The Saccharomyces cerevisiae FAT1 Gene Encodes an Acyl-CoA Synthetase That Is Required for Maintenance of Very Long Chain Fatty Acid Levels*

Jae-Yeon Choi and Charles E. MartinDagger

From the Division of Life Sciences, the Bureau of Biological Research, Nelson Laboratories, Rutgers University, Piscataway, New Jersey 08854-808

    ABSTRACT
Top
Abstract
Introduction
References

The Saccharomyces cerevisiae FAT1 gene appears to encode an acyl-CoA synthetase that is involved in the regulation of very long chain (C20-C26) fatty acids. Fat1p, has homology to a rat peroxisomal very long chain fatty acyl-CoA synthetase. Very long chain acyl-CoA synthetase activity is reduced in strains containing a disrupted FAT1 gene and is increased when FAT1 is expressed in insect cells under control of a baculovirus promoter. Fat1p accounts for approximately 90% of the C24-specific acyl-CoA synthetase activity in glucose-grown cells and approximately 66% of the total activity in cells grown under peroxisomal induction conditions. Localization of functional Fat1p:green fluorescent protein gene fusions and subcellular fractionation of C24 acyl-CoA synthetase activities indicate that the majority of Fat1p is located in internal cellular locations. Disruption of the FAT1 gene results in the accumulation of very long chain fatty acids in the sphingolipid and phospholipid fractions. This includes a 10-fold increase in C24 acids and a 6-fold increase in C22 acids. These abnormal accumulations are further increased by perturbation of very long chain fatty acid synthesis. Overexpression of Elo2p, a component of the fatty acid elongation system, in fat1Delta cells causes C20-C26 levels to rise to approximately 20% of the total fatty acids. These data suggest that Fat1p is involved in the maintenance of cellular very long chain fatty acid levels, apparently by facilitating beta -oxidation of excess intermediate length (C20-C24) species. Although fat1Delta cells were reported to grow poorly in oleic acid-supplemented medium when fatty acid synthase activity is inactivated by cerulenin, fatty acid import is not significantly affected in cells containing disrupted alleles of FAT1 and FAS2 (a subunit of fatty acid synthase). These results suggest that the primary cause of the growth-defective phenotype is a failure to metabolize the incorporated fatty acid rather than a defect in fatty acid transport. Certain fatty acyl-CoA synthetase activities, however, do appear to be essential for bulk fatty acid transport in Saccharomyces. Simultaneous disruption of FAA1 and FAA4, which encode long chain (C14-C18) fatty acyl-CoA synthetases, effectively blocks the import of long chain saturated and unsaturated fatty acids.

    INTRODUCTION
Top
Abstract
Introduction
References

In Saccharomyces cerevisiae, most fatty acids (>95%) are C12-C18 species that are found in membrane glycerolipids. These are formed de novo by the soluble cytoplasmic fatty acid synthetase complex. Very long chain (C20-C26) fatty acids (VLCFAs),1 which are predominantly found in sphingolipids, are formed by a separate, ER membrane-bound system that elongates C16 and C18 saturated fatty acids to C26 and C28 species. Most of the very long chain fatty acids found in yeast are 26:02 and hydroxy-26:0. C20-C24 VLCFAs are minor sphingolipid components that are apparently derived from metabolic intermediates in the formation of the 26-carbon species. Our laboratory has recently found two genes, ELO2 and ELO3, that are involved in very long chain fatty acid synthesis (1). Each gene apparently encodes a single component of a system that elongates C16 and C18 acids to C20-C26 VLCFAs. Elo2p is involved in the elongation of fatty acids up to 24 carbons. Elo3p apparently has a broader substrate specificity and is essential for the conversion of 24-carbon acids to 26-carbon species (1).

Debilitating human diseases are associated with the accumulation of VLCFAs. A characteristic of human X-linked adrenoleukodystrophy is an absence or reduction of very long chain acyl-CoA synthetase (VLACS) activity, which leads to the accumulation of saturated VLCFAs. This appears to be caused by the mutations in the X-linked adrenoleukodystrophy gene, which encodes the ABC transporter in peroxisomal membranes (2, 3). The function of the adrenoleukodystrophy protein is not clearly explained but might be involved in the transport of VLCFAs into peroxisomes or in the transport of proteins that are required for the beta -oxidation of VLCFAs.

beta -Oxidation of VLCFAs in yeast and mammals is known to occur primarily in peroxisomes (4, 5). The formation of CoA thioesters from VLCFAs is an initial step for beta -oxidation and is catalyzed by a VLACS. A VLACS was recently purified from the rat liver peroxisomal membranes and microsomes, leading to the identification of its gene (6, 7).

Acyl-CoA synthetase activity is required for the biosynthesis and the catabolic oxidation of fatty acids. Previous studies have identified four fatty acyl-CoA synthetases in Saccharomyces that appear to be involved in the metabolism of long chain (C12-C18) fatty acids (8, 9). We had previously determined that the two major synthetase activities, encoded by the FAA1 and FAA4 genes, were not essential for VLCFA synthesis and that disruption of either or both genes did not affect VLCFA composition.

Given the important role of acyl-CoA synthetase activity in the biosynthesis and metabolism of VLCFAs, we attempted to identify a yeast VLACS homologue by data base searching. The amino acid sequence of open reading frame YBR041W was found to be homologous to that of the rat VLACS. Previously, YBR041W had been designated as FAT1 and was proposed to function in long chain fatty acid transport (10). In this report, we show that the protein encoded by the YBR041W sequence is a functional homolog for the rat VLACS and that disruption of the gene can cause large accumulations of aberrant, C20-C24 VLCFAs, suggesting that Fat1p plays an important role in controlling cellular VLCFA levels. Bulk fatty acid transport does not appear to be affected by the loss of Fat1p activity; however, given its multiplicity of cellular locations, it could also play a significant role in the uptake of low levels of nutrient fatty acids. In examining the role of fatty acyl-CoA synthetases in fatty acid import, however, we identified two fatty acyl-CoA synthetases, Faa1p and Faa4p, that are essential for the bulk import of nutrient fatty acids.

    EXPERIMENTAL PROCEDURES

Strains and Growth Medium

Yeast strains constructed for this study and their genotypes are presented in Table I. Strains containing faa1Delta , faa4Delta , and faa1Delta /faa4Delta gene disruptions were obtained from J. Gordon and were previously described (11). Plasmids constructed for this study are shown in Table II. Standard yeast genetics methods were used for construction of strains bearing the appropriate mutations. Yeast cells were grown at 30 °C in YPAD (1% Bacto-yeast extract, 2% Bacto-peptone, 2% glucose, and 2 mg/liter adenine), YPADt (YPAD plus 1% tergitol nonionic detergent Nonidet P-40 (to disperse fatty acids), CM (complete synthetic medium (12)), or CMt (CM plus 1% tergitol) medium. Tergitol is used in the medium to disperse fatty acids. Unlike Brij and Tween compounds, which are derived from fatty alcohols or fatty acids, tergitol is a nonylphenol polyethylene oxide polymer that is apparently not metabolized. Alternative carbon sources (2% galactose; 2% raffinose; or 3% glycerol, 0.1% (3.54 mM) oleic acid, 0.25% Tween 40) were used by replacing glucose in appropriate media. Fatty acids were obtained from Sigma or Nu-Chek Prep (Elysian, MN). Escherichia coli strain DH5alpha was obtained from Life Technologies, Inc. 1-14C-Labeled 24:0 was obtained from American Radiolabeled Chemicals, Inc. (St. Louis, MO).

                              
View this table:
[in this window]
[in a new window]
 
Table I
S. cerevisiae strains used in this study

                              
View this table:
[in this window]
[in a new window]
 
Table II
Plasmids used in this study

Cloning and Disruption of FAT1, FAT2, and FAA2

Gene-disrupted cells were constructed by standard yeast methods using cloned gene sequences. Strains containing disrupted genes are shown in Table I. All gene disruptions were verified by PCR using genomic DNA from the disrupted cells as a template.

Disruption of FAT1-- A 2.3-kb DNA fragment containing the FAT1 coding sequence was derived by PCR using DTY10a genomic DNA as templates and PCR primers FAT1-1 and FAT1-2 in Table III. Plasmid pCRSK-FAT1 (Table II), containing the amplified PCR product, was used to create pCRSK-fat1Delta ::LEU2 or pCRSKfat1Delta ::HIS3 (Table II) in which 1.8 kb of the FAT1 coding region was replaced with the corresponding LEU2 or HIS3 marker genes. Linear fat1Delta ::LEU2 and fat1Delta ::HIS3 disruption cassettes were derived from those plasmids by digestion with BamHI and HpaI and transformed into strain DTY10a to create the fat1Delta deletion strains. A linear fat1Delta ::LEU2 sequence was also transformed into the elo3Delta ::HIS3 strain to create fat1Delta /elo3Delta double deletion stains. A linear fat1Delta ::HIS3 sequence was also transformed into fas2Delta ::LEU2 strains to create the fat1Delta /fas2Delta double deletion strains.

