From the University of Cambridge, Departments of Clinical Biochemistry and Medicine, Addenbrooke's Hospital, Cambridge, United Kingdom CB2 2QR
Received for publication, October 27, 2000, and in revised form, December 18, 2000
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Exposure of insulin-sensitive tissues to free
fatty acids can impair glucose disposal through inhibition of
carbohydrate oxidation and glucose transport. However, certain fatty
acids and their derivatives can also act as endogenous ligands for
peroxisome proliferator-activated receptor A full understanding of the control of glucose homeostasis and its
dysregulation in diabetes mellitus will require a better comprehension
of the integration of carbohydrate and lipid metabolism (1). Free fatty
acids are considered to play a pivotal role in the pathogenesis of
diabetes and may be involved in the early events leading to insulin
resistance in man (2). The classic studies of Randle and colleagues (3)
demonstrated that oxidation of fatty acids could inhibit glucose
oxidation through the effects of acetyl-CoA on pyruvate dehydrogenase
activity. More recently it has been shown that increased delivery of
free fatty acids in vivo can impair insulin-mediated glucose
disposal through inhibition of glucose uptake (4-7). Furthermore,
several long chain fatty acids, both saturated and unsaturated, have
been demonstrated to decrease mRNA levels of the insulin-responsive
glucose transporter GLUT4 in vitro (8).
Insulin sensitivity in whole animals can also be influenced by the
manipulation of the dietary fatty acid profile. A diet high in
saturated, monounsaturated (n-9) or polyunsaturated
(n-6) fatty acids
(PUFA)1 leads to insulin
resistance in rodents (9, 10). Substituting a small percentage of fatty
acids in the n-6 PUFA diet with n-3 PUFA
normalizes insulin action (9, 10). Similar dietary intervention studies
in humans have been inconclusive (11). However, in both humans and
rodents, PUFA (particularly n-3) levels in skeletal muscle
membrane phospholipids are positively correlated with insulin sensitivity (10, 12, 13).
In addition to their roles as metabolic fuel, certain fatty acids can
act as precursors for signaling molecules which may influence insulin
action and glucose metabolism. Interest in these activities has been
greatly enhanced by the recent recognition of the importance of the
nuclear hormone receptor PPAR Grunfeld et al. (30) originally reported that exposure of
3T3-L1 adipocytes to certain unsaturated fatty acids during
differentiation could enhance basal glucose uptake. These workers
suggested that relatively nonspecific effects on membrane fluidity
might underlie these effects. More recently, a similar effect on basal
glucose uptake has been observed in fully differentiated 3T3-L1
adipocytes supplemented with the n-6 PUFA, arachidonic acid
(AA) (31, 32). In light of recent advances in the understanding of the
possible roles of lipid mediators in the control of glucose homeostasis we have re-examined the effects of a range of fatty acids on glucose uptake in 3T3-L1 adipocytes. Having established that AA had the most
marked effects on enhancement of glucose disposal we have attempted to
dissect the molecular mechanisms underlying these effects.
Materials--
2-Deoxy-D-[2,6-3H]glucose,
[1-14C]AA, and the enhanced chemiluminescence (ECL) kit
were purchased from Amersham Pharmacia Biotech. Analytical grade
solvents were obtained from BDH Biochemicals (Dorset, United Kingdom).
Rabbit anti-GLUT4 antibody was a gift from Prof. G. Gould (University
of Glasgow), rabbit anti-GLUT1 was a gift from Prof. S. Baldwin
(University of Leeds), and rabbit anti-PPAR Tissue Culture--
3T3-L1 fibroblasts (ATCC) were maintained at
no higher than 70% confluency in DMEM containing 10% newborn calf
serum, 25 mM glucose, 2 mM glutamine, and
antibiotics (DMEM/NBCS). For differentiation they were grown 2 days
post-confluence in DMEM/NBCS and then for 2 days in medium containing
fetal bovine serum instead of newborn calf serum (DMEM/FBS)
supplemented with 0.83 µM insulin, 0.25 µM
dexamethasone, and 0.5 mM isobutylmethylxanthine. The
medium was then changed to DMEM/FBS supplemented only with 0.83 µM insulin for 2 days and then to DMEM/FBS alone for a
further 3-5 days. Differentiated cells were only used when at least
95% of the cells showed an adipocyte phenotype by accumulation of
lipid droplets.
Fatty Acid Solutions--
Fatty acid-supplemented media were
prepared as described previously (30). In short, a 0.2 M
stock solution of the fatty acid in ethanol was diluted 1:25 into 20%
fatty acid-free bovine serum albumin in Dulbecco's
phosphate-buffered saline (PBS) at 60 °C with shaking. This solution
was diluted 1:10 into serum-free DMEM (containing 25 mM
glucose and 2 mM glutamine) for studies with up to 4-h
incubation times, or into DMEM/FBS for studies with >4-h incubation
times. This gives a final concentration of 8 × 10 Glucose Uptake--
Adipocytes (at day 9 after initiation of
differentiation) in 6- (or 24-) well plates were incubated for 4 h
in fatty acid-supplemented DMEM. In the time course experiment,
incubations of 48 and 24 h were commenced on days 7 and 8 of
differentiation, respectively, and on day 9 cells were serum-starved in
DMEM (containing 25 mM glucose and 2 mM
glutamine) for 2 h prior to the assay. For the 1-h time point,
cells were starved for 1 h prior to incubation in the AA medium.
Glucose uptake assays were performed as described previously (33).
Western Blotting--
Treated cells (adipocytes/pre-adipocytes)
were solubilized by scraping and passing 10 times through a 25-gauge
needle in lysis buffer (50 mM Hepes, 150 mM
NaCl, 1 mM EDTA, 30 mM NaF, 1% Triton X-100, 1 mM Na3VO4, 10 mM
Na2P2O7, 2.5 mM
benzamidine, 1 µg/ml antipain, 1 µg/ml leupeptin, 1 µg/ml
pepstatin A). The lysate was clarified by centrifugation at 13,500 × g for 10 min at 4 °C. Crude cell extracts were
resolved by SDS-PAGE before electroblotting to polyvinylidene
difluoride membranes (Millipore). Membranes were blocked in 1% bovine
serum albumin and specific proteins were detected by incubation with
appropriate primary and secondary (horseradish peroxidase-conjugated)
antibodies in TBST (150 mM NaCl, 50 mM Tris,
0.1% Tween 20). Proteins were visualized using an ECL kit.
