Nutrient-dependent phosphorylation channels lipid synthesis to regulate PPARα.

Peroxisome proliferator-activated receptor (PPAR)α is a nuclear receptor that coordinates liver metabolism during fasting. Fatty acid synthase (FAS) is an enzyme that stores excess calories as fat during feeding, but it also activates hepatic PPARα by promoting synthesis of an endogenous ligand. Here we show that the mechanism underlying this paradoxical relationship involves the differential regulation of FAS in at least two distinct subcellular pools: cytoplasmic and membrane-associated. In mouse liver and cultured hepatoma cells, the ratio of cytoplasmic to membrane FAS-specific activity was increased with fasting, indicating higher cytoplasmic FAS activity under conditions associated with PPARα activation. This effect was due to a nutrient-dependent and compartment-selective covalent modification of FAS. Cytoplasmic FAS was preferentially phosphorylated during feeding or insulin treatment at Thr-1029 and Thr-1033, which flank a dehydratase domain catalytic residue. Mutating these sites to alanines promoted PPARα target gene expression. Rapamycin-induced inhibition of mammalian/mechanistic target of rapamycin complex 1 (mTORC1), a mediator of the feeding/insulin signal to induce lipogenesis, reduced FAS phosphorylation, increased cytoplasmic FAS enzyme activity, and increased PPARα target gene expression. Rapamycin-mediated induction of the same gene was abrogated with FAS knockdown. These findings suggest that hepatic FAS channels lipid synthesis through specific subcellular compartments that allow differential gene expression based on nutritional status.


FAS solubility
Solubility assays were performed as previously described ( 15 ) with minor modifi cations. Membranes were isolated from mouse liver by ultracentrifugation and resuspended in buffer containing 20 mM HEPES buffer (pH 7.4), 1 mM EDTA, and 255 mM sucrose. The membrane fraction was subjected to treatment with various solvents (1 M NaCl, 0.1 M Na 2 CO 3 at pH 11.5, 1% SDS or 1% Triton X-100) and then centrifuged once more (4°C, 180,000 g , 30 min). The resulting pellets and supernatants were analyzed by western blotting.

Cell culture
Hepa1-6 and Hek293T cells were maintained in DMEM + 10% FBS. Prior to insulin treatment for FAS activity assays, Hepa1-6 cells were cultured in DMEM + 0.5% FBS for 6 h. All insulin treatments were performed in DMEM + 10% FBS.

Pulse-chase study
Confl uent Hepa1-6 cells in 6 cm dishes were incubated in methionine-free media for 30 min. The cells were then pulsed with 500 µCi of 35 S-methionine per dish. After 1 h, cells for the "0" time point were harvested. For subsequent time points, cells were washed with PBS, chased with nonradioactive complete media, and incubated for an additional 45, 90, or 180 min before harvesting. Cells were fractionated into cytoplasm and membrane as described above. FAS was immunoprecipitated from each fraction, samples were subjected to SDS-PAGE, the gel was transferred onto PVDF membrane, and the bands corresponding to labeled FAS were visualized by autoradiography. Autoradiograms were then analyzed by densitometry.

RT-PCR
Total RNA was extracted with TRIzol reagent (Invitrogen) and reverse transcribed using an iScript™ cDNA synthesis kit (Invitrogen). Semiquantitative RT-PCR was performed using SYBR® Green reagent (Applied Biosystems) with an ABI Prism 7700 PCR instrument.

Mutagenesis and plasmid construction
A retroviral plasmid, pBABE-Puro, containing human FAS ( 16 ) generated by Max Loda (Dana Farber) was utilized to generate FAS phosphosite mutants. A 3.4 kb fragment of FAS/pBABE-Puro, including the two putative phosphorylation sites (hFAS S1028 and T1032) and two fl anking BsrGI sites, was amplifi ed by PCR and subcloned into an intermediate Topo vector. Site-directed mutagenesis of the Topo-FAS plasmid changed the codons corresponding to S1028 and T1032 to alanines, yielding two single mutants. The S1028A/T1032A double mutant was made by sequential mutagenesis, using the S1028A mutant as a template. Mutated FAS fragments were then excised and cloned back into pBabe-Puro using the two BsrGI sites to generate mutant, fulllength FAS cDNAs. Mutations as well as correct orientation of the reinserted FAS fragments were verifi ed by DNA sequencing.
Green fl uorescent protein (GFP)-tagged FAS was generated by amplifying the cDNA encoding FAS from pBabe-Puro-FAS by RT-PCR, adding restriction sites for XhoI and EcoRI on the 5 ′ and 3 ′ ends, respectively. The amplifi ed product was cloned into pEGFP-C3 using the XhoI and EcoRI sites, yielding an N-terminal GFP-tagged FAS construct.

