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Originally published In Press as doi:10.1074/jbc.M413362200 on January 11, 2005

J. Biol. Chem., Vol. 280, Issue 11, 9802-9812, March 18, 2005
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Breakdown of 2-Hydroxylated Straight Chain Fatty Acids via Peroxisomal 2-Hydroxyphytanoyl-CoA Lyase

A REVISED PATHWAY FOR THE {alpha}-OXIDATION OF STRAIGHT CHAIN FATTY ACIDS*

Veerle Foulon{ddagger}§, Mieke Sniekers{ddagger}§, Els Huysmans, Stanny Asselberghs, Vincent Mahieu, Guy P. Mannaerts, Paul P. Van Veldhoven, and Minne Casteels||

From the Afdeling Farmacologie, Departement Celbiologie, Katholieke Universiteit Leuven, Campus Gasthuisberg, 3000 Leuven, Belgium

Received for publication, November 29, 2004 , and in revised form, January 6, 2005.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
2-Hydroxyfatty acids, constituents of brain cerebrosides and sulfatides, were previously reported to be degraded by an {alpha}-oxidation system, generating fatty acids shortened by one carbon atom. In the current study we used labeled and unlabeled 2-hydroxyoctadecanoic acid to reinvestigate the degradation of this class of lipids. Both in intact and broken cell systems formate was identified as a main reaction product. Furthermore, the generation of an n–1 aldehyde was demonstrated. In permeabilized rat hepatocytes and liver homogenates, studies on cofactor requirements revealed a dependence on ATP, CoA, Mg2+, thiamine pyrophosphate, and NAD+. Together with subcellular fractionation data and studies on recombinant enzymes, this led to the following picture. In a first step, the 2-hydroxyfatty acid is activated to an acyl-CoA; subsequently, the 2-hydroxy fatty acyl-CoA is cleaved by 2-hydroxyphytanoyl-CoA lyase, to formyl-CoA and an n–1 aldehyde. The severe inhibition of formate generation by oxythiamin treatment of intact fibroblasts indicates that cleavage through the thiamine pyrophosphate-dependent 2-hydroxyphytanoyl-CoA lyase is the main pathway for the degradation of 2-hydroxyfatty acids. The latter protein was initially characterized as an essential enzyme in the peroxisomal {alpha}-oxidation of 3-methyl-branched fatty acids such as phytanic acid. Our findings point to a new role for peroxisomes in mammals, i.e. the breakdown of 2-hydroxyfatty acids, at least the long chain 2-hydroxyfatty acids. Most likely, the more abundant very long chain 2-hydroxyfatty acids are degraded in a similar manner.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
In mammals, 2-hydroxyfatty acids (2-OH-FA)1 are present in several tissues but are most abundant in brain, where they represent ~6% of the total fatty acids. The 2-hydroxy derivatives of C18 to C26 straight chain saturated and/or mono-unsaturated fatty acids appear to be present exclusively in cerebrosides, cerebroside sulfates, and ceramides, most of which are found in myelin (1). Furthermore, in brain cerebrosides even more than half of the fatty acids are 2-OH-FA, and also odd-numbered fatty acids are present in an unusually large proportion (2). The ratio of 2-OH-FA to normal fatty acids increases during myelination, whereas the percentage of odd-numbered fatty acids continues to increase up to the age of 10–15 years (3).

In 1964 Levis and Mead (4) reported for the first time the existence of an {alpha}-oxidation system for the degradation of the C20 to C26 straight chain fatty acids of rat brain sphingolipids. It was postulated that this pathway would consist of two steps, generating first 2-hydroxy even-numbered fatty acids, and subsequently odd-numbered fatty acids one carbon atom shorter. Later, it was reported that in rat brain the decarboxylation reaction was performed by a microsomal enzyme and that a 2-keto fatty acid, found only in small amounts, was formed as an intermediate (5). {alpha}-Oxidation of straight chain fatty acids has also been studied in plants, and the formation of an n – 1 aldehyde was reported by several authors (6–8). {alpha}-Oxidation has also been described in yeast (9) and in protozoa (10).

More recently, the involvement of peroxisomes in the {alpha}-oxidation of cerebronic acid (2-hydroxytetracosanoic acid) was described. The decarboxylation of 2-hydroxytetracosanoic acid was apparently independent of the preceding formation of an acyl-CoA and was supposed to be distinct from the {alpha}-oxidation of 3-methyl-branched fatty acids such as phytanic acid (11). The latter pathway is currently thought to proceed as follows; 1) activation to a CoA ester, 2) hydroxylation of carbon 2 by phytanoyl-CoA hydroxylase (PAHX), and 3) cleavage of the hydroxylated CoA-ester by 2-hydroxyphytanoyl-CoA lyase (2-HPCL) to formyl-CoA (12) and a 2-methyl-branched fatty aldehyde (13) in a TPP-dependent manner (14). Both PAHX and 2-HPCL are peroxisomal enzymes.

According to our data PAHX does not act on straight chain fatty acids or their CoA esters (15, 16) and, hence, cannot be involved in the formation of 2-hydroxyfatty acids; others claim, however, that PAHX can hydroxylate straight chain acyl-CoAs (17). Regardless of this discrepancy, a recently described fatty acid 2-hydroxylase, highly abundant in brain and encoded by the FA2H gene (18), is likely responsible for the formation of 2-hydroxyfatty acids in man.

The current study was undertaken to elucidate the degradation of 2-hydroxyfatty acids and to highlight a possible role of 2-HPCL in this process. Hereby, we made use of 2-hydroxyoctadecanoic acid, labeled and unlabeled, and its unlabeled CoA ester. These substrates are easier to synthesize and manipulate than the more abundant very long chain 2-hydroxyfatty acids. Although 2-hydroxyoctadecanoic acid is a less abundant 2-hydroxyfatty acid, it forms a more than negligible fraction of the 2-hydroxyfatty acids in brain cerebrosides (4% in newborn, 1% in adult brain) and sulfatides (15 and 1%, respectively) (3).


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Fatty Acids and Derivatives—[1-14C]Hexadecanoic acid was from PerkinElmer Life Sciences. 2-Hydroxyhexadecanoic and 2-hydroxyeicosanoic acid were from Larodan. 2-Keto-octanoic acid was obtained from Serva. The synthesis of most fatty acids/derivatives has been described before: 2-hydroxy-3-methyl[1-14C]hexadecanoic acid, labeled and unlabeled 3-methylhexadecanoic acid, 3-methylhexadecanoyl-CoA, and 2-hydroxy-3-methylhexadecanoyl-CoA (19); 2-methylhexadecanoic acid and 2-methylhexadecanoyl-CoA (20); 3-hydroxy-3-methylhexadecanoyl-CoA (14), 3-hydroxy-2-methylhexadecanoyl-CoA (21); tetradecanal, 2-methylpentadecanal, and heptadecanal (13).