Plasma Membrane Lawn Assay--
3T3-L1 adipocytes (day 9), grown
on collagen-coated glass coverslips, were treated for 4 h with AA
medium. Cells were incubated ± 10 nM insulin for 30 min and a modified version of the plasma membrane lawn assay (33) was
performed. Cells were washed twice in ice-cold buffer A (50 mM Hepes, 10 mM NaCl, pH 7.2), twice in
ice-cold buffer B (20 mM Hepes, 10 mM KCl, 2 mM CaCl2, 1 mM MgCl2,
pH 7.2), and sonicated using a probe sonicator (Kontes, Vineland, NJ)
to generate a lawn of plasma membrane fragments attached to the
coverslip. The membranes were washed twice again in ice-cold buffer B
and fixed to the coverslips for 15 min using freshly prepared 3%
paraformaldehyde. Membranes were then serially washed: 3 times in PBS,
3 times in 50 mM NH4Cl in PBS over 10 min, 3 times in PBS, 3 times in PBS-gelatin (PBS containing 0.2% gelatin and
1 µl/ml goat serum) over 5 min, and finally 3 times in PBS. Membranes
were incubated in either anti-GLUT4 or anti-GLUT1 antibody (1:100
dilution in PBS-gelatin) for 1 h at room temperature. After
washing 3 times in PBS-gelatin and 3 times in PBS, the coverslips were
incubated with the secondary antibody, fluorescein
isothiocyanate-conjugated donkey anti-rabbit IgG for 1 h at room
temperature, washed 3 times in PBS-gelatin, and 3 times in PBS and
mounted on glass slides. Coverslips were viewed using a ×60 objective
lens on a Nikon Optiphot-2/Bio-Rad MRC-1000 microscope operated in
laser scanning confocal mode. Samples were illuminated at 488 nm, and
images were collected at 510 nm. Duplicate coverslips were prepared at
each experimental condition and eight random images of plasma membrane
lawn were collected from each. The images were quantified using Bio-Rad MRC-1000 confocal microscope operating software (CoMOS, version 6.05.8), on an AST premmia SE P/60 personal computer.
[1-14C]Arachidonic Acid Uptake--
0.2
M AA, containing a trace of [1-14C]AA, was
diluted into 20% bovine serum albumin and serum-free DMEM to a final
concentration of 8 × 10 Membrane Preparation, Phospholipid Extraction, and Fatty Acid
Analysis--
3T3-L1 adipocytes (day 9) in 3 × T175 tissue
culture flasks were treated with 8 × 10 Membrane Fluidity Assay--
3T3-L1 adipocytes (day 9) in 2 × 6-well plates were treated with 8 × 10 Inhibitor Studies--
Fresh stock solutions of inhibitors were
prepared in absolute ethanol (ibuprofen, nordihydroguaiaretic acid, and
cycloheximide) or dimethyl sulfoxide (piroxicam and 6-MNA). 3T3-L1
adipocytes (day 9) in 6-well plates were given 1.8 ml of serum-free
DMEM and 20 µl of inhibitor solution for 30 min. 0.2 M AA
in ethanol was diluted 1:25 into 20% fatty acid-free bovine serum
albumin in PBS as before and 200 µl of this solution was added per
well for 4 h. This gives a final 1:100 dilution of the original
inhibitor stock solution. The glucose uptake assay was performed as above.
Enzyme Immunoassay for PGE2--
Cells were treated
±AA/inhibitor for 4 h. 1-ml aliquots of medium were taken
immediately prior to the glucose uptake assay and analyzed for
PGE2 content using an enzyme immunoassay kit for
PGE2 (Cayman Chemical) based on the enzyme-linked
immunosorbent assay method.
Adenovirus Expression--
Recombinant adenoviruses were
generated as described previously (36), expressing GFP (Ad-GFP) or GFP
and full-length L468A/E471A human PPAR RNA Extraction/Quantitative Reverse
Transcriptase-PCR--
Virally infected pre-adipocytes/day 2 adipocytes were scraped and total RNA was extracted using the RNeasy
mini kit from Qiagen. Adipocyte P2 (aP2) gene expression was quantified
using real time quantitative reverse transcriptase-PCR. Briefly,
cDNA was prepared from 100 ng of RNA using 200 units of Moloney
murine leukemia virus reverse transcriptase (Promega). Real time
quantitative PCR was performed using an ABI-PRISM 7700 Sequence
Detection System instrument and software (PE Applied Biosystems, Inc.,
Foster City, CA) as described (37). The primers and probes for
aP2 were: forward, CACCGCAGACGACAGGAAG; reverse, GCACCTGCACCAGGG;
probe, TGAAGAGCATCAAACCCTAGATGGCGG (all 5'-3'). Results were
normalized to the endogenous control, glyceraldehyde-3-phosphate dehydrogenase.
Statistical Analysis--
Data are presented as mean ± S.E. Statistical significance of treatments was determined using the
paired Student's t test (*, p < 0.05; **,
p < 0.01; ***, p < 0.001).
Effect of Fatty Acids on Glucose Uptake in 3T3-L1
Adipocytes--
Fully differentiated 3T3-L1 adipocytes were incubated
with 8 × 10 Effect of Arachidonic Acid on the Expression and Cellular
Localization of Glucose Transporters--
Arachidonic acid had no
effect on total cellular levels of GLUT1 and GLUT4 as assessed by
Western blotting (Fig. 2A). To
examine whether AA might have an effect on levels of GLUT1 or GLUT4 at the plasma membrane the plasma membrane lawn assay was used. Insulin treatment of 3T3-L1 cells produced an ~2-fold increase in GLUT1 and a
5-fold increase in GLUT4 plasma membrane expression, results which are
consistent with previous published studies (33). AA increased levels of
GLUT1 at the plasma membrane in the absence and presence of insulin by
2- (p < 0.05) and 1.3-fold (p < 0.001), respectively, and increased levels of GLUT4 at the plasma
membrane in the absence and presence of insulin by 1.5-fold
(p < 0.01) and 1.4-fold (p < 0.05),
respectively (Fig. 2B).
Effect of Arachidonic Acid on Membrane Phospholipid Composition and
Fluidity--
The rapidity of the effects of AA on glucose uptake
raised the question of whether the uptake of AA into the cell and its incorporation into membrane phospholipid could be accomplished significantly within 4 h. 3T3-L1 adipocytes were incubated with [1-14C]AA. Within 4 h, the cells had taken up
approximately half of the labeled AA (Fig.
3A). 4 h incubation in
8 × 10 Effects of the Inhibition of Protein Synthesis--
The protein
synthesis inhibitor, cycloheximide, has previously been shown to
influence glucose uptake in 3T3-L1 adipocytes (38, 39) and it was
therefore important to choose a concentration which had no effect on
glucose uptake in the absence of AA. While 2.5 µM
cycloheximide had no effect on glucose uptake in control cells (Fig.
4A), it completely inhibited
the AA potentiation of basal glucose uptake (p < 0.001) and partially inhibited the potentiation of insulin-stimulated
glucose uptake (p < 0.05) (Fig. 4B). Thus, despite the fact that the effects of AA on glucose uptake are rapid (4 h), these appear to be highly dependent on de novo protein synthesis.