Lentiviral shRNA-mediated knockdown and human FAS expression
A plasmid encoding a mouse FAS shRNA (TRCN0000075703) was obtained from Open Biosystems. The packaging vector psPAX2 (12260) and envelope vector pMD2.G (12259) were obtained from Addgene. Hek293T cells at 70% confl uence in a 15 cm dish were transfected using Lipofectamine 2000 with 8 µg we demonstrate that FAS at two separate subcellular locations is differentially regulated by nutrients and insulin, that this regulation involves preferential dehydratase domain phosphorylation for the FAS pool that regulates PPAR ␣ , and that the effects of the kinase mammalian/mechanistic target of rapamycin complex 1 (mTORC1) on PPAR ␣ activity require FAS.

Animals
Male C57BL/6J mice at eight weeks of age were provided ad libitum access to chow diet (Purina #5053) or fasted for 18 h. All mice were kept on Aspen bedding and had free access to water. Protocols were approved by the Washington University Animal Studies Committee.

FAS enzyme activity assay
Using a modifi cation of a previously described assay ( 14 ), 20 µl of sample at 1 µg protein/µl was added to 70 µl of assay buffer [0.14 M potassium phosphate buffer (pH 7.0), 1.4 mM EDTA (pH 8.0), 1.4 mM DTT, 0.24 mM NADPH, 0.1 mM acetyl-CoA]. The rate of NADPH oxidation was monitored at 340 nm at baseline and again after adding 10 µl of 0.85 mg/ml malonyl-CoA (Sigma). The substrate-dependent rate was determined by subtracting the baseline NADPH oxidation rate from the rate after addition of malonyl-CoA. The rate of NADPH oxidation was normalized to FAS protein levels as determined by western blotting and densitometry to determine specifi c activity.

Subcellular fractionation
Perfused liver from C57BL/6J mice was homogenized in 20 mM HEPES buffer (pH 7.4) and centrifuged at 100 g for 30 min, and then the pellet was discarded. The supernatant was centrifuged at 500 g for 60 min; 1,200 g for 20 min; 10,000 g for 20 min; 20,000 g for 30 min; 40,000 g for 30 min; 70,000 g for 30 min; 100,000 g for 60 min; and 179,000 g for 75 min. After each spin, the pellet was washed and resuspended, while the supernatant was centrifuged again. All spins were done at 4°C. To obtain crude membrane and cytoplasmic fractions from mouse liver, freshly isolated perfused liver was homogenized in HEPES buffer and centrifuged at 10,000 g for 45 min at 4°C. The resulting pellet was discarded, and the supernatant centrifuged at 179,000 g for 180 min at 4°C. The supernatant (cytoplasm) and pellet (crude membrane) were collected, and the pellet was washed and resuspended in HEPES buffer. To obtain membrane and cytoplasmic extracts from Hepa1-6 cells, a Subcellular Protein Fractionation Kit for Cultured Cells (78840) from Thermo Fisher Scientifi c was used according to the manufacturer's protocol.
Immobilized metal ion affi nity chromatography (IMAC) was used to enrich the sample for phosphopeptides. The sample was incubated with IMAC beads for 1 h at room temperature. Peptides were eluted from the beads in IMAC buffer, and the sample was diluted with 0.1% FA in 3% CH 3 CN. Samples were then analyzed by LC-MS/MS with a NanoLC-LTQ-Orbitrap mass spectrometer (Thermo Fisher Scientifi c) in data-dependent mode. Acquired spectra were searched against Swiss-Prot database through Mascot server to identify the protein and its posttranslational modifi cations. Nonenriched samples were also run to allow a universal search for protein modifi cations as well as to search for acetyl modifi cations.
To identify FAS modifi cations specifi c to membrane-associated FAS and cytoplasmic FAS, membrane and cytoplasmic fractions were isolated from C57BL/6J mice as described above. FAS was immunoprecipitated from equal amounts of membrane and cytoplasmic lysates (1-10 mg/each) by overnight incubation using a polyclonal rabbit anti-FAS antibody. The samples were then subjected to SDS-PAGE and analyzed as described above.

