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Journal of Lipid Research, Vol. 48, 77-85, January 2007
Copyright © 2007 by American Society for Biochemistry and Molecular Biology
in its regulation
Laboratory of Genetic Metabolic Diseases, Departments of Clinical Chemistry and Pediatrics, Emma's Children's Hospital, Academic Medical Center, University of Amsterdam, 1105 AZ Amsterdam, The Netherlands
Published, JLR Papers in Press, October 2, 2006.
1 To whom correspondence should be addressed. e-mail: s.ferdinandusse{at}amc.uva.nl
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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(PPAR). To investigate the metabolism of phytol and the role of PPAR in its regulation, wild-type and PPAR knockout (PPAR–/–) mice were fed a phytol-enriched diet or, for comparison, a diet enriched with Wy-14,643, a synthetic PPAR agonist. After the phytol-enriched diet, phytol could only be detected in small intestine, the site of uptake, and liver. Upon longer duration of the diet, the level of the (E)-isomer of phytol increased significantly in the liver of PPAR–/– mice compared with wild-type mice. Activity measurements of the enzymes involved in phytol metabolism showed that treatment with a PPAR agonist resulted in a PPAR-dependent induction of at least two steps of the phytol degradation pathway in liver. Furthermore, the enzymes involved showed a higher activity toward the (E)-isomer than the (Z)-isomer of their respective substrates, indicating a stereospecificity toward the metabolism of (E)-phytol. In conclusion, the results described here show that the conversion of phytol to phytanic acid is regulated via PPAR and is specific for the breakdown of (E)-phytol.
Supplementary key words peroxisome proliferator-activated receptor • fatty aldehyde dehydrogenase • branched-chain fatty acids
Abbreviations: FALDH, fatty aldehyde dehydrogenase; MBP, maltose binding protein; MTBSTFA, N-methyl-N-(tert-butyldimethylsilyl)trifluoroacetamide; PPAR, peroxisome proliferator-activated receptor
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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-oxidation. This results in the formation of pristanic acid, which is one carbon atom shorter than phytanic acid and can be normally ß-oxidized (10). A deficiency in -oxidation, such as in Refsum disease, leads to increased levels of phytanic acid in plasma and tissues of patients, and this is thought to cause the main clinical symptoms of this disorder: retinitis pigmentosa, peripheral neuropathy, and cerebellar ataxia (3, 4). Because the breakdown of phytol will contribute to the phytanic and pristanic acid levels in these patients, it is important to study its metabolism and regulation.
In many animal studies, phytol is used as a precursor of phytanic acid. Addition of phytol to the diet results in an increase of phytol metabolites in tissues and plasma (6, 11–13). This has been used as a model to study the effects of the accumulation of phytol metabolites on fatty acid metabolism, in particular via the activation of the nuclear hormone receptor peroxisome proliferator-activated receptor (PPAR), which is an important transcription factor in the regulation of fatty acid metabolism. Both phytanic and pristanic acid have been shown to activate PPAR in vitro (14, 15), and more recently, PPAR was also shown to be activated in vivo in mice fed a phytol-enriched diet (13).
Figure 1 shows a schematic representation of the metabolism of phytol to phytanic acid and the enzymes involved. First, phytol is converted into phytenal via a yet unknown alcohol dehydrogenase, and subsequently, phytenal is converted into phytenic acid by the microsomal enzyme fatty aldehyde dehydrogenase (FALDH), which is encoded by the Aldehyde Dehydrogenase 3A2 gene (16). Recently, it was shown that phytenic acid is activated to its corresponding CoA ester, either at the endoplasmic reticulum or at the peroxisome, which is then reduced by a reductase converting phytenoyl-CoA to phytanoyl-CoA either in the peroxisome or at the mitochondrion (17).