Effect of Inhibitors of Cyclooxygenase and Lipoxygenase on the
Arachidonic Acid Potentiation of Glucose Uptake--
Arachidonic acid
is rapidly converted to a number of eicosanoids. To identify
involvement of either cyclooxygenase (COX) or lipoxygenase (LOX) in the
AA potentiation of glucose uptake, inhibitors of the two enzyme systems
were used. Cyclooxygenase, or prostaglandin H synthase, exists as two
isozymes, COX1, which is constitutively expressed, and COX2, which is
inducible (40). Ibuprofen inhibits both isozymes nonselectively with an
IC50 of 10 µM (41). Piroxicam and 6-MNA
selectively inhibit COX1 and COX2 with IC50 values of 18 and 15-55 µM, respectively (42, 41). None of these COX inhibitors significantly inhibited either the AA potentiation of basal
or insulin-stimulated glucose uptake (Fig.
5A). To confirm that the COX
enzyme activity had indeed been inhibited in these experiments, samples
of media were taken prior to the studies of glucose uptake and tested
for the presence of prostaglandin E2, a major metabolite of
AA in the adipocyte (43). All three compounds significantly inhibited
the production of PGE2 from the AA-supplemented cells (Fig.
5B).
Nordihydroguaiaretic acid (NDGA) is a selective inhibitor of the
lipoxygenases (IC50 = 0.2 µM for 5-LOX and 30 µM for 12- and 15-LOX (44)). NDGA significantly inhibited
the AA potentiation of basal glucose uptake at both 30 µM
(p < 0.05) and 60 µM (p < 0.01) and showed a similar trend to inhibit the potentiation of
insulin-stimulated uptake (p = 0.08) (Fig.
5A).
Effect of Dominant-negative PPAR
To investigate the effect of the PPAR A role for fatty acids in the modulation of insulin sensitivity
in vivo and in vitro is well established (11),
although the precise nature of the mechanisms underlying these effects is unknown. The recently established fact that certain fatty acids and
their metabolites can act as endogenous ligands for the nuclear hormone
receptor PPAR The systematic study of the effects of various fatty acids on glucose
uptake in adipocytes was first reported by Grunfeld et al.
(30). These workers showed that incubation of 3T3-L1 cells, during the
process of differentiation, with a range of saturated and
monounsaturated fatty acids increased basal glucose uptake. Saturated
fatty acids decreased insulin-stimulated uptake, whereas
monounsaturated fatty acids had no effect. Our studies investigated the
effect of 10 mainly polyunsaturated, fatty acids on glucose uptake in
fully differentiated 3T3-L1 adipocytes. Differentiated cells were used
to eliminate confounding effects of fatty acids on the adipogenic
process itself. In agreement with Grunfeld et al. (30) there
appeared to be a relatively nonspecific effect of all fatty acids to
increase basal glucose uptake. However, all fatty acids studied, except
the saturated palmitic acid (16:0), increased insulin-stimulated
glucose uptake and this effect did not appear to be specific to fatty
acid series. The most marked effects on glucose uptake were observed
with AA, which increased basal and insulin-stimulated glucose uptake at
all time points studied. Two previous studies have shown a similar
effect of AA on basal glucose uptake in 3T3-L1 adipocytes. Fong
et al. (32) observed an increased basal glucose uptake with
0.2 mM AA at all time points from 1 to 8 h (32).
Similarly, Tebbey et al. (31) reported such an effect with
50 µM AA following 24 and 72 h incubation. Neither
study, however, observed a significant effect of AA on insulin-stimulated glucose uptake. Consistent with the observations of
Fong et al. (32) who demonstrated that the potentiating
effects of 8 h exposure to AA on glucose uptake were partially
inhibited by cycloheximide, we found that the effects of 4 h of AA
were also sensitive to the inhibition of protein synthesis suggesting the involvement of a rapidly synthesized protein intermediate in the
mediation of this response.
Having established the effect of AA on glucose uptake, we examined
whether this was mediated through increased expression or altered
cellular location of glucose transporters. Previous studies have
demonstrated a decrease in total cellular and plasma membrane GLUT4 in
3T3-L1 adipocytes supplemented with AA for 24 and 48 h (31, 8).
Conversely, 3T3-L1 adipocytes supplemented with AA for 8 and 48 h
exhibited increased total cellular and plasma membrane levels of GLUT1
(32, 31). This effect was not, however, observed with 2 h AA
incubation (32). In our experiments, 4 h AA incubation had no
effect on total cellular levels of GLUT1 or GLUT4. However, a highly
significant increase in the plasma membrane levels of both
transporters, in the absence and presence of insulin was observed. Thus
AA appears to enhance either the translocation of both major adipocyte
glucose transporters to the plasma membrane or reduce their rate of internalization.
It has been suggested that fatty acids may have nonspecific effects on
cellular membranes which could secondarily alter the intrinsic activity
of transmembrane glucose transporters (12, 30). In particular,
alterations in the fatty acid composition of membrane phospholipids can
affect membrane fluidity (45). It was unclear, however, whether as
short an exposure as 4 h would be sufficient to lead to such
changes in plasma membrane composition. We observed that, within 4 h, at least half of the available AA had entered the cells and that
there was a highly significant increase in the AA content of membrane
phospholipids. However, this did not have any discernible effect on
membrane fluidity. Therefore, although we have not excluded an effect
of AA on the intrinsic activity of membrane glucose transporters any
such effect is unlikely to be mediated by changes in membrane fluidity.
Arachidonic acid is capable of being metabolized by both the
cyclooxygenase and the lipoxygenase enzyme systems. The resulting metabolites have been implicated in the control of a wide range of
physiological and pathological processes. Inhibitors of the individual
enzyme systems were employed to examine which, if either, of these
systems was involved in the generation of an AA metabolite that might
mediate the effects on glucose uptake. Cyclooxygenase, or prostaglandin
H synthase, catalyzes the conversion of AA to the prostanoids and the
thromboxanes. The enzyme exists as two isozymes, COX1, which is
constitutively expressed, and COX2, which is inducible (40). We
inhibited the COX enzyme activity in 3T3-L1 adipocytes for 30 min prior
to addition of AA using ibuprofen (nonselective), piroxicam (COX1), or
6-MNA (COX2). None of these compounds inhibited the AA potentiation of
either basal or insulin-stimulated glucose uptake in our system,
despite the demonstration that the COX enzyme activity had indeed been
inhibited. These results contrast with those of Fong et al.