Statistics
Data are presented as mean ± standard error of the mean. Comparisons between two groups were performed using an unpaired, two-tailed t -test. ANOVA was used for comparisons involving more than two groups.

Hepatic FAS is present in subcellular compartments
FAS synthesizes palmitate, and FAS defi ciency in liver decreases PPAR ␣ target genes. If the effect of FAS deficiency on PPAR ␣ simply refl ects palmitate availability, then exogenous palmitate should rescue the effect. It did not. Treatment of Hepa1-6 cells with 50 µM palmitate failed to rescue expression of the PPAR ␣ target gene ACO following FAS knockdown ( Fig. 1A ). Higher concentrations of palmitate (125-500 µM) were toxic (data not shown).
Since the FAS knockdown effect was not rescued with exogenous palmitate, it is plausible that not only the product of the FAS reaction but also the location of its synthesis mediates downstream effects. Dogma holds that FAS is a cytoplasmic enzyme. To determine whether FAS is also present at other sites, we fractioned mouse liver FAS by ultracentrifugation ( Fig. 1B ). FAS cofractionated with the cytoplasmic marker S6K but also with markers for several organelles. Immunofl uorescent staining for FAS in murine Hepa1-6 liver cells demonstrated colocalization of FAS with endoplasmic reticulum (ER) and Golgi markers psPAX2, 2.25 µg pMD2.G, and 9 µg shRNA. After 48 h, media was collected and fi ltered through 0.45 µm syringe fi lters. Polybrene was added and the media used to treat 50-70% confl uent He-pa1-6 cells. After 24 h, the media was aspirated and replaced with media containing retroviral particles encoding human FAS (see below). Forty-eight hours after addition of the retroviral media, cells were selected with puromycin. After another 48 h, cells were harvested and knockdown of mouse FAS as well as expression of human FAS were assessed.
To generate retroviral particles encoding human FAS, Hek293T cells in 10 cm dishes were transfected using Lipofectamine 2000 with 3 µg FAS plasmid and 3 µg A helper plasmid. After 48 h, media were collected, fi ltered using 0.45 µM syringe fi lters, then polybrene was added, and the media was used to treat 50-70% confl uent Hepa1-6 cells. After 48 h, 2 µg/ml puromycin was added, and after an additional 48 h, cells were harvested.
In experiments assessing PPAR ␣ target gene expression in cells expressing mutant FAS, the endogenous murine FAS of Hepa1-6 cells was knocked down prior to retroviral expression of human FAS as described above.

PPRE-luciferase reporter assay
Media containing lentiviral particles encoding shRNA for murine FAS and media containing retroviral particles encoding wild-type or S1028A/T1032A double-mutant human FAS were prepared as described above. Seventy percent confl uent Hepa1-6 cells in 10 cm dishes were treated with retroviral media for either wild-type or S1028A/T1032A FAS for 24 h, after which the media was aspirated and replaced with lentiviral media. After another 24 h, the media was again aspirated and replaced with fresh media containing puromycin.
After two days of puromycin selection, the media was aspirated, replaced with charcoal-stripped media, and incubated for 1 h. Charcoal-stripped media was also used for subsequent steps. Hepa1-6 cells were transfected with plasmids encoding 3× PPREluciferase and Renilla luciferase by electroporation. The electroporation for each 10 cm dish of cells was done as follows: 5 µg of PPRE-luciferase plasmid and 5 µg of Renilla luciferase plasmid were added to the bottom of a cuvette. Cells were harvested by trypsinization and spun after adding media. The media was aspirated, and cells were washed once with PBS. The PBS was aspirated, and cells were resuspended in 0.5 ml PBS and transferred to the cuvette followed by electroporation at 360 V and 250 µF (time constant of 4.5-5 s Ϫ 1 ). One milliliter of media was added to the cuvette, cells were transferred to a 15 ml tube, and media containing puromycin was added up to 6 ml. Cells were allowed to recover for 10 min, then plated. One day following transfection, cells were harvested by scraping, washed with room-temperature PBS three times, resuspended in PBS, and plated on a 96-well plate. Luminescence from fi refl y luciferase and Renilla luciferase was then measured using the Dual-Glo Luciferase Assay System (Promega) according to the manufacturer's instructions. PPRE-luciferase activity was calculated as the ratio of fi refl y luciferase to Renilla luciferase luminescence.

