Molecular characterization of peroxisomal multifunctional...

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MOLECULAR CHARACTERIZATION OF PEROXISOMAL MULTIFUNCTIONAL 2-ENOYL-COA HYDRATASE 2/(3R)- HYDROXYACYL-COA DEHYDROGENASE (MFE TYPE 2) FROM MAMMALS AND YEAST YONG-MEI QIN Biocenter Oulu 1999

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MOLECULAR CHARACTERIZATION OF PEROXISOMAL MULTIFUNCTIONAL 2-ENOYL-COA HYDRATASE 2/(3R)-HYDROXYACYL-COA DEHYDROGENASE (MFE TYPE 2) FROM MAMMALS AND YEAST

YONG-MEIQIN

Biocenter Oulu

1999

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OULUN YLIOPISTO, OULU 1999

MOLECULAR CHARACTERIZATION OF PEROXISOMAL MULTIFUNCTIONAL 2-ENOYL-COA HYDRATASE 2/(3R)-HYDROXYACYL-COA DEHYDROGENASE (MFE TYPE 2) FROM MAMMALS AND YEAST

YONG-MEI QIN

Academic Dissertation to be presented with the assent of the Faculty of Medicine, University of Oulu, for public discussion in Raahensali (Auditorium L 10), Linnanmaa, on August 13th, 1999, at 12 noon.

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Copyright © 1999Oulu University Library, 1999

OULU UNIVERSITY LIBRARYOULU 1999

ALSO AVAILABLE IN PRINTED FORMAT

Manuscript received 23.6.1999Accepted 24.6.1999

Communicated by Professor Pekka MäenpääProfessor Juhani Syväoja

ISBN 951-42-5337-X(URL: http://herkules.oulu.fi/isbn951425337X/)

ISBN 951-42-5318-3ISSN 0355-3221 (URL: http://herkules.oulu.fi/issn03553221/)

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Qin, Yong-M ei, M olecular characterization of peroxisomal multifunctional2-enoyl-CoA hydratase 2/(3R)-hydroxyacyl-CoA dehydrogenase (M FE type 2)from mammals and yeastBiocenter Oulu and Departments of Biochemistry and Medical Biochemistry, Universityof Oulu, FIN-90570 Oulu, FinlandActa Univ. Oul. D 537, 1999Oulu, Finland(Manuscript received 23 June 1999)

Abstract

Fatty acid degradation in living organisms occurs mainly via the β-oxidation pathway. Whenthis work was started, it was known that the hydration and dehydrogenation reactions in mammalianperoxisomal β-oxidation were catalyzed by only multifunctional enzyme type 1 (MFE-1; ∆2-∆3-enoyl-CoA isomerase/2-enoyl-CoA hydratase 1/(3S)-hydroxyacyl-CoA dehydrogenase) via the S-specific pathway, whereas in the yeast peroxisomes via the R-specific pathway by multifunctionalenzyme type 2 (MFE-2; 2-enoyl-CoA hydratase 2/(3R)-hydroxyacyl-CoA dehydrogenase).

The work started with the molecular cloning of the rat 2-enoy-CoA hydratase 2 (hydratase 2).The isolated cDNA (2205 bp) encodes a polypeptide with a predicted molecular mass of 79.3 kDa,which contains a potential peroxisomal targeting signal (AKL) in the carboxyl terminus. Thehydratase 2 is an integral part of the cloned polypeptide, which is assigned to be anovel mammalianperoxisomal MFE-2.

The physiological role of the mammalian hydratase 2 was investigated with the recombinanthydratase 2 domain derived from rat MFE-2. The protein hydrates a physiological intermediate(24E)-3α, 7α, 12α-trihydroxy-5β-cholest-24-enoyl-CoA to (24R, 25R)-3α, 7α, 12α, 24-tetrahydroxy-5β-cholestanoyl-CoA in bile acid synthesis.

The sequence alignment of human MFE-2 with MFE-2(s) of different species reveals 12conserved protic amino acid residues, which are potential candidates for catalysis of the hydratase2. Each of these residues was replaced by alanine. Complementation of Saccharomyces cerevisiaefox-2 (devoid of endogenous MFE-2) with human MFE-2 provided a model system for examing thein vivo function of the variants. Two protic residues, Glu366 and Asp510, of the hydratase 2 domainof human MFE-2 have been identified and are proposed to act as a base and an acid in catalysis.

Mammalian MFE-2 has a (3R)-hydroxyacyl-CoA dehydrogenase domain, whereas the yeastMFE-2 has two dehydrogenase domains, A and B. The present work, applying site-directedmutagenesis to dissect the two domains, shows that the growth rates of fox-2 cells expressing asingle functional domain are lower than those of cells expressing S. cerevisiae MFE-2. Kineticexperiments with the purified proteins demonstrate that domain A is more active than domain B incatalysis of medium- and long-chain (3R)-hydroxyacyl-CoA, whereas domain B is solelyresponsible for metabolism of short-chain substrates. Both domains are required when yeast cellsutilize fatty acids as the carbon source.

Keywords: β-oxidation, fatty acid, bile acid synthesis, enzymology

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Acknowledgements

The present study was carried out at Biocenter Oulu, the Department of Biochemistry andDepartment of Medical Biochemistry, University of Oulu during the years 1993-1999. Theaccomplishment of this work would be impossible without the help of so many people.

I wish to express my deepest gratitude to my supervisor, Professor Kalervo Hiltunen,for his hospitality to give me the opportunity to do my thesis work as a member of hisresearch group. His everlasting enthusiasm, open mind and never-failing spirit in sciencehave indefinitely encouraged me to fulfil l this work. I also wish to thank Professors TainaPihlajaniemi, Ilmo Hassinen, Kari Kivirikko , Raili Myllylä, Seppo Vainio, Rik Wierengaand Johan Kemmink for providing excellent working facilities and creating a stimulatingatmosphere in our weekly seminars at the both Departments.

I wish to acknowledge Professors Juhani Syväoja and Pekka Mänepää for their criticalappraisal of the manuscript, and Dr. Sidney Higley for her careful revision of thelanguage. Dmitry Novikov, Ph.D., deserves my warmest thanks for his critical view on thesubject and a pleasant collaboration. I am indebted to my co-authors, Ulf Hellman, Ph.D.,Dean Cuebas, Ph.D., Werner Schmitz, Ph.D., Matti Poutanen, Ph.D., Tuomo Glumoff,Ph.D., Kirsi Siivari, M.Sc., Antti Haapalainen, M.Sc, and Mari Marttila, B.Sc. for theirtremendous contribution to this work. Former and present members of "KH team",especially, Sirpa Filppula, Heli Helander, Kari Koivuranta, Tiila-Riikka Kiema, TanjaKokko, Ahmed Yagi, and Tiina Kotti are appreciated for their friendship and fruitfuldiscussion during and after the work.

I wish to extend my sincere thanks to whole staff of the Departments. I am muchobliged to Aila Holappa, Irma Vuoti, Anja Mattila, Raija Pietilä, Tanja Kokko, VilleRatas and Marika Kamps for their skillful technical assistance, Anna-Leena Hietajärvi formaking our precious substrates, Eeva-Liisa Stefanius for synthesizing oligonucleotides,Maire Jarva for doing the automatic sequencing, Jaakko Keskitalo and Kyösti Keränen fordeveloping the photos and fixing Schimadzu spectrophotometer. Special thanks also go toSeppo Kilpeläinen, M.Sc., for setting up an excellent computer network, Auli Kinnunen,Marja-Leena Kivelä, Anneli Kaattari and Tuula Koret for their kind help with secretarialand many practical tasks.

I would like to thank my father, mother and brothers for encouraging me to studyabroad and their support in each step of my life. I am also grateful to my parents and sisterin-law for their help whenever I need. Finally, I owe my heartfelt thanks to my husband

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Qiang for his support in all matters of life and our daughter Mandi who always keeps mein touch with the true life.

This work was financially supported by grants from the Sigrid Jusélius foundation,Academy of Finland and University of Oulu foundation.

Oulu, June 1999 Yong-Mei Qin

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Abbreviations

ABC ATP-binding cassetteX-ALD X-linked adrenoleukodystrophyATP adenosine triphosphatebp base pair(s)cDNA complementary DNACoA coenzyme AFAD flavine adenine dinucleotideFADH2 flavine adenine dinucleotide (reduced)LCAD long-chain acyl-CoA dehydrogenasekDa kilodalton(s)MFE multifunctional enzymeMFE-1 peroxisomal multifunctional ∆3-∆2-enoyl-CoA isomerase/2-enoyl-CoA

hydratase 1/(3S)-hydroxyacyl-CoA dehydrogenaseMFE-2 peroxisomal multifunctional 2-enoyl-CoA hydratase 2/(3R)-

hydroxyacyl-CoA dehydrogenasemRNA messenger RNANAD+ nicotinamide adenine dinucleotide (oxidized)NADH nicotinamide adenine dinucleotide (reduced)NADPH nicotinamide adenine dinucleotide phosphate (reduced)PCR polymerase chain reactionperoxin protein involved in peroxisome biogenesispI isoelectric pointPPAR peroxisomal proliferator activated receptorPPRE peroxisomal proliferator responsive elementPTS peroxisomal targeting signalRACE rapid amplification of cDNA endsRXR retinoid X receptorSDR short-chain alcohol dehydrogenase/reductase familySDS-PAGE sodium dodecyl sulphate polyacrylamide gel electrophoresisTHCA 3α, 7α, 12α-trihydroxy-5β-cholestanoic acidXaa any amino acid

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List of original articles

This thesis is based on the following articles, which are referred to the text by their Romannumerals:

I Qin Y-M, Poutanen MH, Helander HM, Kvist A-P, Siivari KM, Schmitz W,Conzelmann E, Hellman U & Hiltunen JK (1997) Peroxisomal multifunctional enzyme ofβ-oxidation metabolizing D-3-hydroxyacyl-CoA esters in rat liver: molecular cloning,expression and characterization. Biochem J 321:21-28.

II Qin Y-M, Haapalainen AM, Conry D, Cuebas DA, Hiltunen JK & Novikov DK(1997) Recombinant 2-enoyl-CoA hydratase derived from rat peroxisomal multifunctionalenzyme 2: role of the hydratase reaction in bile acid synthesis. Biochem J 328:377-382.

III Qin Y-M, Marttila MS, Haapalainen AM, Glumoff T, Novikov DK & HiltunenJK Human peroxisomal multifunctional enzyme type 2: site-directed mutagenesis studiesshow the importance of two protic residues for 2-enoyl-CoA hydratase 2 activity.Submitted.

IV Qin Y-M, Marttila, MS, Haapalainen AM, Siivari KM, Glumoff T & Hiltunen JKYeast peroxisomal multifunctional enzyme: (3R)-hydroxyacyl-CoA dehydrogenasedomains A and B are required for optimal growth on oleic acid. Submitted.

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Contents

AbstractAcknowledgementsAbbreviationsList of original articles1. Introduction………………………………………………………………………….. 122. Review of the literature………………………...……………………………………. 14

2.1. Fatty acids……………………………………………………………………… 142.1.1. Uptake and transport of fatty acids into the mammalian cells.………… 142.1.2. Pathways for fatty acid oxidation.……………………………………… 15

2.2. Peroxisomes……………………………………………………………………. 162.2.1. Peroxisomal proliferation in mammalian cell..………………………… 162.2.2. Induction of genes encoding peroxisomal proteins in yeast…………….182.2.3. Activation and transportation of fatty acids into peroxisomes…………. 182.2.4. The peroxisomal β-oxidation spiral……………………………………. 19

2.2.4.1. Acyl-CoA oxidation……………………………...…………… 192.2.4.2. Hydration of trans-2-enoyl-CoA and subsequent

dehydrogenation…………………………………………..…. 212.2.4.3. Cleavage of 3-ketoacyl-CoA…………………………………. 23

2.2.5. Oxidation of the cholesterol side chain………………………………… 242.3. Auxiliary enzymes involved in β-oxidation …..………………………………. 26

2.3.1. 2, 4-Dienoyl-CoA reductase……………………………………………. 262.3.2. ∆3-∆2-Enoyl-CoA isomerase………………………………………...… 262.3.3. ∆3,5-∆2,4-Dienoyl-CoA isomerase……………………………………..282.3.4. 2-Methylacyl-CoA racemase……………………………...……….…… 29

2.4. Multifunctional enzymes (MFEs) of β-oxidation……………………………… 292.4.1. Bacterial β-oxidation multienzyme complex……………….………….. 302.4.2. Fungal multifunctional enzymes……………………………………….. 302.4.3. Plant multifunctional enzymes.………………………………………… 322.4.4. Mammalian multifunctional enzymes………………………………….. 32

3. Outline of the present study……………………………………………..…………… 344. Materials and methods……………………………………………………..………… 35

4.1. Biological resources……………………………………………………….…… 35

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4.2. Culture media……………..…………………………………………………... 354.3. Synthesis of substrates………..……………………….……...…………….…. 36

4.3.1. Synthesis of straight-chain fatty acyl-CoA esters……………………… 364.3.2. Synthesis of 2-pristenic acid..………………………………………….. 364.3.3. Synthesis of 3α, 7α, 12α, 24ξ-tetrahydroxy-5β-cholestanoic acid

and (24E, 24Z)-3α, 7α, 12α-trihydroxy-5β-cholest-24-enoic acid…… 364.4. Enzyme assays………..…………….…………………………………………. 36

4.4.1. Enzymes of fatty acidβ-oxidation...………………………………….... 364.4.2. Hydration and dehydration between (24E)-3α, 7α, 12α-

trihydroxy-5β-cholest-24-enoyl-CoA and 24-hydroxy,3α, 7α, 12α-trihydroxy-5β-cholest-24-enoyl-CoA.…………………... 37

4.4.3. Enzyme kinetics………………………………….…………………….. 374.5. Isolation of peroxisomes…………………….....…………………………….... 374.6. Isolation and characterization of cDNA clones...…..……………………….… 384.7. Northern blot analysis………..……………………………………………….. 384.8. Expression of recombinant proteins.………………………………..………… 38