                              
View this table:
[in this window]
[in a new window]
 
Table III
PCR primers used in this study

Disruption of FAT2-- An 1840-base pair DNA fragment containing the FAT2 coding sequence was derived by PCR using DTY10a genomic DNA as templates and the PCR primers FAT2-1 and FAT2-2 in Table III. The pCRSK-FAT2 plasmid (Table II) containing amplified PCR product was used to create plasmid pCRSKfat2)::HIS3 in which 0.4 kb of the FAT2 coding part was released and replaced with the corresponding HIS3 marker gene. Linear fat2Delta ::HIS3 disruption cassettes were derived from the plasmid by digestion with BstEII and transformed into strain DTY10a to create the fat2Delta strains. A linear fat2)::HIS3 sequence was also transformed into the fat1Delta ::LEU2 strain to create the fat1Delta /fat2Delta double deletion strains.

Disruption of FAA2-- A 2.3-kb DNA fragment containing the FAA2 coding sequence was derived from PCR using DTY10a genomic DNA as templates and PCR primers FAA2-1 and FAA2-2 in Table III. Plasmid pCRSK-FAA2 (Table II) containing the amplified PCR product was used to create pCRSKfaa)::URA3 in Table II in which 0.5 kb of FAA2 protein coding sequence was released and replaced with the URA3 gene. A linear faa2Delta ::URA3 disruption cassette was derived from the plasmid by digestion with ClaI and transformed into strain DTY10a to create the faa2Delta deletion strain shown in Table I. A linear faa2Delta ::URA3 sequence was also transformed into the fat1Delta ::LEU2 strain and fat1Delta ::LEU2/fat2Delta ::HIS3 to create the fat1Delta /faa2Delta double disruption stain and the fat1Delta /fat2Delta /faa2Delta triple disruption strain.

Construction of GAL1-FAT1 Vectors

2.3 kb of DNA sequence containing FAT1 coding sequence and 3'-untranslated region was released by BamHI and HpaI digestion of the pCRSK-FAT1 (Table II). This element was inserted into the linearized and blunt ended YCpGAL1URA vector by BamHI, HindIIIj, and Klenow enzyme digestion to create vector pFAT1. That vector was transformed into fat1Delta ::LEU2 strains, and transformants were selected on CM plates without uracil and leucine.

Construction of FAT1-GFP Fusion Vectors

Protein coding elements of the FAT1 gene were fused in frame to the green fluorescent protein (GFP) sequences by insertion into centromeric plasmid vectors pTS395 and pTS408. These contain the GFP sequence linked to the GAL1 promoter and the 3' terminator sequence from the ACT1 gene. Vector pTS395 was used to fuse GFP to the C terminus, and vector pTS495 was used to fuse GFP to the N terminus of Fat1p. To create vector pTS395FAT1, a DNA sequence containing codons 1-669 of the FAT1 coding sequence was derived from PCR using DTY10a genomic DNA as templates and primers 5'-ATAGGATCCATGTCTCCCATACATGTTGTTGTC-3' (forward) and 5'-TTCTAGATAATTTAATTGTTTGTGCATCG-3' (reverse). To create the vector pTS408FAT1 construct, a DNA sequence containing the same codons was derived from DTY10a genomic DNA and primers 5'-ATAGGATCCATGTCTCCCATACATGTTGTTGTC-3' (forward) and 5'-TTCTAGACTATAATTTAATTGTTTGTGCATCG-3' (reverse). The amplified PCR products were cut with BamHI and XbaI and inserted to vectors digested with the same restriction enzymes.

Fluorescence Microscopy of Cells Containing FAT1-GFP Gene Fusions

To induce expression the FAT1-GFP fusion gene under control of the GAL1 promoter, fat1Delta strains harboring GFP fusion vectors were grown overnight in 5 ml of uracil and leucine drop-out CM containing 2% raffinose. Cells were induced for 4 h by resuspension in the same media containing 2% galactose. For oleic acid and galactose induction, cells were grown overnight in uracil and leucine drop-out CM containing 3% glycerol and induced for 4 h in the same medium containing 2% galactose, 0.1% oleic acid, 0.25% Tween 40. One µg/ml 4',6-diamidino-2-phenylindole was added to the media during the last 1.5 h of galactose induction. After induction, cells were washed with 1 ml of phosphate-buffered saline buffer (pH 7.4) three times and then resuspended in 100 µl of phosphate-buffered saline buffer. In a humid chamber, cells were dropped onto a Superfrost Plus slide (25 × 75 × 1 mm; VWR Scientific) and allowed to settle for 10 min. Excess cells were flushed from the slides with 100 µl of phosphate-buffered saline buffer and mounted in 0.1% sea plaque agar under a coverslip. All samples were viewed on a Nikon Diaphot 300 inverted microscope equipped with a Chroma Endow GFP filter set 41017 (excite HQ470/40; emitter HQ525/50; beam splitter Q495 LP). Images were captured using the Acquire software program.

Lipid Extraction and Fatty Acid Analysis

Fatty acid methyl esters were prepared by HCl methanolysis as described previously (13). Gas chromatography was performed on a Varian 3400CX chromatograph using a Supelcowax TM10 30 m × 0.32 mm column (Supelco) temperature programmed from 70 to 240 °C at 40 °C/min. Data were collected and analyzed using the Class-VP Chromatography Data System version 4.1 (Shimadzu Scientific Instruments) software.

Sphingolipid and glycerolipid fatty acids were fractionated from logarithmic phase cells as described by Pinto et al. (14). 5% trichloroacetic acid-washed cell pellets were subjected to mild alkaline hydrolysis. The methanolic-KOH extract, which contains saponifiable fatty acids from glycerolipids, was acidified and extracted with petroleum ether as described previously (15). Extracted fatty acids were dried under nitrogen, and methyl esters were prepared by HCl methanolysis. Sphingolipids were extracted from the saponified cell pellets with an ethanol/water/diethylether/pyridine/NH4OH (15:15:5:1:0.018 by volume) solvent. After drying under nitrogen, sphingolipid fatty acid methyl esters were prepared by HCl methanolysis.

Preparation of Protein Extracts for VLACS Assay

Glucose-grown cells were cultured in 200 ml of YPAD culture to 3 × 107 cells/ml. For oleic acid induced cells, cultures were pregrown in 50 ml of YPAD medium to 3 × 107 cells/ml and washed with YP medium containing 3% glycerol. Cells were resuspended in 200 ml of YP medium containing 3% glycerol, 0.1% oleic acid, and 0.25% Tween 40 at a density of 4 × 106 cell/ml and grown for 18 h to 3 × 107 cells/ml. Formation of spheroplasts, homogenization, and differential centrifugation were performed according to Thieringer et al. (16) with the following modifications.

Cells were washed twice with ice-cold water and incubated in Zymolase buffer (50 mM Tris-Cl, pH 7.5, 10 mM MgCl2,M sorbitol, 30 mM dithiothreitol) for 20 min at room temperature. The cells were then resuspended in Zymolase buffer and incubated with Zymolase (0.1 mg/g of cells (wet weight) for glucose-grown cells and 2 mg/g for oleic acid-induced cells) for 60 min at 30 °C with shaking at 50 rpm.

Spheroplasts were washed by resuspension of centrifuged pellets in 2 volumes of ice-cold Zymolase buffer three times, followed by resuspension in 2 volumes of lysis buffer (5 mM MES (pH 6.0), 0.6 M sorbitol, 1 mM KCl, 0.5 mM EDTA (pH 8.0), 0.1% ethanol (v/v), 1 mM phenylmethylsulfonyl fluoride, 4 mM benzamidine, 5 µg/ml leupeptin, 5 µg/ml pepstatin). Spheroplasts were disrupted at 4 °C in a Potter-Elvehjem homogenizer with 50 strokes. Homogenates were centrifuged at 3,500 rpm (1,500 × g) for 5 min in a Sorvall SS-34 rotor to remove nuclei and cell debris. The pellet was resuspended in lysis buffer, rehomogenized, centrifuged, and combined with the postnuclear supernatant. Aliquots consisting of 20% of the total volume were quickly frozen to -80 °C. The remaining fraction was centrifuged for 15 min at 25,000 × g. The supernatants (25,000 × g supernatant) were frozen and saved. The resulting pellet (25,000 × g pellets) was gently resuspended in lysis buffer and centrifuged at 600 × g for 5 min to remove aggregated material prior to enzyme assay. Previously frozen supernatant fractions were centrifuged at 80,0000 rpm (256,000 × g) for 40 min in a Beckman TL100 microultracentrifuge to fractionate microsomes. Protein concentrations in cell fractions were measured by the Bradford assay method (17).

Very Long Chain Fatty Acyl-CoA Synthetase Assay

Preparation of the fatty acid substrate and assay of VLACS activity followed the method of Wanders et al. (18). The fatty acid substrate [1-14C]lignocerate was prepared as a 100 µM stock solution. Solubilized substrate was obtained by dissolving the dried fatty acid in 100 mM Tris-HCl (pH 8.5) containing 10 mg/ml alpha -cyclodextrin and incubating for 30 min in a sonicating water bath at room temperature.