Mass spectrometry
To identify posttranslational modifi cations in hepatic FAS, perfused C57BL/6J mouse livers were homogenized in lysis buffer containing 1% Triton X-100. The lysate was spun at 10,000 g for 45 min, and the pellet was discarded. FAS was immunoprecipitated from 10 mg of the lysate by overnight incubation using a polyclonal rabbit anti-FAS antibody. IP beads were washed, boiled in sample buffer, and subjected to SDS-PAGE. The gel was stained with Coomassie, the gel segment corresponding to FAS Collectively, these results suggest that the enzyme activities of cytoplasmic and membrane-associated FAS are differentially regulated, a phenomenon that does not appear to be due to intracellular traffi cking of the protein or differences in its primary structure.

Cytoplasmic FAS is preferentially phosphorylated
To address the possibility that differential regulation of cytoplasmic and membrane-associated FAS is caused by a covalent modifi cation, we immunoprecipitated hepatic FAS from fasting and fed mice, and then tested different fractions for the presence of phosphothreonine by western blotting. Cytoplasmic FAS in fed mice was strongly threonine phosphorylated, a modifi cation that was almost undetectable in fasted mice ( Fig. 4A ). Phosphorylation of membrane-associated FAS was low under both conditions ( Fig. 4A ). In Hepa1-6 cells, insulin treatment (a mimic of feeding) stimulated threonine phosphorylation of cytoplasmic but not membrane-associated FAS ( Fig. 4B ).
Analysis of FAS protein from unfractionated mouse liver by mass spectrometry revealed only a single peptide that was threonine phosphorylated. This modifi cation was detected at two residues, Thr-1029 and Thr-1033 (a representative spectrum is shown in Fig. 5A ). When liver FAS was separated into cytoplasmic and membrane-associated fractions and subjected to the same analysis, the phosphorylated peptide was found predominantly in the cytoplasm ( Fig. 5B ) despite similar total amounts of the peptide in both fractions (data not shown). These results suggest that the phosphorylated FAS species detected in the cytoplasm with feeding or insulin ( Fig. 4A, B ) is modifi ed at Thr-1029 and Thr-1033.
These residues are in the dehydratase domain of FAS. The function of this domain requires two catalytic residues, His-878 and Asp-1032, and a third residue, Gln-1036, that maintains the orientation of the catalytic residues ( 21 ). The phosphorylated residues we identifi ed (denoted by * in Fig.  5C ) are in close proximity to the catalytic residue D1032 and the structural residue Q1036 (denoted by # in Fig. 5C ). Sequence alignment of the dehydratase regions from different species revealed that in addition to strict conservation of the active site residues D1032 and Q1036 (denoted by #), the phosphoresidues we identifi ed are also conserved as either serines or threonines in humans, mice, rats, D. melanogaster , and C. elegans (boxes in Fig. 5D ).
Since the evolutionary conservation of these phosphorylation sites suggests involvement in FAS function, we mutated S1028 and T1032 in human FAS (corresponding to the T1029 and T1033 in murine FAS) to alanines, generating two single mutants (S1028A and T1032A) and one double mutant (S1028A/T1032A) ( Fig. 5E , mutated sites but not peroxisomal or mitochondrial markers ( Fig. 1C ). FAS did not appear in the nucleus ( Fig. 1C ).

