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3 Stumpf, P.K. and James, A.T. (1963) Biochim. Biophys. Acta 70 ... 5 Harington, A., Herbert, C.J., Tung, B., Getz, G.S. and Slonimski, P.P. (1993). Mol. Microbiol.
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Mitochondrial fatty acid synthesis and maintenance of respiratory competent mitochondria in yeast J.K. Hiltunen1 , F. Okubo, V.A.S. Kursu, K.J. Autio and A.J. Kastaniotis Biocenter Oulu and Department of Biochemistry, P.O.B. 3000, University of Oulu, FIN-90014 Oulu, Finland

Abstract Mitochondrial FAS (fatty acid synthesis) of type II is a widely conserved process in eukaryotic organisms, with particular importance for respiratory competence and mitochondrial morphology maintenance in Saccharomyces cerevisiae. The recent characterization of three missing enzymes completes the pathway. Etr1p (enoyl thioester reductase) was identified via purification of the protein followed by molecular cloning. To study the link between FAS and cell respiration further, we also created a yeast strain that has FabI enoylACP (acyl-carrier protein) reductase gene from Escherichia coli engineered to carry a mitochondrial targeting sequence in the genome, replacing the endogenous ETR1 gene. This strain is respiratory competent, but unlike the ETR1 wild-type strain, it is sensitive to triclosan on media containing only non-fermentable carbon source. A colony-colour-sectoring screen was applied for cloning of YHR067w/RMD12, the gene encoding mitochondrial 3-hydroxyacyl-ACP dehydratase (Htd2/Yhr067p), the last missing component of the mitochondrial FAS. Finally, Hfa1p was shown to be the mitochondrial acetyl-CoA carboxylase.

Background The FAS (fatty acid synthesis) type II pathway has been typically associated with bacterial FAS and the components involved in the Escherichia coli FAS pathway have been extensively characterized [1,2]. This has not been the case with eukaryotic FAS type II, even though it has been known for decades that plant plastids harbour a FAS activity distinct from the cytosolic FAS [3]. The uncovering of a mitochondrial type FAS pathway in Saccharomyces cerevisiae and Neurospora crassa [4–6] is a comparatively recent achievement and the fact that this pathway exists has only been slowly creeping into the realm of common knowledge [7]. As evidence is mounting which suggests that this process is not only taking place in the mitochondria of fungi, but is indeed conserved throughout eukaryotes [8–11], more interest in this matter has been generated. Mitochondrial FAS follows the prokaryotic-dissociated type II pathway and several enzymes of this pathway in S. cerevisiae (Scheme 1) show great similarity to corresponding E. coli proteins (Table 1). Only enoyl reductase and dehydratase have resisted identification by bioinformatics approaches. Functional mitochondrial FAS is a prerequisite for proper mitochondrial function in yeast. Deletion of any of the known genes coding for mitochondrial FAS genes results in a respiratory deficient pet phenotype in yeast and these mutants contain only Key words: acetyl-CoA carboxylase (ACC), 2-enoyl-thioester reductase, 3-hydroxyacyl-thioester dehydratase, mitochondrial fatty acid synthesis, respiratory competent mitochondrion, yeast. Abbreviations used: ACC, acetyl-CoA carboxylase; ACP, acyl-carrier protein; ARS, autonomously replicating sequence; Etr1p, enoyl thioester reductase; FAS, fatty acid synthesis; MFE-2, multifunctional enzyme type 2; Mrf1 p, mitochondrial respiratory factor 1; ORF, open reading frame. 1 To whom correspondence should be addressed (email kalervo.hiltunen@oulu.fi).