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in its regulation. To this end, phytol concentrations were determined and the activities of the enzymes involved in phytol metabolism were analyzed in different tissues of wild-type and PPAR-deficient (PPAR–/–) mice fed a phytol-enriched diet. To further investigate the role of PPAR, the activities of the enzymes involved in phytol metabolism were analyzed in different tissues of wild-type and PPAR–/– mice fed a diet enriched with Wy-14,643, a well-known synthetic PPAR agonist. |
MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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cells and Trizol were obtained from Invitrogen (Carlsbad, CA). The pGEM-T vector was from Promega (Madison, WI). The pMAL-C2X vector and amylose-resin columns were purchased from New England Biolabs (Beverly, MA). Freund's complete and incomplete adjuvant were from Difco Laboratories (Detroit, MI). Goat anti-rabbit antibodies conjugated with alkaline phosphatase were obtained from Bio-Rad (Hercules, CA). Completemini tablets containing a cocktail of protease inhibitors were purchased from Roche (Basel, Switzerland). Nitrocellulose membrane was from Schleicher and Shuell (Keene, NH). Protifar was obtained from Nutricia (Zoetermeer, The Netherlands). All other chemicals were of analytical grade.
Animals
Male wild-type and PPAR–/– mice on a Sv/129 genetic background were used to study the effects of phytol and Wy-14,643 treatment (20), and male wild-type mice on a Swiss genetic background were used to study the effects of fenofibrate treatment. Six week old wild-type and PPAR–/– mice were fed pelleted mouse chow containing no (control) or 0.5% (w/w) phytol for 1 or 8 weeks. Each group consisted of three animals. Nine week old wild-type and PPAR–/– mice were fed pelleted mouse chow containing no or 0.1% (w/w) Wy-14,643 for 2 weeks. Each group consisted of three animals. Eight week old wild-type mice were fed pelleted mouse chow containing no or 0.1% (w/w) fenofibrate for 5 weeks. Each group consisted of five animals. At the end of the experiments, mice were anesthetized using isoflurane, and tissues were harvested. The animals were always euthanized at the same time of day, and they had free access to water and food until that moment. Tissues were snap-frozen in liquid nitrogen and stored at –80°C until further analysis. All animal experiments were approved by the University of Amsterdam Animals Experiments Committee.
Synthesis of phytenal
Phytenal was synthesized from its precursor phytol essentially as described previously (18). Briefly, phytol [a mixture of (E)- and (Z)-isomers] was oxidized to the corresponding aldehydes with CrO3-pyridine in dry methylene chloride under a flow of nitrogen to exclude water and oxygen. The oxidant was prepared in 30 ml of dichloromethane (dried over anhydrous Na2SO4) by addition of 24 mmol of anhydrous pyridine, followed by 12 mmol of CrO3 in small batches under magnetic stirring. After clearing of the initial turbidity, 2 mmol of phytol in 4 ml of dichloromethane was added. The mixture was stirred for 1 h. Subsequently, the solution was transferred into a separatory funnel and washed with aqueous 5% NaOH for elimination of the excess CrO3, 5% HCl for the elimination of pyridine, 5% NaHCO3 for neutralization, and finally saturated NaCl. After drying over anhydrous Na2SO4, the organic solvent was filtrated and evaporated under a flow of nitrogen. The residue was then purified over a silica gel column and eluted with chloroform, which was subsequently evaporated under a flow of nitrogen. Phytenal was stored in aliquots under argon at –80°C until use. GC-MS analysis of the product showed two peaks corresponding to the (E)- and (Z)-isomers of phytenal with a molecular ion at m/z 294 and a fragmentation pattern specific for aldehydes.
Phytol analysis in tissue homogenates
Tissue phytol levels were determined as described (13). Briefly, tissues were homogenized in PBS. C19-OH (10 µl of a 50 µM solution dissolved in ethanol) was added to 100 µl of tissue homogenate (5 mg/ml), and the sample was subjected to alkaline hydrolysis by adding 2 ml of 1 M NaOH in methanol for 45 min at 110°C. After cooling to room temperature, the pH was decreased by adding 480 µl of 37% HCl. Subsequently, phytol was extracted with 2 ml of hexane, and the organic layer was evaporated to dryness under nitrogen at 40°C. Samples were dissolved in 0.5 ml of heptane and purified on a silica gel column using 92:8 (v/v) heptane-diethyl ether as eluent. The eluate was evaporated to dryness under nitrogen at 40°C and derivatized with MTBSTFA and pyridine (50 µl each) at 80°C for 30 min. Samples were evaporated to dryness under nitrogen at 40°C, dissolved in 100 µl of hexane, and subjected to GC-MS analysis, essentially as described previously (21). The metabolites were quantified using a calibration curve of phytol.
