Email: ssmith@chori.org
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RESEARCH AREAS

  1. The Cytosolic Fatty Acid Synthase (FAS)


    Figure 1.  Reaction sequence for biosynthesis of fatty acids de novo by the animal FAS.  In the chain initiation step, a primer acetyl moiety is transferred sequentially from its CoA thioester via the nucleophilic serine residue of the acyl­transferase domain to the phosphopantetheine thiol of the acyl carrier protein domain and finally to the active-site cyteine of the ß-ketoacyl synthase domain.  The chain extender malonyl moiety is then loaded onto the vacant phosphopantetheine thiol, again via the acyltransferase domain.  Formation of a new C—C bond, by condensation of the acetyl and malonyl moieties, is coupled with an energetically favorable decarb­oxylation, so that the carbon originating from CO2, introduced in the reaction catalyzed by acetyl-CoA carboxylase, is recycl­ed.  The ß-ketoacyl condensation product is then reduced to a saturated acyl moiety through the sequential action of a ß-ketoacyl reductase, ß-hydroxyacyl dehydrase and enoyl reductase.  The newly formed saturated acyl moiety is then translocated back to the active site cysteine residue, in a reaction catalyzed by the ß-ketoacyl synthase domain, the phosphopantetheine thiol is reloaded with a malonyl moiety, and a second round of condensation and ß-carbon reduction reactions takes place.  After completion of seven cycles of chain extension and ß-carbon processing the 16-carbon satur­ated acyl moiety is released from the phosphopantetheine thiol through the action of the resident chain-terminating thioester­ase enzyme The stereochemistry of the reaction sequence was deciphered in the laboratories of John Cornforth and Gordon Hammes.  The condensation reaction proceeds with stereo­chemical inversion of the malonyl C-2, the ß-ketoacyl moiety is reduced, by transfer of the prochiral 4S hydrogen from NADPH, to a 3R (i.e. D-ß) hydroxyacyl moiety, which then, by the syn elimination of the pro-2S hydrogen and the 3R hydroxyl as water, is dehydrated to a trans-enoyl moiety; finally the enoyl moiety is reduced to a saturated acyl moiety by transfer of the prochiral 4R hydrogen of NADPH to the pro-3R position, with the simultaneous addition of a solvent proton to the pro-2S position.  Thus, with the exception of the terminal methyl group hydrogens, all of the hydrogens can be traced to a unique source.  The two C atoms at the methyl end of the fatty acid are derived from acety-CoA, the remainder from malonyl-CoA. The entire series of reactions takes approximately 1 second.

The cytosolic FAS (Figure 1) is highly active in tissues such as liver, adipose and lactating mammary glands. The FAS is essential for providing fatty acids required during embryological development and in adults it plays a key role in the conversion of dietary carbohydrate into fat.  Malonyl-CoA, one of the substrates for FAS, is now recognized as an important signaling molecule involved in metabolic fuel sensing and appetite control. Inhibitors of FAS that induce elevated malonyl-CoA levels represent a novel class of reagents for the treatment of obesity. Furthermore, high FAS levels appear to be essential for the survival of many cancer cells so that FAS is also considered a potential target for development of novel anti-cancer drugs. Elucidation of the structure and mechanism of action of the FAS should facilitate the development of these potentially valuable chemotherapeutic agents.

            The component enzymes of this system are covalently linked in a large multi­functional polypeptide that functions as a dimer (Figure 2). Until recently the FAS had been refractory to structural analysis. However, in the last several years, electron microscopy (1) and low-resolution x-ray images (2) of the dimer, high-resolution crystal structures of the thioesterase (3), acyl carrier protein (4) and transferase (5) domains and modeled structures of the remaining catalytic domains have provided new insights into the organization of this megasynthase. These studies confirmed the prediction, based on mutant complementation, mutagenesis and cross-linking experiments in the Smith lab, that the two subunits are orientated head-to-head in a coiled conformation (6-9). However, as yet, it has not been possible to trace the entire individual polypeptides through the structure, neither is it clear how the ACP domains are able to access the ß-ketoacyl synthase and malonyl/acetyl transferase domains of either subunit. A recent study has confirmed that the dehydrase domain is actually a pseudodimer consisting of two subdomains derived from contiguous regions of the same polypeptide (10). The possibility that the ß-ketoacyl reductase might consist of two subdomains derived from regions flanking either side of the enoyl reductase domain, SD and KR, in Figure 2, (11) also needs to be evaluated experimentally.