2-Hydroxyoctadecanoic acid was purchased from Larodan or synthesized as follows (adapted from Sandhir et al. (11) and Ashton et al. (22)). Stearic acid treated with PBr3 was brominated in dry dichloromethane in a closed screw-capped thick-wall vial for 72 h at 70 °C (Hell-Volhard-Zelinski reaction; molar ratio acid/PBr3/Br2, 1/12/36). After removal of the solvent, water was carefully added to hydrolyze the 2-bromooctadecanoylbromide, and the 2-bromooctadecanoic acid (Rf = 0.50; Silica gel 60; hexane/diethyl ether/acetic acid, 60/40/1, v/v; stearic acid Rf = 0.59) was extracted into diethyl ether. The bromo group was converted to a hydroxyl group in two steps. The dried diethyl ether extract was dissolved in acetic acid containing potassium acetate and placed under reflux at 120 °C for 24 h. After dilution with water, the 2-acetoxyoctadecanoic acid (Rf = 0.26) was extracted into diethyl ether, dried, and hydrolyzed in 1.5 N NaOH/methanol (15/85, v/v) for 2 h under reflux. The mixture was acidified, and the formed 2-hydroxyoctadecanoic acid was extracted into diethyl ether and further purified by preparative TLC (Rf = 0.14). Overall yield was 35%. 2-Hydroxy[1-14C]octadecanoic acid and 2-hydroxynonadecanoic acid were synthesized in a similar manner, starting from [1-14C]octadecanoic acid (Moravek Biochemicals, Inc.) and nonadecanoic acid (Fluka), respectively. The CoA esters of 2-OH-FA were prepared by transesterification of their thiophenol ester, prepared with N,N'-dicyclohexylcarbodiimide, in 0.5 M NaHCO3/tetrahydrofuran/ethanol (1/5/2, v/v) and purified on a C18-SPE cartridge (1 g; Supelco). Yield was 66% based on CoA input.

Animals and Cell Lines—Male Wistar rats weighing ~200 g were maintained on a standard laboratory diet and a constant light-dark cycle and were fasted overnight before sacrifice. All studies were approved by the University Ethics committee. Control human skin fibroblasts and fibroblasts from X-linked adrenoleukodystrophy patients were kindly provided by Dr. G. Matthys (Center for Human Genetics, Leuven, Belgium). Skin fibroblasts from patients affected with multiple acyl-CoA dehydrogenase deficiency, medium chain acyl-CoA dehydrogenase deficiency, and Zellweger syndrome were provided by Dr. J. Van Hove (University Hospitals, Leuven). Rhizomelic chondrodysplasia punctata type 1 fibroblasts and rat C6 glial cells were obtained from ATCC (Manassas, VA). The preparation of fibroblasts from Pex5+/– and Pex5–/– mice has been described before (23).

Cells were cultured at 37 °C and 5% CO2 in Dulbecco's modified Eagle's medium containing Glutamax (Invitrogen) and a mixture of antibiotics and antimycotics (Invitrogen) and supplemented with 10% fetal calf serum (Invitrogen) or 2% Ultroser G (BioSepra).

Rat hepatocytes were isolated as described by Mannaerts et al. (24) and permeabilized as described by Croes et al. (25). Mouse hepatocytes were prepared as slightly modified from Honkakoski and Negishi (26), plated in collagen-coated T25 culture flasks at 0.5 x 106 cells, and used after an overnight recovery period.

Preparation of Homogenates, Subcellular Fractions, and Cell Lysates—Homogenates of rat liver and brain were prepared in 0.25 M sucrose containing 5 mM Mops-NaOH, pH 7.2, and 0.1% (v/v) ethanol (homogenization medium). Subcellular fractionation of rat liver and brain was done essentially as described previously (27) for rat liver, with some slight modifications for brain; the pellet obtained after centrifugation of a rat brain postmitochondrial supernatant at 13,000 x g (L-fraction) was resuspended in 0.85 M sucrose containing 5 mM Mops-NaOH, pH 7.2, and 0.1% (v/v) ethanol, overlaid with an equal volume of homogenization medium, and centrifuged for 45 min at 108,000 x g (Beckman SW55) to remove most of the myelin (28). Peroxisomal membranes were obtained after subfractionation of the peroxisomal fraction over a Nycodenz gradient and subsequent separation of matrix and membranes by sonication and centrifugation. Marker enzymes and protein were measured as described previously (27, 29). Cells, grown to confluence in 175 cm2 flasks, were harvested by trypsinization, pelleted, washed twice with phosphate-buffered saline, and homogenized by sonication (Branson Sonifer; output 4) in 1 ml of 0.25 M Tris-HCl, pH 7.2.

Fatty Acid Oxidation by Intact and Permeabilized Cells—For measurement of fatty acid oxidation in intact rat hepatocytes, 1.25 x 106 cells were incubated for 10 min in 0.5 ml of Krebs-Henseleit buffer, pH 7.4, containing 100 µM defatted albumin, 20 mM Hepes buffer, and 50 µM concentrations of the appropriate substrate (standard conditions). Incubations with permeabilized and washed hepatocytes were started by adding a 100-µl cell suspension (1.25 x 107 hepatocytes/ml) to 400 µl of incubation mixture containing 50 µM labeled substrate and the appropriate cofactors. Adherent cells (mouse hepatocytes, human and mouse fibroblasts, and rat C6 glial cells) were grown to near confluence in 25-cm2 flasks and incubated (mouse hepatocytes for 6 h (30), other cells for 24 h) with 4 µM labeled substrate in culture medium in the presence of 0.2% Ultroser (31).

Oxidation rates in homogenates and subcellular fractions were measured in 20 mM Hepes-NaOH, pH 7.2, 25 µM BSA, 50 µM radioactive substrate, and the appropriate cofactors. All reactions were terminated after 10 min by adding HClO4 to a final concentration of 2% unless otherwise specified. The released 14CO2 and the amounts of [14C]formate and 14C-labeled acid-soluble material were determined as described before (19).