Enzyme activity measurements
All enzyme activity measurements were performed in freshly prepared tissue homogenates. FALDH activity measurements in liver were performed essentially as described previously (21). Briefly, FALDH activity was measured with both phytol and phytenal as substrates. The reaction mixture consisted of liver protein (20 and 50 µg for phytol and phytenal incubations, respectively), 50 mM glycine buffer, pH 9.2, 1 mM NAD+, 0.1% sodium cholate, and 1 mg/ml methyl-ß-cyclodextrin in a total volume of 500 µl. Reactions were performed at 37°C and initiated by the addition of substrate in final concentrations of 200 and 150 µM for phytol and phytenal, respectively. After 10 min, the incubations were terminated by the addition of 50 µl of 2 M HCl. As internal standard, 50 pmol of [2H3]phytanic acid, dissolved in toluene, was added. Then, 2 ml of hexane was added, and after vigorous vortexing, the samples were centrifuged, after which the organic layer was evaporated to dryness under a stream of nitrogen at 40°C. After derivatization with MTBSTFA and pyridine (50 µl each) at 80°C for 30 min, the samples were evaporated to dryness under nitrogen at 40°C, dissolved in 200 µl of hexane, and subjected to GC-MS analysis as described previously (21).
Enzymatic activation of phytenic acid to its CoA ester was measured essentially as described previously (17). Briefly, tissue homogenate (8 µg of protein for liver homogenates, 10 µg of protein for kidney and heart homogenates, and 25 µg of protein for small intestine and brain homogenates) was incubated with a mixture containing 50 mM Tris, pH 8.0, 10 mM ATP, 10 mM MgCl2, 0.5 mM CoA, 1 mg/ml methyl-ß-cyclodextrin, and 100 µM phytenic acid [a mixture of (E)- and (Z)-phytenic acid] in a total volume of 100 µl. The incubation was allowed to proceed for 30 min when the activity was measured in liver and for 60 min in all other tissues. Reactions were terminated by the addition of 100 µl of acetonitrile and placed on ice. Samples were centrifuged for 10 min at 10,000 g at 4°C, and supernatants were analyzed by HPLC.
Reduction of phytenoyl-CoA to phytanoyl-CoA was determined essentially as described previously (17). Briefly, the reaction mixture contained 50 mM HEPES, pH 7.0, 1 mM NADPH, 40 µM phytenoyl-CoA [a mixture of (E)- and (Z)-phytenoyl-CoA], and tissue homogenate (25 µg for kidney homogenates, 100 µg for liver homogenates, and 200 µg for small intestine, heart, and brain homogenates) in a total volume of 100 µl. Reactions were started by the addition of substrate and incubated at 37°C for 15 min for liver homogenates and for 30 min for all other tissue homogenates. Reactions were terminated by the addition of 100 µl of acetonitrile and placed on ice. Samples were centrifuged for 10 min at 10,000 g at 4°C, and supernatants were analyzed by HPLC.
Quantitative real-time RT-PCR analysis
Total RNA was isolated from RNAlater-treated mouse liver and kidney samples using Trizol extraction, after which cDNA was prepared using a first-strand cDNA synthesis kit for RT-PCR. Quantitative real-time PCR analysis of FALDH and ß-actin in liver and kidney was performed using the LightCycler FastStart DNA Master SYBR Green I kit. The following primers were used for FALDH: forward, 5'-CATACTTACAGATGTTGATCC-3'; reverse, 5'-CTCCAAAGGGCAGAGAATTA-3'. Primers for ß-actin were used as described (22). Melting curve analysis was carried out to confirm the generation of a single product. Amplification of a single product of the correct size was also confirmed by agarose gel electrophoresis. Duplicate analyses were performed for all samples. Data were analyzed using linear regression calculations as described by Ramakers et al. (23). To adjust for variations in the amount of input RNA, the values for FALDH were normalized against the values for the housekeeping gene ß-actin.