Membrane-associated and cytoplasmic FAS are differentially regulated
FAS is induced by insulin and nutrients ( 12 ). Surprisingly, the specifi c activity of mouse liver cytoplasmic FAS was not increased in the fed state when insulin levels are high ( Fig. 2A ). Membrane-associated, FAS-specifi c activity was increased with feeding ( Fig. 2B ). The cytoplasmic/ membrane activity ratio in liver was increased with fasting, when PPAR ␣ is activated ( Fig. 2C ). In Hepa1-6 cells, a transformed liver cell line, insulin signifi cantly decreased cytoplasmic FAS activity ( Fig. 2D ), an effect that was not seen in the membrane fraction ( Fig. 2E ). As with mouse liver, the cytoplasmic/membrane activity ratio in Hepa1-6 cells was increased in the absence of added insulin ( Fig. 2F ), a mimic of fasting.
To begin to address the possibility that membrane-associated FAS is an artifact of preparation, we treated isolated fractions with different solvents. Membrane-associated FAS resisted solubilization by 1 M NaCl, remaining in the pelleted fraction, but it was largely solubilized by 0.1 M Na 2 CO 3 ( Fig. 3A ). Treatment with detergent (1% SDS or 1% Triton X-100) solubilized most FAS protein ( Fig. 3A ). These results suggest (17)(18)(19) that FAS manifests a strong peripheral membrane interaction.
A pulse-chase study showed that radiolabeled FAS decreased over time in the membrane-associated and cytoplasmic compartments ( Fig. 3B ), suggesting that there was no ordered fl ux of protein from one compartment to another over the time course of this experiment. There was no discernible change in the distribution of FAS between membrane and cytoplasm when cells were treated with insulin ( Fig. 3C ).
Given the presence of a putative open reading frame (with a potential alternative start codon) 5 ′ to the published fi rst exon of both mouse and human FAS, we considered the possibility that compartmentalized FAS represented differential splicing leading to nonidentical protein isoforms, only one of which is membrane-targeted. However, mass spectrometric analysis of FAS in membrane and cytoplasm failed to detect the predicted alternative amino acids at the N-terminus, and it identified the published FAS protein sequence as being N-terminally acetylated ( Fig. 3D ). This modifi cation, which marks the N-terminus of most eukaryotic proteins ( 20 ), was present in membrane and cytoplasmic fractions of FAS, precluding the existence of an additional N-terminal sequence. All regions of the FAS protein were similarly represented in each fraction, decreasing the possibility that compartment location was determined by altered protein protein in mouse liver by differential centrifugation followed by western blotting. Organelle markers: S6K = P70/S6 kinase (cytoplasmic marker), GM130 = Golgi Matrix protein 130 (Golgi marker), Cav1 = Caveolin1 (caveolae marker), PDI = protein disulfi de isomerase (endoplasmic reticulum marker), Na + /K + ATPase (plasma membrane marker), PMP70 = peroxisomal membrane protein 70 (peroxisomal marker), and COXIV = cytochrome C oxidase IV (mitochondrial marker). (C) Immunofl uorescent staining of FAS and expression of GFPtagged organelle markers in murine Hepa1-6 cells. Nuclei stained with DAPI are presented on the far left, GFP images are presented second from left, FAS images are presented second from right, and merged GFP/FAS images are presented on the far right. compared with wild-type FAS ( Fig. 5I ), suggesting that effects of the FAS mutant on PPAR ␣ target genes are mediated by PPAR ␣ transcriptional activity. One interpretation of these data is that the inability to phosphorylate FAS disinhibits FAS enzyme activity to promote PPAR ␣ transcription.

mTORC1 phosphorylates and inactivates FAS and inhibits PPAR ␣ activity
mTORC1 was recently identifi ed as a physiologically important negative regulator of hepatic PPAR ␣ ( 22 ). mTOR, the kinase component of mTORC1, is a serine/ threonine kinase that preferentially phosphorylates sites with hydrophobic residues at the +1 position ( 23 ). Since the phosphorylated residues we identifi ed have the highly hydrophobic phenylalanine (F1030) and methionine (M1034) at the +1 positions, we addressed a role for are indicated by boxes and the active site residues by #). Wild-type or mutant human FAS was then expressed in Hepa1-6 cells following knockdown of endogenous mouse FAS. Compared with cells expressing wild-type human FAS, cells expressing the S1028A mutation had increased levels of the PPAR ␣ target gene CPT1 ( Fig. 5F ), whereas cells expressing the T1032A mutation did not show changes in PPAR ␣ target genes ( Fig. 5G ). However, expression of the double-mutant S1028A/T1032A was associated with increased levels of both ACO and CPT1 ( Fig. 5H ). To implicate PPAR transcriptional activity in this effect, we performed a PPRE-luciferase reporter assay. After expression of wild-type or S1028A/T1032A double-mutant FAS and knockdown of endogenous mouse FAS, cells were transfected with a plasmid encoding three tandem PPREs fused to a fi refl y luciferase reporter gene. Luciferase activity was increased in cells expressing the S1028A/T1032A double-mutant FAS