Enzyme Activity Measurements—The production of acyl-CoA esters was measured in 20 mM Hepes-NaOH, pH 7.2, 12.5 µM BSA, 4 mM ATP, 0.5 mM CoA, and 50 µM 1-14C-labeled substrate (250 µl final volume). Reactions were terminated by the addition of 2 ml of isopropanol/0.1 N HCl (1/1, v/v); fatty acids were extracted with 4 ml of heptane, and an aliquot of the water layer containing the CoA-esters was counted in a liquid scintillation counter (PerkinElmer Life Sciences).

When using 1-14C-labeled substrates 2-hydroxyphytanoyl-CoA lyase activity was quantified by measuring [14C]formate and its oxidation product, 14CO2 (32), since the primary 2-HPCL product, [14C]formyl-CoA, is quickly hydrolyzed to formate (12). Incubations (37 °C) were performed in a final volume of 250 µl containing 50 mM Tris-HCl, pH 7.5, 6.6 µM BSA, 0.8 mM MgCl2, and 20 µM TPP (referred to as standard conditions) with 40 µM substrate (2-hydroxy[1-14C]octadecanoic acid, 2-hydroxy-3-methyl[1-14C]hexadecanoyl-CoA, 2-hydroxy-3-methyl[1-14C]hexadecanoic acid, 3-methyl[1-14C]hexadecanoyl-CoA, 3-methyl[1-14C]hexadecanoic acid, [1-14C]hexadecanoyl-CoA, or [1-14C]hexadecanoic acid). For the competition experiments a substrate concentration of 10 µM was used, and a 50 µM concentration of a related compound was added. When using unlabeled substrates (2-hydroxyoctadecanoyl-CoA, 2-hydroxy-3-methylhexadecanoyl-CoA), 2-HPCL activity was quantified by measuring the formation of the n – 1 aldehyde.

Lipid Analysis—The identification of the CoA esters was based on the formation of fluorescent acyl etheno-CoA derivatives by bromoacetaldehyde (prepared by refluxing bromodiethylacetal under acidic conditions and brought to pH 4.6 with sodium acetate, up to a final concentration of 100 mM acetate) (adapted from Larson and Graham (33)). Briefly, standard incubations (250 µl final volume) containing 50 µM unlabeled substrate were stopped by the addition of 25 µl of 1 N H2SO4. After the addition of 25 nmol of the appropriate internal standard, the samples were extracted with 1.2 ml of isopropanol/heptane (4/1, v/v), and the upper phase was evaporated under N2. The residue was reconstituted in 100 µl of water and transferred to a derivatization vial. Bromoacetaldehyde reagent (200 µl; ~250 mM, pH 4.6) was added, the samples were kept in the dark, placed at 80 °C for 15 min, and immediately afterward put on ice. An aliquot of the derivatization mixture was injected onto a C18 column (Symmetry; 150 x 4.6 mm; 5 µm; 100 Å, Waters) on a Waters 1525 HPLC. The acyl-CoA esters were eluted with a gradient of acetonitrile in 0.25 M ammonium acetate buffer, pH 5.0: linear gradient 10–66%, 15 min; linear gradient 66–80%, 2 min; isocratic 80%, 2 min; linear gradient 80–10%, 2 min; isocratic 10%, 6 min. Detection was performed on a Waters 2475 fluorescence detector (excitation 230 nm; emission 420 nm).

For the identification and analysis of aldehydes, reactions were stopped with 125 µl of 2 N HCl. After the addition of 2.5 nmol of the appropriate internal standard and 125 µl of a solution of 2,4-dinitrophenylhydrazine (3.75 mg in 5 ml of 2 N HCl), the samples were incubated for 30 min at 50 °C (in this and subsequent steps samples were protected from light). After adding 0.5 ml of methanol, the hydrazones were extracted into 2 ml of hexane. 1.75 ml of the upper phase was evaporated, the residue was dissolved in 88 µl of acetonitrile, and 50 µl (1.25 nmol of internal standard) was injected on a C18 column (Symmetry 150 x 4.6 mm; 5 µm, 100 Å Waters; Waters 1525 HPLC system) eluted with acetonitrile under isocratic conditions. Detection was done at 360 nm (Waters 484 tunable absorbance detector).

Generation and Purification of Recombinant Hs 2-HPCL—Human 2-HPCL cDNA was amplified from a human liver cDNA library with primers Hs 2-HPCL-F8 (5'-CGCGGATCCGATGCCGGACAGTAAACTTCGC) and Hs 2-HPCL-R8 (5'-AATGCATGCTTACATATTAGAGCGGGTC), and after digestion with BamHI and SphI, the corresponding PCR product was subcloned in pJR233 (34) (construct yVF3). Plasmid yVF3 was transformed into competent Saccharomyces cerevisiae CB80 cells. Transformants were selected and grown on minimal essential medium containing 0.67% (w/v) yeast nitrogen base without amino acids, 2% (w/v) glucose, and a supplement of bases and amino acids (20–150 µg/ml) as required (Sc-ura). Positive colonies were grown at 33 °C in 50 ml Sc-ura containing 0.5% glucose as the sole carbon source. After 12–15 h, cells from 15 ml of culture were harvested by centrifugation, washed, and broken with glass beads in 400 µl of lysis buffer (50 mM potassium phosphate buffer, pH 7.4, containing 1 mM EDTA, 10 mM MgCl2, 1 mM TPP, and a mix of protease inhibitors). Cell debris was removed by centrifugation, and lyase activity was measured on 50-µl samples.