Preparation of antibodies
A polyclonal antiserum was raised against an N-terminal peptide corresponding to amino acids 1–96 of human FALDH, expressed as a fusion protein with maltose binding protein (MBP). The coding region for this peptide (bp 1–288) was amplified from human liver cDNA by PCR using the following primers: a BamHI-tagged forward primer (5'-ggatccATGGAGCTCGAAGTCCGG-3') and a SalI-tagged reverse primer (5'-gtcgacTTAGGCCTCATCCAGCATGGTGAG-3') (restriction sites are underlined). The resulting PCR product was ligated into the pGEM-T vector, and sequence analysis was performed to exclude sequence errors introduced by Taq polymerase. The fragment was subsequently cloned into the pMAL-C2X vector to create pMAL-FALDH-Nterm, using the BamHI and SalI restriction sites downstream of the isopropyl-1-thio-ß-D-galactopyranoside-inducible promoter. The E. coli strain INV was transformed with the expression plasmid pMAL-FALDH-Nterm. Transformed cells were grown at 37°C in 100 ml of Luria-Bertani medium supplemented with 0.2% (w/v) glucose and 100 µg/ml ampicillin to an optical density at 600 nm of 
0.6, and isopropyl-1-thio-ß-D-galactopyranoside was added to a final concentration of 1 mM to induce expression of the fusion protein. Cells were lysed by sonication on ice (twice for 15 s at an output of 8 W, with an interval of 1 min). The MBP fusion protein was purified on amylose-resin columns according to the protocol of the manufacturer (New England Biolabs). MBP-FALDH-Nterm was subjected to preparative SDS-PAGE, isolated from the gel, and subsequently used to raise antibodies. To this end, a female New Zealand White rabbit was injected subcutaneously with 100 µg of the antigen mixed with an equal volume of Freund's complete adjuvant. After 1 month, the immunization was continued by booster injections (each containing 100 µg of antigen in Freund's incomplete adjuvant) until a satisfactory antibody titer was obtained.
Immunoblot analysis
Small pieces of tissue were homogenized in PBS containing a cocktail of protease inhibitors. Twenty micrograms of protein was separated on a 10% (w/v) SDS-polyacrylamide gel and transferred to a nitrocellulose membrane (24). After blocking of nonspecific binding sites with 30 g/l Protifar and 10 g/l BSA in 1 g/l Tween-20/PBS, the blot was incubated for 2 h with rabbit polyclonal antibodies raised against FALDH diluted 1:1,500 in 4% BSA in PBS. Goat anti-rabbit IgG antibodies conjugated to alkaline phosphatase were used for detection.
Statistical analyses
Data are expressed as means ± SD. Statistical significance was evaluated using an unpaired Student's t-test. The results were considered significant at P < 0.01.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In Fig. 2
, the amounts of (Z)- and (E)-phytol detected in liver and small intestine of wild-type and PPAR–/– mice are shown. The phytol levels in small intestine varied considerably between individual mice, most likely attributable to the fact that the animals were fed ad libitum until euthanized. Despite this high variance, the amount of (E)-phytol was higher in PPAR–/– mice than in wild-type animals (Fig. 2C).
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–/– mice was more substantial than in wild-type animals (13). In this study, we determined the levels of both (Z)- and (E)-phytol in liver. After 1 week on a phytol-enriched diet, there were no significant differences in phytol levels between wild-type and PPAR–/– mice (Fig. 2). After 8 weeks on the phytol-enriched diet, however, the amount of (E)-phytol, in contrast to the (Z)-isomer, was increased significantly in PPAR–/– mice compared with wild-type animals. In wild-type mice, the phytol level did not increase with time, but it did increase in PPAR–/– mice.
Phytol-degrading enzymes in the liver
Because FALDH only converts (E)-phytenal into (E)-phytenic acid, the predominant accumulation of (E)-phytol in liver of PPAR–/– mice in combination with lower hepatic levels of phytanic acid in PPAR–/– mice compared with wild-type animals (13) suggests a decreased capacity of the phytol degradation pathway in PPAR–/– mice. To investigate this, we measured the activities of the enzymes of this pathway in liver (Fig. 3
).