DISCUSSION
FAS synthesizes lipid for energy storage and participates in the generation of a lipid ligand involved in the activation of fatty acid oxidation. Energy storage occurs with feeding, and activation of fatty acid oxidation occurs with fasting. To clarify how the same enzyme mediates both processes, we pursued the possibility that distinct pools of FAS are differentially regulated in the liver.
We found FAS in the cytosol, but we also localized FAS to organelles ( Fig. 1 ) through a strong peripheral membrane interaction ( Fig. 3A ). FAS-specifi c activity was relatively higher with feeding/insulin in membranes and relatively higher with fasting in the cytosol ( Fig. 2 ). This effect did not appear to involve movement of FAS between compartments or primary sequence differences between these pools of FAS. Instead, this activity difference was associated with preferential phosphorylation of cytoplasmic (but not membrane) FAS with feeding ( Fig. 4 ) at conserved sites within a catalytic domain ( Fig. 5 ). Mutation of these sites increased endogenous PPAR ␣ target gene expression as well as activity of a PPRE-dependent reporter gene ( Fig. 5 ), consistent with disinhibition of FAS in the mTORC1 in FAS phosphorylation. Treating Hepa1-6 cells with the mTORC1 inhibitor rapamycin for 30 min abolished the insulin-induced increase in cytoplasmic FAS threonine phosphorylation ( Fig. 6A ) and was associated with an increase in cytoplasmic FAS-specific activity ( Fig. 6B ). Treatment of these cells with Torin 1 at 250 nM also abolished insulin-induced FAS phosphorylation (data not shown). Treating Hepa1-6 cells with rapamycin for 24 h (a suffi cient time to reach a new steady state for mRNA levels) increased expression of the PPAR ␣ target gene CPT1 ( Fig. 6C ). These fi ndings confi rm those made in a different system ( 22 ) and extend that work by implicating FAS in the mTORC1-PPAR ␣ axis.
To better defi ne the interaction between mTORC1, FAS, and PPAR ␣ , FAS was knocked down in Hepa1-6 cells followed by rapamycin treatment. FAS knockdown, confi rmed in the presence of rapamycin ( Fig. 6D ), decreased CPT1 expression ( Fig. 6E ). The induction of CPT1 levels with rapamycin occurring with FAS expression ( Fig. 6C ) was lost with FAS knockdown ( Fig. 6E , solid bar). These results suggest that in this cell line under these conditions, the induction of the PPAR ␣ target gene CPT1 caused by inhibition of mTORC1 is FAS-dependent. birds. Using pigeon liver as a model and exclusively studying FAS in the cytoplasm, Qureshi and colleagues found that feeding induced 32 P incorporation into FAS, which was associated with a loss of enzyme activity ( 24 ). In vitro treatment with phosphatases dephosphorylated FAS and restored enzyme activity. The authors of this study did not identify a physiological role for this covalent modifi cation, and it is not known whether the phosphosites we found are conserved in pigeon FAS due to the unavailability of sequence data for this species. Regardless, our work suggests that the phosphorylation state of cytoplasmic FAS may channel lipid fl ow to impact phospholipids inducing gene expression in the nucleus. absence of phosphorylation. Inhibition of mTORC1 with rapamycin decreased FAS phosphorylation, increased cytosolic FAS enzyme activity, and increased expression of the PPAR ␣ target gene CPT1, an effect that was FAS-dependent ( Fig. 6 ). One interpretation of these fi ndings is that hepatic FAS exists in at least two differentially regulated subcellular pools, cytoplasmic and membrane-associated ( Fig. 7 ). Cytoplasmic FAS is phosphorylated with feeding to limit PPAR ␣ activation, and it is dephosphorylated with fasting to promote PPAR ␣ activation.