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small rudimentary mitochondria lacking spectrally detectable cytochromes. Some of these deletion mutants have been reported to experience mitochondrial DNA loss [4,6,12]. That it is indeed a fatty acid that is required to maintain mitochondrial function was demonstrated by the isolation of a mutation in the peroxisomal Faa2p fatty-acyl-CoA ligase that suppressed the respiratory deficient phenotype of a cem1 mutation [13]. This suppressor allele, dubbed FAM1-1, resulted in mislocalization of the fatty-acyl-CoA ligase to the mitochondria. The cause for the suppression effect could only be explained by the ability of the FAM1-1 mutant gene product to introduce an external fatty acid into the mitochondria, bypassing the need for endogenous mitochondrial FAS. It is remarkable that the known eukaryotic FAS and breakdown systems are strongly compartmentalized. General FAS takes place in the cytosol or mitochondria, elongation to very long-chain fatty acids occurs in the endoplasmic reticulum, while breakdown of fatty acid is restricted to peroxisomes in yeast. Higher eukaryotes also break down fatty acids in the mitochondria, but a metabolic rather than physical compartmentalization is still maintained in this case, as synthesis occurs as an ACP (acyl-carrier protein) adduct, while CoA-activated fatty acids are the substrate for βoxidation.

Enoyl thioester reductase When working on NADPH-dependent reductases from Candida tropicalis, we identified a novel mitochondrial reductase belonging to the medium-chain alcohol dehydrogenase/reductase superfamily [14]. The enzyme catalysed the reduction of the trans-2 double bonds in the enoyl thioesters

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Scheme 1 Reactions of the mitochondrial FAS type II in S. cerevisiae X denotes either CoA or ACP, as it is not known which activated form of acetic acid is used for priming.

and the enzyme was named Etr1p (enoyl thioester reductase). A database search revealed Mrf1 p (mitochondrial respiratory factor 1) from S. cerevisiae as the closest homologue. Mrf1 p was originally isolated as a protein binding to a singlestranded core sequence of ARS (autonomously replicating

sequence) and the protein was reported to be localized to the nucleus [15]. The mrf1 strain had lost mitochondrial cytochromes, was respiratory deficient and was unable to grow on non-fermentable carbon sources. When we reinvestigated the localization, it was found that most of the Mrf1 p was localized to the mitochondria. The recombinant purified Mrf1 p was also an enzymatically active protein catalysing an NADPH-dependent reduction of 2-enoyl thioesters. These observations raised the questions of whether mitochondrial or nuclear localizations of Mrf1 p were required for the maintenance of respiratory competent mitochondria and what role was played by the binding of the Mrf1 p to an ARS sequence or enzyme activity. When the nucleotide binding properties of Etr1p were tested by surface plasmon resonance method, the Etr1/Mrf1 p from S. cerevisiae bound to the ARS from S. cerevisiae, but C. tropicalis Etr1p did not, although the expression of any of these proteins rescued the respiratory-deficient phenotype of mrf1 . The crystal structure of Etr1p from C. tropicalis was solved and Tyr79 (Tyr73 ) in C. tropicalis (in S. cerevisiae) was identified as an amino acid residue interacting with NADP(H) [16]. Replacement of this tyrosine residue by asparagine via site-directed mutagenesis resulted in approx. 0.1% of Etr1p catalytic activity remaining, although the protein was properly folded. These variants failed to rescue the respiratory-deficient phenotype of the null mutant strains, indicating that enzymatic activity is required for complementation.

Table 1 Components of mitochondrial FAS type II in S. cerevisiae Systematic name

Protein/molecular function

FAS HFA1/YMR207C MCT1/YOR221C

Bacterial orthologue or analogous protein

Reference

Mitochondrial ACC

E. coli AccA/carboxyltransferase subunit

[24]

Predicted malonyl-CoA:ACP transferase/ putative component of a type-II

E. coli FabD/malonyl-CoA:ACP-transferase

[6]

E. coli AcpS/ACP synthase

[5]

E. coli FabG/β-keto-acyl-ACP reductase

[6]

mitochondrial fatty acid synthase that produces intermediates for phospholipid remodelling CEM1/YER061C

Homology with β-keto-acyl synthases; protein homologous to β-keto-acyl synthase/3-oxoacyl-[ACP]

OAR1/YKL055C

synthase activity Mitochondrial 3-oxoacyl-[ACP] reductase/ may comprise a type II

HTD2/YHR067W

mitochondrial fatty acid synthase Mitochondrial 3-hydroxyacyl-thioester dehydratase

E.coli FabA/β-OH-decanoyl-thioesterdehydratase

[21]

2-Enoyl thioester reductase Mitochondrial matrix ACP

E. coli FabI/enoyl-ACP reductase acpP/ACP

[14] [12]