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–/– mice. Under the influence of the phytol-enriched diet, both assays revealed significantly increased (E)-phytenic acid formation in wild-type mice (Fig. 3A). Because phytol metabolites are ligands for PPAR, this suggests that PPAR plays a role in the regulation of FALDH expression and might also be involved in the regulation of the expression of the alcohol dehydrogenase. Because the fold induction of the activity measured with phytol and phytenal as substrate was the same, it is probable that the reaction catalyzed by FALDH is the rate-limiting step. Interestingly, a small but significant induction of (E)-phytenic acid formation also was observed with both substrates in PPAR–/– mice, although this was much less pronounced compared with wild-type mice, indicating that there are both PPAR-dependent and -independent effects. Immunoblot analysis of FALDH showed a clear induction of FALDH protein in wild-type mice fed a phytol diet compared with the control diet, whereas no induction was observed in PPAR–/– mice (Fig. 3A). To substantiate that FALDH expression is under the control of PPAR, FALDH expression was studied in wild-type and PPAR–/– mice treated with the PPAR ligand Wy-14,643. Figure 3B shows that induction of FALDH protein indeed occurred in wild-type mice, but not in PPAR–/– animals, fed a diet enriched with Wy-14,643. Investigation of liver mRNA levels using quantitative RT-PCR showed a 3.0-fold induction of FALDH mRNA after Wy-14,643 treatment in wild-type mice. The induction of FALDH mRNA is in agreement with the mode of action of Wy-14,643, which activates PPAR to induce the transcription of its target genes.
Second, the activation of (E)-phytenic acid to its CoA ester was significantly increased in liver from phytol-fed and Wy-14,643-treated wild-type animals compared with animals on a control diet (Fig. 3C). The specific activity for the formation of (Z)-phytenoyl-CoA was 17-fold lower than for (E)-phytenoyl-CoA (Table 1
), and although it was slightly higher in phytol-fed and Wy-14,643-treated wild-type mice, this difference was not statistically significant (data not shown). The increase in (E)-phytenoyl-CoA formation was not observed in PPAR–/– mice on a diet enriched with phytol or Wy-14,643, indicating that also the induction of phytenoyl-CoA synthetase is PPAR-dependent.
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–/– mice. This suggests that, in addition to phytenoyl-CoA synthetase, phytenoyl-CoA reductase is regulated in a PPAR-dependent manner. However, no induction of phytenoyl-CoA reductase activity was observed after feeding a Wy-14,643-enriched diet. The difference in phytenoyl-CoA reductase regulation between the phytol diet and Wy-14,643 treatment could be attributable to ligand-specific activation of PPAR or to the fact that phytenoyl-CoA reductase is not induced directly via PPAR activation but is induced in phytol-fed animals as a consequence of accumulating metabolites from the phytol breakdown pathway. To further investigate this, the effect of fenofibrate, another synthetic PPAR agonist, on phytenoyl-CoA reductase activity was investigated. Like Wy-14,643 treatment, fenofibrate treatment did not result in an increase of phytenoyl-CoA reductase activity in wild-type mice. The mean specific activity (±SD) measured in liver from five wild-type mice was 0.43 ± 0.12 nmol/mg/min, and the specific activity after fenofibrate treatment was 0.34 ± 0.03 nmol/mg/min. These results show that phytenoyl-CoA reductase is not induced directly via PPAR activation but that accumulating metabolites of the phytol breakdown pathway, which is upregulated after PPAR activation, cause an increase in phytenoyl-CoA reductase activity. When studying the stereochemistry of the reductase reaction, the specific activity for the reduction of (E)-phytenoyl-CoA was found to be 2.8-fold higher than that for the purified (Z)-isomer (data not shown).
Site of phytol degradation
To gain more insight into the site of phytol metabolism in the body, we investigated the activities of the enzymes involved in this pathway in different mouse tissues. The tissue distribution of FALDH was described previously (25). Highest expression and activity were found in liver, followed by lung, kidney, small intestine, and stomach. Low but detectable FALDH expression and activity were also present in brain, skin, heart, and muscle. Immunoblot analysis of FALDH in kidney, heart, and brain homogenates is depicted in Fig. 4A
. In heart and brain, FALDH protein levels are detectable, but no induction of FALDH protein was observed after the Wy-14,643-enriched diet. In kidney, however, FALDH protein was significantly induced in wild-type mice after the Wy-14,643-enriched diet, indicating regulation via PPAR. Analysis of FALDH mRNA levels in kidney (Fig. 4B) showed an induction of FALDH mRNA in wild-type mice after Wy-14,643 treatment but not in PPAR–/– mice, which is in agreement with the immunoblot results.