4.8.1. Expression of recombinant proteins inE. coli………...………….……. 384.8.2. Expression of recombinant proteins inP. pastoris…...………………... 38

4.9. Site-directed mutagenesis………………..…………………..…………..……. 394.10. Protein purification…..……………………………………………………..… 394.11. Analysis of proteins…………………………………………………………... 394.12. Complementation ofS. cerevisiae fox-2with

MFE-2(s)and their variants………….……….………………………….…… 404.13. Immunoblotting………………………………………………………………. 404.14. Software……………………………………………………………………… 41

5. Results………………………………………………………………………….……. 425.1. A novel peroxisomal multifunctional enzyme type 2…..………………………425.2. Recombinant 2-enoyl-CoA hydratase 2 derived from rat MFE-2………..….…435.3. Site-directed mutagenesis of the 2-enoyl-CoA hydratase 2..………………….. 435.4. Analysis of the two catalytic domains of (3R)-hydroxyacyl-CoA

dehydrogenase of yeast peroxisomal MFE-2……….………………………… 436. Discussion………………………………………………………………………….… 46

6.1. A novel peroxisomal multifunctional enzyme type 2..…………………………466.2. Recombinant 2-enoyl-CoA hydratase 2 derived from rat MFE-2…...………... 476.3. Site-directed mutagenesis of the 2-enoyl-CoA hydratase 2…………………… 476.4. Analysis of the two catalytic domains of (3R)-hydroxyacyl-CoA

dehydrogenase of yeast peroxisomal MFE-2…………………..……….…….. 497. Conclusions…………………………….……………………………………….….…508. References……………………………….…………………………………………....51

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1.Introduction

The name peroxisome was proposed in 1965 by De Duve who discovered a subcellularcompartment containing oxidases and catalase that participate in the production andconsumption of hydrogen peroxide. This far, over 50 enzymes have been found to beinvolved in a variety of peroxisomal metabolic processes, including the β-oxidation offatty acids and fatty acid derivatives; β-oxidation of the side chain of cholesterol thatresults in the formation of bile acids; fatty acid elongation; synthesis of plasmalogen,cholesterol and dolichols; catabolism of amino acids, purines and polyamines; metabolismof glyoxylate; degradation of glucose via the hexose-monophosphate pathway;inactivation of reactive oxygen species and the process of α-oxidation (for a review, seeMannaerts & Van Veldhoven 1996).

The peroxisomal β-oxidation pathway in mammals was discovered by Lazarow and DeDuve in 1976. The co-existence of both mitochondrial and peroxisomal β-oxidation andthe consequences of β-oxidation deficiencies have revealed the significance of fatty acidoxidation in mammals (Kunau et al. 1988). Peroxisomal β-oxidation plays a key role inthe metabolism of polyunsaturated fatty acids (for a review, see Hiltunen et al.1996). The(2E)-enoyl-CoA ester is the only unsaturated intermediate in the β-oxidation spiral andtherefore, double bonds of unsaturated fatty acids mainly in the cis configurations requireadditional auxiliary enzymes in order to be removed.

The present work is concerned with the molecular characterization of the 2-enoyl-CoAhydratase 2 (hydratase 2) from rat liver that was previously found to participate in theepimerization of 3-hydroxyacyl-CoA ester between the R- and S-form (Hiltunen et al.1989, Smeland et al. 1989). According to the original hypothesis, this epimerization wassupposed to be involved in the β-oxidation of cis-2-enoyl-CoA esters (Stoffel et al. 1964)and at that time hydratase 2 was listed as an auxiliary enzyme of β-oxidation. Thephysiological function of hydratase 2 in yeast peroxisomes was elucidated to be in thecatabolism of (2E)-enoyl-CoA to a 3-keto complex via a (3R)-hydroxyacyl-CoAintermediate catalyzed by a multifunctional enzyme (MFE) possessing (3R)-hydroxyacyl-CoA dehydrogenase and hydratase 2 activities (Hiltunen et al. 1992). The processing of(2E)-enoyl-CoA to the 3-keto complex occurs in mammalian peroxisomes via a (3S)-hydroxyacyl-CoA intermediate. It is carried out by a peroxisomal multifunctional enzymewith three activities: (3S)-hydroxyacyl-CoA dehydrogenase, 2-enoyl-CoA hydratase 1

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(Osumi & Hashimoto 1979), and∆3-∆2-enoyl-CoA isomerase (Palosaari & Hiltunen1990). However, the function of hydratase 2 in mammals remainedunknown. The aim ofthis work was the molecular cloning of mammalian hydratase 2, identification of potentialcatalytic amino acid residues contributing to the hydration reaction, and functionalcomplementation of mammalian hydratase 2 inSaccharomyces cerevisiae. In order tofurther elucidate the function of peroxisomal MFE, yeast (3R)-hydroxyacyl-CoAdehydrogenase was also investigated in the present work.

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2. Review of the literature

2.1. Fatty acids

Fatty acids are simple lipids that have a hydrophilic carboxyl group at one end of ahydrocarbon chain. They are mainly classified as saturated or unsaturated fatty acids withmany important species of the latter form being in the cis-configuration. α-Linolenic(C18:3, n-3) and linoleic (C18:2, n-6) acids are known as essential fatty acids (EFAs), since theyare important for the well-being of individuals and must be present in the diet. EFAs serveseveral vital functions as nutrients, an energy source, structural components of cells andprecursors of prostanoids and leukotrienes. In polyunsaturated fatty acids, the doublebonds usually alternate between odd- and even-numbered carbon atoms and are separatedby a methylene group.

2.1.1. Uptake and transport of fatty acids into the mammalian cell

The mechanism of the transport of long-chain fatty acids across the cellular membrane hasbeen proposed as being either a passive or carrier-mediated transmembrane translocation.The passive mechanism is a non-saturable diffusion governed by the transmembranegradient of fatty acid concentration (De Grella & Light 1980). The carrier-mediatedprocess of migration of fatty acid through the plasma membrane is either mediated by aNa+/fatty acid co-transport system (Stremmel 1987) or by membrane-associated fatty acid-binding proteins (mFABPs). The intracellular transport of fatty acids from the plasmamembrane either to the sites of metabolic conversion or to the subcellular target isfacilitated by cytoplasmic fatty acid binding proteins (cFABPs) (for a review, see Glatz etal. 1997). cFABPs are abundant low-molecular-mass proteins (14-16 kDa), whose levelsare responsible for nutritional, endocrine and a variety of pathological states.

To date, several mFABPs have been identified and these are implicated intransmembrane transport of long-chain fatty acids. In general, three classes of mFABPshave been described, and these include fatty acid translocase (FAT; Abumrad et al. 1993),membrane-bound fatty acid-binding proteins (FABPpm; Stremmel et al. 1985) and fatty

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acid transport protein (FATP; Schaffer & Lodish, 1994). The expression of the genesencoding FAT and FATP is regulated by peroxisome proliferator activated receptors(PPARs), indicating that in addition to other PPAR function discussed below, peroxisomeproliferators influence the cellular uptake of fatty acids (Motojimaet al. 1998, Frohnertetal. 1999). InS. cerevisiae, the gene encoding a putative membrane-bound long-chain fattyacid transport protein (FAT1) was identified (Færgemanet al. 1997) and possessed verylong-chain acyl-CoA synthetase activity (see Section 2.2.4.; Watkinset al. 1998). FABPswere proposed to solubilize and compartmentalize fatty acid, to facilitate the cellularuptake and intracellular trafficking of fatty acids, as well as to modulate cell growth anddifferentiation (for a review, see Glatzet al. 1997).

2.1.2. Pathways for fatty acid oxidation

There are three different pathways for oxidizing fatty acids and they are described asα-,β-, andω-oxidation. Quantitatively, the major pathway isβ-oxidation which is located inboth mitochondria and peroxisomes of mammalian cell. In contrast, in lower eukaryotessuch as yeast and filamentous fungi, as well as in most plants,β-oxidation occurs solely inperoxisomes (Kunauet al. 1988). Fatty acidα-oxidation is a minor pathway that isinvolved in the catabolism of branched isoprenoid-derived 3-methyl substituted fattyacids, i.e. phytanic acid (3,7,11,15-tetramethylhexadecanoic acid). It includes fourreactions, namely activation, hydroxylation, release of formyl-CoA resulting in formationof pristanal followed by a dehydrogenation reaction yielding pristanic acid (Croeset al.1996). 2-Methyl branched fatty acid is then furtherβ-oxidized predominantly inperoxisomes (Scheperset al. 1990, Vanhoveet al. 1991). The importance ofα-oxidationin humans has been recognized by Refsum’s disease, which is characterized by anaccumulation of large amounts of phytanic acid. The intracellular site ofα-oxidation is notyet clear. Evidence suggests that it occurs either in peroxisomes (Mihaliket al. 1995,Croeset al. 1996), in mitochondria (Skjeldal & Stokke 1987, Watkins & Mihalik 1990),in the endoplasmic reticulum (ER) (Huanget al. 1992), or in both mitochondria andperoxisomes (Fingerhutet. al. 1993). Ordinary straight-chain fatty acids may also undergoω-oxidation, resulting in the formation of medium- and long-chain dicarboxylates (VanHoof et al. 1988). Monocarboxylic acids are firstω-hydroxylated to form ω-hydroxymonocarboxylic acids by a lauric acidω-hydroxylase (E.C. 1.14.14.1.). Theseintermediates can be eitherω-oxidized in mitochondria and peroxisomes or furtheroxidized to ω-oxomonocarboxylic acids and dicarboxylic acids by cytosolic alcoholdehydrogenase (E.C. 11.1.1.71) and aldehyde dehydrogenase (E.C. 1.2.1.3), respectively.On the other hand, dicarboxylic acids are activated to their CoA esters in microsomes andsubsequentlyβ-oxidized to succinate (for a review, see Osmundsenet al. 1991). Forxenobiotic fatty acids that can not beβ-oxidized, i.e. 3-thia fatty acids,ω-oxidation is themain catabolic pathway (for a review, see Skredeet al. 1997).

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2.2. Peroxisomes

A peroxisome is spherical or ovoid with a diameter of 0.1 to 1.5µm and consists of asingle membrane delimiting a granular matrix. Peroxisomes are versatile in mosteukaryotic cells, and depending on the organism, participate in a variety of metabolicprocesses, including lipid metabolism; H2O2-based respiration; oxidation of unsaturatedvery long-chain fatty acids, branched fatty acids and dicarboxylic acids; synthesis ofplasmalogens, cholesterol and bile acids; and catabolism of purines, polyamines, andamino acids (for a review, see Mannaerts & Van Veldhoven 1996).

Peroxisome biogenesis consists of three aspects: membrane synthesis, import of matrixproteins, and proliferation. Over 20 peroxins (proteins involved in peroxisomal import,biogenesis, proliferation and inheritance) have been identified, and the correspondingPEXgenes have been cloned and characterized (for reviews, see Kunau & Erdmann 1998,Titorenko & Rachubinski 1998). The view of peroxisome biogenesis has reached acrossroad. The orginal model proposed that peroxisomes arose through budding from theER (Novikoff & Shinn, 1964). By the end of the 1980s, the ‘growth and division’ modelbecame accepted which postulated that new peroxisomes were formed by the fission ofpre-existing ones, and that this process is accomplished by the post-translational import ofmembrane and matrix proteins from the cytosol (Lazarow & Fujiki 1985). Recentobservations that the ER plays a direct role both in transport of membrane proteins toperoxisomes and in supplying peroxisomes with phopholipids via ER-derived vesicleshave challenged the current theory of peroxisome biogenesis (for reviews, see Kunau &Erdmann 1998, Titorenko & Rachubinski 1998). Peroxisome biogenesis in the absence ofpre-existing organelles was also addressed by determining the function of Pex16p, whenoverexpressed, restored thede novoformation of peroxisomes in cells of a Zellwegersyndrome patient (South & Gould 1999). Other important progress has been made byidentifying specific signal sequences and the receptors responsible for targeting proteins tothe peroxisomes, but this will not be discussed here (for a review, see Subramani 1998).

2.2.1. Peroxisomal proliferation in mammalian cells

An outstanding feature of peroxisomalβ-oxidation is its powerful induction by a numberof peroxisome proliferators. More than ten-fold induction in the number and size ofperoxisomes is observed in rodents in response to peroxisomal proliferators (Lazarow &De Duve 1976, Lazarow 1977). Peroxisomal proliferators include a broad group ofchemicals such as fibrate hypolipidemic drugs, i.e. clofibrate (ethyl-p-chlorophenoxyisobutyrate), nafenopin, Wy 14,643, industrial plasticisers, i.e. di-(2-ethylhexyl) phtalate, herbicides and leukotrene antagonists. In addition, thia fatty acids,which block β-oxidation, are also powerful inducers of peroxisomalβ-oxidation (for areview, see Skredeet al. 1997). Induction of peroxisomes is found primarily in liver andkidney and is species specific, i.e. rodents are more susceptible than other organisms. Inaddition to the increase in the size and number of peroxisomes, most of the peroxisomalenzyme activities are also dramatically induced. Thus, peroxisomal proliferation provides

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a tool for investigation of the biological aspects of peroxisomal function (for reviews, seeOsmundsenet al.1991, Schoonjanset al.1997).

Peroxisome proliferation is a pleiotropic cellular response following the specificbinding moiety is demonstrated to peroxisome proliferators. This receptor-mediatedmechanism for proliferation was first proposed in 1983 by Lalwani and co-workers.Peroxisomal proliferator-activated receptors (PPARs) have been identified as products ofseparate genes from rodents (Issemann & Green 1990, Göttlicheret al. 1992),Xenopuslaevis(Dreyeret al. 1992), and humans (Sheret al. 1993, Greeneet al. 1995). PPARs indifferent vertebrates form a distinct subfamily of nuclear receptors and have beenclassified as PPARα, β (δ) andγ (for a review, see Schoonjanset al. 1997). PPARα ispredominantly expressed in liver, heart, kidney, and the intestinal mucosa, all of whichdisplay high catabolic rates for peroxisomal metabolism of fatty acids (Braissantet al.1996). Knockout mice deficient in PPARα do not respond to peroxisome proliferators andtranscriptional activation of the target genes is not observed in these animals (Leeet al.1995). PPARβ is ubiquitously and highly expressed in the central nervous system,however, its role is not known (Braissantet al. 1996). PPARγ is predominantly expressedin adipose tissue and is responsible, at least in part, for adipocyte differentiation (for areview, see Spiegelman 1998).

