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Originally published In Press as doi:10.1074/jbc.M501861200 on May 2, 2005

J. Biol. Chem., Vol. 280, Issue 29, 26838-26844, July 22, 2005
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Characterization of the Final Step in the Conversion of Phytol into Phytanic Acid*

Daan M. van den Brink{ddagger}§, Joram N. I. van Miert{ddagger}§, Georges Dacremont¶, Jean-François Rontani||, and Ronald J. A. Wanders{ddagger}§**

From the Departments of {ddagger}Clinical Chemistry and §Pediatrics, University of Amsterdam, Academic Medical Center, Emma Children's Hospital, 1105 AZ Amsterdam, The Netherlands, University of Ghent, 9000 Ghent, Belgium, and ||Laboratoire de Microbiologie, Géochimie et Ecologie Marines (UMR 6117), Centre d'Océanologie de Marseille, Campus de Luminy, 13288 Marseille, France

Received for publication, February 18, 2005 , and in revised form, April 22, 2005.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Phytol is a branched-chain fatty alcohol that is a naturally occurring precursor of phytanic acid, a fatty acid involved in the pathogenesis of Refsum disease. The conversion of phytol into phytanic acid is generally believed to take place via three enzymatic steps that involve 1) oxidation to its aldehyde, 2) further oxidation to phytenic acid, and 3) reduction of the double bond at the 2,3 position, yielding phytanic acid. Our recent investigations of this mechanism have elucidated the enzymatic steps leading to phytenic acid production, but the final step of the pathway has not been investigated so far. In this study, we describe the characterization of phytenic acid reduction in rat liver. NADPH-dependent conversion of phytenic acid into phytanic acid was detected, although at a slow rate. However, it was shown that phytenic acid can be activated to its CoA ester and that reduction of phytenoyl-CoA is much more efficient than that of phytenic acid. Furthermore, in rat hepatocytes cultured in the presence of phytol, phytenoyl-CoA could be detected, showing that it is a bona fide intermediate of phytol degradation. Subcellular fractionation experiments revealed that phytenoyl-CoA reductase activity is present in peroxisomes and mitochondria. With these findings, we have accomplished the full elucidation of the mechanism by which phytol is converted into phytanic acid.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The branched-chain fatty alcohol phytol (3,7,11,15-tetramethylhexadec-2-en-1-ol) is abundantly present in nature as part of the chlorophyll molecule (Fig. 1). After its release from chlorophyll, phytol is converted into phytanic acid, a fatty acid that accumulates in a number of metabolic disorders. Phytanic acid has toxic effects when present at high levels in the body and therefore needs to be broken down (1–3). Because of the presence of a methyl group at the 3 position, phytanic acid cannot be degraded by {beta}-oxidation but instead undergoes {alpha}-oxidation. In this process the carbon chain is shortened by one carbon unit to produce the 2-methyl branched-chain fatty acid, pristanic acid, which can be {beta}-oxidized normally (4, 5). A deficiency of {alpha}-oxidation, such as in Refsum disease, leads to markedly elevated levels of phytanic acid in plasma and tissues of patients and is thought to be the direct cause of symptoms observed in the patients, which include retinitis pigmentosa, peripheral neuropathy, and cerebellar ataxia (6, 7).

Remarkably, very few studies have been performed on the contribution of phytol to the total intake of phytanic acid. This is most likely due to the finding that in humans only a small percentage of phytol is released from chlorophyll; this process only occurs effectively in the digestive system of ruminant animals (8). As a result, free phytol is present in some dairy products (9).

Resolution of the metabolic pathway by which phytol is converted into phytanic acid is also important because phytol is often used in in vitro as well as in in vivo studies as a precursor of phytanic acid. Results from such studies have already led to a proposed model for phytol degradation (outlined in Fig. 1). This model is based on the finding that phytol administration to rats leads to increased levels of phytenic acid. According to this model, phytol is first converted to phytenic acid via the action of an alcohol dehydrogenase and an aldehyde dehydrogenase, after which phytenic acid undergoes reduction to phytanic acid.