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As for phytenoyl-CoA synthetase, the specific activity of phytenoyl-CoA reductase was highest in liver. In kidney, the specific activity was approximately half of the value for liver. In all other tissues tested, very low, but detectable, activities of phytenoyl-CoA reductase were measured (Table 1). In the extrahepatic tissues, phytenoyl-CoA reductase activity was not induced by phytol or by Wy-14,643.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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plays an important role in the regulation of the enzymes involved in phytol degradation. In livers from PPAR–/– mice, phytol was found to accumulate to much higher levels compared with livers from wild-type animals upon longer duration of the phytol-enriched diet, suggesting a decreased capacity for phytol metabolism in PPAR–/– mice compared with wild-type animals. Further investigation showed that the regulation of at least two of the four enzymes of this breakdown pathway (i.e., FALDH and phytenoyl-CoA synthetase) is PPAR-dependent, because their activities increase after phytol or Wy-14,643 feeding in livers from wild-type mice but not from PPAR–/– animals. Only for FALDH was it reported previously that expression can be induced by the action of PPAR, although no functional peroxisome proliferator response element has been identified yet (26). Both (Z)- and (E)-phytol are absorbed in the small intestine of the mouse, as shown by the presence of both isomers. Upon longer duration of the phytol-enriched diet, no increase in the accumulation of phytol was observed in small intestine. In contrast, the accumulation of phytol in the liver increased with time, which was mainly attributable to an increase in (E)-phytol levels. The predominant accumulation of the (E)-isomer of phytol strongly suggests that the phytol degradation pathway is stereospecific. Indeed, FALDH, which catalyzes the second step in the breakdown of phytol, exclusively forms (E)-phytenic acid. The third and fourth steps of the phytol degradation pathway, catalyzed by phytenoyl-CoA synthetase and phytenoyl-CoA reductase, respectively, also show a strong preference for (E)-isomers as substrate. The peroxisomal trans-2-enoyl-CoA reductase identified by Das, Uhler, and Hajra (27) was suggested as a putative candidate for the reduction of phytenoyl-CoA to phytanoyl-CoA (17). This reductase has been shown to be stereospecific toward the (E)-form of enoyl-CoA esters (17, 27) and has now been shown to convert (E)-phytenoyl-CoA to (E)-phytanoyl-CoA (28). It remains to be established, however, whether this enzyme is the sole reductase catalyzing the reduction of phytenoyl-CoA, especially because mitochondria, which lack the reductase identified by Das, Uhler, and Hajra (27), are also able to reduce phytenoyl-CoA. In conclusion, our results show that the phytol degradation pathway is stereospecific for the breakdown of (E)-phytol to phytanic acid.
All of the enzymes involved in the breakdown of phytol to phytanic acid displayed high activity in mouse liver. The first step, catalyzed by the alcohol dehydrogenase, cannot be measured directly, because the product of this reaction (i.e., phytenal) is very unstable. At least in liver, this enzyme must be present, because phytol was converted into (E)-phytenic acid in liver homogenates. The enzyme activity measurements of the other enzymes involved in the breakdown of phytol, together with the measurement of phytol levels in different tissues of the mouse, strongly suggest that the main site of phytol degradation is the liver. Phytol itself could be detected in liver and small intestine only after the phytol-enriched diet. Most likely, the phytol levels in small intestine are a reflection of the phytol that has been absorbed from the diet just before euthanasia. As a consequence, no increase in small intestinal phytol levels was observed with time, in contrast to the hepatic phytol concentration, which did increase with time.
In summary, the results described here show that the degradation of phytol to phytanic acid is regulated via PPAR. Furthermore, phytol accumulates almost exclusively in liver, and the enzymes involved in phytol degradation display their highest activities in liver, indicating that the liver is the main site of phytol metabolism in the body. Finally, the pathway shows specificity for the breakdown of (E)-phytol, because this isomer accumulates predominantly in liver of PPAR–/– mice on a phytol-enriched diet and at least three of the enzymes of the pathway have been shown to be stereospecific toward this isomer.
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ACKNOWLEDGMENTS |
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Manuscript received January 31, 2006 and in revised form May 16, 2006 and in re-revised form June 7, 2006 and in re-re-revised form September 27, 2006.
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