expression in neoplastic steatogenesis. Mod Pathol. 2010;23:480–92. 30. Wolins NE, Skinner JR, Schoenfish MJ, et al. Adipocyte protein S3–12 coats nascent lipid droplets. J Biol Chem. 2003;278: 37713–21. 31. Hall AM, Brunt EM, Chen Z, et al. Dynamic and differential regu-lation of proteins that coat lipid droplets in fatty liver dystrophic mice. J Lipid Res. 2010;51:554–63. 32. Dalen KT, Ulven SM, Arntsen BM, et al. PPARalpha activators and fasting induce the expression of adipose differentiation-related protein in liver. J Lipid Res. 2006;47:931–43. 33. Hashimoto T, Cook WS, Qi C, et al. Defect in peroxisome prolif-erator-activated receptor alpha-inducible fatty acid oxidation determines the severity of hepatic steatosis in response to fasting. J Biol Chem. 2000;275:218–2128. 34. Kersten S, Seydoux J, Peters JM, et al. Peroxisome proliferator-activated receptor alpha mediates the adaptive response to fasting. J Clin Invest. 1999;103:14–98. 35. Leone TC, Weinheimer CJ, Kelly DP. A critical role for the peroxi-some proliferator-activated receptor alpha (PPARalpha) in the cel-lular fasting response: the PPARalpha-null mouse as a model of fatty acid oxidation disorders. Proc Natl Acad Sci U S A. 1999;96: 7473–8. 36. Skinner JR, Shew TM, Schwartz DM, et al. Diacylglycerol enrich-ment of endoplasmic reticulum or lipid droplets recruits perilipin 3/TIP47 during lipid storage and mobilization. J Biol Chem. 2009;284:30941–8. 37. Tansey JT, Sztalryd C, Hlavin EM, et al. The central role of per-ilipin a in lipid metabolism and adipocyte lipolysis. IUBMB Life. 2004;56:379–85. 38. Moore HP, Silver RB, Mottillo EP, et al. Perilipin targets a novel pool of lipid droplets for lipolytic attack by hormone-sensitive lipase. J Biol Chem. 2005;280:43109–20. 39. Shen WJ, Patel S, Miyoshi H, et al. Functional interaction of hormone-sensitive lipase and perilipin in lipolysis. J Lipid Res. 2009;50:2306–13. 40. Marcinkiewicz A, Gauthier D, Garcia A, et al. The phosphoryla-tion of serine 492 of perilipin a directs lipid droplet fragmentation and dispersion. J Biol Chem. 2006;281:11901–9. 41. Zhang HH, Souza SC, Muliro KV, et al. Lipase-selective func-tional domains of perilipin A differentially regulate constitutive in the adipocyte lineage suggests a dual role for FSP27 in adipo-cyte metabolism and cell death. Am J Physiol Endocrinol Metab. 2008;294:E654–67. 47. Liu K, Zhou S, Kim JY, et al. Functional analysis of FSP27 protein regions for lipid droplet localization, caspase-dependent apopto-sis, and dimerization with CIDEA. Am J Physiol Endocrinol Metab. 2009;297:E1395–413. 48. Olofsson SO, Boström P, Andersson L, et al. Triglyceride contain-ing lipid droplets and lipid droplet-associated proteins. Curr Opin Lipidol. 2008;19:441–7. 49. Unger RH. Longevity, lipotoxicity and leptin: the adipocyte defense against feasting and famine. Biochimie. 2005;87:57–.50. Wanders RJ, Ferdinandusse S, Brites P, et al. Peroxisomes, lipid metabolism and lipotoxicity. Biochim Biophys Acta. 2010;1801: 272–80. 51. Malhi H, Gores GJ. Molecular mechanisms of lipotoxicity in non-alcoholic fatty liver disease. Semin Liver Dis. 2008;28:360–9.52. Imanishi Y, Gerke V, Palczewski K. Retinosomes: new insights into intracellular managing of hydrophobic substances in lipid bodies. J Cell Biol. 2004;166:447–53. 53. Parks EJ. Changes in fat synthesis influenced by dietary macro-nutrient content. Proc Nutr Soc. 2002;61:281–6. 54. Kahn A. Transcriptional regulation by glucose in the liver. Biochimie. 1997;79:113–8. 55. Jiang G, Zhang BB. Glucagon and regulation of glucose metabo-lism. Am J Physiol Endocrinol Metab. 2003;284:E671–8. 