Activation of PPARs is initiated by lipophilic ligands that bind to a ligand-binding sitein their carboxyl-terminal region. PPARα is activated either by peroxisomal proliferators(Isseman & Green 1990, Dreyeret al.1993) or by a variety of long-chain fatty acid and, inparticular, polyunsaturated fatty acids (Kelleret al. 1993, Bocoset al. 1995). The assaysbased onin vitro binding, conformational change, or receptor-coactivator transactivationindicate that the synthetic peroxisome proliferators and fatty acids, i.e. 18:2 (n-6), 18:3 (n-3 andn-6) and 20:4 (n-6), as well as leukotriene B4 and prostaglandin I2 analogs act asdirect ligands for PPARs (Formanet al. 1997, Klieweret al. 1997, Kreyet al. 1997). Thebinding of fatty acids to the PPAR subtypes reveals that PPARγ interacts more efficientlywith unsaturated fatty acids than with saturated fatty acids (Xuet al. 1999). The crystalstructure of the PPARβ ligand-binding domain demonstrates that the hydrophobic tail ofthe fatty acid adopts different conformations within the large hydrophobic cavity, whichcan explain the propensity for PPARs to interact with a variety of fatty acids (Xuet al.1999).

A cognate DNA sequence that is recognized by and binds to PPARs has beenidentified to be acis-acting peroxisome proliferator-reponsive element (PPRE). It consistsof a degenerate direct repeat of the canonical AGG(A/T)CA sequence separated by onebase pair. PPREs were identified in the 5’-flanking region of genes coding for manyproteins implicated in intra- and extracellular lipid metabolism, and most of thoseinvolved in peroxisomalβ-oxidation, such as, palmitoyl-CoA oxidase (Tugwoodet al.1992), multifunctional enzyme (Zhanget al. 1993) and thiolase (Klieweret al. 1992).Upon activation of the ligand, PPARs dimerize with the 9-cis retinoic acid receptor(RXRα) to form a PPAR-RXRα complex that binds to the PPRE motif to regulate thetranscription of the target genes (for a review, see Schoonjanset al. 1997). Thenucleotides in the flanking region upstream of the PPRE also play an important role inregulation of the affinity of PPAR-RXRα complex to its target site (Juge-Aubryet al.1997).

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2.2.2. Induction of genes encoding peroxisomal proteins in yeast

In S. cerevisiae, fatty acids are the only exogenous substances known to influence thelevels of peroxisomal enzymes and the number and size of peroxisomes (Veenhuiset al.1987), whereas in other yeast species such as the methylotropic yeastPichia pastoris,methanol is a potent inducer of peroxisomes.

Promoters of genes encoding peroxisomal matrix proteins contain a positivecis-actingelement termed the oleate response element (ORE) that mediates the induction of thesegenes in the presence of a fatty acid such as oleic acid [cis-C18:1(9)] (Einerhandet al. 1993,Filipits et al. 1993). The ORE is an imperfect repeat CGG(Xaa)15/18CCG containing twoconserved CGG triplets that are separated by 15-18 nucleotides (Rottensteineret al.1996). In addition, there are conserved A and T nucleotides in the consensus half site 5'-CGGN3T(Xaa)A-3', which are important for ORE function (Einerhandet al. 1993, Filipitset al. 1993). The ORE is the target for transcription factors consisting of Oaf1p (oaf:oleate-activated transcription factor) and Pip2p (pip: peroxisome induction pathway), bothof which possess a Zn2Cys6 DNA-binding motif and bind to the consensus ORE(Karpichevet al. 1997, Luoet al. 1996, Rottensteineret al. 1997). Heterodimerization ofPip2p and Oaf1p regulates induction of peroxisomalβ-oxidation when yeast cells aregrown in oleic acid medium. When yeast cells are grown on glucose, Oaf1p isconstitutively expressed and binds to ORE as a homodimer, whereas Pip2p is specificallyinduced in the presence of oleic acid (Rottensteineret al. 1997). In addition to Pip2p andOaf1p, other transcription factors are involved in the oleate-dependent induction ofperoxisomal proteins, most notably Adr1p (a regulator of alcohol dehydrogenase IIsynthesis; Simonet al. 1991).

2.2.3. Activation and transportation of fatty acids into peroxisomes

Mammalian peroxisomes are involved in theβ-oxidation of long-chain and very long-chain fatty acids (LCFAs and VLCFAs). Initiation of theβ-oxidation pathway starts withactivation of fatty acids to their fatty acyl-CoA thioesters by fatty acyl-CoA synthetases.The peroxisomal membrane contains at least three acyl-CoA synthetases depending onchain length specificity and species. The first one is a long-chain acyl-CoA synthetase(LACS, E.C. 6.2.1.3) which activates straight long-chain fatty acids (Shindo & Hashimoto1978, Krisans et al.1980, Mannaertset al. 1982, Bronfmanet al. 1984). The rat enzymeadditionally activates pristanic acid, a 2-methyl branched fatty acid originating from theα-oxidation of phytanic acid (Vanhoveet al. 1991, Wanderset al. 1992, Watkinset al.1996). The activity of the enzyme was also found in the membrane of ER (Kornberg &Pricer 1953) as well as in the outer membrane of the mitochondria (Norumet al. 1966).The mammalian gene for this acyl-CoA synthetase has been cloned (Suzukiet al. 1990,Abe et al. 1992) and characterized (Suzukiet al.1995).

The second enzyme is a very long-chain acyl-CoA synthetase (VLACS), whichactivates very-long-chain (C20-C26) fatty acids (Laposataet al. 1985, Singh & Poulos1988). The enzyme is located in the membrane of the peroxisomes and the ER, but not inmitochondria (Singh & Poulos, 1988, Lazoet al., 1990a, Singhet al., 1993). The gene for

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the enzyme was cloned from rat liver (Uchiyamaet al. 1996) and the predicted amino acidsequence had a high sequence similarity to a fatty acid transport protein fromS. cerevisiae(Fat1p, Schaffer & Lodish, 1994). Disruption of theS. cerevisiae FAT1gene decreasesVLACS activity and elevates intracellular concentrations of very long-chain fatty acids(Watkins et al. 1998). FAT1 defective cells were found to grow poorly in mediumsupplemented with oleic acid when the known fatty acid synthases (FAAs) wereinactivated, whereas fatty acid import into the cytosol was not significantly affected (Choi& Martin 1999). This result suggests thatFAT1 codes for peroxisomal VLACS and thatthe primary cause of the growth-defective phenotype is probably due to a failure in fattyacid β-oxidation rather than a defect in fatty acid transport. In humans, a phytanoyl-CoAligase of peroxisomalα-oxidation isolated from fibroblasts is a third acyl-CoA synthetasedistinct from the long-chain- and the very-long-chain- acyl-CoA synthetases. (Pahanet al.1993).

The site of VLCFA activation is still a matter of debate (Lazoet al. 1990b, Lagewegetal., 1991). LCFAs are activated outside peroxisomes, whereas medium-chain fatty acidsare activated inside peroxisomes (Krisanset al. 1980, Mannaertset al. 1982). A newmechanism of activation and transportation of LCFAs inS. cerevisiaehas been proposedfollowing the discovery of two peroxisomal ATP-binding cassete (ABC) proteins, Pat1p(Bossieret al. 1994) and Pat2p (Shaniet al. 1995), that form a heterodimer facilitatingtransportation of cytosolic LACS across peroxisomal membranes (Hettemaet al.1996).One of the peroxisomal disorders, X-linked adrenoleukodystrophy (ALD), characterizedby accumulation of VLCFAs, was previously regarded to be caused by VLACSdeficiency. When the gene for adrenoleukodystropy protein (ALDp) was cloned, itshowed no homology to theVLACs, but instead to the mammalian genePMP70 whoseproduct is a peroxisomal membrane protein belonging to the family of ABC transporterproteins (Mosseret al. 1993). ALDp was reported to assist VLACS to localize inperoxisomes (Yamadaet al. 1999).

2.2.4. The peroxisomalββββ-oxidation spiral

2.2.4.1. Acyl-CoA oxidation

The first reaction of peroxisomalβ-oxidation of fatty acids is the rate-limiting stepcatalyzed by an acyl-CoA oxidase, a FAD-containing enzyme that transfers electrons fromacyl-CoA to oxygen, producing H2O2 as well as desaturation of acyl-CoA to (2E)-enoyl-CoAs (Fig. 1). Rat liver contains at least three peroxisomal acyl-CoA oxidases that

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Fig. 1. The peroxisomalββββ-oxidation cycle consists of two partially parallel pathwaysfollowing reciprocal stereochemistry. The mammalian mitochondrial and bacterialββββ-oxidation pathways occur via (3S)-hydroxyacyl-CoA.

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facilitate the metabolism of a great variety of substrates in peroxisomes. The first enzymeis clofibrate-inducible palmitoyl-CoA oxidase which oxidize the CoA esters of medium-,long- and very long- straight chain fatty acids, medium- and long-chain dicarboxylic fattyacids and eicosanoids (Osumi & Hashimoto 1978, Scheperset al. 1990, Van Veldhovenetal. 1992). The low activity of the enzyme toward short-chain substrates may explain whyβ-oxidaiton of fatty acids in mammalian peroxisomes is not complete (Osumiet al. 1980).The second enzyme is pristanoyl-CoA oxidase which oxidizes the CoA esters of 2-methyl-branched fatty acids and the CoA esters of long chain mono- and dicarboxylic fatty acids,especially very-long-chain monocarboxylic acids (Van Veldhovenet al. 1991 & 1994a).The third acyl-CoA oxidase, trihydroxycoprostanoyl-CoA oxidase is found in hepatictissue (Scheperset al. 1990) and oxidizes the CoA esters of the bile acid intermediates di-and trihydroxycoprostanic acids (Scheperset al. 1989a & 1990, Van Veldhovenet al.1994b). However, only two acyl-CoA oxidases have been detected in human liver. One isa palmitoyl-CoA oxidase with the same function as its rat counterpart. The other is abranched acyl-CoA oxidase, which has both functions of the two other rat oxidases(Casteelset al. 1990, Wanderset al. 1990, Vanhoveet al. 1993).

The cDNAs of palmitoyl-CoA oxidases have been cloned from mammals (Miyazawaet al. 1987, Aoyamaet al. 1994, Fournieret al. 1994, Varanasiet al. 1994). Either the rator human cDNA encodes a protein of 661 amino acid residues, containing a SKL carboxyterminal tripeptide. The corresponding genes have also been characterized and both havebeen shown to be alternatively spliced (Osumiet al. 1987, Fournieret al. 1994, Varanasiet al. 1994). The 5’ untranslated region of both of the genes contains a GC-rich region anda peroxisomal proliferator response element (Osumiet al. 1987, Varanasiet al. 1994 &1996). It has been reported that mice deficient in palmitoyl-CoA oxidase suffer from aliver disorder and spontaneous peroxisome proliferation. This implicates the oxidase as akey regulator of PPARα function and acyl-CoAs as potent biological ligands which areresponsible for the transcriptional activation of PPARα in vivo (Fanet al. 1996 & 1998).

In several yeast species such asCandida tropicalisand Candida maltosa, there aremultiple isoforms of acyl-CoA oxidases (Okazakiet al. 1986 & 1987, Murray &Rachubinski, 1987, Hillet al. 1988). InS. cerevisiae, however, there is only one acyl-CoAoxidase (Pox1p) and the expression of thePOX1gene is induced when the yeast cells aregrown in the presence of oleic acid (Dmochowskaet al. 1990). The regulation of thePOX1gene is mediated by the heterodimerization of Oaf1p and Pip2p, which bind to theORE as described in section 2.2.2. (Rottensteineret al. 1996, Karpichevet al. 1997).

2.2.4.2. Hydration of trans-2-enoyl-CoA and subsequent dehydrogenation

The second reaction in theβ-oxidation is the reversible hydration of (2E)-enoyl-CoA to(3S)-hydroxyacyl-CoA (Fig. 1). In mitochondria, the hydratase reaction catalyzed by amonofunctional 2-enoyl-CoA hydratase 1 (hydratase 1 or crotonase) followssynstereochemistry and is concerted. Two protic amino acid residues are required forcatalysis. In rat hydratase 1, the protonated Glu-164 acts as a catalytic acid providing a

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Fig. 2. The proposed reaction mechanism of rat mitochondrial 2-enoyl-CoAhydratase 1. The hydration and dehydration reactions are catalyzed by amino acidresidues, Glu164 and Glu144, at the active site.

proton to theα-carbon of the (2E)-enoyl-CoA, whereas the deprotonated Glu-144 acceptsa proton from water as a catalytic base and the formed hydroxide anion adds to theβ-carbon of the substrate (Fig. 2.; Engelet al. 1996, Kiemaet al. 1999). The crystalstructure of the protein has been solved and reveals a core domain consisting of repeatingββα-units that form a characteristic right handed spiral structure (Engelet al., 1996).

