Key Event Title
|Level of Biological Organization|
Key Event Components
|fatty acid beta-oxidation||decreased|
Key Event Overview
AOPs Including This Key Event
|AOP Name||Role of event in AOP|
|PPARα antagonism leading to body-weight loss||KeyEvent|
|Adult, reproductively mature||High|
Key Event Description
PPARα acts as a positive transcriptional regulator for many of the genes involved in peroxisomal and mitochondrial fatty acid beta oxidation as well as genes involved in the pre- and post-processing of fatty acids in peroxisomal pathways (Desvergne and Wahili 1999, Kersten 2014). Thus, decreased PPARα nuclear signaling results in decreased transcriptional expression of genes that are regulated by PPARα, and subsequently, decreased expression of the coded proteins and enzymes that ultimately decreased fatty acid metabolism within peroxisomes and mitochondria. In PPARα knock-out mice, fatty acid oxidation capacity was decreased in well fed, rested mice and the reduced fatty acid oxidation capacity was greatly exacerbated during exercise challenge or 24h starvation (Muoio et al 2002).
Genes Involved: The first gene target identified for PPARα was Acyl-CoA oxidase (Acox1, Dreyer et al 1992) which represents the first enzyme in peroxisomal long-chain fatty acid oxidation (Kersten 2014) and is also the rate-limiting enzyme in this pathway (Desvergne and Wahili 1999). In addition to Acox1, a variety of additional enzymes involved in peroxisomal fatty acid metabolism are under transcriptional control of PPARα transactivation including enzymes that facilitate fatty acid uptake into the peroxisome (Abcd1, Abcd2 and Abcd 3), conversion of acyl-CoA/acetyl-CoA to acyl-carnitine/acetyl-carnitine (Crot/Crat), and conversion of acyl-CoAs back to fatty acids via thioesterases (Acots, as reviewed in Kersten 2014). PPARalpha also has transcriptional control over enzymes downstream of Acox1 in the peroxisomal beta-oxidation of acyl-CoA pathway including L-bifunctional enzyme (Ehhadh), D-bifunctional enzyme (Hsd17b4), and peroxisomal 3-ketoacyl-CoA thiolase activity (Acaa1a, Acaa1b, as reviewed in Kersten 2014). All of these genes are potential targets for screening affects within this KE.
As reviewed in Kersten (2014), the genes (and associated functions) regulated by PPARα in the mitochondrial processing of fatty acids include the following: (1) Import of acyl-CoAs into the mitochondria is facilitated by PPARα-induced increases in expression of carnitine palmitoyl-transferases 1a, 1b, and 1 (Cpt1a, Cpt1b, Cpt2) and acyl-carnitine translocase (Slc25a20; Brandt et al 1998, Mascaro et al 1998, Feige et al 2010). (2) The first step of mitochondrial beta-oxidation is catalyzed by length-specific acyl-CoA hydrogenases (Acadvl, Acadl, Acadm, Acads; Aoyama et al 1998, Gulick et al 1994). (3) The three subsequent steps in mitochondrial beta-oxidation that successively release acetyl-CoAs from the hydrocarbon chain are catalyzed by the mitochondrial trifunctional enzyme (Hadha and Hadhb). These enzymes are replaced upon progressive chain shortening by Hadh and Acaa2. (4) the final PPARalpha targets include Eci1, Eci2, Decr1, and Hsd17b10 which convert unsaturated and 2-methlylated aclyl-CoAs into intermediates of beta-oxidation (Sanderson et al 2008, Aoyama et al 1998).
Metabolism Affected: Peroxisomes participate in a variety of lipid metabolic pathways including the beta-oxidation of very long-straight chain (<20 C in length) or branched –chain acyl-CoAs (Lazarow 1978, Kersten 2014). The peroxisomal beta-oxidation pathway is not directly coupled to the electron transport chain and oxidative phosporylation, therefore the first oxidation reaction loses energy to heat (H2O2 production) while in the second step, energy is captured in the metabolically accessible form of high-energy electrons in NADH (Mannaerts and Van Veldhoven 1993, Desvergne and Wahli 1999). The peroxisomal beta-oxidation pathway provides fatty acid chain shortening where two carbons are removed in each round of oxidation in the form of acetyl-CoA (Desvergne and Wahli 1999). The acetyl-CoA monomers serve as fundamental units for metabolic energy production (ATP) via the citric acid cycle followed by electron-transport chain mediated oxidative phosphorylation (Nelson and Cox, 2000A) as well as serve as the fundamental units for energy storage via gluconeogenesis (Nelson and Cox, 2000B) and lipogenesis (Nelson and Cox, 2000C). The shortened chain fatty acids (<20C) can then be transported to the mitochondria to undergo mitochondrial beta-oxidation for complete metabolism of the carbon substrate for cellular energy production (Desvergne and Wahli 1999).