(46) who reported that indomethacin (a nonselective COX inhibitor)
inhibited the 8-h AA potentiation of basal glucose uptake. The reasons
for this discrepancy are unclear but may be explained by the differing
experimental conditions of the two studies, specifically the time
points and particular COX inhibitors studied. Several groups have
demonstrated that certain COX metabolites of AA, particularly the J
series of prostaglandins, can activate PPAR Lipoxygenase enzymes catalyze the conversion of AA to the leukotrienes
and the hydroxyeicosatetraenoic acid. The enzyme exists as three
isozymes, 5-LOX, 12-LOX, and 15-LOX. We inhibited the LOX enzyme
activity in 3T3-L1 adipocytes for 30 min prior to addition of AA using
NDGA, a nonselective LOX inhibitor. We observed a significant
inhibition of the AA potentiation of basal glucose uptake with NDGA.
Furthermore, there was a trend for NDGA to inhibit the AA potentiation
of insulin-stimulated uptake. These results suggest that a LOX
metabolite(s) of AA may be involved in its effect on glucose uptake.
Indeed, it has been shown that the 15-LOX metabolites of both AA and
linoleic acid, 15-hydroxyeicosatetraenoic acid and 9- and
13-hydroxyoctadecadienoic acid, respectively, can function as
micromolar PPAR The nuclear hormone receptor, PPAR In summary, we have undertaken the most comprehensive study to date of
the effects of free fatty acids on basal and insulin-stimulated glucose
uptake in an insulin-responsive cell type. The potentiating effects of
even short-term exposure to AA are at least partially dependent on new
protein synthesis and involve enhanced expression of both GLUT1 and
GLUT4 at the plasma membrane. While activation of PPAR (PPAR
), a
nuclear receptor that positively modulates insulin sensitivity. To
clarify the effects of externally delivered fatty acids on glucose
uptake in an insulin-responsive cell type, we systematically examined the effects of a range of fatty acids on glucose uptake in 3T3-L1 adipocytes. Of the fatty acids examined, arachidonic acid (AA) had the
greatest positive effects, significantly increasing basal and
insulin-stimulated glucose uptake by 1.8- and 2-fold, respectively, with effects being maximal at 4 h at which time membrane
phospholipid content of AA was markedly increased. The effects of AA
were sensitive to the inhibition of protein synthesis but were
unrelated to changes in membrane fluidity. AA had no effect on total
cellular levels of glucose transporters, but significantly increased
levels of GLUT1 and GLUT4 at the plasma membrane. While the effects of
AA were insensitive to cyclooxygenase inhibition, the lipoxygenase inhibitor, nordihydroguaiaretic acid, substantially blocked the AA
effect on basal glucose uptake. Furthermore, adenoviral expression of a
dominant-negative PPAR
mutant attenuated the AA potentiation of
basal glucose uptake. Thus, AA potentiates basal and insulin-stimulated glucose uptake in 3T3-L1 adipocytes by a cyclooxygenase-independent mechanism that increases the levels of both GLUT1 and GLUT4 at the
plasma membrane. These effects are at least partly dependent on
de novo protein synthesis, an intact lipoxygenase pathway
and the activation of PPAR
with these pathways having a greater role in the absence than in the presence of insulin.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
in the control of insulin sensitivity
(14, 15). Thus, pharmacological agonists for PPAR
can enhance
glucose disposal both in vitro (16-18) and in
vivo (19) and loss of function mutations in human PPAR
result
in extreme insulin resistance and diabetes mellitus (20). While the
true endogenous ligand(s) for PPAR
have not been established with
certainty, the receptor can be activated in vitro by a
variety of lipophilic ligands including certain polyunsaturated fatty
acids (21-25) and their metabolites (21, 24, 26-29).
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
was a gift from Dr. M. Lazar (University of Pennsylvania School of Medicine). The
PGE2 monoclonal enzyme immunoassay kit was purchased from
Cayman Chemical. Ibuprofen, 6-methoxy-2-napthylacetic acid (6-MNA), and
piroxicam were purchased from Biomol Research Laboratories. The RNeasy
total RNA kit was from Qiagen. Rosiglitazone was provided by Dr. S. Smith (SmithKline Beecham Pharmaceuticals). Reverse transcription
reagents were obtained from Promega, UK, and TaqMan reagents were from
PE Biosystems. All other reagents were from Sigma.
4
M fatty acid and 0.4% ethanol.
4 M as described
above. 3T3-L1 adipocytes (day 9) in six-well plates were supplemented
with 2 ml of medium containing 0.05 µCi of [1-14C]AA
per well for 2, 4, 6, 8, and 10 h. At these time points, media was
removed and cells were washed twice with PBS. Cells were solubilized in
1 ml of 0.1 M NaOH per well and quenched using 50 µl of
concentrated HCl. Radioactivity in media and cell samples was
determined by liquid scintillation counting.
4
M AA for 4 h. Cells were washed twice with PBS and
solubilized by scraping in 4 ml of PBS/flask and passing 10 times
through a 25-gauge needle. Cell lysates were spun at 10,000 × g for 10 min at 4 °C to remove nuclei and mitochondria
and at 100,000 × g for 1 h at 4 °C to pellet
membranes. The supernatant, including the floating triglyceride layer,
was removed and the pellet resuspended in 1 ml of PBS by passing 10 times through a 27-gauge needle. Total lipids were extracted and
phospholipids separated and transmethylated as described previously
(34). The resulting fatty acid methyl esters were separated and
measured on a Unicam 610 gas chromatograph with flame ionization
detection and a 30 m × 0.53-mm Supelco SP2380 megabore column.
Helium was used as the carrier gas at a flow rate of 19.4 ml/min. A
temperature gradient program was used with an initial temperature of
70 °C increasing at 1 °C/min to 180 °C, then 5 °C/min to
200 °C and remaining at 200 °C for a further 6 min.
Identification of fatty acid methyl esters was made by comparison with
retention times of standard mixtures.
4
M AA for 4 h. Cells were washed and solubilized in PBS
and membranes were prepared as described above. Fluidity of the
membranes was assessed by measurement of steady state fluorescence
polarization of 1,6-diphenyl-1,3,5-hexatriene incorporated into the
hydrophobic core of the membrane bilayer. A modified version of the
previously described assay (35) was performed. Briefly, a stock
solution of 250 µM 1,6-diphenyl-1,3,5-hexatriene in
tetrahydrofuran was prepared. 0.9 mg of membrane in 3 ml of PBS was
incubated for 30 min in the dark with 0.25 µM
1,6-diphenyl-1,3,5-hexatriene at 37 °C. Fluorescence anisotropy
measurements were performed in a PerkinElmer Life Sciences (LS-5B)
luminescence spectrometer at 37 °C, with excitation and emission
wavelengths of 360 and 430 nm, respectively, and slit widths of 10 nm.
Intensities were corrected for intrinsic membrane fluorescence by using
the tetrahydrofuran vehicle. Fluorescence anisotropy is inversely
proportional to membrane fluidity.