To generate a polyhistidine fusion product of Hs 2-HPCL, two oligos (Hs 2-HPCL-H1, 5'-CAAGATGGGCATCATCATCATCATCATCGG, and Hs 2-HPCL-H2, 5'-ATCCCGATGATGATGATGATGATGCCCCATCTTGGTAC) were allowed to hybridize to generate a small linker. The resulting adaptor sequence was cloned between the KpnI-BamHI sites of yVF3 (yVF7), and S. cerevisiae CB80 cells, transformed with yVF7, were grown for 18 h in 400 ml of Sc-ura containing 0.5% (w/v) glucose. The fusion protein was purified from cell lysate (prepared as described before in 8 ml of lysis buffer) on nickel nitrilotriacetic acid-agarose essentially as described for the purification of phytanoyl-CoA hydroxylase (16). Analysis of the purified fraction by SDS-PAGE (12% polyacrylamide, w/v) and subsequent immunoblotting with anti-His antibody (Clontech) revealed one single polyhistidine-tagged protein with a molecular mass of ~63 kDa. The yield was 77–400 µg of protein with a specific activity of about 26 milliunits/mg of protein (substrate, 40 µM 2-hydroxy-3-methyl[1-14C]hexadecanoyl-CoA).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Degradation of 2-OH-FA in Intact and Permeabilized Cells; Indication for a Role of Peroxisomes and 2-HPCL
Analysis of the medium of control human fibroblasts and rat C6 glial cells incubated with 2-hydroxy[1-14C]octadecanoic acid for different possible labeled degradation products revealed formate as the major acid-soluble oxidation product (Table I). In intact isolated rat and cultured mouse hepatocytes, on the other hand, the amount of acid-soluble material was markedly higher than the amount of formate generated (Table I), suggesting that in these cells, apart from formate, another metabolite was generated as the primary reaction product or that formate/CO2 was subsequently converted into another acid-soluble metabolite. When intact rat hepatocytes were incubated in the presence of increasing concentrations of unlabeled formate, the production of 14CO2 from 2-hydroxy[1-14C]octadecanoic acid decreased dramatically (Fig. 1). This decrease was accompanied by a compensatory increase in the generation of [14C]formate (Fig. 1), as was shown earlier for 3-methylhexadecanoic acid (Ref. 35; see also Fig. 1), whereas the {beta}-oxidation of straight chain fatty acids was unaffected throughout the whole formate concentration range (Fig. 1). Hence, these results support the contention that 2-hydroxyfatty acids are shortened by a process resembling the {alpha}-oxidation of phytanic acid whereby not CO2, but formyl-CoA/formate is the primary oxidation product that is subsequently converted into CO2 and possibly other metabolites.


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  		TABLE I
Oxidation of 2-hydroxyoctadecanoic acid in intact cells

CO2, formate, and acid-soluble material (ASM, including formate, which is retained in the water phase) production was measured in the indicated cells incubated with 2-hydroxy[1-14C]octadecanoic acid, as described under "Experimental procedures." Results are expressed as the mean ± S.E. of two or three experiments.

 



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  		FIG. 1.
Breakdown products of 2-hydroxy[1-14C]octadecanoic acid, 3-methyl[1-14C]hexadecanoic acid, and [1-14C]hexadecanoic acid in isolated intact rat hepatocytes. Isolated rat hepatocytes were incubated with the indicated fatty acids (200 µM for 2-hydroxy[1-14C]octadecanoic acid and 3-methyl[1-14C]hexadecanoic acid; 50 µM for [1-14C]hexadecanoic acid) in the presence of increasing concentrations of unlabeled formate. Incubations were stopped after 10 min with HClO4, and the amount of 14CO2 (white bars), [14C]formate (black bars), and [14C] acid-soluble material (gray bars) was measured.

 
To obtain a first insight in the organelle(s)/enzymes involved, the oxidation of 2-hydroxy[1-14C]octadecanoic acid was analyzed in fibroblasts from patients affected with various fatty acid oxidation disorders. In patients with the mitochondrial fatty acid oxidation disorders medium chain acyl-CoA dehydrogenase deficiency or multiple acyl-CoA dehydrogenase deficiency, characterized by a deficiency in the medium chain or multiple acyl-CoA dehydrogenases, respectively, no decrease in oxidation rates was observed (Table II). Hence, these data exclude a significant role of these mitochondrial dehydrogenases in the degradation of 2-hydroxylated straight chain fatty acids. Furthermore the oxidation rates of the 2-hydroxyfatty acids were not reduced either in cells from X-linked adrenoleukodystrophy patients, characterized by a deficiency in the peroxisomal activation/import of very long chain fatty acids, or in those from a rhizomelic chondrodysplasia punctata type 1 patient with a proven deficiency of Pex7p, the import receptor for peroxisomal matrix proteins containing a peroxisomal targeting signal-2 (Table II). Only in fibroblasts from a patient with Zellweger syndrome, with a demonstrated defect in the peroxisomal import of proteins containing a peroxisomal targeting signal-1, was a decrease (43%) in oxidation rate noticed. A similar decrease was seen in fibroblasts from Pex5–/– mice, a mouse model for Zellweger syndrome (Table II). These data suggest that peroxisomes are important and that peroxisomal enzymes with a peroxisomal targeting signal-1 signal are probably involved in the degradation of 2-OH fatty acids. The addition to the culture medium of etomoxir or tetradecylglycidic acid, inhibitors of carnitine palmitoyl-CoA transferase and, hence, of fatty acyl-CoA entry into the mitochondria, did not significantly affect the oxidation of 2-hydroxy[1-14C]octadecanoic acid (results not shown), another indication that mitochondria are likely not involved.


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  		TABLE II
Oxidation of 2-hydroxyoctadecanoic acid in human and murine fibroblasts

Production of CO2 and formate was measured in fibroblasts incubated with 4 µM 2-hydroxy[1-14C]octadecanoic acid, as described under "Experimental Procedures." Results were calculated as the mean values of two determinations per cell line. MADD, multiple acyl-CoA dehydrogenase deficiency; MCAD, medium chain acyl-CoA dehydrogenase deficiency; RCDP, rhizomelic chondrodysplasia punctata; X-ALD, X-linked adrenoleukodystrophy.

 
To study the necessary cofactors, rat hepatocytes, permeabilized with {alpha}-toxin to deplete the cytosolic cofactors (25), were incubated with different sets of cofactors. In these hepatocytes the oxidation of 2-hydroxy[1-14C]octadecanoic acid was strictly dependent on ATP, Mg2+, and CoA and was enhanced by TPP (Table III). Extra addition of NAD+ resulted in a further 20% increase of total oxidation.


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  		TABLE III
Cofactor requirements for oxidation of 2-hydroxyoctadecanoic acid in permeabilized hepatocytes

CO2 and formate production was measured in permeabilized rat hepatocytes incubated with 50 µM 2-hydroxy[1-14C]octadecanoic acid. Concentrations of the cofactors tested: ATP, 4 mM; CoA, 0.5 mM; Mg2+, 2.4 mM; TPP, 0.02 mM; NAD+, 2 mM. Results are expressed as % of the rates observed under standard conditions (ATP, CoA, Mg2+, and TPP; CO2, 5.10 nmol/108 cells/min, and formate, 25.48 nmol/108 cells/min) and are mean values of two experiments, except for ATP, CoA, Mg2+; ATP, CoA, Mg2+, NAD+; and NAD+, which was a single measurement.