Phosphopantetheine:protein transferase/ activates mitochondrial ACP (Acp1p) by

Brevibacterium ammoniagenes Ppt1 phosphopantetheine:protein transferase

[30]

ETR1/YBR026C ACP1/YKL192C Others PPT2/YPL148C

phosphopantetheinylation

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The role of subcellular localization of the Etr1p on the cellular respiratory competence was studied by expressing Etr1p variants either with an N-terminally linked nuclear localization signal from 40 simian-virus-40 large T-antigen or truncated for an N-terminal mitochondrial targeting sequence [14]. Neither of them could complement mrf1 null strains, although they were well expressed as shown by Etr1p enzyme activity measurements. We therefore concluded that mitochondrially targeted active Etr1p was indispensable for the growth of the yeast cells on non-fermentable carbon sources. This conclusion was supported further by restoring the growth of the etr1 strain by expressing mitochondrially targeted FabI, the Etr1p from E. coli. FabI is a homotetrameric protein of short-chain alcohol dehydrogenase/reductase superfamily and does not bear any structural resemblance to the Etr1p dimer. The FabI is a target of triclosan inhibition, whereas the yeast Etr1 is not. Subsequent experiments have demonstrated that replacing the yeast endogenous Etr1p by the bacterial FabI introduces a triclosan sensitivity to yeast under growth on non-fermentable carbon sources [17].

The mitochondrial 3-hydroxyacyl-ACP dehydratase The isolation of mitochondrial 3-hydroxyacyl-ACP dehydratase was hampered by similar problems such as the isolation of the enoyl reductase, as database searches using known prokaryotic dehydratases or the hydratase-2 footprint [18] do not produce any good candidates for this function ([6], and K.J. Autio, unpublished work). Although the activity can be measured in yeast mitochondrial extracts, it is lost during the attempt of purification (K.J. Autio, unpublished work). As mitochondrially localized FabI complements the etr1 deletion phenotype [14], we reasoned that a similar construct employing a mitochondrially localized variant of an E. coli hydroxyacyl-ACP dehydratase (FabA) could be used to find the corresponding yeast mitochondrial enzyme in a genetic ‘cloning by function’ approach [19]. In essence, a redundancy of function is created this way, which allows for a mutation to be generated in the endogenous dehydratase without causing lethality on non-fermentable carbon source, as the function can still be provided by the artificial chimaera. These mutations, however, render the yeast cells dependent on the plasmid carrying the chimaera on non-fermentable carbon source media. This dependence can be screened by using a colony-colour sectoring based assay [20]. Using this approach, we identified mutants unable to lose our plasmid construct and demonstrated the mutations to be in the yeast ORF (open reading frame) YHR067w, which we subsequently named HTD2 (3-hydroxyacyl-thioester dehydratase of type 2) [21]. A deletion of this ORF caused respiratory deficiency and loss of cytochromes, phenotypes identical with those of other mitochondrial FAS mutants. We demonstrated that the protein is localized to the mitochondria and showed that a mutation in this gene resulted in abnormal mitochondrial morphology, while overexpression of Htd2p caused mitochondrial expansion similar to overexpression of Etr1p. Induced expression of the protein also  C 2005

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resulted in increased mitochondrial hydratase-2 activity and an alignment to hydratase-2 protein portions of MFE-2 (multifunctional enzyme type 2) indeed revealed the presence of a hydratase-2 motif in Htd2p. Similar to the MFE-2 hydratase-2 domain [22], the protein is about twice the size of prokaryotic hydratase-2 proteins, suggesting it to be the result of a partial gene duplication event. This is supported by the observation that the Htd2p N- and C-terminal portions share a 44% identity on the nucleotide level, even though this only translates into 9% identity of amino acid residues. Surprisingly, searching the human protein database did not result in the identification of a close homologue and this enzyme will therefore have to be isolated by genetic or biochemical means.

