(Received for publication, April 12, 1995; and in revised form, August 4, 1995)
From the
Peroxisome proliferator-activated receptors (PPARs) are nuclear
hormone receptors that regulate gene transcription in response to
peroxisome proliferators and fatty acids. PPARs also play an important
role in the regulation of adipocyte differentiation. It is unclear,
however, what naturally occurring compounds activate each of the PPAR
subtypes. To address this issue, a screening assay was established
using heterologous fusions of the bacterial tetracycline repressor to
several members of the peroxisome proliferator-activated receptor
(PPAR) family. This assay was employed to compare the activation of
PPAR family members by known PPAR activators including peroxisome
proliferators and fatty acids. Interestingly, the activation of PPARs
by fatty acids was partially inhibited by the cyclooxygenase inhibitor
indomethacin, which prevents prostaglandin synthesis. Indeed,
prostaglandins PGA1 and 2, PGD1 and 2, and PGJ2-activated PPARs, while
a number of other prostaglandins had no effect. We also screened a
variety of hydroxyeicosatetraenoic acids (HETEs) for the ability to
activate PPARs. 8(S)-HETE, but not other (S)-HETEs,
was a strong activator of PPAR. Remarkably, PPAR activation by
8(S)-HETE was stereoselective. In addition, 8(S)-HETE
was able to induce differentiation of 3T3-L1 preadipocytes. These
results indicate that PPARs are differentially activated by naturally
occurring eicosanoids and related molecules.
The cloning and characterization of nuclear receptors has greatly enhanced our understanding of gene regulation by lipophilic hormones such as steroids, vitamin D, thyroxine, and retinoids. These receptors comprise a superfamily of transcription factors containing highly related DNA-binding domains(1, 2) . This family includes multiple subtypes of receptors for thyroxine and retinoids, encoded by distinct genes which are regulated quite differently during development and in the adult. There is evidence that retinoid receptor subtypes differentially bind retinoids(3, 4) . In addition, there is a large number of ``orphan'' receptors that have important roles in the development of species as diverse as invertebrates and mammals(5) . It is not known whether all of the orphan receptors will prove to be activated directly by small lipophilic molecules. However, at least one class of nuclear receptor, the retinoid X receptor, was initially an orphan member of the family (6) but later proved to bind and activate transcription in response to a naturally occurring retinoid, 9-cis-retinoic acid(7, 8) .
Peroxisome proliferator-activated
receptors (PPAR) ()were initially cloned as orphan receptors
and were subsequently found to be activated by peroxisome
proliferators. These include compounds such as clofibrate and Wy-14,643
which have been used clinically to treat hyperlipidemia, as well as by
plasticizers which may be carcinogenic for mammals(9) . There
are multiple subtypes of PPAR, called
,
(or NUC-I), and
in mammals. Studies from several investigators have suggested
that these subtypes are differentially activated by various
agents(10, 11, 12, 13, 14) .
PPAR
is most abundant in liver, while the tissue distribution of
PPAR
is more widespread. In contrast, expression of PPAR
is
limited to adipose tissue (15, 16) and, indeed,
activators of PPAR can suffice to induce adipose conversion of
preadipocyte cell lines(17, 18) . Moreover, ectopic
expression of PPAR
causes fibroblast cell lines to differentiate
into adipocytes in the presence of PPAR activators(19) . The
role of PPARs in adipocyte differentiation is likely to be complex,
since other PPARs are induced during adipocyte
differentiation(18, 20) .
Because none of the PPAR
activating compounds have been demonstrated to bind directly to PPAR, a
number of groups have searched for an endogenous ligand. These studies
revealed fatty acids to be activators of PPAR at high micromolar
concentrations(21, 22) . It remains unclear, however,
whether fatty acids are physiological activators of one or more PPAR
subtypes. Therefore, we devised a screen for PPAR activators and
applied it to known and potential activating compounds. PPAR,
, and
had highly divergent properties with respect to
activation by peroxisome proliferators and fatty acids. In addition, we
found that prostaglandins A, D, and J differentially activated PPAR
subtypes. Moreover, the naturally occurring,
12-O-tetradecanyolphorbol-13-acetate-inducible eicosanoid
8(S)-hydroxyeicosatetraenoic acid (8(S)-HETE)
activated PPAR
with greater effectiveness than other known
compounds. Activation by 8-HETE was stereoselective, and other (S)-HETEs were ineffective. In addition to activating
PPAR-mediated transcription, 8(S)-HETE induced adipogenic
differentiation of 3T3-L1 preadipocytes. Together, our results confirm
that PPAR subtypes are pharmacologically distinct and suggest that
certain naturally occurring eicosanoids are PPAR activators.