1 (Ad
m). 3T3-L1
preadipocyte (2 days post-confluence) or day 7 adipocyte cultures in
24-well plates were infected with recombinant virus by addition of
1 × 109 plaque-forming units/well. 12 h later
medium containing free virus was removed and appropriate experimental
medium was added. Comparable viral infection efficiency was verified by
microscopy using a Zeiss axiovert 135 inverted fluorescence microscope.
Only cells with >70% infectivity were used in experiments.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
4 M free fatty acids for
4 h and basal and insulin-stimulated glucose uptake was compared
with cells incubated in vehicle alone. Ten different fatty acids were
studied, including several members of the n-3 and
n-6 series.2 Basal
glucose uptake was significantly increased with 16:1(n-7), 20:3(n-6), 20:4(n-6), 18:3(n-3), and
22:6(n-3) with all others showing a nonsignificant trend in
the same direction (Fig. 1A). All fatty acids, except 16:0, significantly increased
insulin-stimulated glucose uptake. The largest effect on glucose uptake
was observed with 20:4(n-6) (AA). This increased basal
glucose uptake by 1.8-fold and insulin-stimulated glucose uptake
2-fold. To establish the time course of these effects, cells were
incubated with AA for 1, 2, 4, 24, and 48 h. Significant
enhancement of glucose uptake, both basal and insulin stimulated, was
seen from 4 h onward, at which time point maximum effects were
seen (Fig. 1B). In repeat experiments, occasional cellular
toxicity was noted at 24- and 48-h exposures. Concentrations of AA
lower than 5 × 10
4 M had no discernible
effect (data not shown). All further experiments were undertaken using
8 × 10
4 M AA at the 4-h time point.
View larger version (25K):
[in a new window]
Fig. 1.
Glucose uptake. A, effect of
fatty acids. Day 9 adipocytes in six-well plates were supplemented with
8 × 10 4 M fatty acid in serum-free DMEM
for 4 h and stimulated for 30 min ± 10 nM
insulin (open columns, unstimulated; filled
columns, stimulated). 2-Deoxyglucose uptake was measured over 5 min as described under "Experimental Procedures." Data are mean
uptakes ± S.E. from four or more independent experiments
performed in triplicate, normalized to vehicle insulin-stimulated
uptake (mean insulin responses, 21,630 dpm/well). B, time
course of AA supplementation. Days 7-9 adipocytes in six-well plates
were supplemented with 8 × 10
4 M AA in
serum-free DMEM (1, 2, and 4 h) or in DMEM/FBS followed by 2 h in serum-free DMEM (24 and 48 h) as described under
"Experimental Procedures." 2-Deoxyglucose (DOG) uptake
was measured over 5 min following stimulation ± 10 nM
insulin for 30 min. Data are mean uptakes ± S.E. from four or
more independent experiments performed in triplicate, normalized to
vehicle insulin-stimulated uptake at each time point (mean
insulin-responses, 17,000 dpm/well). Open columns, vehicle;
filled columns, AA.
View larger version (31K):
[in a new window]
Fig. 2.
Effect of arachidonic acid on glucose
transporters. Day 9 adipocytes were supplemented with 8 × 10 4 M AA in serum-free DMEM for 4 h.
A, Western blotting. Cells were lysed as described under
"Experimental Procedures." 40 µg of total protein was resolved by
SDS-PAGE and immunoblotted with anti-GLUT1 or GLUT4 antibody. A
representative gel is shown for each data set. Numerical data are
percentage mean ± S.E. obtained by quantitation of gels from four
independent experiments, normalized to transporter levels in
vehicle-supplemented cells. Open columns, vehicle;
filled columns, AA. B, plasma membrane lawn
assay. Cells were stimulated ± 10 nM insulin for 30 min before preparation of plasma membrane lawns and assay of glucose
transporter translocation. Representative images from a typical
experiment are shown. Data from each experiment, utilizing 16 fields
for each condition, were quantified as described under "Experimental
Procedures" and overall results are shown as mean ± S.E. from
five independent experiments. Open columns, unstimulated;
filled columns, 10 nM insulin.
4 M AA also had a highly
significant effect on the fatty acid composition of cellular membranes
with a 4.5-fold increase in the AA content of membrane phospholipid
(p < 0.001) (Fig. 3B). The positive effects of AA on glucose uptake did not appear to be mediated by nonspecific effects on membrane fluidity which was unchanged after 4 h of incubation with AA (Fig. 3C).
View larger version (24K):
[in a new window]
Fig. 3.
Effect of arachidonic acid on membrane
phospholipid composition and fluidity. A,
[1-14C]AA uptake. Day 9 adipocytes in six-well plates
were supplemented with 8 × 10 4 M AA,
including a trace of [1-14C]AA, in serum-free DMEM for 2, 4, 6, 8, and 10 h. Radioactivity in media and cell lysates was
measured. Data are mean dpm/well ± S.E. from six wells in one
representative experiment. Circles, medium;
triangles, cell lysate. B, membrane phospholipid
fatty acid composition. Day 9 adipocytes in 3 × T175 flasks were
supplemented with 8 × 10
4 M AA in
serum-free DMEM for 4 h. Membranes were prepared, phospholipids
extracted, and their fatty acid composition analyzed as described under
"Experimental Procedures." Data are mean percentage fatty acid
composition of membrane phospholipids ± S.E. from four
independent experiments. C, membrane fluidity. Day 9 adipocytes in six-well plates were supplemented with 8 × 10
4 M AA in serum-free DMEM for 4 h.
Membranes were prepared and the fluidity assay performed as described
under "Experimental Procedures." Data are mean fluorescence
anisotropy "r" ± S.E. from eight independent experiments performed
in duplicate.
View larger version (15K):
[in a new window]
Fig. 4.
Effect of cycloheximide on the arachidonic
acid potentiation of glucose uptake. Day 9 adipocytes were
supplemented with 2.5 µM cycloheximide for 30 min and
with 8 × 10 4 M AA for a further 4 h as described under "Experimental Procedures." Cells were
stimulated for 30 min ± 10 nM insulin and
2-deoxyglucose uptake was measured over 5 min. A, data are
mean 2-deoxyglucose (DOG) uptake ± S.E. from seven
independent experiments performed in triplicate, normalized to nil
cycloheximide (CHX), vehicle insulin-stimulated uptake (mean
insulin responses 19,800 dpm/well). Open columns, vehicle;
filled columns, AA. B, data from A is
expressed as mean percentage difference in glucose uptake (AA,
cf. fatty acid vehicle) ± S.E.
View larger version (21K):
[in a new window]
Fig. 5.
Effect of metabolic inhibitors on the
arachidonic acid potentiation of glucose uptake. A,
glucose uptake. Day 9 adipocytes were supplemented with the indicated
concentrations of inhibitors for 30 min and with 8 × 10 4 M AA for a further 4 h as described
under "Experimental Procedures." Cells were stimulated for 30 min ± 10 nM insulin and 2-deoxyglucose uptake was
measured over 5 min. Data are mean percentage difference in
2-deoxyglucose uptake (AA, cf. fatty acid vehicle) ± S.E. from four or more independent experiments performed in triplicate,
normalized to inhibitor control (mean control values, basal, 57%;
insulin, 128%). B, PGE2 enzyme immunoassay.