 
The dependence on TPP was further investigated by culturing fibroblasts and C6 glial cells in the presence of the thiamine antimetabolite oxythiamin. The addition of this compound (1 mM) to the culture medium for several cell divisions reduced the overall oxidation rates of 2-hydroxyoctadecanoic acid by 44, 39, and 60% in control fibroblasts, rhizomelic chondrodysplasia punctata fibroblasts, and C6 glial cells, respectively. These data confirm the involvement of a TPP-dependent enzyme and are similar to the effect of oxythiamin on {alpha}-oxidation of 3-methylhexadecanoic acid. The addition of oxythiamin had no effect on the {beta}-oxidation of long chain fatty acids.2

The finding that ATP and CoA are essential suggests that the oxidation of 2-hydroxyfatty acids involves an activation reaction. The dependence of the pathway on TPP points to a reaction similar to that of the TPP-dependent cleavage of 2-hydroxy-3-methylacyl-CoA esters during the {alpha}-oxidation of 3-methyl-branched fatty acids.

Degradation of 2-OH-FA in Homogenates; Identification of Intermediates and End Products, Subcellular Localization, and Enzymes Involved
In rat liver homogenates similar results as those in permeabilized hepatocytes were obtained (results not shown): breakdown of 2-hydroxy[1-14C]octadecanoic acid in whole rat liver homogenates was dependent on the presence of CoA, ATP, Mg2+, and TPP. Although the oxidation rates in the presence of ADP were about 65% of those obtained with ATP, no oxidation was detected upon the addition of GTP or AMP. Furthermore, CoA could not be replaced by desulfo-CoA or dephospho-CoA.

In homogenates the main product was formate (>90% of total products). The addition of KCN did not affect the oxidation of 2-hydroxy[1-14C]octadecanoic acid (results not shown), again ruling out mitochondria as a key player.

Identification of Intermediates and End Products of 2-Hydroxyoctadecanoic Acid Oxidation—When rat liver homogenates were incubated with 2-hydroxy[1-14C]octadecanoic acid in the presence of ATP, CoA, and Mg2+, labeled acyl-CoA esters were generated. HPLC analysis of the etheno-CoA esters revealed a peak with significant fluorescence at 18.7 min coeluting with the 2-hydroxyoctadecanoyl-CoA standard (Fig. 2, A–C). The amount of 2-hydroxyoctadecanoyl-CoA (0.630 nmol/mg of protein/min) based on the internal standard, agreed well with the total amount of labeled CoA ester formed, based on extraction procedures (0.746 nmol/mg of protein/min) (mean of two measurements). Additionally, the generation of a 2-hydroxyoctadecanoyl-CoA intermediate was also demonstrated in rat brain homogenates and in lysates from rat C6 glial cells.



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  		FIG. 2.
Identification of 2-hydroyxoctadecanoyl-CoA (A–C) and heptadecanal (D–F) in incubations of rat liver homogenate with 2-hydroxyoctadecanoic acid and 2-hydroxyoctadecanoyl-CoA, respectively. HPLC analysis of acyl etheno-CoA esters (left panel) and aldehydes (right panel) formed during incubations of rat liver homogenate with 2-hydroxyoctadecanoic acid (100 µM) and 2-hydroxyoctadecanoyl-CoA (40 µM), respectively. Homogenates were incubated for 0 min (A and D) and 10 min (B and E), and reaction products were derivatized after the addition of the appropriate internal standard (IS) as described under "Experimental Procedures." Elution of standards is shown in C and F.

 
In the presence of Mg2+ and TPP, heptadecanal was formed by rat liver homogenates (Fig. 2, D–F) incubated with 2-hydroxyoctadecanoyl-CoA at rates of 0.014 nmol/mg of protein/min (n = 2). Overall, these data indicate that 2-hydroxyacyl-CoA esters are intermediates in the degradation of 2-hydroxy straight chain fatty acids. Unfortunately, due to the presence of interfering peaks, the formation of the n – 1 aldehyde could not be demonstrated in incubations with brain homogenate.

Characterization of the acyl-CoA Synthetase, Catalyzing the Formation of 2-Hydroxyacyl-CoA Esters—In rat liver homogenates the activation of 2-hydroxyoctadecanoic acid was shown to be linear for up to 3 min (Fig. 3A). The formation of 2-hydroxyoctadecanoyl-CoA reached an optimum at a substrate/albumin ratio ({nu}) of 4 (Fig. 3B). Measuring the generation of 2-hydroxyoctadecanoyl-CoA at increasing substrate concentrations (at {nu} = 4) resulted in a plateau from 100 µM onward; transformation of the data according to Lineweaver-Burk allowed the calculation of an apparent Km of 19.5 µM (Fig. 3, C–D). Subsequent experiments in rat liver homogenates and subcellular fractions were, therefore, performed at a substrate concentration of 50 µM and a {nu} ratio of 4 and were terminated after 3 min. For measurement of synthetase activity in lysates of rat C6 glial cells, optimum conditions were 100 µM substrate, a {nu} ratio of 2, and an incubation time of 10 min.



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  		FIG. 3.
Kinetics of the activation of 2-hydroxyoctadecanoic acid in rat liver homogenates. Generation of labeled acyl-CoA was measured in incubations of 2-hydroxy[1-14C]octadecanoic acid (50 µM in A and B; varying substrate concentrations are in C and D) with rat liver homogenates. Reactions were stopped after 3 min except in panel A. The velocity (V, expressed in nmol/min/g) versus substrate concentration (S, expressed in µM) curve, as shown in panel C, was linearly transformed according to the method of Lineweaver-Burk as shown in D.

 
The activation of 2-hydroxyoctadecanoic acid in subcellular fractions of rat liver showed a bimodal distribution coinciding with the light mitochondrial fraction, enriched in peroxisomes, and the microsomal fraction (Fig. 4, Fractionation A). In the light mitochondrial fraction the activity appeared to be predominantly membrane-associated. Further separation of the light mitochondrial fraction on a Nycodenz gradient revealed an association with peroxisomes (data not shown). In rat brain a bimodal distribution was observed as well, the microsomal localization being most prominent (results not shown).