Support for this model came from measurements in isolated rat livers (10) and, more recently, from our own studies in cultured human fibroblasts that provided evidence for the involvement of the microsomal fatty aldehyde dehydrogenase catalyzing the conversion of E-phytenal into E-phytenic acid (11).

In this work, we have studied the last step of the phytol degradation pathway, in which phytenic acid is converted into phytanic acid. Interestingly, our studies show that in contrast to the proposed pathway, phytenic acid is first activated to its CoA ester before the double bond is reduced. In addition, we show that NADPH-dependent reductase activity is located in mitochondria and peroxisomes.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Differential and Density Gradient Centrifugation—All animal experiments were approved by the Institutional Animal Ethical Committee of the University of Amsterdam. For subcellular fractionation studies, male Wistar rats were fed ad libitum with a standard laboratory diet supplemented with 1% (w/w) diethylhexylphthalate for 9 days, after which livers were isolated, washed in ice-cold phosphate-buffered saline, and homogenized on ice in a buffer containing 250 mM sucrose, 2.5 mM EDTA, and 5 mM morpholinopropanesulfonic acid (final pH 7.4). A post-nuclear supernatant was obtained by centrifugation at 600 x g for 10 min at 4 °C, which was subsequently subjected to differential centrifugation essentially as described previously (12). Glutamate dehydrogenase (mitochondria), esterase (microsomes), catalase (peroxisomes), and phospho-glucose isomerase (cytosol) were used as marker enzymes for the different subcellular compartments indicated in parentheses and measured as described elsewhere (12, 13). Amounts of protein were determined using the bicinchoninic acid method according to the manufacturer's instructions (Sigma). Subsequently, the light mitochondrial fraction was subjected to equilibrium density gradient centrifugation in a linear Nycodenz gradient as described previously (13).



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  		FIG. 1.
The conversion of phytol into phytanic acid. According to the model in the literature, phytol is converted into phytanic acid via the action of three different enzyme reactions, including 1) an alcohol dehydrogenase, 2) an aldehyde dehydrogenase, and 3) a reductase reaction. We have previously shown that the aldehyde dehydrogenase involved in the conversion of E-phytenal to E-phytenic acid is the microsomal enzyme fatty aldehyde dehydrogenase (FALDH).

 
Phytenic Acid Reductase Assay—A mixture of Z- and E-phytenic acids was synthesized as described previously (14). Rat livers were derived from male Wistar rats fed ad libitum with a diet supplemented with 1% (w/w) diethylhexylphthalate. After sacrifice, livers were isolated, washed with ice-cold phosphate-buffered saline, and stored at –80 °C prior to use. Homogenates were made by cutting off a piece of the liver, which was homogenized on ice in phosphate-buffered saline and subsequently sonicated on ice (3 cycles of 10 s at 70 W).

Reduction of phytenic acid to phytanic acid was determined in an assay mixture consisting of 50 mM Hepes (pH 7.7), 1 mM NADPH, 1 mg/ml methyl-{beta}-cyclodextrin, 100 µM phytenic acid (dissolved in dimethyl sulfoxide, 1% final concentration in the assay), and 0.2 mg/ml rat liver homogenate in a total volume of 500 µl. The reactions were started by addition of homogenate and allowed to proceed for 30 min at 37 °C, after which they were terminated by addition of 2 M HCl. For analysis of the reaction products by gas chromatography-mass spectrometry, first 49.3 pmol of [2H3]phytanic acid dissolved in toluene (purchased from Dr. H. J. ten Brink, Free University Hospital, Amsterdam, The Netherlands) was added as internal standard. Then 2 ml of hexane was added, after which the organic layer was evaporated to dryness under nitrogen at 40 °C. The sample was then derivatized with N-tert-butyldimethylsilyl-N-methyl-trifluoroacetamide (Aldrich) and pyridine (50 µl each) at 80 °C for 30 min, dried under nitrogen, and taken up in hexane. Analysis of the sample was performed on an Agilent Technologies model 5890/5973 gas chromatography-mass spectrometry system equipped with a CPsil 19CB capillary column (internal diameter, 25 m x 0.25 mm; film thickness, 0.2 µm; Varian) as described previously (15, 16). Mass spectrometry acquisition was performed in the single ion monitoring mode, specifically the [M-57]+ ions of the various compounds (corresponding to 369.3 and 372.3 for phytanic acid and the internal standard, respectively). Calibration curves were generated in a concentration range of 0–100 µM, and concentrations of phytanic acid were calculated using a linear fit.