Aliquots of medium were removed immediately prior to the glucose uptake
assay and tested for the presence of PGE2. Data are mean
picomole of PGE2/ml ± S.E. for AA-treated cells ± inhibitor from four independent experiments performed in triplicate.
Fatty acid vehicle-treated cells gave values below the level of
detection of the EIA kit (4.25 pmol/ml).
on the Arachidonic Acid
Potentiation of Glucose Uptake--
The L468A/E471A human PPAR
1
mutant is a powerful dominant-negative inhibitor of PPAR
action
(36). A recombinant adenovirus (Ad
m) which coexpresses
this mutant PPAR
receptor and green fluorescent protein (GFP) has
previously been demonstrated to markedly inhibit
thiazolidinedione-induced differentiation of human pre-adipocytes (36).
We first established that this virus could infect both 3T3-L1
pre-adipocytes and differentiated 3T3-L1 adipocytes with high
efficiency, resulting in an increase in levels of PPAR
1 expression
(Fig. 6A and Fig.
7, A and B).
Adenoviral infection with a control virus expressing only GFP (Ad-GFP)
had no effect on endogenous PPAR
levels (Figs. 6A and
7B). Expression of the dominant-negative PPAR
in 3T3-L1
pre-adipocytes significantly inhibited the induction of the
differentiation marker aP2 by rosiglitazone (p < 0.05)
(Fig. 6B) and also inhibited adipocyte differentiation in
response to a standard differentiation mixture (p < 0.05) (Fig. 6C). No such effects were seen with Ad-GFP (data
not shown). Thus, these studies confirmed that the human PPAR
mutant
receptor could inhibit the function of endogenous PPAR
activity in a
murine pre-adipocyte line.
View larger version (18K):
[in a new window]
Fig. 6.
Transduction of 3T3-L1 pre-adipocytes with
dominant-negative PPAR . Confluent
pre-adipocytes in 24-well plates were infected for 12 h with
1 × 109 pfu/well Ad-GFP or Ad
m. Medium
was changed to DMEM/FBS with differentiation mixture or DMEM/NBCS ± 1 × 10
7 M rosiglitazone for 48 h.
A, Western blotting. Infected pre-adipocytes (minus
rosiglitazone) and day 2 adipocytes were scraped and lysed as described
under "Experimental Procedures." 10 µg of total protein was
resolved by SDS-PAGE and immunoblotted with anti-PPAR
antibody.
B, aP2 induction by rosiglitazone in pre-adipocytes.
Infected pre-adipocytes ± 1 × 10
7
M rosiglitazone were lysed and RNA extracted and reverse
transcribed. aP2 gene expression was quantified using real time
quantitative PCR as described under "Experimental Procedures." Data
are mean fold rosiglitazone induction of aP2 ± S.E. C,
aP2 induction during differentiation. Infected pre-adipocytes (minus
rosiglitazone) and day 2 adipocytes were lysed and RNA extracted and
reverse transcribed. aP2 gene expression was quantified using real time
quantitative PCR. Data are mean fold induction of aP2 during
differentiation ± S.E. For experiments B and C, data are from
four independent experiments performed in triplicate, normalized to the
endogenous control, glyceraldehyde-3-phosphate dehydrogenase. Ad-GFP
data are representative of data from uninfected cells.
View larger version (37K):
[in a new window]
Fig. 7.
Transduction of 3T3-L1 adipocytes with
dominant-negative PPAR . Day 7 adipocytes in 24-well plates were infected for 12 h with 1 × 109 pfu/well Ad-GFP or Ad
m.
Medium was changed to DMEM/FBS for 48 h. A, microscopy.
Infection efficiency was estimated using fluorescence microscopy.
Representative images at ×320 magnification from a typical experiment
are shown. B, Western blotting. Infected adipocytes were
scraped and lysed as described under "Experimental Procedures." 10 µg of total protein was resolved by SDS-PAGE and immunoblotted with
anti-PPAR
antibody. C and D, glucose uptake.
Infected adipocytes were supplemented with 8 × 10
4
M AA in serum-free DMEM for 4 h and stimulated for 30 min ± 10 nM insulin. 2-Deoxyglucose uptake was
measured over 5 min. C, data are mean 2-deoxyglucose
uptake ± S.E. from four independent experiments performed in
triplicate, normalized to Ad-GFP, vehicle insulin-stimulated uptake
(mean insulin responses 11,310 dpm/well). Open columns,
vehicle; filled columns, AA. D, data from
C is expressed as mean percentage difference in glucose
uptake (AA, cf. fatty acid vehicle) ± S.E.
mutant on the AA potentiation
of glucose uptake, 3T3-L1 adipocytes were infected on day 7 of
differentiation and the glucose uptake studies were undertaken 2 days
later. Transduction efficiencies were ascertained by fluorescence microscopy (Fig. 7A) and plates with <70% infectivity were
discarded. The high-level expression of mutant PPAR
1 in
Ad
m-infected adipocytes was confirmed by Western
blotting (Fig. 7B). In these cells Ad
m significantly inhibited the AA potentiation of basal, but not insulin-stimulated, glucose uptake compared with cells infected with
the control virus (p < 0.01) (Fig. 7D). It
is also noteworthy that there was a trend for Ad
m to
reduce basal and insulin-stimulated glucose uptake in the absence of AA
(Fig. 7C).
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
provides a potential mechanism (15). With this in
mind, we investigated the effect of a range of fatty acids on glucose
uptake in 3T3-L1 adipocytes. Having established that AA had the most
marked effect on the enhancement of glucose uptake, we have attempted
to dissect the molecular events underlying this effect. Our studies
have found that 4 h of exposure to AA potentiates basal and
insulin-stimulated glucose uptake by increasing levels of GLUT1 and
GLUT4 at the plasma membrane. The potentiation of glucose uptake is
independent of cyclooxygenase but at least partly dependent on an
intact lipoxygenase pathway, de novo protein synthesis, and
PPAR
activation. Our results also suggest that the molecular
mechanisms whereby AA potentiates basal versus
insulin-stimulated glucose uptake may differ slightly.
in vitro and
could therefore act as endogenous ligands (21, 26, 27). Our results
would suggest that the AA potentiation of glucose uptake is not
mediated via the action of a COX metabolite on PPAR
. However, the
involvement of PPAR
in mediating the AA effect on glucose uptake is
not excluded. It is possible that other metabolites, or AA itself,
could act as PPAR
activators in this system. It should also be noted
that COX inhibitors themselves have also been demonstrated to act as ligands at PPAR
(47). Ibuprofen was shown to induce PPAR
activity 20-fold at 10
4 M but piroxicam had a
negligible effect. The effects of 6-MNA have not been reported.
agonists (29).