Reactions were started by adding 20 µl of yeast extract to the assay solution (50 mM Tris-HCl (pH 8.5), 150 µM coenzyme A, 300 µM dithiothreitol, 10 mM ATP, 10 mM MgCl2, 0.01% (v/v) Triton X-100, 10 µM [1-14C]lignocerate dissolved in alpha -cyclodextrin) to the final volume of 200 µl at protein concentrations of 50-100 µg/ml. After 30 min at 37 °C, reactions were terminated by transferring 150 µl of the incubation mixture to a glass tube containing 750 µl of Dole's reagent (isopropyl alcohol, heptane, 1 M H2SO4; 40:10:1 by volume). After vigorous mixing, the protein was removed by centrifugation, and 650 µl of the supernatant was extracted with 350 µl of heptane plus 190 µl of a solution containing 400 nM Mops-NaOH (pH 6.5). The lower (aqueous) layer was washed three times with 400 µl of heptane, and the radioactivity in that fraction was measured by scintillation counting. The radioactivity measured from the boiled protein control extracts was subtracted from the corresponding assay to determine the amount of acyl-CoA product.

Fatty Acid Transport Assays

Cells were pregrown in 40 ml of CM to midlogarithmic phase, collected by centrifugation, and reinoculated at a concentration that would yield 2-3 × 107 ml at the end of the experiment in 40 ml of fresh CMt containing 0.5 or 1.0 mM fatty acids. Following incubation with fatty acids, cells were washed with 1% tergitol solution three times and distilled water three times. Aliquots of cells subjected to this procedure were plated on YPD medium and found to show no significant changes in viability compared with unwashed control cells.

To measure fatty acid incorporation in FAT1-overexpressing cells, strains were grown in CMt (2% raffinose) medium, reinoculated in fresh CMt galactose medium, and grown for 4 h to induce the GAL1 promoter. 1 mM 18:2 was then added to these cultures for 1 h.

To compare uptake over short times in fatty acid synthesis-defective strains, fas2Delta and fat1Delta /fas2Delta cells were grown in CMt supplemented with a mixture of odd numbered fatty acids (13:0, 15:0, and 17:0; 0.2, 0.4, and 0.2 mM, respectively). After 72 h of growth, the native, even chain length, fatty acids are replaced with odd chain species (Table VII), permitting detection of imported even numbered species (18:1, 14:0, and 16:0) that would normally be masked by their endogenous counterparts. Import was monitored by the addition of 0.5 mM even chain fatty acids to the growth medium.

For incorporation assays in faaDelta strains, aliquots of cells grown to late log phase in CMt glucose medium were diluted to 6 × 105 cells/ml in 40 ml of fresh medium and grown to 2 × 107 cells/ml prior to the addition of the test fatty acid. In experiments of more than 30-min duration, the initial cell concentration was modified so that the final culture density would be 2 × 107 cells/ml.

RNA Isolation and Northern Blot Analysis

Total yeast RNA was isolated as described previously (19). Equal amounts (10 µg) of total RNA from each time point of an experiment were analyzed by Northern blots using 1% formaldehyde gels. RNA from the gels was transferred to Zeta ProbeTM membranes (Bio-Rad) using a vacuum blotter (Bio-Rad). Northern blots were quantified using a PhosphorImager (Molecular Dynamics).

Heterologous Expression of the FAT1 Gene in Sf9 Insect Cells

Cloning FAT1 Gene into a Baculovirus Transfer Vector-- A 2.3-kb DNA fragment containing the FAT1 protein coding sequence and 3'-untranslated region was released by BamHI and NotI digestion of vector pCRSK-FAT1 (Table II). The element was inserted into the pVL1393 vector (Pharmingen) that was linearized by BamHI and NotI digestion, placing the FAT1 sequence under control of the baculovirus polyhedrin promoter. Two vectors (pVLFAT1-1 and pVLFAT1-2) derived from independent E. coli transformation were used for transfection.

Generating Recombinant Baculoviruses by Co-transfection-- All reagents, plasmid vectors, and insect cells for baculovirus expression were purchased from Pharmingen, and experiments were done according to the manufacturer's protocols. Uninfected Spodoptera frugiperda (Sf9) cells were maintained by subculturing 1:3 dilutions into 15 ml of TNM-FH medium upon confluency. Cells were incubated at 26-28 °C. For co-transfection, a mixture of 0.5 µg of BaculoGold DNA (consisting of Autographa californica nuclear polyhedrosis virus DNA (Pharmingen, catalog no. 21100D)) and 5 µg of either the recombinant pVLFAT1-1 or -2 vector or the pVL1393 control vector were mixed by gentle vortexing and allowed to sit for 5 min before adding 1 ml of transfection buffer B (25 mM Hepes, pH 7.1, 125 mM CaCl2, 140 mM NaCl). Three drops of healthy Sf9 cells were dropped onto each 60-mm tissue culture plate containing 3 ml of fresh TNM-FH medium. The plates were allowed to sit for 5 min to allow cells to attach. The medium was then aspirated and replaced with 1 ml of transfection buffer A (Grace's medium supplemented with 10% fetal bovine serum). One ml of transfection buffer B/DNA solution was added drop by drop into each of the remaining plates for co-transfection. Plates were incubated at 27 °C for 4 h, washed with 3 ml of fresh TNM-FH medium, and then replenished with 3 ml of fresh TNM-FH medium. The plates were incubated at 27 °C for 4-5 days.

Amplification of Baculovirus Vectors by Serial Infection-- After 5 days of transfection, 0.5 ml of the supernatants were inoculated to the plates containing healthy insect cells growing in 3 ml of fresh TNM-FH medium. Cells infected with viral stocks were grown for 5 days. 0.8 ml of recombinant viral supernatant from the infection plate was transferred to healthy Sf9 cells in 10 ml of TNM-FH medium. If necessary, detached Sf9 cells from the infection plate were cleared from the supernatant by microcentrifugation before transfer.

Preparation of Insect Cellular Extract for the Very Long Chain Acyl-CoA Synthetase Assay-- Cells grown for 48 h after infection were transferred to sterile 15-ml centrifuge tubes and collected by centrifugation at 2,500 × g for 5 min. Cell pellets were resuspended in 1.5 ml of phosphate-buffered saline solution by gentle pipetting and washed three times with the same solution. Cells were resuspended in lysis buffer (5 mM MES (pH 6.0), 0.6 M sorbitol, 1 mM KCl, 0.5 mM EDTA (pH 8.0), 0.1% ethanol (v/v), 1 mM phenylmethylsulfonyl fluoride, 4 mM benzamidine, 5 µg/ml leupeptin, 5 µg/ml pepstatin) and disrupted by three 10-s sonications with 1 min intervals on ice. Cell debris was removed by centrifugation at 1,400 × g for 5 min. The supernatant was assayed for very long chain acyl-CoA synthetase activity using the procedure described for the analysis of yeast membrane fractions.

    RESULTS

The Amino Acid Sequence of the S. cerevisiae FAT1 Gene Is Highly Homologous to Those of Mammalian Very Long Chain Fatty Acyl-CoA Synthetases-- A homology search revealed that FAT1 is related to a rat gene that encodes a very long chain fatty acyl-CoA synthetase. After PCR cloning and DNA sequencing, we determined that the GenBankTM sequences YBR041W that describe the FAT1 sequence contained an error in codon 619. Correction of the sequence yielded an open reading frame encoding a 669-amino acid polypeptide with a significant improvement in homology to the rat protein (Fig. 1). A revised version of the FAT1 gene sequence noting the same error was provided to the sequence data base (GenBankTM number AF065148) by another laboratory while this work was in preparation. The amino acid homology and identity between two proteins are 50 and 33% over 570 amino acids, respectively. By comparison, a human homologue in the data base (GenBankTM number D88308) is 93% identical to the rat protein and 33% identical to Fat1p. In addition to the sequence similarity, Fat1p and the mammalian VLACS share several common characteristics. Both proteins are very close in amino acid length (669 and 620 residues), in theoretical pI (8.54 and 8.82), and in calculated molecular mass (77141 and 70694 daltons). Analysis by the Psort method, which predicts protein localization sites, indicates that Fat1p might be located in the ER membrane, peroxisomes, or plasma membrane. The rat VLACS protein is known to be localized to peroxisomes and microsomes. Both proteins contain an AMP-binding motif that is commonly found in acyl-CoA synthetases, and hydropathy analysis using the Kyte and Doolittle algorithm revealed that the two proteins have four hydrophobic potential membrane-spanning domains located in similar positions.


View larger version (93K):
[in this window]
[in a new window]
 
Fig. 1.   Comparison of the amino acid sequence of yeast Fat1p, human VLACS, and rat VLACS. Shaded and black backgrounds indicate similarities and identities, respectively, among the Fat1p and VLACSs. Protein sequences were aligned with GCG software and shaded by Boxshade 3.2.1 (available on the World Wide Web at ).

Disruption of FAT1 and Measurement of VLACS Activity in Deletion Strains-- Disruption of the FAT1 gene by homologous recombination produced viable haploid cells with normal growth characteristics, indicating that the gene is not essential under normal culture conditions. To verify that Fat1p functions as a very long chain acyl-CoA synthetase, VLACS activity was measured in cell extracts of the fat1Delta and wild type strains using 24:0 as a substrate (Table IV). The specific enzyme activity in the postnuclear extract was 2-fold greater in wild type cells that were grown on glycerol and oleic acid (used to induce peroxisome maturation) than on cells grown with glucose as a carbon source. Total enzyme activity toward the 24:0 substrate was reduced to <FR><NU>1</NU><DE>10</DE></FR> wild type levels in glucose-grown fat1Delta cells and to <FR><NU>1</NU><DE>3</DE></FR> the wild type levels in oleic acid-grown cells.