In addition to 2-enoyl-CoA hydratase 1, 2-enoyl-CoA hydratase 2 in rat liver catalyzesreversible hydration/dehydration between (2E)-enoyl-CoA and (3R)-hydroxyacyl-CoA,which is supposed to arise from a minor pathway associated with the metabolism ofpolyunsaturated fatty acids withcis double bonds, i.e.trans-2, cis-4-decadienoyl-CoA(Stoffel et al. 1964). The “epimerase-pathway” is only present in peroxisomes and wouldbe essential for preventing the accumulation of (3R)-hydroxyoctanoyl-CoA (Chu & Schulz1985, Yanget al. 1986). The 3-hydroxyacyl-CoA epimerase reaction in mammalian cellsis not catalyzed by a single enzyme, but by cooperation of two stereospecific hydratases:2-enoyl-CoA hydratase 1 and a 2-enoyl-CoA hydratase 2 (Hiltunenet al. 1989). Theepimerization mechanism in a bacterial system also confirms that 3-hydroxyacyl-CoAesters are epimerized by theβ-oxidation complex ofEscherichia colivia a dehydration-hydration mechanism (Smelandet al. 1991). It is still not clear whether monofunctionalepimerase exists. However, the cloning and expression of a plant glyoxysomalmultifunctional protein (MFE) demonstrated that the (3R)-hydroxyacyl-CoA convertingactivity was actually an epimerase rather than a part of a combinend water eliminating andwater attaching system (Preisig-Mülleret al. 1994). Inhibition studies showed that thehydratase 2 activity was inhibited by a sulfhydryl inhibitor (N-methylmaleimide) andacetoacetyl-CoA. The studies of the chain length specificity of the enzyme showed that theratio of hydration rates (C10/C4) was 14.4, indicating the low activity toward short-chainsubstrates. At least two hydratase 2 species have been purified from rat liver. The isoforms

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of 44 kDa and 31.5 kDa were found to be located in peroxisomes (Liet al. 1990) andmicrosomes (Malilaet al 1993), respectively.

The third reaction is the dehydrogenation of (3S)-hydroxyacyl-CoA to ketoacyl-CoA by(3S)-hydroxyacyl-CoA dehydrogenase (E.C. 1.1.35). In mammalian peroxisomes, thesecond and third reactions ofβ-oxidation are catalyzed by a single multifunctional protein(MFE-1) which has 2-enoyl-CoA hydratase 1, (3S)-hydroxyacyl-CoA dehydrogenase and∆3-∆2-enoyl-CoA isomerase activities (Osumi & Hashimoto 1979, Palosaari & Hiltunen1990). In the yeast peroxisomes, the second and third reactions ofβ-oxidation proceed viaa (3R)-hydroxyacyl-CoA intermediate and these two reactions are catalyzed by yeastperoxisomal MFE (MFE-2) possessing (3R)-hydroxyacyl-CoA dehydrogenase andhydratase 2 activities (Hiltunenet al.1992).

2.2.4.3. Cleavage of 3-ketoacyl-CoA

The fourth reaction ofβ-oxidation is thiolytic cleavage of a 3-ketoacyl-CoA thioester,which produces an acyl-CoA shortened by two carbon atoms and acetyl-CoA (Fig.1). Atleast three peroxisomal 3-ketoacyl-CoA thiolase genes exist. They are thiolase A (encodedby gene A) containing a pre-sequence of 36 amino acids (Hijikataet al. 1987 & 1990),thiolase B (encoded by gene B) containing a pre-sequence of 26 amino acid residues(Miyazawaet al. 1980) and a sterol carrier protein 2/3-ketoacyl-CoA thiolase (Seedorfetal. 1994, Antonenkovet al. 1997). The pre-sequences at the amino terminus of both ratliver thiolases harbor the peroxisomal targeting signal PTS-2 (Osumiet al. 1991, Swinkelset al. 1991, Miuraet al. 1994, Tsukamotoet al. 1994). Gene B was markedly activated bytreatment with peroxisome proliferators whereas gene A was only slightly or not activated(Hijikata et al. 1990). Thiolase A and thiolase B isolated from rat liver are virtuallyidentical in their substrate specificity for the thiolytic cleavage of straight-chain 3-ketoacyl-CoAs (Antonenkovet al. 1997 & 1999). In humans, only one peroxisomal 3-ketoacyl-CoA thiolase has been identified and comparison with two rat thiolases reveals78% identity at the amino acid level (Boutet al. 1988, Fairbarin & Tanner 1989, Bondaret al.1990).

The mRNA encoding sterol carrier protein 2/3-ketoacyl-CoA thiolase, has two initiationsites giving polypeptides with molecular masses of 58 kDa (SCPx) and 15 kDa (pro-SCP2). The open reading frame encodes a fusion protein containing a C-terminal 143-amino acid lipid transfer domain (SCP2 domain) and an N-terminal domain of 404 aminoacids, displaying thiolase activity toward 2-methyl-branched 3-ketoacyl-CoA and the bileacid intermediate (Antonenkovet al. 1997). In line with this observation, catabolism ofmethyl-branched fatty acyl CoAs is found to be impaired inSCP2 (-/-) mice, exemplifiedby the accumulation of the tetramethyl-branched fatty acid phytanic acid due to defectivethiolytic cleavage of 3-ketopristanoyl-CoA (Seedorfet al. 1998).

In S. cerevisiae, 3-ketoacyl-CoA thiolase has been isolated (Erdman & Kunau, 1994)and its structure has been determined to 1.8Å resolution (Mathieuet al. 1994).Peroxisomal acetoacetyl-CoA thiolase (thiolase I) and 3-ketoacyl-CoA thiolase (thiolaseII) were found and purified fromC. tropicalis (Kuriharaet al. 1989, Tanakaet al. 1995).The kinetic studies showed that the long-chain substrates were degraded exclusively by

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thiolase II, while acetoacetyl-CoA was degraded preferentially by thiolase I. ThePOT1gene encoding 3-ketoacyl-CoA-thiolase fromYarrowia lipolytica has been isolated andcharacterized (Berningeret al. 1993). As there are no other catabolic thiolases inY.lipolytica, the inability of this yeast to grow on oleic acid when thePOT1 gene isdisrupted implies that Pot1p palys an important role inβ-oxidation.

2.2.5. Oxidation of the cholesterol side chain

Primary bile acids are synthesized from cholesterol in the liver by the actions of at least 14different enzymes (for reviews, see Russel & Setchell 1992, Perdersen 1993). Thesequence of reactions involved in the formation of bile acids may be seen as a series ofdetoxification steps transforming the highly hydrophobic cholesterol molecules into water-soluble products. The reactions are divided into two main stages: i) oxidation andmodification of the ring system; and ii) oxidation and cleavage of the side chain followedby conjugation with glycine or taurine.

The terminal steps of side-chain oxidation take place mainly in the peroxisome and arethought to follow a pathway similar to that of theβ-oxidation of fatty acids in thisorganelle (Fig. 2.). The 3α, 7α, 12α-trihydroxy-5β-cholestanoic acid (THCA) is initiallyactivated to 3α, 7α, 12α-trihydroxy-5β-cholestanoyl-CoA (THCA-CoA) by atrihydroxycoprostanoyl-CoA synthetase (Scheperset al. 1989b), which is subsequentlyacted upon by an acyl-CoA oxidase to yield (24E)-∆24-THCA-CoA. Followed byhydration and dehydrogenation catalyzed by a peroxisomal MFE, 24-keto-THCA-CoA isformed and thiolytically cleaved, yielding a propionyl-CoA and the formation of a bileacid-CoA product that is shortened by three carbons.

For the first reaction of peroxisomalβ-oxidation, three different acyl-CoA oxidaseshave been described in rat and humans (see section 2.2.5.1; Van Veldhovenet al. 1991 &1992, Scheperset al. 1990). Whereas, only rat trihydroxycoprostanoyl-CoA oxidase orhuman branched acyl-CoA oxidase is responsible for oxidation of THCA-CoA to (24E)-∆24-THCA-CoA (Casteelset al. 1990, Scheperset al. 1990). The substrate specificity ofthe oxidase indicates that the 25S isomer of THCA-CoA is a preferential substrate,implying that 2-methylacyl-CoA rasemase exists in peroxisomes (Ikegawaet al. 1995).Previously, it was believed that the mammalian MFE-1 catalyzed the hydration anddehydrogenation of (24E)-∆24-THCA-CoA to a 24-keto-THCA-CoA. However, Xu &Cuebas (1996) found that the product of hydration of (24E)-∆24-THCA-CoA by MFE-1was (24S, 25S)-24-OH-THCA-CoA, which was not a proper substrate for thedehydrogenase component of the enzyme. Thus, there is a paucity of data regarding whichenzyme can act on these two reactions leading to biosynthesis of bile acid in theperoxisomes.

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Fig. 3. The terminal steps in oxidation of a cholesterol side chain.

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2.3. Auxiliary enzymes involved inββββ-oxidation

The only double bond of unsaturated acyl-CoA that is catabolized via normalβ-oxidationis at the∆2-position and in atrans configuration. Therefore, the removal of other doublebonds requires the auxiliary enzymes, including 2, 4-dienoyl-CoA reductase,∆3-∆2-enoyl-CoA isomerase and∆3,5-∆2,4-dienoyl-CoA isomerase (Fig. 4.). Fatty acids with a doublebond at even-numbered position yield 2, 4-dienoyl-CoAs, which can be converted to (2E)-enoyl-CoAs by 2, 4-dienoyl-CoA reductase and∆3-∆2-enoyl-CoA isomerase. In contrast,2, 5-dienoyl-CoAs, which arise from fatty acids with double bonds at odd-numberedpositions, enterβ-oxidation by two distinct pathways (see Section 2.3.3.).

α-Methyl-acyl-CoA racemase is a recently discovered auxiliary enzyme required forβ-oxidation of branched fatty acids and bile acid intermediates (Schmitzet al. 1994,Ikegawaet al.1995).

2.3.1. 2, 4-Dienoyl-CoA reductase

An NADPH-dependent 2, 4-dienoyl-CoA reductase (E.C. 1.3.1.34) participates in theβ-oxidation of fatty acids having double bonds at even-numbered positions (Fig. 4.; Kunau& Dommes 1978, Dommes & Kunau 1984a). It catalyzes the reduction of 2, 4-dienoyl-CoAs totrans-3-enoyl-CoAs in eukaryotes andtrans-2-enoyl-CoAs inE. coli (Dommes &Kunau 1984b).Trans-3-enoyl-CoAs are converted to (2E)-enoyl-CoAs by∆3-∆2-enoyl-CoA isomerase (Cuebas & Schulz 1982, Osmundsenet al. 1982, Hiltunenet al. 1983)with the final product entering theβ-oxidation spiral (Fig. 4.). Peroxisomes areimpermeable to NADPH (Van Roermundet al. 1995), which is, at least in yeast, providedby an NADP+-dependent isocitrate dehydrogenase (Idp3) functioning in an NADP+ redoxshuttle across the peroxisomal membrane (Van Roermundet al. 1998, Henkeet al. 1998).Mammals possess at least two isoforms of mitochondrial 2, 4-dienoyl-CoA reductaseswith different molecular masses (120 kDa and 60 kDa) and a third peroxisomal one(Dommeset al. 1981, Hakkola & Hiltunen 1993). The 120-kDa isoform has been clonedfrom rat and humans, (Hiroseet al. 1990, Koivurantaet al. 1994), and the human gene hasbeen characterized and assigned to the chromosome 8q21.3 (Helanderet al. 1997). InS.cerevisiae, the product of theSPS19gene (Coeet al. 1994) was identified to be aperoxisomal 2, 4-dienoyl-CoA reductase with a molecular mass of 69 kDa (Gurvitzet al.1997).

2.3.2.∆∆∆∆3-∆∆∆∆2-Enoyl-CoA isomerase

Fatty acids with a double bond extending from an odd-numbered carbon atom werethought to proceed stepwise viaβ-oxidation until the∆3-double bond was introduced.∆3-∆2-Enoyl-CoA isomerase (E.C. 5.3.3.8) converts bothcis- andtrans-3-enoyl-CoA esters totheir trans-2-counterparts, which are metabolized further by regular fatty acidβ-oxidation(Stoffel et al 1964). The known isomerases belong to the low-similarity

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Fig. 4. Auxiliary enzymes are required for theββββ-oxidation of unsaturated fatty acids.The bacterial 2, 4-dienoyl-CoA reductase hastrans-2-enoyl-CoA as an end product.∆∆∆∆3, 5-∆∆∆∆2, 4-Dienoyl-CoA isomerase activity has not yet been found in prokaryotes.

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isomerase/hydratase protein superfamily with a conserved fingerprint –Val-Ser-Xaa-Ile-Asn-Gly-Xaa-Xaa-Xaa-Ala-Gly-Gly-Xaa-Leu-Xaa-Xaa-Xaa-Xaa-Cys-Asp-Tyr- (Müller-Newen & Stoffel 1993). Members include isomerases and hydratases acting an either the∆2 or the ∆3-double bond, and additionally include 4-chlorobenzoyl-CoA dehalogenase(Babbittet al. 1992), naphthoate synthase (Sharmaet al. 1992),β-hydroxyisobutyryl-CoAhydrolase (Haweset al. 1996), ∆3,5-∆2,4-dienoyl-CoA isomerase (Filppulaet al. 1998),carnitine racemase (Eichleret al. 1994) and other uncharacterized proteins. They arethought to have evolved from a common ancestor since they were found in a variety ofspecies including bacteria and mammals. Interestingly, hydratase 1 has been reported tocontain an intrinsic isomerase activity in addition to its known hydratase activity (Kiemaet al. 1999). There are two separate mitochondrial short-chain and long-chain isomerasesin rat liver (Palosaariet al. 1990, Kilponenet al. 1990). The mitochondrial short-chainisomerase has been charaterized (Stoffel & Grol 1978, Palosaariet al 1990 & 1991,Tomiokaet al. 1991 & 1992, Müller-Newenet al. 1991, Stoffelet al. 1993, Janssenet al.1994) and crystalized (Zeelenet al. 1992). The isomerase activity is markedly induced bythe peroxisome proliferators. Theoretically, two protic amino acid residues are required toparticipate in catalysis, as in rat mitochondrial hydratase 1 (see Section 2.2.5.2). However,only one conserved protic residue in the isomerase, Glu165, was identified to beresponsible for the isomerase activity (Müller-Newenet al. 1995). The second proticresidue is unknown and could be a water molecule or side chain of an unidentified proticresidue. In addition to the two mitochondrial isomerases, a third isoform of mitochondrialisomerase, that does not show any substrate length specificity, has been characterized inhumans (Kilponen & Hiltunen 1993, Kilponenet al. 1994). The peroxisomal∆3-∆2-enoyl-CoA isomerase is a part of the peroxisomal MFE-1 in mammals (Palosaari & Hiltunen1990). An oleate-inducible peroxisomal monofunctional isomerase has been identified inS. cerevisaeand is also a member of the isomerase/hydratase superfamily (Gurvitzet al.1998, Geisbrechtet al. 1998). This protein does not contain a conserved glutamate (Glu165) at its active site and therefore lacks the conserved fingerprint. The significance ofthis divergence remains to be studied.