56. Wakil SJ, Abu-Elheiga LA. Fatty acid metabolism: target for met-abolic syndrome. J Lipid Res. 2009;50:S138–43. 57. Chang SI, Hammes GG. Structure and mechanism of action of a multifunctional enzyme: fatty acid synthase. Acc Chem Res. 1990;23:363–9. 58. Rioux V, Catheline D, Legrand P. In rat hepatocytes, myristic acid occurs through lipogenesis, palmitic acid shortening and lauric acid elongation. Animal. 2007;1:820–6. 59. Wang Y, Jones Voy B, Urs S, et al. The human fatty acid synthase gene and de novo lipogenesis are coordinately regulated in human adipose tissue. J Nutr. 2004;134:1032–8. 60. Gutiérrez-Juárez R, Pocai A, Mulas C, et al. Critical role of stearoyl-CoA desaturase-1 (SCD1) in the onset of diet-induced hepatic insulin resistance. J Clin Invest. 2006;116:1686–95. 61. Aarsland A, Wolfe RR. Hepatic secretion of VLDL fatty acids dur-ing stimulated lipogenesis in men. J Lipid Res. 1998;39:1280–6.62. Leonard AE, Pereira SL, Sprecher H, et al. Elongation of long-chain fatty acids. Prog Lipid Res. 2004;43:36–54. 63. Jump DB. Mammalian fatty acid elongases. Methods Mol Biol. 2009;579:375–. . Guillou H, Zadravec D, Martin PG, et al. The key roles of elon-gases and desaturases in mammalian fatty acid metabolism: insights from transgenic mice. Prog Lipid Res. 2010;49:186–99.65. Jakobsson A, Westerberg R, Jacobsson A. Fatty acid elongases in mammals: their regulation and roles in metabolism. Prog Lipid Res. 2006;45:237–49. 10 Hepatic Lipid Metabolism 66. Stein Y, Shapiro B. Uptake and metabolism of triglycerides by the rat liver. J Lipid Res. 1960;1:326–31. 67. Davidson NO, Shelness GS. APOLIPOPROTEIN B: mRNA edit-ing, lipoprotein assembly, and presecretory degradation. Annu Rev Nutr. 2000;20:169–93. 68. Anant S, Davidson NO. Identification and regulation of protein components of the apolipoprotein B mRNA editing enzyme. A complex event. Trends Cardiovasc Med. 2002;12:311–7. 69. Cano A, Ciaffoni F, Safwat GM, et al. Hepatic VLDL assembly is disturbed in a rat model of nonalcoholic fatty liver disease: is there a role for dietary coenzyme Q? J Appl Physiol. 2009;107:707–17. 70. Shelness GS, Sellers JA. Very-low-density lipoprotein assembly and secretion. Curr Opin Lipidol. 2001;12:151–7. 145 88. Steinberg SJ, Morgenthaler J, Heinzer AK, et al. Very long-chain acyl-CoA synthetases. Human \"bubblegum\" represents a new family of proteins capable of activating very long-chain fatty acids. J Biol Chem. 2000;275:35162–9. . Baes M, Huyghe S, Carmeliet P, et al. Inactivation of the peroxi-somal multifunctional protein-2 in mice impedes the degradation of not only 2-methyl-branched fatty acids and bile acid intermedi-ates but also of very long chain fatty acids. J Biol Chem. 2000;275:16329–36. 90. Pyper S, Reddy JK. PPARa(alpha): energy combustion, hypo-lipidemia, inflammation and cancer. Nucl Recept Signal. 2010;8:e002. 91. Yu S, Rao S, Reddy JK. Peroxisome proliferator-activated recep- 71. Ginsberg HN, Fisher EA. The ever-expanding role of degradation in the regulation of apolipoprotein B metabolism. J Lipid Res. 2009;50:S162–6. 72. Hussain MM, Iqbal J, Anwar K, et al. Microsomal triglyceride transfer protein: a multifunctional protein. Front Biosci. 2003;8:S500–6. 73. Gibbons GF, Wiggins D, Brown AM, et al. Synthesis and function of hepatic very-low-density lipoprotein. Biochem Soc Trans. 2004;32:59–. 74. Rustaeus S, Lindberg K, Stillemark P, et al. Assembly of very low density lipoprotein: a two-step process of apolipoprotein B core lipidation. J Nutr. 1999;129:463S–6. 75. Hebbachi AM, Gibbons GF. Microsomal membrane-associated apoB is the direct precursor of secreted VLDL in primary cultures of rat hepatocytes. J Lipid Res. 2001;42:1609–17. 