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  		FIG. 4.
Subcellular distribution of acyl-CoA synthetase activity (Fractionation A) and the production of heptadecanal from 2-hydroxyoctadecanoyl-CoA (Fractionation B) in rat liver. In two separate experiments fresh liver homogenates were fractionated by differential centrifugation into nuclear (N), heavy mitochondrial (M), light mitochondrial (L), microsomal (P), and cytosolic (S) fractions. Fractionation A, fractions were incubated with 50 µM 2-hydroxy[1-14C]octadecanoic acid, and the production of the CoA ester (synthetase) was measured as described under "Experimental Procedures." Fractionation B, the production of heptadecanal from 2-hydroxyoctadecanoyl-CoA (lyase) was measured in each fraction as described under "Experimental Procedures." Marker enzymes were determined in each fraction: carboxylesterase (endoplasmic reticulum), glutamate dehydrogenase (GDH; mitochondria), catalase (peroxisomal matrix), and lactate dehydrogenase (LDH; cytosol). Results are expressed as relative specific activities versus percentage of total protein. Relative specific activity is defined as the percentage of total recovered activity present in a particular fraction divided by the corresponding percentage of protein. Recovery for synthetase activity was 89%; recovery for heptadecanal production was 109%; recoveries for marker enzymes were between 86 and 119%.

 
To further characterize the acyl-CoA synthetase responsible for the activation of 2-hydroxy straight chain fatty acids, incubations of lysates from C6 glial cells with 2-hydroxy[1-14C]octadecanoic acid were performed with and without the addition of a range of unlabeled fatty acids. Whereas 3-methylhexadecanoic acid, 2-methylhexadecanoic acid, eicosatetraenoic, and tetracosanoic acid had no major inhibitory effect on the activation of 2-hydroxyoctadecanoic acid, hexadecanoic acid and octadecanoic acid markedly reduced the activation rates (results not shown), indicating that the substrate spectrum of the activating enzyme involved certainly covers long chain fatty acids.

Identification of 2-HPCL as the Enzyme Catalyzing the Cleavage Reaction—Investigation of the formation of heptadecanal from 2-hydroxyoctadecanoyl-CoA in subcellular fractions of rat liver indicated that the responsible enzyme has a peroxisomal localization (Fig. 4, Fractionation B). Both this subcellular distribution and the marked thiamine dependence point toward 2-HPCL as the enzyme catalyzing the cleavage reaction.

In the presence of ATP, CoA, Mg2+, and TPP, incubations of the purified recombinant polyhistidine-fused human 2-HPCL with 2-hydroxyoctadecanoyl-CoA yielded heptadecanal, in analogy with the formation of 2-methylpentadecanal from 2-hydroxy-3-methylhexadecanoyl-CoA. The reaction kinetics of recombinant 2-HPCL toward these substrates were compared by measuring the generation of 2-methylpentadecanal and heptadecanal. The cleavage rate of the branched substrate was linear for up to 10 min, whereas for the straight chain substrate it was linear for up to at least 30 min (results not shown). At increasing substrate concentrations a plateau from 40 µM onward was reached for 2-hydroxy-3-methylhexadecanoyl-CoA, whereas for 2-hydroxyoctadecanoyl-CoA a plateau was reached from 20 µM onward (Fig. 5, A and B); apparent Km values of 15.8 and 6.3 µM, respectively, could be calculated. Furthermore, the amount of aldehyde generated with both substrates increased linearly with the amount of recombinant protein up to at least 10 µg.



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  		FIG. 5.
Characterization of the cleavage reaction of 2-hydroxy-3-methylhexadecanoyl-CoA and 2-hydroxyoctadecanoyl-CoA by recombinant human 2-HPCL. Aldehyde production was measured in incubations of 2-hydroxy-3-methylhexadecanoyl-CoA (A) or 2-hydroxyoctadecanoyl-CoA (B) (varying substrate concentrations; {nu} = 6) with recombinant human 2-HPCL (2 µg of protein). Reactions were stopped after 10 min. The velocity (V, expressed in nmol/assay) versus substrate concentration (S, expressed in µM) curves were linearly transformed according to the method of Lineweaver-Burk.

 
The substrate spectrum of recombinant, polyhistidine-fused human 2-HPCL was further investigated using labeled substrates. Incubation of the recombinant 2-HPCL with 2-hydroxy-3-methyl[1-14C]hexadecanoyl-CoA under standard conditions resulted in production of [14C]formyl-CoA/[14C]formate at rates of 26.06 ± 1.17 milliunits/mg of protein (mean ± S.E.; n = 4), but the lyase showed no activity toward [1-14C]-labeled 3-methylhexadecanoyl-CoA, hexadecanoyl-CoA, 3-hydroxyhexadecanoyl-CoA, octadecanoyl-CoA, or toward 2-hydroxy-3-methylhexadecanoic acid, 3-methylhexadecanoic acid, hexadecanoic acid, and 2-hydroxyhexadecanoic acid. Furthermore, the production of [14C]formyl-CoA/[14C]formate from 2-hydroxy-3-methyl[1-14C]hexadecanoyl-CoA was not reduced in the presence of 2-keto-octanoate, 2-hydroxyhexadecanoic acid, 2-methylhexadecanoyl-CoA, 2-methylhexadecanoic acid, 3-methylhexadecanoyl-CoA, 3-hydroxy-3-methylhexadecanoyl-CoA, or 3-hydroxy-2-methylhexadecanoyl-CoA (Fig. 6). The addition of 50 µM 2-hydroxyhexadecanoyl-CoA or 2-hydroxyoctadecanoyl-CoA to incubations with 10 µM 2-hydroxy-3-methyl[1-14C]hexadecanoyl-CoA, however, reduced the cleavage rates to a major extent (Fig. 6), indicating that 2-hydroxy straight chain acyl-CoA esters compete with 2-hydroxy-3-methyl-branched acyl-CoAs for cleavage by 2-HPCL.