Mitochondrial processing of fatty acids involves: (1) Import of short, medium and long chain fatty acids (<C20) acyl-CoAs into the mitochondria by carnitine palmitoyl-transferases 1a, 1b, and 1 (Cpt1a, Cpt1b, Cpt2) and acyl-carnitine translocase (Slc25a20, Brandt et al 1998; Mascaro et al 1998, Kersten et al 2014). (2) The first step of beta-oxidation catalyzed by the length-specific acyl-CoA hydrogenases (Acadvl, Acadl, Acadm, Acads; Aoyama et al 1998, Gulick et al 1994, Kersten et al 2014). (3) The three subsequent steps in mitochondrial beta-oxidation that successively release acetyl-CoAs from the hydrocarbon chain are catalyzed by the mitochondrial trifunctional enzyme (Hadha and Hadhb, Kersten et al 2014). These enzymes are replaced upon progressive chain shortening by Hadh and Acaa2 (Kersten et al 2014). (4) The conversion of unsaturated and 2-methylated acetyl-CoAs into intermediates of beta-oxidation are catalyzed by Eci1, Eci2, Decr1, and Hsd17b10 (Sanderson et al 2008, Aoyama et al 1998, Kersten et al 2014). Interestingly, tissue-specific differences in sensitivity to PPARα knock out have been observed in mice where fatty acid oxidation was markedly impaired in liver and heart tissues whereas skeletal muscle was largely unaffected due to seven-fold increased abundance of PPARδ in skeletal muscle tissue (Muoio et al 2007). Redundancy in regulation of genes involved in fatty acid metabolism across PPARα, PPARβ, PPARδ, and PPARγ are known in canonical signaling pathways (KEGG Pathway map03320) where metabolic responses are coordinated across the iso-types with whole-organism-level effects on insulin sensitivity, body fat, and body weight (Harrington et al 2007).
How It Is Measured or Detected
Beta oxidation of fatty acids in mitochondria was measured using mouse liver homogenates where a radio-labeled fatty acid substrate was reacted for 30 minutes and then centrifuged to separate reaction products for fractional radioactivity measurements in Aoyama et al (1998). Comparative measures of reaction products were also measured where potassium cyanide was added to the reaction mixture to inhibit mitochondrial beta oxidation activity to normalize the contribution of mitochondrial enzymatic reactions to the overall reaction product (Aoyama et al 1998). In Muoio et al (2002), capacity for beta oxidation of fatty acids in serum was determined by measuring the nonesterified fatty acids and transcript expression was quantified using RT-qPCR. Harrington et al (2007) administered agonists for the various PPARs and PPARpan (affecting all PPARs simultaneously) used ex vivo quantification of PPAR-induced fatty acid oxidation of 14C-labeled fatty acids to CO2.
A variety of transcript expression assays have been used to demonstrate the effect of PPARα signaling inhibition on downstream transcript expression (see literature cited above for specific methods within each investigation). Investigation of PPARα transcriptional targets (especially those involved in fatty acid metabolism) have been conducted via variety of methods, with RT-qPCR being the benchmark standard (Kersten 2014 and Feige et al 2010). Spectroscopic analysis of the characteristic absorption bands for fatty acid substrates and fatty acid beta oxidation products were examined for peroxisomal fractions purified from rat livers by differential and of equilibrium density centrifugation (Lazarow 1978). Additionally, NAD reduction assays were conducted for acyl-CoA substrates with varying chain lengths where increased oxidation was observed for substrates with long chain length relative to short chain acyl-CoAs (Lazarow 1978).
Brandt et al (1998) investigated concentration response effects of Oleate, Decanoate and Hexanoate fatty acid chains on mitochondrial carnitine palmitoyl-transferases I (M-CPT I) expression using promoter-reporter plasmid MCPT.Luc.1025 reporter transfected into rat neonate cardiac myocytes. Human M-CPT I was investigate using an analogous method (Brandt et al 1998). Expression of human medium chain acyl-CoA dehydrogenase (MCAD) was investigated using a MCAD.luc.1054 reporter transfected into HepG2 cells in response to fatty acids with various chain lengths (Gulick et al 1994). Investigation of various enzymes involved in hepatic fatty acid metabolism described in Aoyama et al (1998) were investigated using Western immunoblot quantitiation.
Domain of Applicability
Human (as reviewed in Brandt et al 1998, Evans et al 2004, Gulick et al 1994, Kersten 2014 and Desvergne and Wahli 1999; various product labels for fibrate drugs available at the US FDA). Rat (as measured by Lazarow 1978). Mouse (as reviewed in Kersten 2014 and Desvergne and Wahli 1999). Fatty acid oxidation in response to various PPAR agonists administered in mouse and human skeletal muscle tissues using in vitro assays showed the same response profiles among species (Muoio et al 2002). Results from Rakshanderhroo et al (2009) showed that PPARα transactivation caused unique transcriptional expression profiles between human and mouse, however expression for genes involved in lipid metabolism tended to be the most conserved among species. (IMPORTANT NOTE: The results from Rakshanderhroo et al (2009) should be viewed with caution given a potential an N=1 for all of the mouse strains where primary hepatocytes were obtained from 1 mouse per strain.) Interestingly, PPARα-humanized mice exposed to the PPARα agonist Wy-14643 showed no hepatocellular proliferation compared to wild type mice, but both humanized and wild types showed the same capacity for induction of peroxisomal and mitochondrial fatty acid metabolizing enzymes and resultant decreases of blood-serum trigycerides (Cheung et al 2004). (IMPORTANT NOTE: The use of wild type versus mice with humanized PPARα for extrapolating species-to-species differences should be viewed with caution. Humanized receptor is not likely to interact with the same cofactors in mice relative to humans and the regulatory grammars may differ between among species that may further complicate the biochemistry.)