                              
View this table:
[in this window]
[in a new window]
 
Table IV
Comparison of VLACS activities in FAT1 and fat1Delta cells grown on glucose-containing or glycerol-oleic acid-containing (peroxisomal induction) medium
Cells were grown to 3 × 107 cells/ml, converted to spheroplasts, homogenized, and fractionated as described under "Experimental Procedures." Enzyme activities were assayed using [1-14C]lignoceric acid (24:0). Enzyme activities from samples boiled 10 min were used as controls for each reaction. Each data set represents the means ± S.D. from at least three independent experiments, except for the fat1Delta /fat2Delta strain, which was assayed one time. Each assay was done in triplicate.

Heterologous Expression of the Yeast FAT1 Gene in Sf9 Cells-- To further verify that the FAT1 gene encodes a very long chain acyl-CoA synthetase, the gene was expressed in a heterologous insect cell system. Results presented in Fig. 2 show that, using a 24-carbon fatty acid substrate, the acyl-CoA synthetase activity in Sf9 cells infected with the FAT1-containing baculovirus was about 4-fold greater than the activity in uninfected cells and 5-fold greater in cells infected with wild type virus. Taken together, the data strongly support the hypothesis that the FAT1 gene encodes a very long chain acyl-CoA synthetase enzyme.


View larger version (43K):
[in this window]
[in a new window]
 
Fig. 2.   Very long chain acyl-CoA synthetase activities in insect cells expressing the FAT1 gene. Sf9 cells infected with recombinant virus particles containing the S. cerevisiae FAT1 gene were assayed using [1-14C]lignoceric acid (24:0) as a substrate as described under "Experimental Procedures." Each data set represents the means ± S.D. from three independent experiments. Each assay was done in duplicate. Activity is expressed in terms of pmol 24:0 CoA formed/mg of protein/min. a, uninfected cells; b, wild type virus; c, Fat1-1 virus isolate; d, Fat1-2 virus isolate.

Intracellular Location of Fa1p-- Reports of subcellular fractionation of the yeast Pichia pastoris and mammalian cells has shown that VLACS activities are primarily found in peroxisomal and microsome fractions (5). VLACS activities in S. cerevisiae were found in similar locations (Table IV). In oleic acid-induced cultures, the total activity was distributed between the 25,000 × g organelle pellet (approximately 30%), and its supernatant fractions (approximately 70%). Fractionation of the 25,000 × g supernatant into a microsomal pellet and 250,000 × g supernatant fraction revealed that approximately 25% of that activity was associated with the microsomes and 75% remained in the supernatant fraction. The latter contains high levels of a low density protein fraction that is apparently associated with oil bodies produced under the oleic acid induced culture conditions. No significant low density fraction was seen in the 250,000 × g supernatant fraction of extracts from glucose grown cells. These results indicate that under oleic acid-induced conditions, the Fat1p-dependent VLACS activity is not peroxisome-specific but is distributed in several cytoplasmic and organellar fractions.

Fat1p Directs Green Fluorescent Protein Chimeras to ER and Peroxisome-like Structures-- To further determine the cellular location of Fat1p, the GFP coding sequence was fused to either the C terminus or the N terminus of FAT1 protein coding sequence under the control of the GAL1 promoter. Both types of fusion proteins were functional and restored the wild type very long chain fatty acid composition in galactose-induced fat1Delta cells. In cells grown only on galactose, most of the GFP fluorescence surrounded the 4',6-diamidino-2-phenylindole-stained nucleus (Fig. 3, A and B), indicating that the fusion proteins are mainly localized to ER. In many cells, GFP fluorescence was also associated with one or two small, membrane structures in close proximity to the plasma membrane. These are similar in size and location to microbodies (peroxisomes) previously identified by serial electron microscope sections and immunofluorescence-stained cells of Saccharomyces (20, 21) grown in fatty acid-deficient medium conditions. In cells induced with galactose plus oleic acid, the fluorescence is associated with more numerous and larger intracellular bodies that are more irregularly shaped than those in galactose-grown cells. This is also consistent with EM and fluorescence observations of oleic acid-induced cells, which undergo peroxisomal proliferation and the formation of large oil bodies (20, 21). The location of the fluorescence in cells grown under both conditions was distinct from the 4',6-diamidino-2-phenylindole-stained mitochondria.


View larger version (80K):
[in this window]
[in a new window]
 
Fig. 3.   Localization of Fat1-GFP fusion protein in the fat1Delta ::LEU2 deletion strains. Cells containing the GFP:Fat1p fusion vector, pTS408FAT1, were induced by galactose (A and B) or oleic acid/galactose (C and D) as described under "Experimental Procedures." The figure shows fluorescence (A and C), 4',6-diamidino-2-phenylindole stain (B), and phase contrast views (D) of cells. Similar results were obtained in cells harboring the Fat1p:GFP fusion vector pTS395 (data not shown).

FAA2, Not FAT2, Contributes Most of the Residual VLACS Activity in fat1Delta Cells-- The increased levels of residual VLACS enzyme activity detected in fat1Delta strains grown in oleic acid-containing medium suggest that an additional VLACS enzyme is induced by the fatty acid. Two genes that encode oleic acid-induced peroxisomal AMP-binding proteins were identified that might be the source of the inducible residual activity. PcsP60/FAT2 (21) and FAA2 (22) were previously identified as oleic acid-inducible genes that encode peroxisomal proteins. Although its function is not clearly understood, Fat2p contains an ATP-dependent AMP binding site and shows a high degree of similarity to the E. coli long chain acyl-CoA synthetase. FAA2 has been previously shown to be a fatty acyl-CoA synthetase (9). Faa2p is reported to be a peroxisomal fatty acyl-CoA synthetase that is localized to the matrix side of peroxisomal membrane, where it is involved in the conversion of medium and long chain fatty acids to their CoA derivatives (22).

To examine the roles of Faa2p and Fat2p in VLCFA activation, we constructed strains containing combinations of disrupted FAT1, FAT2, and FAA2 genes. VLACS activity levels in fat1Delta /fat2Delta cells grown on glycerol-oleic acid medium did not differ from that observed in the fat1Delta disruption strain (Table IV), indicating that Fat2p is not the source of the residual activity. VLACS activity in the oleic acid-induced fat1Delta /faa2Delta strain, however, was reduced in the postnuclear supernatant fraction by 80% compared with the fat1Delta strain. Those cells contained less than 5% of the total VLACS activity found in wild type. As expected, due to the peroxisomal matrix location of Faa2p, the loss of residual activity was primarily associated with the organelle pellet fraction (Table IV). These data indicate that Fat1p and Faa2p constitute the major VLACS activities in yeast, with Fat1p being the primary activity. Although previous reports suggest that Faa2p is involved in medium and long chain fatty acid activation (22), these data indicate that it also exhibits activity toward very long chain fatty acid substrates. This is consistent with reports on the rat enzymes, which showed that the long chain acyl-CoA synthetase in rat peroxisome exhibits VLACS activity, although its activity was much stronger (359-fold) toward long chain fatty acid 16:0 than toward 24:0. By contrast, the purified rat peroxisomal VLACS enzyme was only 1.5-fold more active toward the 16:0 substrate than the 24:0 substrate (6).

FAT1 Expression Is Not Induced by Fatty Acids-- The FAA2 gene has been shown to contain oleate-responsive elements in its promoter and to be strongly induced in cells grown on glycerol-oleate medium (22) in a manner similar to that of peroxisomal protein genes that are repressed by glucose, de-repressed by glycerol, and activated by fatty acids (23, 24). Examination of the FAT1 promoter region did not reveal sequences that were homologous to oleate-responsive elements. To see if the FAT1 gene is regulated by similar conditions, quantitative measurements of its mRNA levels were performed by Northern blot analysis using the constitutively expressed ACT1 (actin) gene as an internal standard.

No differences were observed in FAT1 mRNA levels in wild type cells grown in glucose medium with or without fatty acids (data not shown). FAT1 mRNA levels are approximately 2 times higher in glucose grown cells than in glycerol-grown cells that are cultured in fatty acid-deficient medium (Fig. 4a, lanes 1 and 2). The addition of 0.5 mM 16:0 to cells grown in glycerol medium elevates FAT1 mRNA levels 2-fold to the same level as in glucose-grown cells (Fig. 4a, lanes 4, 6, and 8). Identical results were observed when the fatty acid 18:2 was added to the growth medium. Under similar induction conditions, expression of peroxisomal genes is increased 10-250-fold (16, 23, 25). To test the response of FAT1 when cells were dependent on the import of nutrient fatty acids, fatty acid synthetase activity was inhibited by cerulenin (Fig. 4a, lanes 3, 5, 7, and 9). No significant changes in FAT1 expression were found under those conditions. Overexpression of Elo2p, a condition that we found to cause an increase in VLCFAs (Fig. 4a), also did not increase the expression of FAT1 (Fig. 4b). These data indicate that FAT1 does not follow the same pattern of expression seen with peroxisomal proteins that are induced by exogenous fatty acids. This may be linked to a requirement for FAT1p function in nonperoxisomal membrane fractions that are insensitive to fatty acid induction.