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  		FIG. 2.
Characterization of the reduction of phytenic acid. A, production of phytanic acid upon incubation of rat liver homogenate in the presence of NADPH, NADP+, or NADH (all at 1 mM). B, incubations with various concentrations of NADPH. From a Michaelis-Menten plot (inset), a Km of 25 µM was calculated. C, determination of the pH optimum using a composite buffer consisting of citrate, MES, MOPS, and Bicine (35 mM each). D, dependence on the concentration of phytenic acid. Values represent the mean ± S.D. of duplicates expressed in nmol/h/mg protein.

 
Phytenoyl-CoA Reductase Assay—Reduction of phytenoyl-CoA to phytanoyl-CoA was determined in an assay mixture containing 50 mM Hepes (pH 7.0), 1 mM NADPH, 1 mg/ml bovine serum albumin, 100 µM phytenoyl-CoA, and 0.025 mg/ml rat liver homogenate in a final volume of 100 µl. Reactions were started by addition of substrate and incubated for 30 min at 37 °C, after which they were terminated by the addition of 100 µl of acetonitrile and placed on ice. Samples were spun for 10 min at 10,000 x g at 4 °C, supernatants were analyzed by high performance liquid chromatography (HPLC),1 and CoA esters were quantified using UV detection. To this end, 100-µl samples were injected on a C18 reverse-phase column (Supelcosil LC-18-D8; internal diameter, 25 cm x 4.6 mm; particle size, 5 µM; Supelco) and eluted with a linear gradient from 16.9 mM sodium phosphate (pH 6.9) dissolved in 46–64% acetonitrile over 20 min, at a flow rate of 1 ml/min. Absorbance of the eluate was monitored on a Shimadzu UV-visible detector (SPD-10Avp) at a wavelength of 260 nm. To determine HPLC retention times, standards of phytanoyl-CoA and phytenoyl-CoA were produced from the corresponding free acids using the acid anhydride method (17).

Acyl-CoA Synthetase Assay—Enzymatic activation of phytenic acid to its acyl-CoA ester was investigated essentially as described for phytanic acid (18). Briefly, 0.2 mg/ml rat liver homogenate was incubated in a mixture containing 50 mM Tris (pH 8.0), 10 mM MgCl2, 0.6 mM CoA, 10 mM ATP, 100 µM phytenic acid, and 1 mg/ml methyl-{beta}-cyclodextrin in a total volume of 100 µl for 30 min at 37 °C, after which the reaction was terminated, and amounts of phytenoyl-CoA were determined as described for the phytenoyl-CoA reductase assay.

Detection of Phytenoyl-CoA in Cultured Rat Hepatocytes—Cultured rat hepatocytes (CRL-1601 cells) were grown in Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum (Invitrogen), penicillin (100 units/ml), and streptomycin (100 µg/ml) (all from Invitrogen) at 37 °C with 5% CO2, in the presence or absence of 50 µM phytol (mixture of Z- and E-isomers; Merck). After 4 days, the cells were harvested using trypsin and lysed on ice by the addition of 1 mg/ml digitonin. Analysis of CoA esters was performed using tandem mass spectrometry (tMS) as described elsewhere.2 Briefly, lysates were subjected to Dole's extraction (19), whereby the aqueous phase containing the CoA esters was dried under a flow of N2 and taken up in water. Samples were injected on a HPLC tMS system (Finnigan Surveyor TSQuantum AM; Thermofinnigan) and analyzed for HPLC retention time and mass determinations. Detection by tMS was performed by selecting for double-charged parent ions (508.5, 528.5, and 529.5 for heptadecanoyl-CoA, phytenoyl-CoA, and phytanoyl-CoA, respectively).