The mitochondrial ACC (acetyl-CoA carboxylase) For a long time, the origin of mitochondrial malonyl-CoA was uncertain. Hoja et al. [23] initially proposed that the essential yeast cytosolic ACC1 was responsible for the production of malonyl-CoA, but such a role was unlikely because the mitochondrial membranes are probably impermeable for this molecule. The same group recently reported the yeast Hfa1 protein to be the enzyme carrying out this reaction [24]. It had been known for a while that HFA1 was highly similar to ACC1 [25], but it was unclear for a long time whether the former encoded a functional enzyme, as the homology of HFA1 to ACC1 extends dozens of 5 base pairs of the reported initiation ATG of HFA1. However, it was shown subsequently that a disruption of HFA1 causes a respiratory-deficient phenotype very similar to mitochondrial FAS gene deletions. Remarkably, at least 72 codons upstream of the predicted start codon were required for complementation of the null mutant [24]. These 72 triplets code for the additional homologous portion of Hfa1p to Acc1p and also for a putative mitochondrial targeting sequence that appears to be functional, as the protein was demonstrated to be localized to the mitochondria. A construct lacking the mitochondrial targeting sequence did not complement the hfa1 disruption mutant. In contrast, the same construct was capable of complementing a deletion mutation of ACC1, which cannot be accomplished with native Hfa1p. It is possible that HFA1 represents a rare case, where yeast uses an initiation codon other than AUG for the start of translation. Other more exotic explanations could be programmed frame shifting, which occurs in a few yeast genes [26,27], or the mRNA editing. An enzyme capable of carrying out this latter process has been found in yeast, although no substrate has been identified yet [28]. The particulars of HFA1 expression warrant further investigation, as it is tempting to speculate that the entire pathway may be controlled by the abundance of Hfa1p, which carries out the initial reaction of the FAS pathway, and regulation of Hfa1p production could occur via translation start site selection. A first step to solve the puzzle of HFA1 expression would be the identification of the in vivo translation start site.

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Products of mitochondrial FAS Plants, fungi and humans have mitochondrial lipoic acid synthase. It has been shown that deletion of mitochondrial ACP1 in yeast results in strongly reduced lipoic acid content [4]. This observation suggests that the physiological function of mitochondrial FAS is to provide octanoyl-ACP to be used for lipoic acid synthesis. However, comparison of the phenotypes of yeast strains inactivated in the FAS type II or lipoic acid synthesis shows that the decreased lipoic acid is not sufficient to explain all the mitochondrial dysfunction observed in the yeast strains with disrupted mitochondrial FAS. In line with the idea that mitochondrial FAS is generating more than C8 end products, the C16 and C18 acyl groups have been found to be linked to ACP in mitochondria from plant and N. crassa [29]. Furthermore, the recent studies on kinetics of isolated FAS enzymes reveal that human CEMp (KAS; ketoacyl synthase) can catalyse a condensation reaction at least up to C14 –C16 substrates [11] and the ETR1p from human mitochondria accepts C16 substrates [8]. All these observations suggest that the mitochondrial FAS can generate fatty acids longer than C8 in chain length.

Future Analysis of the mammalian genome confirms that mammalian cells also contain bacterial FAS type II enzymes in mitochondria, in addition to cytosolic FAS type I. The occurrence of both FAS and degradation in mitochondria appears to be a surprise. However, the conservation of the FAS type II during evolution suggests an indispensable physiological role in eukaryotic mitochondria. The role could be linked, for instance, to mitochondrial phospholipid metabolism, mitochondrial protein acylation or generation of currently unknown specific lipid molecules. Other topics of further studies will be the flow of information from mitochondria to nucleus under conditions of disturbed flux through the mitochondrial FAS and ultimately identification of inborn errors of FAS systems in humans. Our original work was supported by grants from the Academy of Finland and Sigrid Juselius Foundation.

References 1 Rock, C.O. and Cronan, J.E. (1996) Biochim. Biophys. Acta 1302, 1–16 2 White, S.W., Zheng, J., Zhang, Y.M. and Rock, C.O. (2005) Annu. Rev. Biochem., doi:10.1146/annurev.biochem.74.082803.133524