2.3.3. ∆∆∆∆3, 5-∆∆∆∆2, 4-Dienoyl-CoA isomerase

The β-oxidation of unsaturated fatty acids with double bonds in odd-numbered positionsproceeds to producetrans-2-enoyl-CoA via a ∆3-intermediate by∆3-∆2-enoyl-CoAisomerase (Fig. 4.; reductase-independent pathway; Stoffelet al. 1964). An additionalNADPH-dependent reductive metabolism pathway of unsaturated fatty acids with doublebonds at the∆5 position has been described (Tserng & Jing 1991), and occurs through anintermediate oftrans-2-∆5-dienoyl-CoA (Fig. 4.; reductase-dependent pathway; Smelandet al. 1992). In the proposed sequence of reactions,∆5-enoyl-CoA is dehydrogenated totrans-2, ∆5 dienoyl-CoA, which is isomerized to∆3-∆5-dienoyl-CoA by∆3-∆2-enoyl-CoAisomerase or peroxisomal MFE-1. The∆3-∆5-dienoyl-CoA is subsequently isomerized totrans-2, trans-4-dienoyl-CoA by∆3,5-∆2,4-dienoyl-CoA isomerase. (Luoet al. 1994, Chenet al. 1994, Luthriaet al. 1995). The metabolic sequence fromtrans-2, trans-4-dienoyl-CoA operates by the NADPH-dependent pathway as proposed for unsaturated fatty acids

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with even-numbered double bonds. The contribution of these two pathways inmitochondria is still a debate (Tsernget al. 1996, Shoukry & Schulz 1998).∆3,5-∆2,4-Dienoyl-CoA isomerase has been purified from rat liver and is localized in bothmitochondria (Smelandet al. 1992) and peroxisomes (Heet al. 1995). Hiltunen’s group(Filppula et al. 1998) expressed a cDNA encoding a protein of 36 kDa, rECH1(FitzPatrick et al. 1995), which belongs to the hydratase/isomerase superfamily. Theexpressed recombinant protein shows∆3,5-∆2,4-dienoyl-CoA isomerase activity.Subcellular fractionation and immunoelectronmicroscopy of rat liver localizes thepolypeptide to both mitochondria and peroxisomes. Consistent with these observations,the open reading frame of rECH1 has a potential N-terminal mitochondrial targeting signalas well as a C-terminal peroxisomal PTS-1. The crystal structure of rat∆3,5-∆2,4-dienoyl-CoA isomerase has been solved at 1.5 Å resolution and it resembles that of 2-enoyl-CoAhydratase 1 and 4-chlorobenzoyl-CoA dehalogenase. The amino acid residue Asp204 hasbeen revealed to be essential for catalysis (Modiset al. 1998). Recently, a peroxisomal∆3,5-∆2,4-dienoyl-CoA isomerase has been identified fromS. cerevisiae(Gurvitz et al.1999), but the physiological role of its participation in the reductase-dependent pathway inoxidation of unsaturated fatty acids with double bonds at odd-numbered position seemsnot important.

2.3.4. 2-Methylacyl-CoA racemase

Branched fatty acids arise from the metabolism of isoprenoids. For example, accumulationof phytanic acid (3,7,11,15-tetramethylhexadecanoic acid) is observed in Refsum’sdisease and in a number of peroxisomal diseases with defects inα-oxidation. Phytanicacid is a 3-methyl-branched fatty acid and is formed from phytol, which is present inchlorophyll in an esterified form. Phytanic acid is converted to pristanic acid (2-methyl-branched fatty acid, 2,6,10,14-tetramethylpentadecanoic acid) viaα-oxidation, whichshortens it by one carbon atom, and allows further degradation byβ-oxidation. Theresulting pristanic acid is a racemic mixture of 2S- and 2R-enantiomers. The racemaseallows theβ-oxidation of both enantiomers of 2-methyl-branched fatty acids by converting(2R)-enantiomer to (2S)-enantiomer, which is accepted as the substrate for mitochondrialacyl-CoA dehydrogenases or peroxisomal oxidases (Schmitz & Conzelmann 1997, VanVeldhovenet al. 1996). The bile acid intermediate, (25R)-trihydroxycoprostanoyl-CoA isrequired to be racemized to the (25S)-enantiomer by the racemase in bile acid formation.The cDNA of the racemaces from rat or mouse does not resemble any other known aminoacid sequences ofβ-oxidation proteins (Schmitzet al. 1997). The racemaces have beenshown to be present in mitochondria, peroxisomes and the cytosol of human and rat livers(Schmitzet al. 1995, Van Veldhovenet al. 1997).

2.4. Multifunctional enzymes (MFEs) ofββββ-oxidation

Multifunctional enzymes (MFEs) are associated with all knownβ-oxidation systems

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characterized thus far and comprise the second and the third reactions of theβ-oxidationspiral. In mammalian mitochondria, these two reactions can also be catalyzed bymonofunctional enzymes. MFEs have been purified from bacteria, fungi, plants andmammals, and their activities depend on the subcellular location and vary from onespecies to another (Table 1; for a review, see Hiltunenet al. 1993).

2.4.1. Bacterialββββ-oxidation multienzyme complex

E. coli is used as a model system for studying fatty acid degradation. It contains aβ-oxidation complex with a native molecular mass of 240 kDa, which is composed of twodifferent subunits,α andβ, that form a tetramerα2β2. α andβ are encoded by thefadBAoperon (Binstocket al. 1977, Binstock & Schulz 1981, Pawar & Schulz 1981). 2-Enoyl-CoA hydratase 1, (3S)-hydroxyacyl-CoA dehydrogenase and∆3-∆2-enoyl-CoA isomeraseactivities are catalyzed by the 79 kDa-α subunit encoded by thefadBgene and 3-ketoacyl-CoA thiolase, encoded byfadA gene, is located in the 42 kDaβ subunit (Yang & Schulz1983, Yanget al. 1988, Yanget al. 1991). The epimerization reaction occurs via ahydration/dehydration mechanism (Smelandet al. 1991), which is the same as that foundin mammals (see Section 2.2.5.2; Hiltunenet al. 1989). It has been proposed thatE. colihydratase 1 might function as a 3-hydroxyacyl-CoA epimerase possessing hydratase 2 andhydratase 1 activities (Yang & Elzinga 1993). Site-directed mutagenesis of the amino-terminal domain of theα-subunit and kinetic studies of the expressed recombinantproteins have revealed that both Glu119 and Glu139 are the catalytic residues for thehydration reaction following an acid-base mechanism (Yanget al. 1995, He & Yang1997), which has been elucidated from rat mitochondrial 2-enoyl-CoA hydratase 1 (Kiemaet al. 1999). His450 was assigned to be the catalytic residue of (3S)-hydroxyacyl-CoAdehydrogenase (He & Yang 1996) and this observation confirms that theE. colidehydrogenase has a proposed NAD+-binding site in its aminoterminal domain andstructurally resembles the porcine heart dehydrogenase (EC 1.1.1.35; Yanget al. 1991).

2.4.2. Fungal multifunctional enzymes

Alkane-assimilating yeast has been proposed to be an excellent model with which toinvestigate fatty acid degradation in mammalian cells. The capacity to degrade fatty acidsis confined to peroxisomes rather than mitochondria. In yeast peroxisomes, unlike in thoseof mammals and bacteria,β-oxidation of fatty acids proceeds onlyvia R-hydroxyintermediates catalyzed by a dimeric MFE-2 (Hiltunenet al. 1992). The genes encodingthe MFE-2 in yeastC. tropicalisandS. cerevisiaehave been characterized

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Fig. 5. The proposed reaction mechanism of (3R)-hydroxyacyl-CoA dehydrogenaseof yeast peroxisomal MFE-2.

(Nuttley et al. 1988, Hiltunenet al. 1992). Amino acid and nucleotide sequence analysisof the yeast MFE-2 suggested a division of the polypeptide into aminoterminal, middleand carboxyterminal domains and a partial gene duplication in the first two domainsduring the evolution (Nuttleyet al. 1988). A truncated version of the MFE-2 lacking 271amino acids in carboxylterminal domain retains only the dehydrogeanse activity,suggesting that the catalytic domains of the dehydrogenase are located in theaminoterminus, whereas the hydratase domain is located in the carboxylterminus (Hiltunenet al. 1992). Limited proteolysis of MFE-2 isolated from bothC. tropicalis andNeurospora crassa(Thieringer & Kunau 1991) yields two major fragmentation products,supporting the occurrence of two superdomain structures.

The duplicated dehydrogenase domains of MFE-2 belong to a short-chain alcoholdehydrogenase/reductase family (SDR), which includes phylogenetically related groups ofenzymes acting on as diverse substrates as sugars, steroids, prostaglandins, aromatichydrocarbons, antibiotics and compounds involved in nitrogen metabolism (for a review,see Jörnvallet al. 1995). A characteristic segment of the nucleotide [NAD(H) orNADP(H)] binding fold Gly-Xaa-Xaa-Xaa-Gly-Xaa-Gly ("Rossmann fold") is largelyconserved in the SDR family. Sequence analysis has revealed that the members of theSDR family contain a highly conserved pentapeptide motif of Tyr-Xaa-Xaa-Xaa-Lys. Thetyrosine’s deprotonated phenolic group is proposed to catalyze hydride transfer with theaid of the positively charged lysine residue that decreases the pKa of the tyrosine’sphenolic group from 10 to about 7 (Fig. 5.). However, the significance of the existence oftwo dehydrogenase domains in yeast MFE-2 is unknown.

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2.4.3. Plant multifunctional enzymes

Linoleic acid and other highly unsaturated fatty acids amount to up to 70% of the fattyacid moieties in plant lipids. In plants, the fatty acids are mainly degraded in glyoxysomesor peroxisomes. Mitochondrialβ-oxidation is generally regarded to be absent or to haveonly a minor contribution. The peroxisomal chain shortening of acyl-CoA requires threeenzymes: acyl-CoA oxidase, MFEs and thiolase. In the plantβ-oxidation pathway, (3R)-hydroxyacyl-CoA is an intermediate in the conversion betweencis-2-enoyl-CoA andtrans-2-enoyl-CoA by (3R)-hydroxyacyl-CoA hydrolase (Engeland & Kindl 1991). Threeisoenzymes of the MFEs have been found and they differ in the number of active sites,catalytic properties, size and other molecular characteristics (Gühnemann-Schäfer & Kindl1995). One of these isoenzymes has been found to have a molecular mass of 79 kDa andcontains the four activities of (3S)-hydroxyacyl-CoA hydrolyase, (3S)-hydroxyacyl-CoAdehydrogenase, (3R)-hydroxyacyl-CoA epimerase, and∆3-∆2-enoyl-CoA isomerase(Preisig-Mülleret al. 1994).

2.4.4. Mammalian multifunctional enzymes

The peroxisomal MFE-1 was induced by peroxisomal proliferators and purified from rat(Osumi & Hashimoto 1979) and humans (Reddyet al. 1987). Palosaari and Hiltunen(1990) showed that the enzyme possesses∆3-∆2-enoyl-CoA isomerase activity in additionto 2-enoyl-CoA hydratase 1 and (3S)-hydroxyacyl-CoA dehydrogenase activities. The ratMFE-1 gene encodes 722 amino acid residues (Osumiet al. 1985), whereas the humangene encodes 723 amino acid residues (Hoefleret al. 1994) and both contain SKLperoxisomal targeting signals at the carboxy terminus. The amino acid identity betweenthe rat and human enzyme sequences is 78%. Sequence comparison of MFE-1 withmitochondrial 3-hydroxyacyl-CoA dehydrogenase (Minami-Ishiiet al. 1989) andE. colifadB (Yang et al. 1991) reveals that the hydratase and isomerase reside in theaminoterminal domain and share the same CoA binding site (Palosaariet al. 1991),whereas the dehydrogenase is located in the carboxyterminus (Osumi & Hashimoto 1979).TheMFE-1 gene has been characterized from rat (Ishiiet al. 1987) and a PPRE has beenidentified in its promoter region (Zhanget al. 1992).

In addition to peroxisomal MFE-1, there is a mitochondrial trifunctional enzyme thathas a structure ofα4β4 and catalyzes the second, third and fourth reaction of theβ-oxidation spiral (Uchidaet al. 1992). Amino acid sequence analysis of the enzymeindicates thatα-subunit harbors a 2-enoyl-CoA hydratase in the aminoterminal domainand a 3-hydroxyacyl-CoA dehydrogenase in the carboxylterminal domain, while a 3-ketoacyl-CoA thiolase resides in theβ-subunit (Kamijoet al. 1993).