, has recently been shown to have
an important role in the control of insulin sensitivity (14, 15). Loss
of function mutations in human PPAR
result in extreme insulin
resistance and diabetes mellitus (20). As previously mentioned, several
metabolites of AA have been recognized as potential endogenous ligands
for PPAR
. Such metabolites include 15d-PGJ2 (26, 27),
12-PGJ2 (28), and PGD2 (21). It has also
been demonstrated that AA itself can act as a PPAR
ligand (22-25).
To investigate the role of PPAR
in our system, we inhibited endogenous activity by transducing fully differentiated adipocytes with
a dominant-negative hPPAR
mutant receptor. This mutant form of
hPPAR
has previously been shown to inhibit co-transfected wild-type
receptor action and to inhibit the thiazolidinedione-induced differentiation of human pre-adipocytes (36). We demonstrated that the
transduced hPPAR
mutant receptor was highly expressed in murine
3T3-L1 pre-adipocytes and adipocytes and inhibited endogenous wild-type
activity. We then showed that the AA potentiation of basal glucose
uptake was partially, but significantly, inhibited in cells transduced
with the mutant receptor. There was, however, no significant effect of
the mutant receptor on the AA potentiation of insulin-stimulated
glucose uptake. These results suggest that PPAR
activation is at
least partly required for the AA effect on glucose uptake. While the
dominant-negative mutant is clearly capable of specifically inhibiting
PPAR
function some caveats of this system should be mentioned.
First, although high levels of infectivity were observed, 3T3-L1
adipocytes are notoriously difficult to transduce with high efficiency
using adenoviruses and full inhibition of PPAR
function in all cells
may not have been achieved. It is also theoretically possible that this
dominant-negative mutant might, at high concentrations, occupy and
inhibit transcription through response elements other than the true
native peroxisome proliferator response elements. Such effects,
however, have not actually been demonstrated.
appears to be
necessary, particularly in the absence of insulin, this does not appear
to involve cyclooxygenase metabolites of AA but, somewhat surprisingly,
our results suggest a greater role for lipoxygenase metabolites in the
mediation of this important metabolic effect.
![]() |
FOOTNOTES |
---|
* This work was supportd in part by grants from the Wellcome Trust (to S. O. R., J. B. P., J. P. W., J. M. W., and V. K. K. C.).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.
Recipient of a studentship from Diabetes UK.
§ To whom correspondence should be addressed: Dept. of Medicine, University of Cambridge, Addenbrooke's Hospital, Box 157, Cambridge, United Kingdom CB2 2QR. Tel.: 1223-336855; Fax: 1223-330160; E-mail: sorahill@hgmp.mrc.ac.uk.
Published, JBC Papers in Press, December 21, 2000, DOI 10.1074/jbc.M009817200
2
The following fatty acid nomenclature was used:
16:0, palmitic acid; 16:1(n-7), palmitoleic acid;
18:2(n-6), linoleic acid; 18:3(n-6),
-linolenic acid; 20:3(n-6), dihomo-
-linolenic acid (DGLA); 20:4(n-6), arachidonic acid; 22:4(n-6),
adrenic acid; 18:3(n-3),
-linolenic acid;
20:5(n-3), EPA; and 22:6(n-3), DHA.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
PUFA, polyunsaturated fatty acid;
PPAR, peroxisome proliferator-activated
receptor
;
AA, arachidonic acid;
NBCS, newborn calf serum;
6-MNA, 6-methoxy-2-napthylacetic acid;
NDGA, nordihydroguaiaretic acid;
GFP, green fluorescent protein;
aP2, adipocyte P2;
PGE2, prostaglandin E2;
DMEM, Dulbecco's modified Eagle's
medium;
FBS, fetal bovine serum;
PBS, phosphate-buffered saline;
PAGE, polyacrylamide gel electrophoresis;
PCR, polymerase chain reaction;
COX, cyclooxygenase;
LOX, lipoxygenase.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | McGarry, J. D. (1992) Science 258, 766-770[Medline] [Order article via Infotrieve] |
2. | Boden, G. (1997) Diabetes 46, 3-10[Abstract] |
3. | Randle, P. J., Hales, C. N., Garland, P. B., and Newsholme, E. A. (1963) The Lancet April 13, 785-789 |
4. | Boden, G., Chen, X., Ruiz, J., White, J. V., and Rossetti, L. (1994) J. Clin. Invest. 93, 2438-2446[Medline] [Order article via Infotrieve] |
5. | Boden, G., and Chen, X. (1995) J. Clin. Invest. 96, 1261-1268[Medline] [Order article via Infotrieve] |
6. |
Roden, M.,
Krssak, M.,
Stingl, H.,
Gruber, S.,
Hofer, A.,
Furnsinn, C.,
Moser, E.,
and Waldhausl, W.
(1999)
Diabetes
48,
358-364 |
7. |
Hansen, P. A.,
Han, D. H.,
Marshall, B. A.,
Nolte, L. A.,
Chen, M. M.,
Mueckler, M.,
and Holloszy, J. O.
(1998)
J. Biol. Chem.
273,
26157-26163 |
8. |
Long, S. D.,
and Pekala, P. H.
(1996)
J. Biol. Chem.
271,
1138-1144 |
9. | Storlien, L. H., Kraegen, E. W., Chisholm, D. J., Ford, G. L., Bruce, D. G., and Pascoe, W. S. (1987) Science 237, 885-888[Medline] [Order article via Infotrieve] |
10. | Storlien, L. H., Jenkins, A. B., Chisholm, D. J., Pascoe, W. S., Khouri, S., and Kraegen, E. W. (1991) Diabetes 40, 280-289[Abstract] |
11. | Storlien, L. H., Baur, L. A., Kriketos, A. D., Pan, D. A., Cooney, G. J., Jenkins, A. B., Calvert, G. D., and Campbell, L. V. (1996) Diabetologia 39, 621-631[CrossRef][Medline] [Order article via Infotrieve] |
12. |
Borkman, M.,
Storlien, L. H.,
Pan, D. A.,
Jenkins, A. B.,
Chisholm, D. J.,
and Campbell, L. V.