76. Blasiole DA, Davis RA, Attie AD. The physiological and molecu-lar regulation of lipoprotein assembly and secretion. Mol Biosyst. 2007;3:608–19. 77. Reddy JK. Peroxisome proliferators and peroxisome proliferator-activated receptor alpha: biotic and xenobiotic sensing. Am J Pathol. 2004;1:2305–21. 78. Hashimoto T, Fujita T, Usuda N, et al. Peroxisomal and mitochon-drial fatty acid beta-oxidation in mice nullizygous for both peroxi-some proliferator-activated receptor alpha and peroxisomal fatty acyl-CoA oxidase. Genotype correlation with fatty liver pheno-type. J Biol Chem. 1999;274:19228–36. 79. Bartlett K, Eaton S. Mitochondrial beta-oxidation. Eur J Biochem. 2004;271:462–9. 80. Wanders RJ, van Grunsven EG, Jansen GA. Lipid metabolism in peroxisomes: enzymology, functions and dysfunctions of the fatty acid alpha- and beta-oxidation systems in humans. Biochem Soc Trans. 2000;28:141–9. 81. Peluso G, Petillo O, Margarucci S, et al. Differential carnitine/ acylcarnitine translocase expression defines distinct metabolic sig-natures in skeletal muscle cells. J Cell Physiol. 2005;203:439–46. 82. Ramsay RR. The carnitine acyltransferases: modulators of acyl-CoA-dependent reactions. Biochem Soc Trans. 2000;28:182–6. 83. Modre-Osprian R, Osprian I, Tilg B, et al. Dynamic simulations on the mitochondrial fatty acid beta-oxidation network. BMC Syst Biol. 2009;3:1–15. 84. Laffel L. Ketone bodies: a review of physiology, pathophysiology and application of monitoring to diabetes. Diabetes Metab Res Rev. 1999;15:412–26. 85. Kamijo T, Indo Y, Souri M, et al. Medium chain 3-ketoacyl-coen-zyme A thiolase deficiency: a new disorder of mitochondrial fatty acid beta-oxidation. Pediatr Res. 1997;42:569–76. 86. Zhang Z, Zhou Y, Mendelsohn NJ, et al. Regulation of the human long chain acyl-CoA dehydrogenase gene by nuclear hormone receptor transcription factors. Biochim Biophys Acta. 1997;1350: 53–. 87. Rakheja D, Bennett MJ, Rogers BB. Long-chain L-3-hydroxyacyl-coenzyme a dehydrogenase deficiency: a molecular and biochemi-cal review. Lab Invest. 2002;82:815–24. tors, fatty acid oxidation, steatohepatitis and hepatocarcinogenesis. Curr Mol Med. 2003;3:561–72. 92. Reddy JK, Rao MS. Lipid metabolism and liver inflammation. II. Fatty liver disease and fatty acid oxidation. Am J Physiol Gastrointest Liver Physiol. 2006;290:G852–8. 93. Hunt MC, Alexson SE. Novel functions of acyl-CoA thioesterases and acyltransferases as auxiliary enzymes in peroxisomal lipid metabolism. Prog Lipid Res. 2008;47:405–21. 94. Yeldandi AV, Rao MS, Reddy JK. Hydrogen peroxide generation in peroxisome proliferator-induced oncogenesis. Mutat Res. 2000;448:159–77. 95. Dansen TB, Kops GJ, Denis S, et al. Regulation of sterol carrier protein gene expression by the forkhead transcription factor FOXO3a. J Lipid Res. 2004;45:81–8. 96. Dhar M, Sepkovic DW, Hirani V, et al. Omega oxidation of 3-hydroxy fatty acids by the human CYP4F gene subfamily enzyme CYP4F11. J Lipid Res. 2008;49:612–24. 97. Savas U, Hsu MH, Johnson EF. Differential regulation of human CYP4A genes by peroxisome proliferators and dexamethasone. Arch Biochem Biophys. 2003;409:212–20. 98. Sanders RJ, Ofman R, Duran M, et al. Omega-oxidation of very long-chain fatty acids in human liver microsomes. Implications for X-linked adrenoleukodystrophy. J Biol Chem. 2006;281:13180–7.99. Mortensen PB. Formation and degradation of dicarboxylic acids in relation to alterations in fatty acid oxidation in rats. Biochim Biophys Acta. 1992;1124:71–9. 00. Wierzbicki AS. Peroxisomal disorders affecting phytanic acid alpha-oxidation: a review. Biochem Soc Trans. 