Evidence for Perturbation by Stressor
Aoyama, T., Peters, J.M., Iritani, N., Nakajima, T., Furihata, K., Hashimoto, T., et al., 1998. Altered constitutive expression of fatty acid-metabolizing enzymes in mice lacking the peroxisome proliferator-activated receptor alpha (PPARalpha). Journal of Biological Chemistry 273:5678e5684.
Brandt, J.M., Djouadi, F., Kelly, D.P., 1998. Fatty acids activate transcription of the muscle carnitine palmitoyltransferase I gene in cardiac myocytes via the peroxisome proliferator-activated receptor alpha. Journal of Biological Chemistry 273:23786e23792.
Cheung, C., Akiyama, T.E., Ward, J.M., Nicol, C.J., Feigenbaum, L., Vinson, C., Gonzalez, F.J., 2004. Diminished hepatocellular proliferation in mice humanized for the nuclear receptor peroxisome proliferator-activated receptor alpha. Cancer Res. 64, 3849-3854.
Desvergne B, Wahli W (1999) Peroxisome proliferator-activated receptors: nuclear control of metabolism. Endocrine Reviews 20(5): 649-688.
Gulick, T., Cresci, S., Caira, T., Moore, D.D., Kelly, D.P., 1994. The peroxisome proliferator-activated receptor regulates mitochondrial fatty acid oxidative enzyme gene expression. Proceedings of the National Academy of Sciences of the United States of America 91:11012e11016.
Harrington, W.W., C, S.B., J, G.W., N, O.M., J, G.B., D, C.L., W, R.O., M, C.L., D, M.I., 2007. The Effect of PPARalpha, PPARdelta, PPARgamma, and PPARpan Agonists on Body Weight, Body Mass, and Serum Lipid Profiles in Diet-Induced Obese AKR/J Mice. PPAR research 2007, 97125.
Kersten S. 2014. Integrated physiology and systems biology of PPARα. Molecular Metabolism 2014, 3(4):354-371.
Lazarow PB: Rat liver peroxisomes catalyze the beta oxidation of fatty acids. J Biol Chem 1978, 253(5):1522-1528.
Mannaerts GP, Van Veldhoven PP 1993 Metabolic role of mammalian peroxisomes. In: Gibson G, Lake B (eds) Peroxisomes: Biology and Importance in Toxicology and Medicine. Taylor & Francis, London, pp 19–62.
Mascaro, C., Acosta, E., Ortiz, J.A., Marrero, P.F., Hegardt, F.G., Haro, D., 1998. Control of human muscle-type carnitine palmitoyltransferase I gene transcription by peroxisome proliferator-activated receptor. Journal of Biological Chemistry 273:8560e8563.
Muoio, D.M., MacLean, P.S., Lang, D.B., Li, S., Houmard, J.A., Way, J.M., Winegar, D.A., Corton, J.C., Dohm, G.L., Kraus, W.E., 2002. Fatty acid homeostasis and induction of lipid regulatory genes in skeletal muscles of peroxisome proliferator-activated receptor (PPAR) alpha knock-out mice. Evidence for compensatory regulation by PPAR delta. J. Biol. Chem. 277, 26089-26097.
Nelson DL, Cox MM 2000A. The Citric Acid Cycle. Lehninger Principles of Biochemistry. 3rd Edition. Worth Publishers. New York, NY. p567-592.
Nelson DL, Cox MM 2000B. Carbohydrate Biosynthesis. Lehninger Principles of Biochemistry. 3rd Edition. Worth Publishers. New York, NY. p722-764.
Nelson DL, Cox MM 2000C. Lipid Biosynthesis. Lehninger Principles of Biochemistry. 3rd Edition. Worth Publishers. New York, NY. p770-814.
Rakhshandehroo, M., Hooiveld, G., Muller, M., Kersten, S., 2009. Comparative analysis of gene regulation by the transcription factor PPARalpha between mouse and human. PLoS One 4, e6796.
Sanderson, L.M., de Groot, P.J., Hooiveld, G.J., Koppen, A., Kalkhoven, E., Muller, M., et al., 2008. Effect of synthetic dietary triglycerides: a novel research paradigm for nutrigenomics. PLoS One 3:e1681.