fig_2
Figure 2. Structural organization of FAS domains. Upper panel shows the linear arrangement of the functional domains in the polypeptide. KS, ß-ketoacyl synthase; MAT, malonyl/acetyl transferase; DH1 and DH2, subdomains of the dehydrase; SD, structural domain; ER, enoyl reductase; KR, ß-ketoacyl reductase; ACP, acyl carrier protein; TE, thioesterase.  The role of the structural domain is unknown but it may be required for activity of the ß-ketoacyl reductase. The lower panel shows, in light gray, the structure of the dimer, as revealed by electron microscopy while the fitting of the functional domains was estimated primarily from the low-resolution X-ray structure of the whole complex. High-resolution x-ray structures have been derived for the TE, ACP and MAT domains, whereas structures of the other domains are derived from modeling in silico. Biochemical studies have revealed that the ACP is able to make functional contacts with all of the domains associated with the same subunit, as well as the KS and MAT domains associated with the companion subunit, as represented by the arrows.

 

II. The Mitochondrial Fatty Acid Synthase
The Smith laboratory at CHORI and the Hiltunen laboratory in Finland have independently identified, cloned, expressed and characterized most of the components of a human mitochondrial FAS system (12-15). Components of the mitochondrial FAS system exist as separate, free­standing proteins that are more closely related to freestanding prokaryotic FAS components than they are to their covalently-linked counterparts in the cytosolic FAS (Figure 3). This finding is consistent with the hypoth­esis that mitochondria orig­inated from free-living pro­karyotic org­anisms. These mitochondrial FAS proteins are nuclear encoded  but contain aminoterminal extension sequences that target the proteins for import into the mitochondria (Figure 4).

Figure 3. Phylogenetic Tree of Mitochondrial ß-Ketoacyl Synthases.  Mitochondrial KS lineages are shown in red and closely related bacterial and plant KSs in green; KSs associated with animal cytosolic FASs are shown in blue.

 

Figure 4. Mitochondrial targeting of FAS components.  The N-terminal 38 residues of the mitochondrial KS, when fused to the N-terminus of the green fluorescent protein directs the chimera into the mitochondria in HeLa cells.

The group is presently evaluating the role of the pathway in mitochondrial function. The working hypothesis is that this FAS likely plays an important role in maintaining mitochondrial integrity and may supply lipoyl moieties essential for the functioning of the alpha-ketoacid dehydrogenases. Reconstitution of the entire pathway in matrix extracts from bovine heart mitochondria has revealed that one of the major products is octanoyl-ACP (16). Furthermore the octanoyl moieties can be translocvated to the lipoylation site on an acceptor protein, the H-protein of the glycine cleavage complex. Future studies will utilize siRNA silencing and knockout mouse models to evaluate the physiological importance of the pathway in mammals.

Although mitochondria contain more than 1500 proteins, the function of most of them has yet to be determined. Failure in mitochondrial function has been implicated in the pathogenesis of late developing neurodegenerative disorders such as Parkinson’s, Alzheimer’s, and Huntington’s diseases. Elucidation of the role of the mitochondrial FAS system in mitochondrial function may aid in understanding the etiology of these disorders.

 

III. Posttranslational Phosphopantetheinylation
Four mammalian proteins have been identified that require posttranslational phosphopantetheinylation for biological activity: the acyl carrier proteins of the cytosolic and mitochondrial fatty acid synthases, aminoadipate semialdehyde dehydrogenase and 10-formyltetrahydrofolate dehydrogenase. Although microorganisms, and even fungi, utilize different phosphopantetheinyl transferases to service different apoproteins, mammals appear to employ a single promiscuous enzyme for modification of all apoproteins (17).