3 Stumpf, P.K. and James, A.T. (1963) Biochim. Biophys. Acta 70, 20–32 4 Brody, S., Oh, C., Hoja, U. and Schweizer, E. (1997) FEBS Lett. 408, 217–220 5 Harington, A., Herbert, C.J., Tung, B., Getz, G.S. and Slonimski, P.P. (1993) Mol. Microbiol. 9, 545–555 6 Schneider, R., Brors, B., Burger, F., Camrath, S. and Weiss, H. (1997) Curr. Genet. 32, 384–388 7 Surolia, N., RamachandraRao, S.P. and Surolia, A. (2002) BioEssays 24, 192–196 8 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, 20154–20161 9 Joshi, A.K., Zhang, L., Rangan, V.S. and Smith, S. (2003) J. Biol. Chem. 278, 33142–33149 10 Zhang, L., Joshi, A.K. and Smith, S. (2003) J. Biol. Chem. 278, 40067–40074 11 Zhang, L., Joshi, A.K., Hofmann, J., Schweizer, E. and Smith, S. (2005) J. Biol. Chem. 280, 12422–12429 12 Schneider, R., Massow, M., Lisowsky, T. and Weiss, H. (1995) Curr. Genet. 29, 10–17 13 Harington, A., Schwarz, E., Slonimski, P.P. and Herbert, C.J. (1994) EMBO J. 13, 5531–5538 14 Torkko, J.M., Koivuranta, K.T., Miinalainen, I.J., Yagi, A.I., Schmitz, W., Kastaniotis, A.J., Airenne, T.T., Gurvitz, A. and Hiltunen, K.J. (2001) Mol. Cell. Biol. 21, 6243–6253 15 Yamazoe, M., Shirahige, K., Rashid, M.B., Kaneko, Y., Nakayama, T., Ogasawara, N. and Yoshikawa, H. (1994) J. Biol. Chem. 269, 15244–15252 16 Airenne, T.T., Torkko, J.M., Van den plas, S., Sormunen, R.T., Kastaniotis, A.J., Wierenga, R.K. and Hiltunen, J.K. (2003) J. Mol. Biol. 327, 47–59 17 Torkko, J.M., Koivuranta, K.T., Kastaniotis, A.J., Airenne, T.T., Glumoff, T., Ilves, M., Hartig, A., Gurvitz, A. and Hiltunen, J.K. (2003) J. Biol. Chem. 278, 41213–41220 18 Qin, Y.M., Haapalainen, A.M., Kilpelainen, S.H., Marttila, M.S., Koski, M.K., Glumoff, T., Novikov, D.K. and Hiltunen, J.K. (2000) J. Biol. Chem. 275, 4965–4972 19 Kranz, J.E. and Holm, C. (1990) Proc. Natl. Acad. Sci. U.S.A. 87, 6629–6633 20 Bender, A. and Pringle, J.R. (1991) Mol. Cell. Biol. 11, 1295–1305 21 Kastaniotis, A.J., Autio, K.J., Sormunen, R.T. and Hiltunen, J.K. (2004) Mol. Microbiol. 53, 1407–1421 22 Koski, M.K., Haapalainen, A.M., Hiltunen, J.K. and Glumoff, T. (2003) Acta Crystallogr. D Biol. Crystallogr. 59, 1302–1305 23 Hoja, U., Wellein, C., Greiner, E. and Schweizer, E. (1998) Eur. J. Biochem. 254, 520–526 24 Hoja, U., Marthol, S., Hofmann, J., Stegner, S., Schulz, R., Meier, S., Greiner, E. and Schweizer, E. (2004) J. Biol. Chem. 279, 21779–21786 25 Kearsey, S.E. (1993) DNA Seq. 4, 69–70 26 Morris, D.K. and Lundblad, V. (1997) Curr. Biol. 7, 969–976 27 Asakura, T., Sasaki, T., Nagano, F., Satoh, A., Obaishi, H., Nishioka, H., Imamura, H., Hotta, K., Tanaka, K., Nakanishi, H. et al. (1998) Oncogene 16, 121–130 28 Dance, G.S., Beemiller, P., Yang, Y., Mater, D.V., Mian, I.S. and Smith, H.C. (2001) Nucleic Acids Res. 29, 1772–1780 29 Mikolajczyk, S. and Brody, S. (1990) Eur. J. Biochem. 187, 431–437 30 Stuible, H.P., Meier, S., Wagner, C., Hannappel, E. and Schweizer, E. (1998) J. Biol. Chem. 273, 22334–22339

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