(1993)
N. Engl. J. Med.
328,
238-244 |
13. | Pan, D. A., Lillioja, S., Milner, M. R., Kriketos, A. D., Baur, L. A., Bogardus, C., and Storlien, L. H. (1995) J. Clin. Invest. 96, 2802-2808[Medline] [Order article via Infotrieve] |
14. | Auwerx, J. (1999) Diabetologia 42, 1033-1049[CrossRef][Medline] [Order article via Infotrieve] |
15. | Willson, T. M., Brown, P. J., Sternbach, D. D., and Henke, B. R. (2000) J. Med. Chem. 43, 527-550[CrossRef][Medline] [Order article via Infotrieve] |
16. | el-Kebbi, I. M., Roser, S., and Pollet, R. J. (1994) Metabolism 43, 953-958[Medline] [Order article via Infotrieve] |
17. | Tafuri, S. R. (1996) Endocrinology 137, 4706-4712[Abstract] |
18. | Shimaya, A., Kurosaki, E., Shioduka, K., Nakano, R., Shibasaki, M., and Shikama, H. (1998) Horm. Metab. Res. 30, 543-548[Medline] [Order article via Infotrieve] |
19. | Day, C. (1999) Diabetic Medicine 16, 179-192[CrossRef][Medline] [Order article via Infotrieve] |
20. | Barroso, I., Gurnell, M., Crowley, V. E., Agostini, M., Schwabe, J. W., Soos, M. A., Maslen, G. L., Williams, T. D., Lewis, H., Schafer, A. J., Chatterjee, V. K., and O'Rahilly, S. (1999) Nature 402, 880-883[CrossRef][Medline] [Order article via Infotrieve] |
21. |
Yu, K.,
Bayona, W.,
Kallen, C. B.,
Harding, H. P.,
Ravera, C. P.,
McMahon, G.,
Brown, M.,
and Lazar, M. A.
(1995)
J. Biol. Chem.
270,
23975-23983 |
22. |
Forman, B. M.,
Chen, J.,
and Evans, R. M.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
4312-4317 |
23. |
Kliewer, S. A.,
Sundseth, S. S.,
Jones, S. A.,
Brown, P. J.,
Wisely, G. B.,
Koble, C. S.,
Devchand, P.,
Wahli, W.,
Willson, T. M.,
Lenhard, J. M.,
and Lehmann, J. M.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
4318-4323 |
24. |
Krey, G.,
Braissant, O.,
L'Horset, F.,
Kalkhoven, E.,
Perroud, M.,
Parker, M. G.,
and Wahli, W.
(1997)
Mol. Endocrinol.
11,
779-791 |
25. | Xu, H. E., Lambert, M. H., Montana, V. G., Parks, D. J., Blanchard, S. G., Brown, P. J., Sternbach, D. D., Lehmann, J. M., Wisely, G. B., Willson, T. M., Kliewer, S. A., and Milburn, M. V. (1999) Mol. Cell 3, 397-403[Medline] [Order article via Infotrieve] |
26. | Kliewer, S. A., Lenhard, J. M., Willson, T. M., Patel, I., Morris, D. C., and Lehmann, J. M. (1995) Cell 83, 813-819[Medline] [Order article via Infotrieve] |
27. | Forman, B. M., Tontonoz, P., Chen, J., Brun, R. P., Spiegelman, B. M., and Evans, R. M. (1995) Cell 83, 803-812[Medline] [Order article via Infotrieve] |
28. |
Ma, H.,
Sprecher, H. W.,
and Kolattukudy, P. E.
(1998)
J. Biol. Chem.
273,
30131-30138 |
29. | Nagy, L., Tontonoz, P., Alvarez, J. G., Chen, H., and Evans, R. M. (1998) Cell 93, 229-240[Medline] [Order article via Infotrieve] |
30. | Grunfeld, C., Baird, K. L., and Kahn, C. R. (1981) Biochem. Biophys. Res. Commun. 103, 219-226[Medline] [Order article via Infotrieve] |
31. |
Tebbey, P. W.,
McGowan, K. M.,
Stephens, J. M.,
Buttke, T. M.,
and Pekala, P. H.
(1994)
J. Biol. Chem.
269,
639-644 |
32. | Fong, J. C., Chen, C. C., Liu, D., Chai, S. P., Tu, M. S., and Chu, K. Y. (1996) Cell. Signal. 8, 179-183[CrossRef][Medline] [Order article via Infotrieve] |
33. |
Urso, B.,
Cope, D. L.,
Kalloo-Hosein, H. E.,
Hayward, A. C.,
Whitehead, J. P.,
O'Rahilly, S.,
and Siddle, K.
(1999)
J. Biol. Chem.
274,
30864-30873 |
34. | Pan, D. A., and Storlien, L. H. (1993) J. Nutr. 123, 512-519[Medline] [Order article via Infotrieve] |
35. | Knodell, R. G., Whitmer, D. I., and Holman, R. T. (1990) Gastroenterology 98, 1320-1325[Medline] [Order article via Infotrieve] |
36. |
Gurnell, M.,
Wentworth, J. M.,
Agostini, M.,
Adams, M.,
Collingwood, T. N.,
Provenzano, C.,
Browne, P. O.,
Rajanayagam, O.,
Burris, T. P.,
Schwabe, J. W.,
Lazar, M. A.,
and Chatterjee, V. K.
(2000)
J. Biol. Chem.
275,
5754-5759 |
37. | Wang, T., and Brown, M. J. (1999) Anal. Biochem. 269, 198-201[CrossRef][Medline] [Order article via Infotrieve] |
38. | Stouthard, J. M., Oude Elferink, R. P., and Sauerwein, H. P. (1996) Biochem. Biophys. Res. Commun. 220, 241-245[CrossRef][Medline] [Order article via Infotrieve] |
39. |
Clancy, B. M.,
Harrison, S. A.,
Buxton, J. M.,
and Czech, M. P.
(1991)
J. Biol. Chem.
266,
10122-10130 |
40. |
Smith, W. L.,
Garavito, R. M.,
and DeWitt, D. L.
(1996)
J. Biol. Chem.
271,
33157-33160 |
41. |
Meade, E. A.,
Smith, W. L.,
and DeWitt, D. L.
(1993)
J. Biol. Chem.
268,
6610-6614 |
42. | Laneuville, O., Breuer, D. K., Dewitt, D. L., Hla, T., Funk, C. D., and Smith, W. L. (1994) J. Pharmacol. Exp. Ther. 271, 927-934[Abstract] |
43. | Richelsen, B. (1987) Biochem. J. 247, 389-394[Medline] [Order article via Infotrieve] |
44. | Tobias, L. D., and Hamilton, J. G. (1979) Lipids 14, 181-193[Medline] [Order article via Infotrieve] |
45. | Stubbs, C. D., and Smith, A. D. (1990) Biochem. Soc. Trans. 18, 779-781[Medline] [Order article via Infotrieve] |
46. | Fong, J. C., Chen, C. C., Liu, D., Tu, M. S., Chai, S. P., and Kao, Y. S. (1999) Cell. Signal. 11, 53-58[CrossRef][Medline] [Order article via Infotrieve] |
47. |
Lehmann, J. M.,
Lenhard, J. M.,
Oliver, B. B.,
Ringold, G. M.,
and Kliewer, S. A.
(1997)
J. Biol. Chem.
272,
3406-3410 |