View larger version (27K):
[in this window]
[in a new window]
 
Fig. 4.   a, effects of carbon source and fatty acid supplements on FAT1 mRNA levels FAT1 mRNA levels were monitored by Northern blot in wild type cells grown in glucose-containing (lane 1) or glycerol-containing (lanes 2-9) medium with no fatty acids (lanes 1-3) or 0.5 mM 16:0 (lanes 4-9) for the indicated times in minutes. 25 µM of the fatty acid synthetase inhibitor cerulenin was added 30 min prior to the fatty acid where indicated. mRNA levels were quantified by PhosphorImager analysis using the Saccharomyces actin gene as an internal standard. b, effects of ELO2 overexpression on FAT1 mRNA levels. FAT1 mRNA levels were monitored as described in DTY10A (wild type) cells without (lanes 1-3) and with (lanes 4-9) plasmid pGALELO2. Cells were grown on raffinose, and the plasmid-borne FAT1 gene was induced by the addition of galactose for the times (in hours) indicated prior to RNA isolation.

FAT1 Deletion Causes an Accumulation of Intermediate Chain Length Saturated VLCFAs-- A characteristic of a human X-linked adrenoleukodystrophy is a reduction of VLACS activity that leads to the accumulation of saturated VLCFAs. We suspected that FAT1 deletion might cause a similar accumulation of VLCFAs. This was confirmed by quantitative gas chromatography of total fatty acids isolated from wild type and fat1Delta cells grown on minimal glucose medium.

No significant differences were seen between the fatty acid compositions of wild type and fat1Delta cells with respect to the major, C14-C18, long chain fatty acid species. The FAT1 deletion, however, caused striking changes in very long chain fatty acid levels (Table V). Wild type cells usually contain less than 5% very long chain fatty acids. Within that class, 26:0 and hydroxy-26:0 are the most abundant species, and intermediate length (C20-C24) VLCFAs usually comprise less than 10% of the total VLCFAs. Disruption of FAT1 dramatically increased the levels of C22-C26 saturated VLCFAs but did not change in the levels of hydroxy-26:0 species (Table V). In logarithmic phase cells, there was a 2-fold increase in 20:0, a 6-fold increase in 22:0, a 10-fold increase in 24:0, and 2-fold increase 26:0. Accumulation of VLCFAs in fat1Delta strains reached even higher levels as the cells approached stationary phase (data not shown). Under those conditions, 24:0 comprises 3.7% of the total fatty acids compared with 0.13% in wild type, and 26:0 is elevated to 6% from 2.6% in wild type strains.

                              
View this table:
[in this window]
[in a new window]
 
Table V
Effects of FAT1 and ELO3 deletions on VLCFA composition
Wild type, fat1Delta , elo3Delta , and fat1Delta /elo3Delta strains were grown to logarithmic phase on glucose medium and subjected to HCl methanolysis to isolate total fatty acids. Fatty acid methyl ester composition was analyzed by gas chromatography as described under "Experimental Procedures." Data shown represent weight percentage of total fatty acids of the species shown ± S.D. for duplicate samples in three independent experiments.

The abnormal accumulation of intermediate chain length fatty acids was found in both the sphingolipid and phospholipid fractions (Table VI). In sphingolipids of the fat1Delta strain, 22:0, 24:0, hydroxy-24:0, and 26:0 were increased 24-, 53-, 5-, and 2.4-fold, respectively. By contrast, 22:0, 24:0, and 26:0 levels in the phospholipid fraction were increased 5.5-, 6.8-, and 1.6-fold. No increase in hydroxy-VLCFAs was observed in the phospholipid fraction, which is consistent with our previous report that Fah1p, a sphingolipid fatty acid hydroxylase, adds an alpha -hydroxyl group to the fatty acid after it is acylated with sphingosine to form a ceramide (15). A small increase in free fatty acid levels was observed in the fat1Delta strain compared with wild type cells (5.4 versus 3.0%, respectively). The increase was not caused by disproportionate accumulations in very long chain fatty acid species, however, and consisted almost exclusively of increases in C12-C18 fatty acids. To verify that VLCFA accumulations were caused by the absence of Fat1p activity, a plasmid containing a copy of the FAT1 gene under control of the GAL1 promoter was transformed into fat1Delta cells. Normal levels of fatty acids were observed when the plasmid-borne gene was expressed by adding galactose to the growth medium (data not shown).

                              
View this table:
[in this window]
[in a new window]
 
Table VI
Effect of FAT1 deletion on VLCFA composition in sphingolipid (SL) and phospholipid (PL) fractions
Cells were grown to logarithmic phase in glucose medium and subjected to mild alkaline hydrolysis to extract fatty acids from phospholipids. Sphingolipids were then extracted from the cell pellet and subjected to HCl methanolysis to isolate the remaining sphingolipid fatty acids as described under "Experimental Procedures." Fatty acid methyl esters were analyzed from each lipid fraction as described in Table V.

Overexpression of Fatty Acid Elongation Enzymes in a fat1 Strain Further Elevates VLCFA Levels-- We recently identified two essential genes, ELO2 and ELO3, that are components of an ER membrane-bound enzyme system that forms VLCFAs (1). Elo2p appears to be involved in the formation of C20-C24 VLCFAs, and Elo3p is involved in the formation of C20-C26 VLCFAs and is essential for the formation of 26:0. Disruption of either gene causes reduced VLCFA levels, the accumulation of intermediate length VLCFAs, and a concomitant reduction in ceramide synthesis and sphingolipid levels. Given the accumulation of intermediate length fatty acids in fat1Delta strains and the cytoplasmic/microsomal location of a fraction of Fat1p, we hypothesized that Fat1p might act to remove intermediate length VLCFAs that accumulate abnormally during the fatty acid elongation process and thus play a role in maintaining VLCFA levels. To test this interaction, we perturbed VLCFA levels by manipulating Elo2p and Elo3p activities.

Elo2p expression under control of the GAL1 promoter was elevated by induction with galactose for 8 h. In wild type cells, overexpression of Elo2p results in a 6-fold increase of the production of C20-C26 saturated VLCFA species (Fig. 5a). When Elo2p was overexpressed in fat1Delta cells, saturated VLCFA levels were 15-fold greater than in wild type cells. The combined effect of Elo2p overexpression and fat1 deletion was to elevate VLCFA levels to 20% of total cell fatty acids. The elevated production of VLCFAs, however, did not affect the levels of hydroxy-VLCFAs, changing the normal 1:1 ratio of 26:0 to hydroxy-26:0 to about 4.5:1.


View larger version (29K):
[in this window]
[in a new window]
 
Fig. 5.   Effects of overexpression of Elo2p or Elo3p on the VLCFA composition in wild type and fat1Delta ::LEU2 strains. DTY10A (wild type) and fat1Delta strains containing either plasmid pGALELO2 (a) or pGALELO3 (b) were grown in raffinose prior to induction of the relevant ELO gene under control of the GAL1 promoter. The culture was split, and one aliquot was induced with galactose for 8 h before harvesting and extraction of total fatty acids by HCl methanolysis. Data shown are the weight percentage of the most abundant saturated VLCFAs in the total fatty acid fraction for each culture condition.

In wild type cells, overexpression of Elo3p in conjunction with fat1Delta produced a strikingly different pattern of VLCFA accumulation. This resulted in a 2-fold increase in 26:0 with no significant changes in intermediate VLCFAs (Fig. 5b). Overexpression of Elo3p in the fat1Delta strains did not increase the elevated levels of 22:0 and 24:0 but caused a greater than 2-fold further increase in the 26:0 levels elevated by either the disruption of FAT1 or the overexpression of Elo3p.

An elo3Delta /fat1Delta Double Disruption Strain Accumulates 22:0, but Not 24:0 or 26:0, with Severely Retarded Growth-- Oh et al. (1) showed that elo3Delta gene-disrupted strains failed to elongate 24:0 to 26:0. This results in the absence of 26:0 and elevated intermediate VLCFA levels. The most abundant of these are 22:0, hydroxy-16:0, and hydroxy-22:0. Elo3 deletion strains also grow very slowly, possibly due to the lack of normal sphingolipid composition. Simultaneous disruption of ELO3 and FAT1 resulted in more severe retardation of growth (data not shown) and further changes to fatty acid composition that included elevation of 20:0 and 22:0 and the loss of accumulated hydroxy-22:0, hydroxy-24:0 and a greater than 2-fold reduction in hydroxy-16:0 (Table V).

Growth of the fat1Delta /fas2Delta Strains Is Severely Retarded on 18:1-containing Medium but Is Similar to Its Parental FAT1/fas2Delta Strain When Grown on Saturated Fatty Acids---Previous reports indicated that FAT1 plays a role in the transport of nutrient fatty acids (10). Disruption of FAT1 was reported to impair the uptake of long chain fatty acids and reduce the rate of growth on fatty acid-supplemented growth medium when fatty acid synthetase activity was inhibited by the antibiotic cerulenin. Although inactivation of fatty acid synthase causes a cellular requirement for saturated fatty acids, reduced growth rates were most pronounced on cerulenin-treated fat1Delta cells that were grown in oleic acid-containing medium.