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Table 1. MFEs of various species

Species MolecularMass

Activities Structure Reference

Escherichiacoli

260,000 H1, H2, I, DS, T α2β2 Yang & Schultz 1983

Pseudomonasfragi

240,000 H, E, I, D, T α2β2 Imamuraet al. 1990

Neurosporacrassa

365,000 H2, DR α4 Fossået al. 1995

Candidatropicalis

186,000 H2, DR α2 De La Garzaet al.1985,Nuttleyet al.1988

Saccharomycescerevisiae

170,000 H2, DR α2 Hiltunenet al. 1992

Cucumissativus

74,000* H, D, E α Gühnemann-Schäferet al.1994

76,000** H, D, I, E α Gühnemann-Schäferet al.1994

81,000*** H1, DS, I, E α Gühnemann-Schäfer & Kindl1995

79,047 H1, DS, I, E α Preisig-Mülleret al. 1994Rat(peroxisomal)

78,511 H1, DS, I α Osumi & Hashimoto 1979,Palosaari & Hiltunen 1990

Rat(mitochondrial)

460,000 H1, DS, T α4β4 Uchidaet al. 1992,Kamijo et al.1993

Mouse(peroxisomal)

5,000 H1, DS, I α Stark & Meijer 1994

Human(peroxisomal)

78,000 H1, DS, I α Reddyet al. 1987, Hoefleretal. 1994, Kilponen &Hiltunen 1993

Human(mitochondrial)

230,000 H1, DS, T α4β4 Carpenteret al. 1992,Kamijo et al. 1994

Abbreviations: H, 2-enoyl-CoA hydratase (stereochemistry of the reacion not specified); H1, 2-enoyl-CoA hydratase 1; H2, 2-enoyl-CoA hydratase 2, D, 3-hydroxyacyl-CoA dehydrogenase(stereochemistry of the reaction not specified); DS, (3S)-hydroxyacyl-CoA dehydrogenase; DR,(3R)-hydroxyacyl-CoA dehydrogenase; I,∆3-∆2-enoyl-CoA isomerase; T, 3-ketoacyl-CoA thiolase;E, 3-hydroxyacyl-CoA epimerase. When present, 3-ketoacyl-CoA thiolase activity is located in theβ-subunit. *MFE I, **MFE II, ***MFE III.

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3. Outlin e of the present study

Although the main pathways of the β-oxidation of fatty acids has been investigatedextensively, many questions still remain. When this work was started, it was assumed thatthe peroxisomal β-oxidation of fatty acids occured only via MFE-1 in mammals and viaMFE-2 in yeast. Rat 2-enoyl-CoA hydratase 2 that hydrates trans-2-enoyl-CoA to (3R)-hydroxyacyl-CoA had been purified from liver (Malila et al. 1993), however, thephysiological function of the enzyme was not known. Furthermore, the reason forexistence of the duplicate dehydrogenase domains in yeast MFE-2 was not clear. Thespecific aims of the present studies were:

1) to clone 2-enoyl-CoA hydratase 2 from rat liver,2) to investigate the physiological role of 2-enoyl-CoA hydratase 2,3) to study the catalytic properties of 2-enoyl-CoA hydratase 2,4) to elucidate the significance of duplication of the gene encoding the (3R)-

hydroxyacyl-CoA dehydrogenase domains of yeast peroxisomal MFE-2.

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4. Material s and methods

Materials and methods are described in detail in the original articles (I-IV).

4.1. Biological sources

The rats used for isolation of 2-enoyl-CoA hydratase 2 were Wistar strains obtained fromthe Laboratory Animal Center of the University of Oulu. In order to study induction ofperoxisomal proliferators (hypolipidaemic agents), the rats were fed on a standard pelleteddiet containing 0.5 % clofibrate (w/w) for two weeks before removal of the livers.

E. coli strain DH5α was used for all plasmid transformations and isolations. The yeastS. cerevisiae wild-type strain UTL-7A is ura3-52, trp1, leu2,3-112 and the fox-2 mutant isura3-52, trp1, leu2,3-112, ade, fox2.

4.2. Cultur e media

Yeast S. cerevisiae strains were cultured in the following media, YPD (1% yeast extract,2% Bacto peptone and 2% D-glucose), synthetic complete medium without uracil(SD/uracil, 0.67% Bacto-yeast nitrogen base without amino acids, 0.2% dropout powderwithout uracil, 2% D-glucose), oleic acid medium, YPOD (0.5% yeast extract, 0.5%Bacto peptone, 0.1% oleic acid, 0.1% glucose, 0.5% Tween 40, pH 7.0) and YNO (0.1%yeast extract, 0.67% yeast nitrogen base with amino acids, 0.1% oleic acid and 0.5%potassium phosphate, pH 6.0).

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4.3. Synthesis of substrates

4.3.1. Synthesis of straight-chain fatty acyl-CoA esters

(3R)-Hydroxybutyric acid, (2E)-hexenoic acid, (2E)-decenoic acid and crotonyl-CoA arecommercially available from Sigma. Acyl-CoA esters oftrans-3-decenoic, (3R, 3S)-hydroxydecanoic and (3R)-hydroxybutyric acids are prepared by the mixed anhydridemethod (Rasmussenet al. 1991) and purified on aµBondapak�C18 column (Waters,Milford, MA) by applying an acetonitrile gradient (15%-55%) for elution of thesubstrates. The structures of the substrates were verified by determining their masses withMALDI-TOF mass spectrometer (Ompact MALDI III, Kratos Analytical, Manchester,UK) and by testing with appropriate enzymes.

4.3.2. Synthesis of 2-pristenic acid

The detailed procedure of the synthesis of 2-pristenic acid and 2-methyltetradec-2-enoicacid is described in Paper I. CoA esters were prepared from the corresponding carboxylicacids by a modified mixed-anhydride method (Rasmussenet al.1991).

4.3.3. Synthesis of 24-OH-3αααα, 7αααα, 12αααα-trihydroxy-5ββββ-cholestanoic acid and (24E, Z)-∆∆∆∆24-3αααα, 7αααα, 12αααα-trihydroxy-5ββββ-cholestanoic acid

A mixture of the four (C24, 25) diastereomers of 3α,7α,12α,24ξ-tetrahydroxy-5β-cholestanoic acid (24-OH-THCA) was prepared using procedures previously described bycondensing 3α,7α,12α-triformyloxy-5β-cholan-24-al with 2-methyl-bromopropionate(Batta.et al. 1982). (24E, Z)-3α,7α,12α-trihydroxy-5β-cholest-24-enoic acid [(24E, Z)-∆24-THCA] was prepared from 3α,7α,12α-triformyloxy-5β-cholan-24-al according to thepublished procedures (Iqbalet al. 1991). CoA derivatives were prepared by the mixedanhydride method and purified by HPLC (Schulz 1974).

4.4. Enzyme assays

4.4.1. Enzymes of fatty acidββββ-oxidation

The activities of 2-enoyl-CoA hydratase 2, 2-enoyl-CoA hydratase 1, (3R)- and (3S)-hydroxyacyl-CoA dehydrogenases, 3-hydroxyacyl-CoA epimerase,∆3-∆2-enoyl-CoAisomerase and combined activities [metabolism of (2E)-enoyl-CoA esters to 3-ketoacyl-CoA esters] were measured as previously described (Hiltunenet al. 1989, Palosaari &

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Hiltunen 1990, Filppulaet al. 1995). The enzyme activities were measured usingShimadzu UV 3000 spectrophotometer.

4.4.2. Hydration and dehydration between (24E)-3αααα, 7αααα, 12αααα-trihydroxy-5ββββ-cholest-24-enoyl-CoA and 24-hydroxy, 3αααα, 7αααα, 12αααα-trihydroxy-5ββββ-cholest-24-enoyl-CoA

The incubation mixture for the reactions catalyzed by the recombinant 46 kDa hydratase 2contained either 45µM (24E)-∆24-THCA or an 80 µM mixture of the (C24, 25)-diastereomers of 24-OH-THCA-CoA as substrate. The reactions were initiated by theaddition of the enzyme. At various time intervals, a 150µl-aliquot of the incubationmixture was withdrawn and the reaction stopped by the addition of 2M HCl, and themixture was subsequently neutralized with 2M KOH. Aliquots of 150µl were analyzedusing HPLC and were performed on a Water dual-pump gradient with a YMC-Pack ODS-A reverse-phase column applying a linear gradient of methanol.

4.4.3. Enzyme kinetics

Kinetic constants (kcat and Km) of dehydrogenase activity were determined towardoxidation of (3R)-hydroxyacyl-CoA and formation of the Mg2+ complex of 3-ketoacyl-CoA was detected at 303 nm. The reaction buffer consisted of 50 mM Tris-HCl (pH 8.0),50 mM KCl, 1 mM NAD+, 1 mM sodium pyruvate, and 2.5 mM MgCl2, and 21 units of L-lactic dehydrogenase (Sigma Chemicals) as an auxiliary enzyme. The measurement wasperformed at different substrate concentrations in a final volume of 0.5 ml at 22°C. Thekinetic constants for hydratase 2 activity were measured with (2E)-decenoyl-CoA by adirect assay method (Binstock & Sculz 1981). The pH-dependence curve of hydratase 2was determined by monitoring the hydration reaction at 263 nm over a broad range of pHvalues in 200 mM potassium phosphate buffer with (2E)-decenoyl-CoA as substrate invarying concentrations.

4.5. Isolation of peroxisomes

Peroxisomes were isolated from the light mitochondrial fraction of rat liver with acontinuous Nycodenz density gradient (15-40%, w/w) as previously described (Ghosh &Hajra 1986).

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4.6. Isolation and characterization of cDNA clones

mRNA from rat liver was isolated with Dynabead oligo (dT)25 (Dynal, Oslo, Norway)according to the manufacture’s instructions. The isolated mRNA was transcribed to thecDNA by reverse transcription. The PCR amplification was performed by usingdegenerate oligonucleotides deduced from the amino acids sequences of the isolatedpeptides as primers, and using the resultant cDNA as the template. The amplified PCRfragment was subcloned into pUC 18 plasmids with a SureClone ligation kit. The plasmidwas used to transformE. coli DH5α. The subcloned fragments were sequenced by thedideoxynucleotide sequencing method.

4.7. Northern blot analysis

mRNA species were fractionated in 1% (w/v) agarose gel in the presence offormaldehyde, transferred to a nylon membrane by capillary blotting overnight and fixedby UV cross-linking. The membrane was hybridized with the labeled ratMFE-2 cDNAand exposed to X-ray film.

4.8. Expression of recombinant proteins

4.8.1. Expression of recombinant proteins in E. coli

The pET system is a powerful system developed for the expression of recombinantproteins inE. coli (Novagen). Target genes were cloned in pET plasmids under the controlof the strong bacteriophage T7 promoter. The resulting constructs were transformed intoan E. coli expression strain (BL21plysS or HMS174plysS) and plated on LB-ampicillinplate. Single colonies from the plates were used to inoculate M9ZB medium containingcarbenicillin (50µg/ml) and chloramphenicol (34µg/ml). The cultures were grown at37°C under aerobic conditions until an OD600 of 0.4-0.6 was reached. The expression ofthe recombinant protein was induced by addition of IPTG to a final concentration of 0.4mM. After 2 hours of induction at 33°-35°C, the cells were harvested and washed withphosphate-buffered saline (10 mM sodium phosphate, 2 mM potassium phosphate, 140mM NaCl, 3 mM KCl, 5 mM β-mercaptoethanol, pH 7.4) and the pellet was stored at -70°C until used for protein purification.

4.8.2. Expression of recombinant proteins in P. pastoris

P. pastoris, representing one of four different genera of methylotrophic yeast, is capableof using methanol as a sole carbon source. Alcohol oxidase catalyzes the oxidation of

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methanol to formaldehyde. Expression of this enzyme, encoded by theAOX1 gene, istightly regulated and induced by methanol to very high levels, typically≥ 30 % of the totalsoluble protein in cells grown with methanol as the carbon source. TheAOX1 gene hasbeen isolated and a plasmid-borne version of theAOX1 promoter is used to driveexpression of a gene of interest for heterologous protein expression. The coding sequencefor a protein of interest was cloned into the intracellular expression vector pHILD2 behindAOX1 promoter (Invitrogen). In addition, the vector, pHILD2, has aHIS4 gene tofacilitate the screening of recombinant strains. The resulting construct was linearized andintegrated into theAOX1 locus by transforming theP. pastoris HIS4-deficient strain,GS115, with the expression vector by electroporation (Gene Pulser, Bio-RadLaboratories) and allowing for homologous recombination to occur. Recombinanttransformants were selected from solid histidine-deficient media and those withAOX1-disrupted were screened by poor growth in the presence of methanol. For expression, cellswere grown on glycerol as a carbon source and were shifted to methanol for induction.

4.9. Site-directed mutagenesis

In vitro site-directed mutagenesis is a widely used technique for studying proteinstructure-function relationships. The QuickChange site-directed mutagenesis kit(Stratagene) was used to introduce separate point mutations into proteins. In this method,PCR reactions were performed using a supercoiled, double-stranded DNA plasmid as atemplate with two synthetic oligonucleotide primers containing the desired mutationencoding a mutated amino acid residue.pfu DNA polymerase was used in PCR reaction toreplicate both DNA strands with high fidelity, according to the manufacture's instructions.

4.10. Protein purification

Various chromatographic methods were applied to purify the enzymes. The columns weregenerally a dye ligand column, Matrex Red A, hydroxyapatite (HA), hydrophobic butyryl-sepharose, anion-exchange DEAE-Sephacel and Resource Q, cation-exchange Resource Sand size exclusion Superdex 200 HR 10/30. The purification protocols are described inmore detail in papers I-IV.

4.11. Analysis of proteins

In-Gel protein digestion was performed as previously described (Hellmanet al. 1995).The stained protein band was cut from the SDS-PAGE gel and digested with trypsin. Theresulting peptides were separated by reversed-phase liquid chromatography using aµRPCC2/C18 SC 2.1/10 column connected to a SMART micropurification system (PharmaciaBiotech). The peptides were eluted in a linear gradient of acetonitrile in 0.05% (v/v)

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trifluoroacetic acid. The selected peptides were subjected to automatic Edman degradationin an Applied Biosystems model 470A sequencer.

The molecular weight of a native protein was determined on a size exclusion columnSuperdex 75 using bovine liver catalase (235 kDa), rabbit muscle lactase dehydrogenase(140 kDa), pig heart malate dehydrogenase (70 kDa) and cytochrome C (12.5 kDa) asstandards. SDS-PAGE was run to determine the subunit molecular size in the presence of6M urea, and the gels were stained with 0.25 % Coomassie Brilliant blue inmethanol/acetic acid/water (5:1:5).