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  		FIG. 3.
Phytenic acid reductase activity in fractions obtained after differential centrifugation of a rat liver. Differential centrifugation of a rat liver homogenate was performed as described under "Materials and Methods," by which a post-nuclear supernatant (PNS), a mitochondrial fraction (M), a light mitochondrial fraction (L), a microsomal fraction (P), and a cytosolic fraction (S) were obtained. In the different fractions, the following activities were measured: A, phytenic acid reduction; B, phytenoyl-CoA reduction; C, glutamate dehydrogenase (mitochondrial marker); D, esterase (microsomal marker); E, catalase (peroxisomal marker); and F, {beta}-hexosaminidase (lysosomal marker). All activities are expressed as a percentage relative to that of the post-nuclear supernatant fraction.

 



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  		FIG. 4.
Phytenoyl-CoA synthetase and reductase activity. Detection of phytenoyl-CoA in incubations of rat liver homogenates in the presence of phytenic acid, CoA, Mg2+, and ATP by HPLC. A, standards for phytanoyl-CoA (dashed line) and Z- and E-phytenoyl-CoA (solid line). Phytenoyl-CoA isomers are separated into two distinct peaks, with the Z-isomer eluting before the E-isomer. Chromatograms were acquired by measuring absorbance at 260 nm as described under "Materials and Methods." B, depicted are typical chromatograms of experiments in which a rat liver homogenate was incubated for t = 0 or 30 min with either phytanic acid (dashed line) or phytenic acid (solid line). Presumably, only E-phytenoyl-CoA isomer is formed. C, reductase activity using either phytenic acid or phytenoyl-CoA as substrate. Rat liver homogenates were incubated as described before with either phytenic acid or phytenoyl-CoA. After termination of the reactions, samples were made alkaline (pH > 11) by addition of KOH and incubated for 60 min at 60 °C to hydrolyze all CoA esters. Free acids were then derivatized with N-tert-butyldimethylsilyl-N-methyl-trifluoroacetamide and detected by gas chromatography-mass spectrometry as described under "Materials and Methods."

 

   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Characterization of the Enzymatic Reduction of Phytenic Acid—To study the conversion of phytenic acid into phytanic acid, rat liver homogenates were incubated with phytenic acid in the presence of a variety of cofactors. Production of phytanic acid was observed only with NADPH present (Fig. 2A). The Km for NADPH was calculated to be ~25 µM (Fig. 2B). As shown in Fig. 2C, a pH optimum was found at pH 7.7. Titration of the substrate phytenic acid resulted in a sharp decrease in activity at concentrations higher than 150 µM (Fig. 2D), which prevented calculation of the Km and Vmax. Based on these results, we selected a substrate concentration of 100 µM phytenic acid for all subsequent assays. The standard reaction medium also contained 1 mg/ml methyl-{beta}-cyclodextrin, which increased activity 3-fold. Under these incubation conditions, formation of phytanic acid was linear with protein up to 0.2 mg/ml and time up to at least 30 min (data not shown).

Subcellular Localization of the Reduction of Phytenic Acid— Next, we studied in which subcellular compartment reduction of phytenic acid to phytanic acid takes place. To this end, a rat liver homogenate was subjected to differential centrifugation to obtain fractions enriched in different cell organelles. The activity profile of the reduction of phytenic acid approximately mimicked that of glutamate dehydrogenase, a mitochondrial marker, whereas reductase activity was also present in the microsome-enriched fraction (Fig. 3).

To establish the subcellular localization more precisely, density gradient centrifugation was performed, using a rat liver light mitochondrial fraction as starting material. However, substantial phytanic acid production could not be measured in any of the fractions (data not shown). Because we had observed earlier that the enzyme activity measured after differential centrifugation was substantially lower than that in non-fractionated rat liver homogenates, it was speculated that the centrifugation procedures might be detrimental to enzyme activity.