We examined this effect further to try to discriminate between fat1Delta -derived fatty acid transport defects and growth defects that might be caused by the failure to metabolize the internalized fatty acids. The requirement for exogenous fatty acids in cerulenin-treated cells does not appear to affect the expression of FAT1. Our tests indicate that FAT1 mRNA levels do not differ between cells grown in glucose (data not shown) or glycerol medium with or without cerulenin and 0.5 mM 16:0 (Fig. 4a). To test the effects of inactivated fatty acid synthetase on transport, we disrupted the FAT1 gene in a previously constructed fas2Delta ::LEU2 strain. FAS2 encodes the beta -subunit of fatty acid synthetase, and its disruption inactivates saturated fatty acid biosynthesis without affecting other enzyme systems that might also be sensitive to cerulenin. The resulting fat1Delta ::HIS3/fas2Delta ::LEU2 disruption strains were isolated on media supplemented with a mixture of 14:0 (0.2 mM), 16:0 (0.4 mM), and 18:0 (0.2 mM).

The fat1Delta /fas2Delta and fas2Delta strains were grown to midlog phase in YPD medium supplemented with a mixture of saturated fatty acids (14:0, 16:0, and 18:0 (0.2, 0.4, and 0.2 mM, respectively)). Washed cells from that culture were used to inoculate, at a density of 105 cells/ml, minimal glucose medium containing no fatty acids, 14:0, 16:0, 18:1, or a mixture of 14:0, 16:0, and 18:0. Growth was monitored by counting cells with a hemocytometer (Fig. 6). Both types of cells failed to grow on medium containing no fatty acids. Active and identical growth patterns were observed for the fas2Delta and fat1Delta /fas2Delta strains on medium containing 16:0 or a mixture of 14:0, 16:0, and 18:0. In the 14:0-supplemented culture, growth of the fat1Delta /fas2Delta strain was slightly slowed compared with that of the FAT1/fas2 strain. By contrast, the growth rate of the fat1Delta /fas2Delta strains in the 18:1-supplemented culture was severely retarded compared with the FAT1/fas2Delta parent (Fig. 6d). Under those conditions, the fat1Delta /fas2Delta cells grew only two generations before growth was arrested.


View larger version (22K):
[in this window]
[in a new window]
 
Fig. 6.   Growth of fas2Delta ::LEU2 (open circles) and fat1Delta ::HIS3/fas2Delta ::LEU2 (black circles) strains on CMt culture medium. Cells were inoculated on CMt medium with supplements as described below. Aliquots were removed at the indicated intervals, and cell density was determined by counting with a hemocytometer. Media supplements were as follows: no fatty acids or a mixture of 0.2 mM 14:0, 0.4 mM 16:0, and 0.2 mM 18:0 (a); 0.5 mM 14:0 (b), 0.5 mM 16:0 (c), or 0.5 mM 18:1 (d).

To examine the incorporation of fatty acids into these strains, the fas2Delta /FAT1 and fas2Delta /fat1Delta strains were grown for 72 h in medium containing the odd chain fatty acids, 13:0, 15:0, and 17:0. As we have previously observed with wild type cells (26), both strains grew on these supplements at a rate similar to that observed with the 14:0/16:0/18:0-supplemented medium. The imported 13-, 15-, and 17-carbon fatty acids are readily elongated to odd chain VLCFAs and converted to monounsaturated acids in similar proportions to those found with endogenous even chain fatty acids in wild type cells. Under those conditions, the odd chain acids replaced virtually all of the even chain species. No significant differences between the fatty acid compositions of the fas2Delta /FAT1 and fas2Delta /fat1Delta strains were observed (Table VII). Furthermore, the odd chain fatty acids 25:0 and hydroxy-25:0 were the most abundant VLCFAs in those strains, indicating that the imported saturated acids can serve as substrates for the fatty acid elongation system. Levels of 25:0 were elevated 9-fold in the fat1Delta /fas2Delta strain compared with the FAT1/fas2Delta , indicating that Fat1p plays a role controlling the levels of VLCFAs derived from imported fatty acids as well as the endogenously derived species.

                              
View this table:
[in this window]
[in a new window]
 
Table VII
Incorporation of odd chain fatty acids in fat1Delta and fat1Delta /fas2Delta strains
The FAT1/fas2Delta and fat1Delta /fas2Delta strains were grown on medium containing a mixture of 13:0 (0.2 mM), 15:0 (0.4 mM), and 17:0 (0.2 mM) for 72 h. Fatty acid methyl esters were prepared from washed cells and analyzed by gas chromatography as described under "Experimental Procedures" and in Table IV.

This fatty acid replacement technique was used to further monitor the transport of 18:1 and related unsaturated fatty acids in the FAT1/fas2Delta and fat1Delta /fas2Delta strains. No significant differences were seen in the uptake of 18:1 or 18:2 over intervals of 10 min (Fig. 7). Similar rates of import between the two strains were also obtained with the even chain saturated species 14:0 and 16:0 (data not shown).


View larger version (21K):
[in this window]
[in a new window]
 
Fig. 7.   Fatty acid import in fas2Delta ::LEU2 and fat1Delta ::HIS3/ fas2Delta ::LEU2 strains. Cells pregrown to 3 × 107 cells/ml on CMt culture were supplemented with a mixture of odd carbon fatty acids (0.2 mM 13:0, 0.4 mM 15:0, and 0.2 mM 17:0) to replace endogenous even chain species. Cells were then collected by centrifugation, washed with CMt twice, and resuspended in fresh CMt. 0.5 mM of oleic acid (A) or linoleic acid (B) was then added to the cultures. At the indicated times, cells were harvested and washed as described above, and the fatty acid composition was analyzed by gas chromatography as described under "Experimental Procedures." The figure shows the level of incorporated fatty acids as weight percentage of total cell fatty acids.

To test for the effects of Fat1p function on long term import, glucose-grown wild type and fat1Delta cells were simply incubated with 18:2, 17:0, 18:1, or 17:1 for 4-6 h prior to analysis (Fig. 8a). No significant differences were observed in the levels of any of the fed fatty acids between the two strains. These tests indicated that the fat1 deletion does not affect the bulk uptake of fatty acids at levels needed to sustain growth. They further indicate that 18:1 and other long chain fatty acids are imported at similar rates to the saturated fatty acids required to sustain growth of the fat1Delta /fas2Delta strain. This suggests that the growth retardation of fat1Delta /fas2Delta strains in 18:1-supplemented medium is caused by the defects in the metabolism of that fatty acid after it is imported.


View larger version (19K):
[in this window]
[in a new window]
 
Fig. 8.   Long term import of fatty acids in glucose-grown wild type (open bars) and fat1Delta cells (solid bars) (a) and galactose-induced wild type, fat1Delta , and pFAT1/fat1Delta strains (b). a, 17:0 (0.5 mM), 17:1 (1 mM), 18:1 (1 mM), or 18:2 (1 mM) were added to the cell cultures. At the indicated times, cells were harvested, washed, and subjected to fatty acid analysis as described under "Experimental Procedures." The percentages of 18:1 shown represent the sum of endogenous and imported fatty acids. The percentages of 17:0 shown are the sum of 17:0 and unsaturated 17:1. b, strains were grown overnight on CMt containing 2% raffinose as a carbon source, harvested, washed, and resuspended in CMt galactose-containing medium for 4 h. 1 mM 18:2 was added to each culture. Cells were harvested after 1 h of incubation and washed as described in the legend to Fig. 7, and fatty acid compositions were analyzed by gas chromatography as described. Data shown in the figure are expressed as the weight percentage of 18:2 incorporated/total cell fatty acids.

As an additional test of the role of Fat1p in fatty acid transport, we examined the import of fatty acids in cells containing a FAT1 gene expressed to high levels under the control of the strong GAL1 promoter (Fig. 8b). Cells were tested for their ability to incorporate 18:2 for 1 h after 4 h of growth on the 2% galactose inducer. In the wild type strain under those conditions, levels of the incorporated 18:2 reach approximately 20% of the total cellular fatty acid mass. There were no significant differences observed in the incorporation of 18:2 between the wild type, fat1Delta , and the fat1Delta strain containing the induced FAT1 gene.

Faa1p and Faa4p Are Required for Bulk Transport of Fatty Acids in Saccharomyces-- A similar but more pronounced response to oleic acid has been previously reported for the two fatty acyl-CoA synthetases, Faa1p and Faa4p. FAA1 and FAA4 are functionally interchangeable genes that account for 99% of the total cellular 14:0-CoA and 16:0-CoA synthetase activities (8) and are responsible for the activation of these imported fatty acids. Simultaneous disruption of the two genes blocks growth and eventually leads to cell death when faa1Delta /faa4Delta cells are grown on cerulenin. We had also previously observed that fatty acid-specific repression of the Saccharomyces OLE1 gene was blocked by simultaneous disruption of the FAA1 and FAA4 genes (11). Previous metabolic radiolabeling studies had suggested that 14:0 and 16:0 were incorporated into the faa1Delta /faa4Delta cells, and the growth defect was attributed to the inability of the cells to activate the imported fatty acids (8).

Given the similarity of this phenotype to the response of fat1Delta -disrupted cells, we examined the ability of faa1Delta /faa4Delta cells to incorporate sufficient quantities of fatty acids to sustain growth in the presence of inactivated fatty acid synthetase activity. Wild type and faa1Delta /faa4Delta gene-disrupted cells were incubated with 1 mM 18:2 for up to 6 h.