The crystal structure of the human phosphopantetheinyl transferase has been solved complexed with the phosphopantetheine donor and acceptor, CoA and acyl carrier protein, respectively (Figure 5). X-ray analysis and mutagenesis experiments have provided insights into the substrate recognition and catalytic mechanisms (4). Human phosphopantetheinyl transferase exhibits an α/β fold and 2-fold pseudosymmetry and although the bound ACP exhibits a typical four-helix structure, its binding is unusual in that it is facilitated predominantly by hydrophobic interactions.

 

IV. Collaborators
Our collaborators for the electron microscopy studies on the cytosolic FAS are Drs. Ed Brignole and Francisco Asturias, at The Scripps Research Institute, La Jolla. X-ray crystallographic analyses are performed by collaboration with Dr. Udo Oppermann and his colleagues at the Structural Genomics Consortium in Oxford, England.

V. Funding
The Smith group is supported by grants DK-16073 and GM-69717 from the National Institutes of Health.

 

VI. References

  1. Asturias, F. J., Chadick, J. Z., Cheung, I. K., Stark, H., Witkowski, A., Joshi, A. K., and Smith, S. (2005) Nat. Struct. Mol. Biol. 12(3), 225-232

  2. Maier, T., Jenni, S., and Ban, N. (2006) Science 311(5765), 1258-1262

  3. Pemble, C. W. t., Johnson, L. C., Kridel, S. J., and Lowther, W. T. (2007) Nat Struct Mol Biol 14(8), 704-709
  4. Bunkoczi, G., Pasta, S., Joshi, A., Wu, X., Kavanagh, K. L., Smith, S., and Oppermann, U. (2007) Chem Biol 14(11), 1243-1253
  5. Bunkoczi, G., Kavanagh, K., Hozjan, V., Rojkova, A., Wu, X., Arrowsmith, C., Edwards, A., Sundstrom, M., Weigelt, J., Smith, S., and Oppermann, U. (2007) Structure of the MAT domain of human FAS. In. PDB, 2JFD
  6. Rangan, V. S., Joshi, A. K., and Smith, S. (2001) Biochemistry 40(36), 10792-10799.
  7. Joshi, A. K., Rangan, V. S., Witkowski, A., and Smith, S. (2003) Chem. Biol. 10(2), 169-173
  8. Witkowski, A., Ghosal, A., Joshi, A. K., Witkowska, H. E., Asturias, F. J., and Smith, S. (2004) Chem. Biol. 11(12), 1667-1676
  9. Witkowski, A., Joshi, A. K., Rangan, V. S., Falick, A. M., Witkowska, H. E., and Smith, S. (1999) J Biol Chem 274(17), 11557-11563
  10. Pasta, S., Witkowski, A., Joshi, A. K., and Smith, S. (2007) Chem Biol 14(12), 1377-1385
  11. Keatinge-Clay, A. T., and Stroud, R. M. (2006) Structure 14(4), 737-748
  12. Zhang, L., Joshi, A. K., and Smith, S. (2003) J Biol Chem 278, 40067-40074
  13. Zhang, L., Joshi, A. K., Hofmann, J., Schweizer, E., and Smith, S. (2005) J Biol Chem 280(13), 12422-12429
  14. Autio, K. J., Kastaniotis, A. J., Pospiech, H., Miinalainen, I. J., Schonauer, M. S., Dieckmann, C. L., and Hiltunen, J. K. (2007) Faseb J
  15. Miinalainen, I. J., Chen, Z. J., Torkko, J. M., Pirila, P. L., Sormunen, R. T., Bergmann, U., Qin, Y. M., and Hiltunen, J. K. (2003) J Biol Chem 278(22),
  16. Witkowski, A., Joshi, A. K., and Smith, S. (2007) J Biol Chem 282(19), 14178-14185
  17. Joshi, A. K., Zhang, L., Rangan, V. S., and Smith, S. (2003) J Biol Chem 278(35), 33142-33149


Revised: Tuesday, January 24, 2012 11:49 AM

 

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