2007;35:881–6.01. Fukao T, Song XQ, Mitchell GA, et al. Enzymes of ketone body utilization in human tissues: protein and messenger RNA levels of succinyl-coenzyme A (CoA): 3-ketoacid CoA transferase and mitochondrial and cytosolic acetoacetyl-CoA thiolases. Pediatr Res. 1997;42:498–502. 02. Guillou H, Martin PG, Pineau T. Transcriptional regulation of hepatic fatty acid metabolism. Subcell Biochem. 2008;49:3–47.103. Anderson N, Borlak J. Molecular mechanisms and therapeutic targets in steatosis and steatohepatitis. Pharmacol Rev. 2008;60:311–57.104. Chakravarthy MV, Lodhi IJ, Yin L, et al. Identification of a physio-logically relevant endogenous ligand for PPARalpha in liver. Cell. 2009;138:476–88. 105. Kliewer SA, Umesono K, Noonan DJ, et al. Convergence of 9-cis retinoic acid and peroxisome proliferator signalling pathways through heterodimer formation of their receptors. Nature. 1992;358:771–4.06. Issemann I, Green S. Activation of a member of the steroid hor-mone receptor superfamily by peroxisome proliferators. Nature. 1990;347:5–50. 07. Barish GD, Evans RM. PPARs and LXRs: atherosclerosis goes nuclear. Trends Endocrinol Metab. 2004;15:158–65. 08. Liu Z, Sall A, Yang D. MicroRNA: an emerging therapeutic target and intervention tool. Int J Mol Sci. 2008;9:978–99. 09. Eberlé D, Hegarty B, Bossard P, et al. SREBP transcription factors: master regulators of lipid homeostasis. Biochimie. 2004; 86:839–48. 1 1 1 1 1 1 1 146 110. Dif N, Euthine V, Gonnet E, et al. Insulin activates human sterol-regulatory-element-binding protein-1c (SREBP-1c) pro-moter through SRE motifs. Biochem J. 2006;400:179–88. 111. Dentin R, Girard J, Postic C. Carbohydrate responsive element binding protein (ChREBP) and sterol regulatory element binding protein-1c (SREBP-1c): two key regulators of glucose metabolism and lipid synthesis in liver. Biochimie. 2005;87:81–6. 112. Abu-Elheiga L, Matzuk MM, Abo-Hashema KA, et al. Continuous fatty acid oxidation and reduced fat storage in mice lacking acetyl-CoA carboxylase 2. Science. 2001;291:2613–6. 113. Yahagi N, Shimano H, Hasty AH, et al. Absence of sterol regula- tory element-binding protein-1 (SREBP-1) ameliorates fatty livers J. Huang et al. 126. Tontonoz P, Speigelman BM. Fat and beyond: the diverse biology of PPAR gamma. Annu Rev Biochem. 2008;77:2–312. 127. Houten SM, Wanders RJ. A general introduction to the biochem- istry of mitochondrial fatty acid beta-oxidation. J Inherit Metab Dis. 2010 [Epub ahead of print]. 128. Fan CY, Pan J, Usuda N, et al. Steatohepatitis, spontaneous peroxi- some proliferation and liver tumors in mice lacking peroxisomal fatty acyl-CoA oxidase. Implications for peroxisome proliferator-activated receptor alpha natural ligand metabolism. J Biol Chem. 1998;273:15639–45. 129. Qi C, Zhu Y, Pan J, et al. Absence of spontaneous peroxisome proliferation in enoyl-CoA Hydratase/L-3-hydroxyacyl-CoA but not obesity or insulin resistance in Lep(ob)/Lep(ob) mice. J Biol Chem. 2002;277:19353–7. 1 14. Denechaud PD, Bossard P, Lobaccaro JM, et al. al; ChREBP, but not LXRs, is required for the induction of glucose-regulated genes in mouse liver. J Clin Invest. 2008;118:956–. 1 15. Postic C, Girard J. The role of the lipogenic pathway in the devel-opment of hepatic steatosis. Diabetes Metab. 2008;34:3–8. 1 16. Bartosch B. Hepatitis C virus and its complex interplay with hepatic glucose and lipid metabolism. J Hepatol. 2009;50:845–7. 117. Denechaud PD, Dentin R, Girard J, et al. Role of ChREBP in hepatic steatosis and insulin resistance. FEBS Lett. 2008;582:68–73. 