Analysis of fatty acid methyl esters derived from the washed cell pellets showed a rapid incorporation of the 18:2 into the wild type cells so that by 6 h the supplemented fatty acid comprised approximately 60% of the total cellular fatty acids (Fig. 9a). Fatty acid import into the faa1Delta /faa4Delta strain was negligible under the same conditions. Similar effects were observed in the transport of saturated fatty acids. Incorporation of 15:0 into wild type cells resulted in its rapid import and either desaturation to 15:1 or elongation to 17:0 and subsequent desaturation of that species to 17:1. The total accumulation of the imported and subsequently modified species in wild type cells over 6 h was greater than 65% of the total cellular fatty acids (Fig. 9b). Under the same condition, negligible incorporation of 15:0 was observed in the faa1Delta /faa4Delta strain. Furthermore, no 15:1, 17:0, or 17:1 was observed in the faa1Delta /faa4Delta cells, indicating that the small amounts of bound 15:0 were not accessible to the microsomal desaturase or elongation systems. The overlapping functions of Faa1p and Faa4p were illustrated by comparison of cells containing single disruptions of each gene with the double disrupted strain (Fig. 9a). Cells containing the single faa4Delta disruption imported fatty acids at slightly higher levels than wild type, whereas faa1Delta cells closely paralleled the wild type incorporation.


View larger version (16K):
[in this window]
[in a new window]
 
Fig. 9.   Import of fatty acids (1 mM 18:2 or 15:0) in cells with deleted fatty acyl-CoA synthetase genes, FAA1 and FAA4. Cells were inoculated in medium containing fatty acid supplements and analyzed for import of fatty acids as described in the legend to Fig. 7 and under "Experimental Procedures." a, cells incubated in medium containing 1 mM 18:2. Open circles, YB332 (wild type); closed squares, YB525 (faa1Delta /faa4Delta ); closed triangles, YB513 (faa1Delta ); inverted closed triangles, YB524 (faa4Delta ). Data shown indicate weight percentage of 18:2 incorporated into washed cells at the times indicated. b, YB332 (wild type) and YB525 (faa1Delta /faa4Delta ) strains incubated in medium containing 1 mM 15:0. When 15:0 is internalized, it can be converted to 15:1 or elongated to 17:0 and then converted to 17:1. Open symbols, wild type; closed circles, faa1Delta /faa4Delta . For wild type cells, the figure shows total incorporated C15 and C17 fatty acids (triangles), 15:0 (squares), and 15:1 (circles). No 15:1 or C17 fatty acids were observed during the course of the experiment in the faa1Delta /faa4Delta strain.


    DISCUSSION

In this report, we show that the yeast FAT1 gene encodes a very long chain fatty acyl-CoA synthetase. Although Fat1p was previously reported to have homology to a presumptive murine fatty acid transport protein (10), further analysis revealed other homologies to members of the acyl-CoA synthetase enzyme family that include an ATP-dependent AMP binding sequence (27) and elements of a recently identified rat very long fatty acyl-CoA synthetase (7). The correction of an error in the FAT1 DNA sequence from the yeast genome data base significantly improved the homology comparison between Fat1p and the mammalian VLACS genes.

A number of fatty acyl-CoA synthetases have been recently identified in yeast. Faa1p and Faa4p, which account for 99% of the total acyl-CoA synthetase activity toward long chain fatty acid substrates, apparently do not catalyze the activation of very long chain fatty acids (9, 28). The role of Fat1p in VLCFA metabolism, however, is readily evident. Disruption of the FAT1 gene causes a severe loss of VLACS activity and the subsequent accumulation of VLCFAs. In glucose-grown cultures, Fat1p accounts for about 90% of the total activity toward C24 fatty acids. In oleic acid-induced cultures, Fat1p is responsible for approximately 70% of the total VLACS activity. Experiments described here indicate that most of the remaining activity toward very long chain substrates in the induced cultures is catalyzed by Faa2p. FAA2 has recently been shown to be an oleic acid-inducible gene that encodes a peroxisomal matrix protein involved in the beta -oxidation of medium chain and long chain fatty acids (22). The ability of Faa2p to form 24:0 CoA in these studies indicates that it has an broader range of substrate specificity than the C6-C20 activity observed when it is expressed as a recombinant protein (28). Faa3p does show activity toward C9-C24 substrates when expressed in E. coli (28), but its activity toward all substrates is very low. It apparently plays a minor, if detectable, role in vivo.

Fractionation studies of Fat1p activity suggest that it is distributed over multiple cellular locations, particularly under growth conditions that induce peroxisomal proliferation. This is consistent with a previous report of VLACS activities in the yeast P. pastoris (5). The intracellular locations of Fat1p in oleic acid-induced and -uninduced cells is further supported by a fusion of green fluorescent protein to elements of Fat1p. In uninduced and induced cells, the most intense fluorescence is found in areas immediately surrounding the nucleus, which is consistent with an ER location. In many uninduced cells, intense fluorescence is also associated with one or two small, spherical regions that are typically located below the plasma membrane. This is consistent with descriptions of microbodies (peroxisomes) found by electron microscopy of serial sections of Saccharomyces grown in fatty acid-depleted medium (20). The same studies revealed that growth under oleic acid-induced conditions produces multiple microbody profiles and the proliferation of lipid bodies. This is consistent with the pattern of fluorescence produced by the Fat1p-GFP chimera in oleic acid-induced cells.

The accumulation of C20-C24 VLCFAs in fat1Delta strains and their further elevation when VLCFA synthesis is perturbed appears to be an important clue to the primary function of Fat1p. Although disruption of FAT1 results in the major loss of very long chain-specific acyl-CoA synthetase activity, it appears unlikely that Fat1p is directly involved in VLCFA biosynthesis. Very long chain fatty acid levels, including the normally abundant C26 species, are actually increased in fat1Delta cells. Taken together, these data suggest that Fat1p primarily plays a catabolic role in VLCFA metabolism, presumably by mobilizing excess very long chain species for degradation via beta -oxidation. This further indicates that other, unidentified, acyl-CoA synthetase activities are employed in the microsomal fatty acid elongation system.

The high levels of Fat1p activity suggest that VLCFA biosynthetic rates may be greater than indicated by steady state levels of cellular very long chain fatty acids. The production of excess acids might, in fact, be a distinctive property of the membrane-bound elongation system. Unlike fatty acid synthetase, which keeps its substrate covalently linked to the multifunctional enzyme complex until the C14-C18 product is released, the VLCFA biosynthetic system elongates fatty acids by a series of reactions that are catalyzed by separate membrane-bound enzymes (29). If transfer of substrates between reaction centers is inefficient, significant amounts of C20-C24 species might be displaced from the system before they were extended to 26:0. This could require high levels of Fat1p as a component of a system that "scavenges" intermediate chain length fatty acids and transfers them to peroxisomes for beta -oxidation. The existence of this system might allow cells to control VLCFA levels primarily through fatty acid degradation to prevent toxic accumulations of VLCFAs. Such accumulations cause lethal conditions such as X-linked adrenoleukodystrophy and similar lipid-mediated disorders. This regulatory mechanism could act on fatty acids prematurely released from the elongation system or in concert with phospholipases that remove excess VLCFAs that have been erroneously acylated into ceramides, triglycerides, or phospholipids. This would also be consistent with the multiple cellular locations observed for Fat1p.

A regulatory role of Fat1p is also consistent with the effects seen in fat1Delta cells when fatty acid elongation is perturbed. Our previous observations (1) indicate that overexpression of ELO2 or ELO3 produces higher levels of VLCFAs than in wild type cells. The large elevations of VLCFAs in fat1Delta cells when ELO genes are either overexpressed or deleted support that hypothesis and suggest that maintenance of optimal VLCFA levels requires a carefully coordinated balance of control between elongation gene expression and Fat1p-mediated VLCFA degradation.

The role that FAT1 plays in fatty acid import is difficult to resolve. The level of homology of Fat1p to the presumptive mouse fatty acid transporter (10) is virtually the same as the level of homology to the mammalian peroxisomal acyl-CoA synthetases. Previous reports indicate that disruption of FAT1 can cause a 2-3-fold decrease in the rate of oleate uptake (10). This was based on carefully done kinetic studies using relatively low levels of radiolabeled fatty acids. The cell fractionation and fluorescence localization studies, however, suggest that the majority of Fat1p is located in intracellular membranes and not concentrated at the cell surface.

We attempted to assess whether Fat1p contributes to the functionally significant import of fatty acids under conditions where cells require them to sustain growth. Under those conditions, we could see no clear differences between fat1Delta and FAT1 strains in the rate of import or the level of incorporation for either saturated or unsaturated long chain fatty acids. We cannot rule out, however, that Fat1p might play some role in importing fatty acids that are present at very low concentrations in the growth medium. The ability of fat1Delta strains to import fatty acids contrasts markedly, however, with the inability of the faa1Delta /faa4Delta strains to transport fatty acids under the same experimental conditions. The loss of import when these two genes are disrupted are consistent with previous reports that fatty acyl-CoA synthetases play a functional role in vectorial transport of fatty acids in E. coli (27, 30) and mammalian cells (31).

Previous reports of fat1Delta and faa1Delta /faa4Delta gene-disrupted strains with inhibited fatty acid synthetase activity indicate that they exhibit similar growth defect phenotypes when they are supplied with 18:1 (8, 10). This could be interpreted as caused by defects either in the transport of 18:1 or in its metabolism after it is internalized. We attempted to replicate these experiments by specifically inactivating fatty acid synthetase through gene disruption. In the case of the faa1Delta /faa4Delta strain, growth inhibition appears to be caused by a simple failure to import sufficient 18:1 (or other saturated and unsaturated fatty acids) to sustain growth. By contrast, the fat1Delta /fas2Delta strain appears to import sufficient saturated and unsaturated fatty acids but is apparently unable to employ the imported 18:1 in some essential metabolic function.