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  		FIG. 6.
Effect of competitors on the production of 14C-formate/14C-formyl-CoA from 2-hydroxy-3-methyl[1-14C]hexadecanoyl-CoA in incubations with recombinant human 2-HPCL. The production of [14C]formate/[14C]formyl-CoA from 2-hydroxy-3-methyl[1-14C]hexadecanoyl-CoA (10 µM) with 4.7 µg of recombinant human 2-HPCL was measured with and without the addition of 50 µM unlabeled substrates. Panel A, 2-hydroxyhexadecanoyl-CoA, 2-hydroxyhexadecanoic acid, 2-methylhexadecanoyl-CoA, 2-methylhexadecanoic acid, or 2-keto-octanoate measured in the presence of 1.67 µM BSA. Panel B, 2-hydroxy-3-methylhexadecanoyl-CoA, 3-methylhexadecanoyl-CoA, 2-hydroxyhexadecanoyl-CoA, 2-hydroxyoctadecanoyl-CoA, 3-hydroxy-3-methylhexadecanoyl-CoA, or 3-hydroxy-2-methylhexadecanoyl-CoA measured in the presence of 10 µM BSA. Results, calculated from single determinations, are represented as the percentage of control activity.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
In the current study 2-hydroxyoctadecanoic acid was used as a substrate to document the degradation of 2-hydroxy straight chain fatty acids. This compound is less abundant than 2-hydroxytetracosanoic acid (cerebronic acid) in brain cerebrosides and sulfatides (3), but it offers advantages over the very long chain substrates both in synthesis and solubility. The fact that 2-hydroxy[1-14C]octadecanoic acid gives rise to the generation of labeled formate in intact as well as in broken cell systems was a first indication that 2-hydroxyfatty acids can be degraded via a process resembling the {alpha}-oxidation of 3-methylbranched fatty acids. This was further supported by the requirement of ATP, Mg2+, CoA, and TPP in permeabilized cells and by the observation that 2-hydroxyoctadecanoyl-CoA serves as a substrate for recombinant 2-HPCL, thereby being converted to heptadecanal. The in vivo significance of the 2-HPCL-dependent pathway was highlighted by the marked inhibitory effect of the thiamine antimetabolite oxythiamin in cultured cells.

Overall, our data led to the following picture (Fig. 7). A 2-hydroxy fatty acid is first activated to its CoA ester in an ATP/Mg2+/CoA-dependent reaction. The generated 2-hydroxyacyl-CoA is then cleaved by 2-HPCL into an n – 1 aldehyde and formyl-CoA. The latter is converted to formate and subsequently to CO2. As for the 2-methyl n – 1 aldehydes formed during {alpha}-oxidation of 3-methyl-branched fatty acids, the n – 1 aldehyde is most probably dehydrogenated to the corresponding odd-numbered fatty acid, which can then be further degraded via {beta}-oxidation. Although we cannot exclude that 2-hydroxyfatty acids can be degraded by other pathways, the inhibition by oxythiamin suggests that the main pathway, at least in the cells and tissues we studied, is via the TPP-dependent 2-HPCL. In addition, we did not find evidence for the formation of a 2-keto fatty acid, formed as an intermediate via an {alpha}-OH acid oxidase (36). In kidney, but not in liver, an L-2-hydroxy long chain acid oxidase has been described (37).



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  		FIG. 7.
{alpha}-Oxidation of 2-hydroxyfatty acids and of 3-methyl-branched fatty acids. The scheme represents the proposed {alpha}-oxidation pathway for 2-hydroxy straight chain fatty acids next to the {alpha}-oxidation pathway for 3-methyl-branched fatty acids. *FA2H, fatty acid 2-hydroxylase (18).

 
2-Hydroxyacyl-CoAs as substrate for {alpha}-oxidation have not been described in plants or bacteria, but in plants n – 1 aldehydes as a product of {alpha}-oxidation have been reported (6). The relevance of 2-HPCL to the {alpha}-oxidation of 2-hydroxy straight chain fatty acids in these organisms is unknown at present.

With regard to the stereochemistry of the process we studied, not all details are yet available. The naturally occurring 2-hydroxyfatty acids possess a D-configuration, but so far racemic 2-hydroxyoctadecanoic acid has been used. The finding that in C6 glial cells within 10 h >70% of the substrate was oxidized suggests that racemization can occur or that both isomers are degraded. Because 2-HPCL is able to cleave all four possible isomers of 3-methyl-2-hydroxyhexadecanoyl-CoA (38), the latter possibility is more likely.

Measurement of the activation of 2-hydroxyoctadecanoic acid in subcellular fractions from rat liver revealed that the activity is associated both with the peroxisomal membrane and with the endoplasmic reticulum. This distribution together with the fact that, among the fatty acids tested, only hexadecanoic acid and octadecanoic acid competitively inhibited the activation of 2-hydroxyoctadecanoic acid, point toward the long chain acyl-CoA synthetase as the activating enzyme. However, because it is well known that the substrate spectra of long chain and very long chain acyl-CoA synthetases overlap (39, 40), no definitive conclusions can be drawn as to the specific enzyme involved. Moreover, it might well be that the chain length of the substrate determines the involved isoform. Hence, long chain 2-hydroxyfatty acids (e.g. 2-hydroxyoctadecanoic acid) might be activated by a long chain acyl-CoA synthetase and very long chain 2-hydroxyfatty acids might be activated by a very long chain acyl-CoA synthetase.

The contention of 2-HPCL being involved in the degradation of 2-hydroxyfatty acids was strengthened by the increased relative specific activities of heptadecanal formation from 2-hydroxyoctadecanoyl-CoA in subcellular fractions enriched in peroxisomes, the organelles that harbor 2-HPCL (14). The marked inhibition of 2-hydroxyoctadecanoic acid degradation by the addition of oxythiamin to cultured cells further corroborates the thiamine dependence of the pathway in the intact cell. The finding that in rat liver homogenates and permeabilized hepatocyte stimulation by the addition of TPP was only limited can be explained by the fact that in these preparations TPP is still partly bound to the enzyme. As shown earlier, 2-HPCL only gradually loses its activity during purification due to the release of bound TPP (14).

The fact that 2-hydroxyfatty acid degradation is decreased by only {approx}50% in fibroblasts from Zellweger patients and Pex5–/– mice, an animal model for Zellweger syndrome (23), might suggest that 2-HPCL, a peroxisomal matrix enzyme (14), remains partially active in the cytosol under conditions where peroxisomal protein import is deficient. This is in contrast with most other peroxisomal matrix enzymes, which are labile in the cytosol when their import is impaired, but in agreement with our previous measurements of 2-HPCL activity in liver homogenates of Zellweger patients and Pex5–/– mice using 2-hydroxy-3-methylhexadecanoyl-CoA as substrate (41). Whether the partial impairment of 2-hydroxyfatty acid degradation leads to an accumulation of 2-hydroxyfatty acids in tissues from Zellweger patients is unknown. Moderate increases in cerebrosides and/or gangliosides have been described in fibroblasts or CHO cells lacking peroxisomes, but their 2-hydroxyfatty acid content was not documented (42, 43). When considering the identification of this second substrate for 2-hydroxyphytanoyl-CoA lyase, one could expect that an isolated deficiency of 2-HPCL, although hitherto not identified, might lead not only to an impaired {alpha}-oxidation of phytanic acid but also to an accumulation of 2-hydroxyfatty acids.