To address this question, we investigated the stability of the enzymatic activity. Studies in a rat liver homogenate revealed that there was minimal deterioration of reductase activity over time. Even after storage for 7 days at room temperature, more than 50% of the initial activity remained (data not shown). The addition of a reducing agent such as dithiothreitol or protease inhibitors did not prevent deterioration of activity.



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  		FIG. 5.
Detection of phytanoyl-CoA and phytenoyl-CoA in rat hepatocytes cultured in the presence of phytol. Rat hepatocytes were cultured for 4 days in the presence of 50 µM phytol, after which CoA esters were extracted by Doles extraction and analyzed by HPLC tMS. A, standards for heptadecanoyl-CoA (used as internal standard), phytanoyl-CoA, and phytenoyl-CoA (mixture of Z- and E-isomers). B, a typical sample from hepatocytes cultured in control medium. No peaks corresponding to either phytanoyl-CoA or phytenoyl-CoA were present, although an additional peak with the same m/z ratio as phytanoyl-CoA was observed eluting at 15.5 min, and an additional peak with the same m/z ratio as phytenoyl-CoA was observed eluting at 17 min. The identity of these peaks is unknown. C, a sample from hepatocytes cultured on phytol-supplemented medium. Peaks corresponding to phytanoyl-CoA and E-phytenoyl-CoA were detected.

 



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  		FIG. 6.
Characterization of the reduction of phytenoyl-CoA. A, incubations of rat liver homogenate with phytenoyl-CoA and various concentrations of NADPH. From a Michaelis-Menten plot (inset), a Km of 9 µM was calculated. B, determination of the pH optimum using a buffer consisting of citrate-MES-MOPS-Bicine (35 mM). C, dependence on substrate concentration. Values represent the mean ± S.D. of duplicates expressed in nmol/h/mg protein.

 
Activation of Phytenic Acid to Its CoA Ester—Because of the extremely low phytenic acid reductase activity, we investigated whether phytenic acid might be converted into phytanic acid via some alternative mechanism. Because phytanic acid is a known substrate for long-chain acyl-CoA synthetase (20) and needs to be activated to its CoA ester before it can be {alpha}-oxidized, we explored whether phytenic acid might also be a substrate for a synthetase. To test this, the rate of activation of phytenic acid to its CoA ester was determined by incubating rat liver homogenate with phytenic acid in the presence of Mg2+, ATP, and CoA. Synthetase activity with phytanic acid as substrate was measured for comparison. In Fig. 4B, it is shown that phytenic acid can indeed be activated to phytenoyl-CoA, and the rates of activation were found to be only slightly lower compared with those observed for phytanic acid. Interestingly, predominantly one isomer was produced, presumably corresponding to E-phytenoyl-CoA based on the elution profile and response ratio of Z- and E-phytenoyl-CoA (14).

Subsequently, we tested whether phytenoyl-CoA could also be a substrate for the reductase reaction. Rat liver homogenates were incubated with either phytenic acid or phytenoyl-CoA, and after termination of the reactions, samples were subjected to alkaline hydrolysis to release the CoA so that all reaction products could be analyzed by gas chromatography-mass spectrometry in the same way. Remarkably, these experiments revealed that the reductase activity was 15 times higher using phytenoyl-CoA as a substrate instead of phytenic acid (Fig. 4C).

To establish that phytenoyl-CoA is a true intermediate of phytol degradation, acyl-CoA esters were analyzed in cultured cells after incubation with phytol. To this end, cultured rat hepatocytes were grown for 4 days in medium supplemented with 50 µM phytol. The cells were then homogenized, and CoA esters were extracted and analyzed by HPLC tMS. In addition to phytanoyl-CoA, E-phytenoyl-CoA was also detected (Fig. 5), suggesting that E-phytenoyl-CoA is indeed a true intermediate of the phytol degradation pathway.