The inability of fas2Delta /fat1Delta strains to metabolize 18:1 suggests that Fat1p has a broad acyl chain length specificity that includes 18-carbon fatty acids and that it plays a role in the metabolism of both long chain and very long chain fatty acids. The homologous rat liver peroxisomal VLACS activity is also reported to be active toward both long (C16) and very long chain (C24) fatty acids (7), indicating that the function of these enzymes may be to modulate a wide range of fatty acid species.

Experiments described in this paper indicate that a primary role Fat1p involves the maintenance of very long chain fatty acid levels by a mechanism that is not clearly understood. One of the most striking effects caused by inactivation of Fat1p activity is the elevation of VLCFA levels under conditions in which very long chain fatty acid biosynthesis rates are perturbed. This suggests that Fat1p-mediated modulation of VLCFA levels is a component of a regulatory system that prevents toxic accumulations of those fatty acids. In humans, loss of control of VLCFA metabolism and accumulation of VLCFAs can be devastating, resulting in progressive degeneration of neural and organ tissue and eventual death. The interactions of the Saccharomyces Fat1p and its VLCFA biosynthetic system may provide a useful model system for understanding complex genetic diseases associated with VLCFA regulation.

    ACKNOWLEDGEMENTS

We are grateful to Dr. Jeffrey Gordon for providing FAA deletion strains and for valuable discussions on fatty acid transport. We also thank Joanne Barbiaz for technical help on fluorescence microscopy and Dr. Chan-Seok Oh for the pGALELO2 and pGALELO3 vectors. We thank Dr. Jon Huibregtse for assistance and advice with the expression of the FAT1 gene in insect cells.

    FOOTNOTES

* This work was supported by National Institutes of Health Grant GM45768.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger To whom correspondence should be addressed: Nelson Biological Laboratories, Bureau of Biological Research, Dept. of Cell Biology and Neuroscience, Rutgers University, Piscataway, NJ 08854-8082. Tel.: 732-445-3972; Fax: 732-445-0644.

    ABBREVIATIONS

The abbreviations used are: VLCFA, very long chain fatty acid; VLACS, very long chain fatty acyl-CoA synthetase; PCR, polymerase chain reaction; kb, kilobase pair(s); ER, endoplasmic reticulum; MES, 4-morpholineethanesulfonic acid; Mops, 4-morpholinepropanesulfonic acid; GFP, Green fluorescent protein; FAT1, gene encoding presumptive VLACS; FAT2, gene encoding a peroxisomal protein with AMP-binding motif; FAA2, gene encoding a peroxisomal fatty acyl-CoA synthetase; FAA1, FAA4 genes encoding fatty acyl-CoA synthetase; FAS2, gene encoding the alpha -subunit of fatty acid synthetase.

2 Fatty acids are denoted by a standard designation that indicates the number of carbons followed by the number of double bonds (e.g. 26:0, hexacosanoic acid, a 26-carbon fatty acid with no double bonds; 18:2, linoleic acid, an 18-carbon fatty acid with two double bonds).

    REFERENCES
Top
Abstract
Introduction
References

  1. Oh, C. S., Toke, D. A., Mandala, S., and Martin, C. E. (1997) J. Biol. Chem. 272, 17376-17384[Abstract/Free Full Text]
  2. Mosser, J., Douar, A. M., Sarde, C. O., Kioschis, P., Feil, R., Moser, H., Poustka, A. M., Mandel, J. L., and Aubourg, P. (1993) Nature 361, 726-730[CrossRef][Medline] [Order article via Infotrieve]
  3. Douar, A. M., Mosser, J., Sarde, C. O., Lopez, J., Mandel, J. L., and Aubourg, P. (1994) Biomed. Pharmacother. 48, 215-218[Medline] [Order article via Infotrieve]
  4. Singh, H., Derwas, N., and Poulos, A. (1987) Arch. Biochem. Biophys. 254, 526-533[CrossRef][Medline] [Order article via Infotrieve]
  5. Kalish, J. E., Chen, C. I., Gould, S. J., and Watkins, P. A. (1995) Biochem. Biophys. Res. Commun. 206, 335-340[CrossRef][Medline] [Order article via Infotrieve]
  6. Uchida, Y., Kondo, N., Orii, T., and Hashimoto, T. (1996) J. Biochem. (Tokyo) 119, 565-571[Abstract]
  7. Uchiyama, A., Aoyama, T., Kamijo, K., Uchida, Y., Kondo, N., Orii, T., and Hashimoto, T. (1996) J. Biol. Chem. 271, 30360-30365[Abstract/Free Full Text]
  8. Knoll, L. J., Johnson, D. R., and Gordon, J. I. (1995) J. Biol. Chem. 270, 10861-10867[Abstract/Free Full Text]
  9. Johnson, D. R., Knoll, L. J., Levin, D. E., and Gordon, J. I. (1994) J. Cell Biol. 127, 751-762[Abstract]
  10. Faergeman, N. J., DiRusso, C. C., Elberger, A., Knudsen, J., and Black, P. N. (1997) J. Biol. Chem. 272, 8531-8538[Abstract/Free Full Text]
  11. Choi, J. Y., Stukey, J., Hwang, S. Y., and Martin, C. E. (1996) J. Biol. Chem. 271, 3581-3589[Abstract/Free Full Text]
  12. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., and Struhl, K. (1993) Current Protocols in Molecular Biology, John Wiley & Sons, Inc., New York
  13. Stukey, J. E., McDonough, V. M., and Martin, C. E. (1989) J. Biol. Chem. 264, 16537-16544[Abstract/Free Full Text]
  14. Pinto, W. J., Wells, G. W., and Lester, R. L. (1992) J. Bacteriol. 174, 2565-2574[Abstract]
  15. Mitchell, A. G., and Martin, C. E. (1997) J. Biol. Chem. 272, 28281-28288[Abstract/Free Full Text]
  16. Thieringer, R., Shio, H., Han, Y. S., Cohen, G., and Lazarow, P. B. (1991) Mol. Cell. Biol. 11, 510-522[Medline] [Order article via Infotrieve]
  17. Bradford, M. M. (1976) Anal. Biochem. 72, 248-254[CrossRef][Medline] [Order article via Infotrieve]
  18. Wanders, R. J., van Roermund, C. W., van Wijland, M. J., Schutgens, R. B., Schram, A. W., van den Bosch, H., and Tager, J. M. (1987) Biochim. Biophys. Acta 919, 21-25[Medline] [Order article via Infotrieve]
  19. Gonzalez, C. I., and Martin, C. E. (1996) J. Biol. Chem. 271, 25801-25809[Abstract/Free Full Text]
  20. Veenhuis, M., Mateblowski, M., Kunau, W. H., and Harder, W. (1987) Yeast 3, 77-84[Medline] [Order article via Infotrieve]
  21. Blobel, F., and Erdmann, R. (1996) Eur. J. Biochem. 240, 468-476[Abstract]
  22. Hettema, E. H., van Roermund, C. W., Distel, B., van den Berg, M., Vilela, C., Rodrigues-Pousada, C., Wanders, R. J., and Tabak, H. F. (1996) EMBO J. 15, 3813-3822[Abstract]
  23. Einerhand, A. W. C., Voorn-Brouwer, T. M., Erdmann, R., Kunau, W.-H., and Tabak, H. F. (1991) Eur. J. Biochem. 200, 113-122[Abstract]
  24. Kos, W., Kal, A. J., van Wilpe, S., and Tabak, H. F. (1995) Biochim. Biophys. Acta 1264, 79-86[Medline] [Order article via Infotrieve]
  25. Swartzman, E. E., Viswanathan, M. N., and Thorner, J. (1996) J. Cell Biol. 132, 549-563[Abstract]
  26. McDonough, V. M., Stukey, J. E., and Martin, C. E. (1992) J. Biol. Chem. 267, 5931-5936[Abstract/Free Full Text]
  27. Black, P. N., Zhang, Q., Weimar, J. D., and DiRusso, C. C. (1997) J. Biol. Chem. 272, 4896-4903[Abstract/Free Full Text]
  28. Knoll, L. J., Johnson, D. R., and Gordon, J. I. (1994) J. Biol. Chem. 269, 16348-16356[Abstract/Free Full Text]
  29. Cinti, D. L., Cook, L., Nagi, M. N., and Suneja, S. K. (1992) Prog. Lipid. Res. 31, 1-51[CrossRef][Medline] [Order article via Infotrieve]
  30. Nunn, W. D., Colburn, R. W., and Black, P. N. (1986) J. Biol. Chem. 261, 167-171[Abstract/Free Full Text]
  31. Schaffer, J. E., and Lodish, H. F. (1994) Cell 79, 427-436[Medline] [Order article via Infotrieve]
  32. Toke, D. A., and Martin, C. E. (1996) J. Biol. Chem. 271, 18413-18422[Abstract/Free Full Text]


Copyright © 1999 by The American Society for Biochemistry and Molecular Biology, Inc.