Dynamic light scattering measurements were performed with a DynaPro-MSTCdynamic light scattering device (Protein Solution, Charlottesville, VA). The measurementwas started after incubation of the sample for 5 minutes at 6°C in the sample cell andhydrodynamic radius was determined.

Circular dichroism (CD) sepctroscopy measurements were carried out at 22°C using aJasco JHO spectropolarimeter. Adsorption at 280 nm was measured and used for fineadjustment of the protein concentration of the sample used for CD spectroscopy. The far-UV spectra of the proteins were measured from 200 to 250 nm in 80 mM potassiumphosphate, pH 7.0, with the following instrument settings: response 1s, sensitivity 100mdeg, speed 50 nm/min, average of 30 scans.

Protein concentrations were determined with the Bio-Rad protein assay reagent, usingbovine serum albumin as the standard.

4.12. Complementation ofS. cerevisiae fox-2with MFE-2(s) andtheir variants

S. cerevisiae fox-2cells (Hiltunenet al. 1992) that are devoid of endogenousMFE-2served as a tool for examining thein vivo function of S. cerevisiaeMFE-2 variants,heterologous human MFE-2 and its variants. The gene of interest was cloned into theS.cerevisiaeexpression vector pYE352 behind the catalase A1 promoter, which contains anORE.fox-2cells were transformed with the resultant construct by a lithium acetate method(Gietz & Schiestl 1995) and selected for on SD/uracil plates. The transformants werestreaked onto solid medium containing oleic acid. Utilization of fatty acids by thetransfomedfox-2cells was observed by zones of clearing on the oleic acid plates.

4.13. Immunoblotting

The proteins from SDS-PAGE were electroblotted onto nitrocellulose filters. Detection ofthe blotted/transferred proteins was performed using the rabbit IgG fraction of theantiserum as the primary antibody and goat antirabbit IgG coupled to horseradishperoxidase as the secondary antibody (Bio-Rad Laboratories). The recognized epitopeswere detected by either 4-chloro-1-naphthol (Sigma Chemicals) or an enhancedchemiluminescence detection system (Amersham Pharmacia Biotech).

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4.14. Software

Using the BLAST network service (Altschulet al. 1990) performed amino acid homologycomparisons with the GenBank, EMBL, PIR and Swiss-Prot data base at the NCBI.Multiple sequence comparisons of polypeptides were carried out with multiple alignmentprogram CLUSTAL_X using the default parameters. The kinetic constants (Km andkcat),pH dependence curves of thekcat/Km for hydratase 2 reaction, pKa values, and their errorswere calculated with the GraFit computer software (Sigma Chemicals).

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5. Results

5.1. A novel peroxisomal multifunctiona l enzyme type 2

2-Enoyl-CoA hydratase 2 with a molecular mass of 31.5 kDa was purified from rat liver(Malila et al. 1993) and was subjected to trypsin digestion. The resulting peptides wereisolated and sequenced (Fig. 2 of paper I). The cloning of hydratase 2 reveals that the fulllength cDNA is 2535 bp that encodes a 735-residue polypeptide containing a potentialperoxisomal targeting signal (AKL) in the C-terminus (Fig. 2 of paper I). The molecularmass of the polypeptide is 79,3 kDa, which is 2.5 fold higher than that predicted for thepurified rat hydratase 2. To explain the apparent discrepancy between the purifiedhydratase 2 and the cloned polypeptide, immunoblotting of rat liver extract and pureperoxisomes was performed using the antibody to the rat hydratase 2 (Malila et al. 1993).The results revealed three bands, 79 kDa, 66 kDa, 46 kDa both in the liver cell extract andthe peroxisomes, and a 31.5 kDa band that was present only in the liver cell extract (Fig. 5of paper I), suggesting that hydratase 2 is actually a fragmentation product from thepolypeptide and cannot target into peroxisomes without a peroxisomal targeting signal.

The enzyme activities catalyzed by the polypeptide were studied with a recombinantprotein obtained via expression of the cDNA in P. pastoris cells. To confirm expression ofthe rat cDNA that was integrated into the P. pastoris genome, Northern blot analysis wasperformed and a 2.9 kb-signal was detected (Fig. 4 of paper I). Immunoblotting showedthat a 79 kDa signal and two smaller signals (66 kDa, 31.5 kDa) were detected withantibody to the rat hydratase 2 (Fig. 4 of paper I). The partially purified recombinantprotein was used for substrate specificity studies (Table 2 of paper I). The proteincatalyzes hydration and dehydrogenation reactions with (2E)-decenoyl-CoA (C10) andcrotonyl-CoA (C4) substrates, the C10/C4 activity ratio being 6.6 and 1.4 for the hydrationand the dehydrogenation respectively. The 2-methyl-branched esters were slowlymetabolized by the protein. The protein did not have detectable activities of 2-enoyl-CoAhydratase 1, (3S)-specific dehydrogenase, ∆3-∆2-enoyl-CoA isomerase and 3-hydroxyacyl-CoA epimerase. The first three activities are associated with the known mammalianperoxisomal MFE-1. Thus, the cloned polypeptide is assigned to be a novel mammalianperoxisomal multifunctional enzyme type 2 (MFE-2).

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Amino acid sequence comparisons of rat MFE-2 revealed that it has 81.3% identitywith porcine 17β-HSD type IV (Leenderset al. 1994). Rat MFE-2 shows a very lowcatalytic velocity for oxidation of estradiol to estrone (Table 2 in paper I). The N-terminaldomain of rat MFE-2 (residues 1-292) shows a high degree of identity with the twodehydrogenase domains of MFE-2 fromC. tropicalis (Fig. 3. of paper I; Nuttleyet al.1988) andS. cerevisiae(Fig. 3. of paper I; Hiltunenet al. 1992). The middle domain ofMFE-2 (residues 315-600) is similar to the hydratase region of yeast MFE-2(s). Theextreme C-terminus of rat MFE-2 (residues 612-731) is similar to rat sterol carrier protein2 (SCP-2; Seedorf & Assman 1991).

5.2. Recombinant 2-enoyl-CoA hydratase 2 derived fromrat MFE-2 (II)

The region of cDNA encoding amino acid residues 318–735 of rat MFE-2 was cloned intotheE. coli expression vector pET-3a, yielding the plasmid pET-Hydr2. The 1279 bp insertin the plasmid was expressed inE. coli BL21 (DE3) plysS cells and the recombinantprotein was purified (Table 1 of paper II). The molecular mass of the protein wasdetermined to be 46 kDa by SDS-PAGE (Fig. 1 of paper II), which agrees with 46.58 kDapredicted from the amino acid sequence. Immunoblotting showed that an antibody to therat hydratase 2 (Malilaet al. 1993) recognizes the 46 kDa-hydratase 2 (Fig. 1 of paper II).

When the 46 kDa-hydratase 2 was incubated with 30 nmol crotonyl-CoA, the productof the hydration reaction serves as a substrate for (3R)-hydroxyacyl-CoA dehydrogenasebut not for the (3S)-hydroxyacl-CoA dehydrogenase. The kinetic constants of the 46 kDa-hydratase 2 with (2E)-enoyl-CoA substrates were determined (Table 2 of paper II). TheKm value with (2E)-decenoyl-CoA (C10) was one tenth of that with crotonyl-CoA (C4).The catalytic efficiency with C10/C4 substrates is 11.3, which was consistent with apreviously reported value (14.1) for rat liver hydratase 2 (Hiltunenet al. 1989). Thespecificity constant (kcat/Km) of the hydratase 2 with crotonyl-CoA was about 104 timeslower than that of mitochondrial short chain 2-enoyl-CoA hydratase 1 (for a review, seeSchulz 1987). The enzyme preparation retained> 90% of the original activity when it wasstored in the buffer containing 200 mM potassium phosphate at pH 7.4 at 4°C for a year.

Since Xu and Cuebas (1996) reported that the product of hydration of (24E)-∆24-THCA-CoA by peroxisomal MFE-1 is not a physiological isomer, hydratase 2 derivedfrom rat MFE-2 was investigated. The protein hydrates (24E)-3α, 7α, 12α, 24-tetrahydroxy-5β-cholestanoyl-CoA [(24E)-∆24-THCA-CoA] to (24R, 25R)-3α, 7α, 12α,24-tetrahydroxy-5β-cholestanoyl-CoA [(24R, 25R)-∆24-OH-THCA-CoA], which has beenpreviously identified to be a physiological intermediate in bile acid synthesis (Fig. 2 ofpaper II).

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5.3. Site-directed mutagenesis of the 2-enoyl-CoA hydratase 2 (III)

To characterize the hydratase 2 reaction, two approaches were applied. One wascomplementation ofS. cerevisiae fox-2cells with a wild-type human MFE-2 (HsMFE-2)and its variantsin vivo. The other was production of a recombinant protein inE. coli tostudy its propertiesin vitro.

Because amino acid sequences of mammalian hydratase 2(s) differ considerably fromthose of mammalian mitochondrial hydratase 1(s) and theE. coli β-oxidation multienzymecomplex (Yanget al. 1991), the putative catalytic amino acid residues were identified bythe sequence alignment of the hydratase 2 domain ofHsMFE-2 with the other knownhydratase 2 molecules (Fig. 2 of paper III). Each of the identified conserved proticresidues was substituted to alanine by site-directed mutagenesis. WhenHsMFE-2 wasexpressed inS. cerevisiae fox-2cells, it was found to complementfox-2 cells and restorethe ability of the cells to grow on oleic acid as a carbon source. In this case,fox-2 cellscould serve as a tool for examining different variants ofHsMFE-2 in vivo. The expressedprotein in soluble yeast extracts was recognized as 79 kDa by an antibody to the rathydratase 2. Five variants (Tyr347Ala, Glu366Ala, Tyr505Ala, Asp510Ala andHis515Ala) did not complementfox-2as indicated by failure to produce zones of clearingon the oleic acid plates (Fig. 3 of paper III). The variants Tyr510Ala and His515Ala lostboth their hydratase and dehydrogenase activities (Table 2 of paper III), suggesting thatthese substitutions probably influence the folding of the protein and were not studiedfurther.

HsMFE-2(dh∆) with a deleted dehydrogenase domain and its variants,HsMFE-2(dh∆E366A) andHsMFE-2(dh∆D510A), were expressed inE. coli and purified (Fig. 4Bof paper III). Native molecular masses ofHsMFE-2(dh∆) and its variants were 83 kDa asdetermined by size-exclusion chromatography (Fig. 4A of paper III). Dynamic lightscattering measurement indicated that the purified recombinant hydratase 2 samples weremonodisperse and had hydrodynamic radii corresponding to a dimer. CD far UV-spectroscopy revealed that the molar ellipticities of the purified proteins were virtuallyidentical (Fig. 5 of paper III). Thekcat value of variant Glu366Ala is almost 100-fold lowerthan that of wild type at pH 5, but the Km value is not significantly affected by thissubstitution (Table 3 of Paper III). The pH-dependence curve ofkcat/Km is bell-shaped,giving pKa1 of 8.2± 0.1 and pKa2 of 9.7± 0.2, whereas the variant Glu366Ala has a lowerpKa1 of 6.5± 0.1 (Fig. 6 of paper III). The variant Asp510Ala is completely inactive.

5.4. Analysis of the two catalytic domains of (3R)-hydroxyacyl-CoAdehydrogenase of yeast peroxisomal MFE-2 (IV)

The yeast peroxisomal MFE-2(s) has two (3R)-hydroxyacyl-CoA dehydrogenase domains,A and B, at the N-terminus, both of which consist of a NAD+ binding site and a catalyticmotif (Fig. 1 of paper IV, Jörnvallet al. 1995). Based on the Gly16Ser mutation identifiedin human MFE-2 deficiency (van Grunsvenet al. 1998), a substitution of glycine to serinewas introduced into the NAD+ binding site ofS. cerevisiaeMFE-2 to dissect the twodehydrogenase domains (Fig. 2 of paper IV), resulting inScMFE-2(a∆), ScMFE-2(b∆)

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and ScMFE-2(a∆b∆) variants. In order to reveal the physiological roles of these twodehydrogenase domains,S. cerevisiae fox-2cells were taken as a model system and testedfor complementation. The results show thatS. cerevisiae fox-2cells transformed with theexpression plasmids containingScMFE-2(a∆) or ScMFE-2(b∆) developed zones ofclearing on oleic acid plates, whereasScMFE-2(a∆b∆) failed to grow (Fig. 3 of paper IV).In liquid oleic acid medium, the growth rates offox-2 cells expressingScMFE-2(a∆) orScMFE-2(b∆) were about 50% of that found in wild typeScMFE-2 (Fig. 3 of paper IV).To verify thatS. cerevisiaeMFE-2 and its variants were expressed infox-2, both 2-enoyl-CoA hydratase 2 and (3R)-hydroxyacyl-CoA dehydrogenase activities were measured inthe soluble yeast extracts (Table 1 of paper IV). With the exception of thefox-2 cells,hydratase 2 activity was detected in all transformants. The dehydrogenase activitydecreased inScMFE-2(a∆) andScMFE-2(b∆), but it was completely absent in theScMFE-2(a∆b∆) variant.

C. tropicalis MFE-2 with a deleted 2-enoyl-CoA hydratase 2 domain [CtMFE-2(h2∆)]plus variants of domains A and B [CtMFE-2(h2∆a∆), CtMFE-2(h2∆b∆) and CtMFE-2(h2∆a∆b∆)] were overexpressed inE. coli, purified and characterized (Table 2 of paperIV). Size-exclusion chromatography (Fig. 4 of paper IV) and CD spectroscopy (Fig. 5 ofpaper IV) showed thatCtMFE-2(h2∆) and variants were dimeric proteins with similarsecondary structural elements. TheCtMFE-2(h2∆) showed the highest catalytic efficiency(kcat/Km) with the substrate (3R)-hydroxydecanoyl-CoA (C10). Interestingly, thedehydrogenase activity ofCtMFE-2(h2∆) broke into two different profiles when thevariants were analyzed (Table 3). The catalytic constant (kcat) of CtMFE-2(h2∆a∆) for C4was the same as that ofCtMFE-2(h2∆) (29 ± 1 s-1 vs 31 ± 2 s-1), whereaskcat of CtMFE-2(h2∆b∆) was below the detection limit, indicating that domain B is solely responsible forthe utilization of C4 substrate. Thekcat values ofCtMFE-2(h2∆a∆) for C10 and C16 were17 ± 1 s-1 and 12± 2 s-1, respectively. Interestingly, thekcat values ofCtMFE-2(h2∆b∆) forC10 and C16 were 33± 2 s-1 and 36± 6 s-1, respectively, suggesting that domain Acontributes more than domain B to the metabolism of medium- and long-chain substrates.The activity ofCtMFE-2(h2∆a∆b∆) toward the substrates tested was below the detectionlimits (<0.001µmol x mg-1 x min-1) of the assay.