Next, we assessed whether phytenic acid reductase activity could be restored in the fractions obtained by differential centrifugation by addition of cofactors CoA, ATP, and Mg2+, which are required for the activation of phytenic acid to phytenoyl-CoA. Activity was measured as described in Fig. 4C to allow a direct comparison between incubations with or without the cofactors. An increased reductase activity in the presence of these cofactors was observed (data not shown), indicating that phytenoyl-CoA rather than phytenic acid might indeed be the true substrate for the reductase. However, because formation of phytanoyl-CoA from phytenic acid requires the action of an acyl-CoA synthetase, which might not be present in some of the fractions or might be limiting, the activity measured in the presence of ATP, Mg2+, and CoA is likely to be an underestimation. For this reason (and because phytenoyl-CoA appears to be the bona fide intermediate of phytol, as concluded from the previous experiments), we used phytenoyl-CoA as substrate for subsequent reductase measurement.



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  		FIG. 7.
Phytenoyl-CoA reductase activity in a density gradient. Organelles from a rat liver homogenate were separated on a continuous density gradient as described under "Materials and Methods." The rat liver was derived from a rat fed a diethylhexylphthalate-supplemented diet to increase the number of peroxisomes and the weight of the liver. The gradient was divided into discrete samples, numbered from 1 at the bottom to 21 at the top, in which phytenoyl-CoA reductase activity was measured (A). To localize different organelles in the gradient, the marker enzymes (B) glutamate dehydrogenase (mitochondria), (C) esterase (microsomes), and (D) catalase (peroxisomes) as well as (E) protein were measured. F, phytenic acid activation to phytenoyl-CoA. All enzymatic activities are expressed as a percentage of total activity.

 
Characterization of the Reduction of Phytenoyl-CoA—Optimal reaction conditions were determined, this time using phytenoyl-CoA as substrate, and samples were analyzed by HPLC. The reaction was dependent on NADPH as cofactor, and the Km for NADPH was 9 µM (Fig. 6A). Maximum activity was found at a pH of 7.0 (Fig. 6B). As had been observed for phytenic acid, titration of phytenoyl-CoA revealed decreasing activities with substrate concentrations exceeding 100 µM, making calculation of Km and Vmax impossible (Fig. 6C). Based on these results, we selected a phytenoyl-CoA concentration of 100 µM in all subsequent reductase measurements. Addition of bovine serum albumin or methyl-{beta}-cyclodextrin to the incubation mixture had no influence on reductase activity (data not shown). Phytanoyl-CoA production was linear with protein up to at least 0.5 mg/ml and linear over time up to at least 60 min (data not shown).

Subcellular Localization of Phytenoyl-CoA Reductase Activity—Reductase activity using phytenoyl-CoA as substrate was measured in the same fractions obtained by differential centrifugation of a rat liver homogenate as used in the experiment shown in Fig. 3. In contrast to the incubations with phytenic acid as substrate, good recovery was found with phytenoyl-CoA as substrate (85%). The relative specific phytenoyl-CoA reductase activity was >1 in both the mitochondria-enriched and lysosome-enriched fractions (Fig. 3B).

To establish the subcellular localization of phytenoyl-CoA reductase more precisely, we measured activity in the different fractions of the density gradient. A bimodal activity profile was found, with peaks corresponding to the peroxisomal marker and the mitochondrial marker (Fig. 7).

In addition, we measured synthetase activity in the same density gradient fractions with phytenic acid as substrate. Phytenoyl-CoA production was found in the peroxisomes and microsomes (Fig. 7F).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
To elucidate the breakdown pathway of phytol to phytanic acid, we previously studied the conversion of phytol to phytenic acid (11). We observed that phytol was converted to its aldehyde, which was then converted into phytenic acid, as had been postulated by Muralidharan and Muralidharan (10, 21). We also found that the production of phytenic acid from phytol was mediated by the enzyme fatty aldehyde dehydrogenase. In this study, we focused on the last step of this pathway, the conversion of phytenic acid into phytanic acid. Enzymatic production of phytanic acid from phytenic acid was indeed observed in rat liver homogenates. However, doubts about the relevance of this reaction arose when hardly any activity was recovered in fractions prepared from rat liver homogenates by means of differential and density gradient centrifugation. As an alternative, activation of phytenic acid to its CoA ester prior to the reduction of the double bond was investigated. We showed that phytenic acid was indeed a substrate for an acyl-CoA synthetase and, moreover, that phytenoyl-CoA was a better substrate for the reductase reaction than phytenic acid. Proof that phytenoyl-CoA is a bona fide intermediate of phytol degradation came from the fact that it could be detected in rat hepatocytes cultured in the presence of phytol. Fig. 8 shows the new, fully elucidated pathway of the conversion of phytol into phytanic acid.