1 18. Abu-Elheiga L, Matzuk MM, Kordari P, et al. Mutant mice lacking acetyl-CoA carboxylase 1 are embryonically lethal. Proc Natl Acad Sci U S A. 2005;102:12011–6. 1 19. Abu-Elheiga L, Oh W, Kordari P, et al. Acetyl-CoA carboxylase 2 mutant mice are protected against obesity and diabetes induced by high-fat/high-carbohydrate diets. Proc Natl Acad Sci U S A. 2003;100:10207–12. 1 20. Choi CS, Savage DB, Abu-Elheiga L, et al. Continuous fat oxida-tion in acetyl-CoA carboxylase 2 knockout mice increases total energy expenditure, reduces fat mass, and improves insulin sensi-tivity. Proc Natl Acad Sci U S A. 2007;104:180–5. 1 21. Chakravarthy MV, Pan Z, Zhu Y, et al. \"New\" hepatic fat activates PPARalpha to maintain glucose, lipid, and cholesterol homeosta-sis. Cell Metab. 2005;1:309–22. 1 22. Sampath H, Ntambi JM. Stearoyl-coenzyme A desaturase 1, sterol regulatory element binding protein-1c and peroxisome prolifera-tor-activated receptor-alpha: independent and interactive roles in the regulation of lipid metabolism. Curr Opin Clin Nutr Metab Care. 2006;9:84–8. 1 23. Miyazaki M, Flowers MT, Sampath H, et al. Hepatic stearoyl-CoA desaturase-1 deficiency protects mice from carbohydrate-induced adiposity and hepatic steatosis. Cell Metab. 2007;6:484–96. 1 24. McEwan IJ. Nuclear receptors: one big family. Methods Mol Biol. 2009;505:3–18. 1 25. Desvergne B, Wahli W. Peroxisome proliferator-activated receptors: nuclear control of metabolism. Endorcri Rev. 1999;20: 9–88. dehydrogenase-deficient mouse liver. Further support for the role of fatty acyl CoA oxidase in PPARalpha ligand metabolism. J Biol Chem. 1999;274:15775–80.130. Lee SS, Pineau T, Drago J, et al. Targeted disruption of the alpha isoform of the peroxisome proliferator-activated receptor gene in mice results in abolishment of the pleiotropic effects of peroxisome proliferators. Mol Cell Biol. 1995;15:3012–22. 31. Aoyama T, Peters JM, Iritani N, et al. Altered constitutive expres-sion of fatty acid metabolizing enzymes in mice lacking the per-oxisome proliferator-activated receptor a(alpha)(PPARa(alpha)). J Bil Chem. 1998;273:5678–94. 32. Qi C, Zhu Y, Reddy JK. Peroxisome proliferator-activated recep-tors, coactivators, and downstream targets. Cell Biochem Biophys. 2000;32:187–204. 33. Nguyen P, Leray V, Diez M, et al. Liver lipid metabolism. J Anim Physiol Anim Nutr (Berl). 2008;92:272–83. 34. Gonzalez FJ, Shah YM. PPARa(alpha): mechanisms of species differences and hepatocarcinogenesis of peroxisome proliferators. Toxicology. 2008;245:2–9. 35. Ip E, Farrell GC, Robertson G, et al. Central role of PPARa(alpha)-dependent hepatic lipid turnover in dietary steatohepatitis in mice. Hepatology. 2003;38:123–32. 36. Sun T, Fu M, Bookout AL, et al. MicroRNA let-7 regulates 3T3-L1 adipogenesis. Mol Endocrinol. 2009;23:925–31. 37. Wilfred BR, Wang WX, Nelson PT. Energizing miRNA research: a review of the role of miRNAs in lipid metabolism, with a predic-tion that miR-103/107 regulates human metabolic pathways. Mol Genet Metab. 2007;91:209–17. 38. Gatfield D, Le Martelot G, Vejnar CE, et al. Integration of microRNA miR-122 in hepatic circadian gene expression. Genes Dev. 2009;23:1313–26. 39. Nakanishi N, Nakagawa Y, Tokushige N, et al. The up-regulation of microRNA-335 is associated with lipid metabolism in liver and white adipose tissue of genetically obese mice. Biochem Biophys Res Commun. 2009;385:492–6. 40. Jin X, Ye YF, Chen SH, et al. MicroRNA expression pattern in different stages of nonalcoholic fatty liver disease. Dig Liver Dis. 2009;41:2–97. 1 1 1 1 1 1 1 1 1 1
因篇幅问题不能全部显示，请点此查看更多更全内容
Copyright © 2019- 91gzw.com 版权所有 湘ICP备2023023988号-2
违法及侵权请联系：TEL:199 18 7713 E-MAIL:2724546146@qq.com
本站由北京市万商天勤律师事务所王兴未律师提供法律服务