Our fi ndings provide molecular defi nition and physiological context to an observation made nearly four decades ago in (F) RT-PCR analyses of PPAR ␣ target gene expression in Hepa1-6 cells expressing wild-type or S1028A mutant FAS. Endogenous FAS was knocked down using lentiviral shRNA for murine FAS. Wild-type or mutant human FAS was expressed using retroviruses. Data are averages of three independent experiments . * P р 0.05. (G) RT-PCR analyses of PPAR ␣ target gene expression in Hepa1-6 cells expressing wild-type or T1032A mutant FAS. Assay performed as in (F). Data are averages of three independent experiments. (H) RT-PCR analyses of PPAR ␣ target gene expression in Hepa1-6 cells expressing wild-type or S1028A/T1032A mutant FAS. Assay performed as in (F). Data are averages of three independent experiments. * P р 0.05. (I) PPRE-luciferase activity in Hepa1-6 cells expressing wild-type or S1028A/T1032A mutant FAS. Wild-type or mutant human FAS was expressed using retroviruses. Endogenous FAS was knocked down using lentiviral shRNA for murine FAS. Cells were cotransfected with plasmids encoding 3× PPRE-fi refl y luciferase and Renilla luciferase. PPRE-luciferase activity is reported as the ratio of fi refl y/ Renilla luciferase luminescence. N = 3-6/group. *** P р 0.0005. preferential phosphorylation depending on cellular location and nutritional state.
There is precedent for compartmentalization in metabolism. Exogenous administration of T3, the active form of thyroid hormone that can be produced locally from its precursor T4, does not rescue gene expression defects in the setting of hypothyroidism. But administration of T4, which is metabolized to generate T3 locally, restores downstream effects ( 32 ). There is also precedent for compartmentalization in lipid signaling. Phosphatidic acid derived from glycerolipid synthesis has effects on mTORC2 that are opposite from those induced by phosphatidic acid derived from membrane lipolysis ( 33 ). These observations are consistent with our model ( Fig. 7 ). In the fed state, cytoplasmic FAS is phosphorylated to limit lipid production resulting in PPAR ␣ activation, while membrane FAS, less susceptible to phosphorylation, likely produces lipids for energy storage or export. Given the rapid demands of lipid synthesis prompted by transition from the fasting to the fed state, the induction of membrane FAS may be predominantly substrate-driven through allosteric Physiological, mass spectrometric, and crystal structure data indicate that phospholipids interact with nuclear receptors ( 6,(25)(26)(27)(28)(29). FAS appears to be linked to PPAR ␣ through phosphatidylcholine synthesis mediated by the Kennedy pathway ( 6 ). Viewed with previous studies showing that phosphorylation regulates the CDP-choline branch of the Kennedy pathway ( 30,31 ), our identification of functionally relevant FAS phosphorylation sites raises the possibility that phosphorylation at several nodes within a cascade of lipid signaling from the cytoplasm to the nucleus coordinates FAS-mediated PPAR ␣ activation.
Palmitate is the direct product of the FAS reaction. If the mere availability of palmitate were required to activate PPAR ␣ , exogenous palmitate would correct FAS deficiency. However, the addition of palmitate to liver cells with FAS defi ciency does not restore defects in PPAR ␣dependent genes ( Fig. 1 ), and elevated serum palmitate levels that accompany inactivation of liver FAS in mice does not rescue impaired activation of PPAR ␣ -dependent genes ( 8 ). Thus, palmitate produced by FAS appears to be compartmentalized, a notion supported by our fi nding of   7. Schematic depiction of insulin/feeding-regulated FAS phosphorylation and FAS-mediated PPAR ␣ activation. In the fed state, mTORC1 promotes phosphorylation of FAS, thus limiting downstream generation of a phosphatidylcholine ligand that activates PPAR ␣ -dependent gene expression. In the fasting state, dephosphorylated FAS in the cytoplasm is permissive for the generation of the ligand activating PPAR ␣ -dependent gene expression. PC, phosphatidylcholine; TAG, triacylglycerol.