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  		FIG. 8.
Revised pathway of the breakdown of phytol to phytanic acid. Phytol is converted by an alcohol dehydrogenase and fatty aldehyde dehydrogenase on the endoplasmic reticulum into E-phytenic acid. A synthetase at the endoplasmic reticulum or the peroxisome produces E-phytenoyl-CoA, which is reduced to phytanoyl-CoA at the mitochondrion or the peroxisome. Phytanoyl-CoA is further broken down by {alpha}-oxidation in the peroxisome.

 
Subsequent characterization of the phytenoyl-CoA synthetase reaction showed that this activity is localized both in microsomes and peroxisomes. Because production of phytenic acid takes place on the outside of the endoplasmic reticulum membrane where fatty aldehyde dehydrogenase is localized, activation of phytenic acid to phytenoyl-CoA could occur either directly at the endoplasmic reticulum membrane or at the peroxisomes. The same finding has been observed for phytanic acid, which was shown to be activated by long-chain acyl-CoA synthetase (20, 22).

Phytenoyl-CoA reductase activity was detected both in mitochondria and in peroxisomes. Because {alpha}-oxidation of phytanoyl-CoA is a peroxisomal process (5, 23), it is tempting to speculate that reduction of phytenoyl-CoA also takes place inside the peroxisome (Fig. 8). Interestingly, in a study by Das et al. (24), the cloning of a peroxisomal 2-enoyl-CoA reductase, an enzyme that is speculated to play a role in the chain elongation pathway, was described. This reductase has affinity for enoyl-CoA esters with chain lengths up to C16 and appears to be ubiquitously expressed with high levels present in liver and kidney. Future work will have to demonstrate whether this enzyme indeed catalyzes the reduction of phytenoyl-CoA.

In summary, the results described in this study show that phytenic acid is first activated to its CoA ester before being reduced to phytanoyl-CoA, which is in contrast to the mechanism that was proposed earlier (10, 21). Phytenoyl-CoA reductase activity was found to be present in mitochondria and peroxisomes, and we hypothesize that under in vivo conditions, phytenoyl-CoA reduction takes place in peroxisomes, where its product, phytanoyl-CoA, can subsequently be {alpha}-oxidized. With these findings, we have achieved full understanding of the phytol degradation pathway.


   FOOTNOTES
 
* This work was supported by a grant from the Meelmeijer Fund and Grant QLG3-2002-00696 from the European Commission. 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

** To whom correspondence should be addressed: Laboratory for Genetic Metabolic Diseases (F0-224), Dept. of Pediatrics, Academic Medical Center, University of Amsterdam, P. O. Box 22700, 1100 DE Amsterdam, The Netherlands. Tel.: 31-205664197; Fax: 31-206962596; E-mail: R.J.Wanders{at}amc.uva.nl.

1 The abbreviations used are: HPLC, high performance liquid chromatography; tMS, tandem mass spectrometry; MES, 2-(N-morpholino)ethanesulfonic acid; MOPS, 3-(N-morpholino)propanesulfonic acid; Bicine, N,N-bis(2-hydroxyethyl)glycine. Back

2 C. W. van Roermund, manuscript in preparation. Back


   ACKNOWLEDGMENTS
 
We thank Dr. Sacha Ferdinandusse for many helpful comments about the results and preparation of this article. We are also grateful to Rob Ofman for work on the centrifugation experiments and to Dr. Carlo van Roermund, Albert Bootsma, and Dr. Wim Kulik for technical assistance with the detection of acyl-CoA esters.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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