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6. Discussion

6.1. A novel peroxisomal multifunctiona l enzyme type 2 (I)

Although mammalian 2-enoyl-CoA hydratase 2 was first found to be involved in theepimerization reaction (Hiltunen et al. 1989, Smeland et al. 1989) and purified from eithermicrosomes or peroxisomes (Malila et al. 1993, Li et al. 1990), its physiological functionin mammals was largely unknown. To investigate its role in fatty acid metabolism ofmammalian cells, the microsomal 31.5 kDa hydratase 2 was cloned from rat liver in thepresent work.

Surprisingly, the results demonstrate that hydratase 2 is an integral part of a novelmammalian peroxisomal MFE-2. The presence of both (R)-specific dehydrogenase andhydratase 2 activities were confirmed by expression of the rat MFE-2 in P. pastoris (Table2 of paper I). Mammalian MFE-2 has a SCP-2 like domain at the C-terminus. It has beenreported that SCP-2 is not only a non-specific lipid transfer protein but also acts as aperoxisomal acyl-CoA binding protein (for a review, see Wirtz et al. 1998). The role ofthe SCP-2 homologue in MFE-2(s) requires further investigation. Mammalian MFE-2shows a pronounced sequence similarity to yeast peroxisomal MFE-2, suggesting that bothmammalian and yeast MFE-2(s) have evolved from a common ancestor and have beenassociated with β-oxidation of fatty acids via a (3R)-hydroxyacyl-CoA intermediate.

In mammalian cells, there is a conventional peroxisomal MFE-1 displaying 2-enoyl-CoA hydratase1/(3S)-hydroxyacyl-CoA dehydrogenase/∆3-∆2-enoyl-CoA isomerase.MFE-1 has a similar function as MFE-2 in β-oxidation of straight chain fatty acids but viaa (3S)-hydroxyacyl-CoA intermediate. Even though MFE-1(s) have not been found inyeast and MFE-1 shares no sequence similarities with mammalian MFE-2(s), rat MFE-1complements the S. cerevisiae fox-2 and restores the ability of fox-2 cells to grow on oleicacid (Filppula et al. 1995). This implies that the two types of peroxisomal MFEs arefunctionally related.

Mammalian MFE-2 is a protein that was previously identified as 17β HSD type IV(Leenders et al. 1994). As a multifunctional enzyme, MFE-2 was cloned and characterizedsimultaneously from mouse (Normand et al. 1995), rat (paper I, Dieuaide-Noubhani et al.1996, Corton et al. 1996), and man (Adamski et al. 1995, Jiang et al. 1997) by fiveindependent research groups.

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The results suggested a new organization of the peroxisomalβ-oxidation system inmammalian peroxisomes, which consists of two parallel pathways following reciprocalstereochemistry. The significance of the two alternative pathways in mammals for theβ-oxidation of fatty acids requires further examination.

6.2. Recombinant 2-enoyl-CoA hydratase 2 derived fromrat MFE-2 (II)

Mammalian peroxisomes are capable ofβ-oxidizing not only straight-chain fatty acids, butalso isoprenoid-derived 2-methyl-branched fatty acids and carboxyl side-chains of the bileacid intermediates di- and trihydroxycoprostanic acids. Since rat MFE-2 slowlymetabolizes 2-methyl-branched substrates pristenoyl-CoA and 2-methyltetradecenoyl-CoAesters (Table 2 of paper I), the question was raised whether this enzyme participates in thechain shortening of cholesterol during the synthesis of primary bile acids. The resultsdemonstrate that the recombinant hydratase 2 hydrates (24E)-∆24-THCA-CoA to (24R,25R)-OH-THCA-CoA, which is a physiological isomer, subsequently dehydrogenated bythe dehydrogenase domain of rat MFE-2 (Dieuaide-Noubhaniet al. 1996). Bile acidprofiles of a patient with MFE-2 deficiency show an accumulation of (24R, 25R)- and(24R, 25S)-THCA, as well as, small amounts of their (24S) counterparts (Uneet al.1997), implying that an alternative pathway for the formation of 24-keto-THCA-CoAwould operate by a combination of MFE-1 andα-methylacyl-CoA racemase (Ikegawaetal. 1995) in the following steps: i) the hydration of (24E)-∆24-THCA-CoA to (24S, 25S)-24-OH-THCA-CoA catalyzed by the hydratase 1 domain of MFE-1, ii) the hypotheticalracemization of theα-methyl group to (24S, 25R)-24-OH-THCA-CoA by anα-methylacyl-CoA racemase, and iii) the dehydrogenation of (24S, 25R)-24-OH-THCA-CoA to 24-keto-THCA-CoA catalyzed by MFE-1 (Scheme 2 of paper II). Althoughα-methylacyl-CoA racemase was purified from rat (Schmitzet al. 1994) and human(Schmitz et al. 1995) livers andα-methyl-THCA-CoA racemase activity was found inperoxisomes (Ikegawaet al. 1995), direct evidence that this racemase is involved in theracemization of theα-methyl group of 24-OH-THCA-CoA diastereomers is still missing.Obviously, since hydratase 1 is commercially available and is widely applied in metabolicstudies, no monofunctional hydratase 2 has been found in nature. Thus, this recombinanthydratase 2 is a useful tool for studying stereospecificities in theβ-oxidation pathway.

6.3. Site-directed mutagenesis of the 2-enoyl-CoA hydratase 2 (III)

Hydratase 2 was first cloned from yeastS. cerevisiaeas a domain of peroxisomal MFE-2(Hiltunen et al. 1992) and subsequently cloned from pig (Leenderset al. 1994), mouse(Normandet al. 1995), humans (Adamskiet al. 1995) and rat (paper I). The physiologicalrole of the mammalian hydratase 2 was revealed to be associated with bile acid synthesis(paper II). However, the amino acid residues participating in catalysis were not yet known.

By complementingS. cerevisiae fox-2with HsMFE-2 and its variantsin vivo, three

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amino acid residues Glu366, Tyr347 and Asp510 were identified to affect only hydratase2 activity. HsMFE-2(dh∆), HsMFE-2(dh∆E366A) and HsMFE-2(dh∆D510A) wereexpressed inE. coli and purified (Fig. 4 of paper III).HsMFE-2(dh∆Y347A) could not bepurified due to the changes in its chromatographic behavior although it was recognized inthe soluble extract by the antibody to the rat hydratase 2. The pH-dependence curve ofkcat/Km of HsMFE-2(dh∆) is bell-shaped (Fig. 6 of paper III), suggesting that two proticresidues participate in catalysis. The Km value ofHsMFE-2(dh∆E366A) is similar to thatof HsMFE-2(dh∆), indicating that the Glu366Ala substitution does not affect substratebinding of the enzyme. However, the pKa1 of HsMFE-2(dh∆E366A) decreases from 8.2to 6.5, showing that the substitution affects the catalytic efficiency of the enzyme at lowpH (Table 3 of Paper III). This suggests that the negatively charged side-chain of Glu366acts as a base in catalysis.HsMFE-2(dh∆D510A) was enzymatically inactive. CD far UV-spectroscopy revealed that the molar ellipticities ofHsMFE-2(dh∆D510A) andHsMFE-2(dh∆) are identical (Fig. 5 of paper III), indicating that the substitution does not affect thecomposition of secondary structural elements. Alignment of the amino acid sequence ofHsMFE-2(dh∆) with other MFE-2 proteins reveals a new fingerprint of hydratase 2, Ala-Ala-(Leu, Ile)-Tyr-Arg-Leu-(Xaa)-Ser-Gly-Asp510-Xaa-Asn-Pro-Leu-His-(Ile, Val)-Asp-Pro-Xaa-(Phe, Leu)-Ala-(Xaa)4-Phe-(Xaa)2-Pro-Ile-Leu-His-Gly-(Leu, Met)-Cys-(Thr,Ser)-Xaa-Gly (Fig. 2 of Paper III). Asp510 is highly conserved in this motif fromprokaryotes to eukaryotes and plays an important role in the hydratase 2 reaction, eitheracting as an acid in the catalysis or affecting the binding of substrate. Basically, thehydratase 2 reaction also follows a general acid-base catalysis, like the hydratase 1reaction (Engelet al. 1996, Kiemaet al. 1999). This implies that both reactions utilize acommon mechanistic strategy for lowering the free energies of the rate-limiting transitionstates. Surprisingly, the R-specific dehydratase ofS. cerevisiaefatty acid synthase (Chiralaet al. 1987) also contains this motif, which implies that the hydratase 2 and thedehydratase converged at a particular point during evolution.

The straight-chain fatty acids are substrates for rat MFE-2in vitro (Table 2 of paper I).In line with this observation,HsMFE-2 complements and restores the ability offox-2cellsto utilize oleic acid (cis-C18:1(9)) as the carbon source, which provides the first evidence onthe ability of MFE-2 to metabolize straight-chain acyl-CoA estersin vivo. In line with thisobservation, the patients with MFE-2 deficiency were characterized by accumulation ofvery-long-chain fatty acids in plasma and fibroblasts (Suzukiet al. 1997, Van Grunsvenetal. 1998). The available gene structure ofHsMFE-2 (Leenderset al. 1998) will help thesequence analysis of such patients. One case of MFE-2 deficiency is caused by a deletionof the cDNA encoding amino acid residues 480-501 in the hydratase 2 domain, resultingin absence of the enzyme as shown by immunofluorescence staining (Van Grunsvenet al.1999). The identification of important amino acid residues (Glu366 and Asp510) willprovide a deep insight into the functional analysis of MFE-2 deficiency in future.

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6.4. Analysis of the two catalytic (3R)-hydroxyacyl-CoAdehydrogenase domains of yeast peroxisomal

MFE-2 (IV)

Yeast peroxisomal MFE-2 provides a good example for studying gene duplication duringevolution. The interesting question is raised of what the physiological functions of the twodehydrogenase domains are and whether they show enzymatic activities. By applying site-directed mutagenesis to inactivate either domain A or domain B, bothin vivo and in vitrostudies indicate that these two domains play important roles in the utilization of fatty acidsas a sole carbon source in yeast cells. This conclusion is further supported by CD far UVspectroscopy ofCtMFE-2(h2∆) and its variants, which show that the composition ofsecondary structural elements are not affected. Conceivably, the differences in the kineticdata are due to the substitutions affecting catalytic sites of each domain rather than overallfolding of the proteins.

It is also worth noting that the amino acid sequence of domain A of yeast MFE-2 ismore homologous to that ofHsMFE-2 (50% identity) than to domain B, suggesting thatthe dehydrogenase domain ofHsMFE-2 may have evolved from the yeast dehydrogenasedomain A.

It was previously suggested that gene duplication has led to the evolution of separateenzymatic activities, such as mammalian acyl-CoA dehydrogenases, which are presentedas several paralogs (Tanakaet al. 1990). An acquisition of two domains with differentchain-length specificities within a single polypeptide is a novel strategy for amultifunctional enzyme to overcome the problems related to the metabolism of a largevariety of substrates.

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7. Conclusions

Cloning of rat 2-enoyl-CoA hydratase 2 reveals that it is an integral domain of a novelmammalian MFE-2. The cDNA of rat MFE-2 has an open reading frame of 2205 bpencoding a 735 amino acid residues with a molecular mass of 79 kDa. Recombinant MFE-2 generated in yeast P. pastoris has 2-enoyl-CoA hydratase 2 and (3R)-hydroxyacyl-CoAdehydrogenase activities. The enzyme catalyzes reactions of several different substrates,including straight-chain (2E)-enoyl-CoA, branched 2-methyltetradecenoyl-CoA,pristenoyl-CoA esters and steroids. It was shown that in addition to the earlier describedMFE-1, a novel MFE-2 exists in rat liver. Both MFE-1 and MFE-2 catalyze the sequentialhydratase and dehydrogenase reactions of β-oxidation but through reciprocalstereochemical courses.

Recombinant hydratase 2 derived from rat MFE-2 was overexpressed in E. coli and thepurified enzyme was found to hydrate (24E)-3α, 7α, 12α-trihydroxy-5β-cholest-24-enoyl-CoA to (24R, 25R)- 3α, 7α, 12α, 24-tetrahydroxy-5β-cholestanoyl-CoA, which is aphysiological intermediate in bile acid synthesis. The result reveals that MFE-2, instead ofMFE-1, is involved in the synthesis of bile acid.

Several amino acid residues in human MFE-2 were shown, by complementation invivo, to be involved either in catalysis reactions of the hydratase by the hydratase domainor to influence folding of the protein. Based on the enzymatic properties, Glu366 andAsp510 were identified to be important amino acid residues for the catalysis reaction ofhydratase 2 in vitro. The pH-dependence curve of kcat/Km for the purified recombinanthuman hydratase 2 is bell-shaped, suggesting that the reaction utilize two protic aminoacid residues.

Analysis of the two catalytic (3R)-hydroxyacyl-CoA dehydrogenase domains of yeastperoxisomal MFE-2 reveals that one of the domains is more active on medium- and long-chain substrates while the other domain is responsible for metabolism of the short-chainsubstrates. The results suggest that both domains are definitely required for propagationof yeast cells in an optimal manner on fatty acids as the carbon source.

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