The identification of 2-HPCL as the cleavage enzyme in the degradation of 2-hydroxyfatty acids is also of interest for its reaction mechanism. It demonstrates that the methyl branch at position 3 is not necessary but that both the hydroxy group at position 2 and the CoA moiety are important. A 2-hydroxy carboxyl compound (instead of a 2-keto compound) is an unusual substrate for TPP-dependent enzymes. Only one other enzyme, N2-(2-carboxyethyl)-L-arginine synthase, catalyzing the condensation of L-arginine and D-glyceraldehyde in the presence of TPP (the first step in the biosynthesis of clavulanic acid), has been reported to show activity toward compounds with a hydroxy group at position 2 (44). In all TPP-dependent cleavage reactions described so far decarboxylation involves the activation of the C2-H of the thiazole ring of TPP to an intermediate carbanion. This is followed by a nucleophilic attack at the carbonyl atom of the substrate (carbon 2) (45). Most likely, the formation of a carbanion is also required for the cleavage of 2-hydroxy-(3-methyl)acyl-CoA esters by 2-HPCL. However, this carbanion will then attack carbon 1 of the substrate, which is highly reactive due to the presence of the thioester bond. Ultimately, this will lead to the formation of formyl-CoA and an n – 1 fatty aldehyde (Fig. 8).



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  		FIG. 8.
Generation of a carbanion in enzyme-bound TPP and the proposed reaction mechanism for 2-HPCL. Reaction of 2-HPCL with the substrate 2-hydroxyoctadecanoyl-CoA involves a nucleophilic attack of the C2-atom of TPP at the carbon 1 of the substrate. A key function herein is the interaction of a conserved glutamate with the N1' atom of the coenzyme, resulting in an increased basicity of its 4' amino group, which facilitates the deprotonation of the C2 (Ref. 45 and references cited herein).

 
The observation that extra addition of NAD+ to the incubation mixture resulted in a further 20% increase of total oxidation (Table III) might be explained by a diminished product inhibition through NAD+-dependent dehydrogenation of the generated aldehyde. Fatty aldehyde dehydrogenase activity is found in peroxisomes and endoplasmic reticula (13, 46). It is currently unknown whether fatty aldehydes generated in the peroxisome are dehydrogenated exclusively by the peroxisomal dehydrogenase or also partially by the endoplasmic reticulum enzyme.

In summary, our present work shows that (the bulk of) 2-hydroxyfatty acids undergo an initial degradation that apparently shares three reactions (activation, cleavage of the C1-C2 bond, aldehyde dehydrogenation) with the {alpha}-oxidation sequence of 3-methyl-branched fatty acids, giving rise to n – 1 odd-numbered fatty acids, which can subsequently be degraded via {beta}-oxidation. It is of interest to note that the second enzyme of the {alpha}-oxidation sequence of 3-methyl-branched fatty acids (the peroxisomal PAHX, which hydroxylates 3-methyl-acyl-CoAs to 2-hydroxy-3-methyl-acyl-CoAs) is likely not involved in the synthesis of 2-hydroxy straight chain fatty acids (see the Introduction). One of the underlying reasons might be that an (imaginary) hydroxylation of straight chain acyl-CoAs by PAHX would lead to the immediate further breakdown of the 2-hydroxylated acyl-CoAs by 2-HPCL within the peroxisome.

To control the levels of 2-hydroxyfatty acids in brain cerebrosides and sulfatides, supposed to play a role in myelinization, a strategy relying on different sets of enzymes for their synthesis and degradation, located at different subcellular sites, might be more beneficial. Moreover, as was reported in older literature and recently discussed by Alderson et al. (18), hydroxylation of straight chain fatty acids might occur only after incorporation in sphingolipids, which would further rule out peroxisomes as a key player in this hydroxylation process. In this context it is of interest to note that 2-hydroxyoctadecanoic acid was hardly incorporated into complex lipids when given to intact cells.3 Overall, the {alpha}-oxidation of straight chain fatty acids, as has been described especially for brain, appears to proceed as follows; 1) hydroxylation of the fatty acid by a fatty acid 2-hydroxylase (see the Introduction), 2) activation of the 2-hydroxyfatty acid to a 2-hydroxyacyl-CoA, 3) cleavage of the CoA ester into formyl-CoA and an n – 1 fatty aldehyde, and 4) dehydrogenation of the aldehyde to the corresponding n – 1 odd-numbered fatty acid.


   FOOTNOTES
 
* This work was supported by Geconcerteerde Onderzoeksacties van de Vlaamse Gemeenschap Grants GOA 99/03-09 and GOA 2004/08, Fonds voor Wetenschappelijk Onderzoek-Vlaanderen Grant G.0115.02, and European Union project RDDPT, Grant QLG3-CT-2002-00696. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

{ddagger} Both authors contributed equally to this work. Back

§ Supported by a fellowship from the Fonds voor Wetenschappelijk Onderzoek-Vlaanderen. Back

To whom correspondence may be addressed: Departement Celbiologie, Afdeling Farmacologie, Faculteit Geneeskunde, Katholieke Universiteit Leuven, Herestraat 49, Box 601, 3000 Leuven, Belgium. Tel.: 32-16-345802; Fax: 32-16-345699; E-mail: paul.vanveldhoven{at}med.kuleuven.ac.be. || To whom correspondence may be addressed: Departement Celbiologie, Afdeling Farmacologie, Faculteit Geneeskunde, Katholieke Universiteit Leuven, Herestraat 49, Box 601, 3000 Leuven, Belgium. Tel.: 32-16-345816; Fax: 32-16-345699; E-mail: minne.casteels{at}med.kuleuven.ac.be.

1 The abbreviations used are: 2-OH-FA, 2-hydroxyfatty acids; 2-HPCL, 2-hydroxyphytanoyl-CoA lyase; {nu}, substrate/BSA ratio; PAHX, phytanoyl-CoA hydroxylase; TPP, thiamine pyrophosphate; Mops, 4-morpholinepropanesulfonic acid; BSA, bovine serum abumin; HPLC, high performance liquid chromatography. Back

2 V. Foulon, M. Sniekers, M. Casteels, and P. P. Van Veldhoven, manuscript in preparation. Back

3 M. Sniekers, V. Foulon, P. P. Van Veldhoven, and M. Casteels, unpublished data. Back


   ACKNOWLEDGMENTS
 
We thank Dr. Myriam Baes for providing Pex5–/– mouse fibroblasts, Ruud Dirkx for providing mouse hepatocytes, and Wendy Geens and Luc Govaert for expert technical assistance.



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 DISCUSSION
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