Figure 1:
Transactivation of pTetOluciferase
reporter gene by TetR-ER and TetR-PPAR fusion receptors. A,
schematic representations of the pTetO-Luc reporter and various
pSG5TetR fusion proteins. B, activation of transcription by
the TetR-PPAR fusions. U2OS cells were transfected then plated in
96-well dishes, as described under ``Materials and Methods,''
and induced with activators (1.0 µM 17--estradiol (
E) for ER, 100 µM Wy-14,643 (Wy)
for PPAR
/LBD and PPAR
, 50 µM DHA for
PPAR
/LBD). The data shown are the means of multiple repetitions of
the experiment (n = 8-12) with standard deviation
of 5-15%. The average luciferase activity for each induction was
shown as the relative -fold activation compared to the M2
SO (DMSO) vehicle control.
48 h after transfection and addition of
activators, media was removed from cells. The 96-well plates were
washed two times with 100 µl/well 1 phosphate-buffered
saline (-Mg
2
, -Ca2
). Cells
were then lysed by 20 µl of 1
luciferase lysis reagent (25
mM Tris-phosphate, pH 7.8, 2 mM dithiothreitol, 2
mM 1,2-diaminocyclohexane-N, N,N`,N`-tetraacetic acid, 10% glycerol, 1% Triton
X-100) for 15 min at room temperature using an orbital shaker. A
96-well format luminometer (ML3000, Dynatech Laboratories) was used for
reading the luciferase assay results. Immediately after addition of 100
µl/well of luciferase assay reagent (25 mM glycylglycine,
15 mM Mg(OAc)
, 0.75 mM ATP, 20
mM dithiothreitol, 0.1 mM EDTA, 0.8 mM luciferin, 0.24 mM acetyl-CoA), the plates were read in
the appropriate mode (e.g. cycle mode). The mean value and its
associated standard deviation for each set of repeated wells were
obtained using BioLink software that was supplied with the luminometer.
The effects of various compounds on activation of PPARs were presented
as ``fold-activation'' relative to the vehicle
(Me
SO or EtOH) control values. Transfection efficiency was
not a variable in these experiments because cells were simultaneously
transfected by electroporation prior to plating and addition of
compounds.
The system
was first tested using the human estrogen receptor (hER). The ER
full-length receptor and LBD sequences were each fused to the TetR and
cotransfected with pTetO-Luc reporter plasmid into U2OS cells, plated
with or without 1 µM 17--estradiol (E2). Fig. 1B shows that E2 induced transcription by the
full-length receptor 50-100-fold (Fig. 1B). The
ER LBD fusion produced very similar results (data not shown). We next
constructed and tested TetR-PPAR
(full-length and LBD),
TetR-PPAR
(full-length), and TetR-PPAR
(LBD) fusion
constructs (Fig. 1B). The TetR-PPAR
(LBD) and
full-length TetR-PPAR
were both activated
20-40-fold by
100 µM Wy-14,643, a known peroxisome proliferator and
activator of PPAR. The PPAR
full-length and LBD constructs gave
very similar results except that the full-length PPAR
fusion had a
relatively higher background luciferase activity in the absence of
added activator (data not shown). The TetR-PPAR
(LBD) fusion was
only weakly activated by Wy-14,643 but was activated up to 30-fold by
50 µM DHA (Fig. 1B and see below).
Figure 2:
Differential activation of PPAR,
PPAR
, and PPAR
by peroxisome proliferators, fatty acids, and
ETYA. U2OS cells were transfected then plated in 96-well dishes as
described under ``Materials and Methods'' then treated with
the following concentrations of the indicated PPAR activators: 100
µM WY-14,643, 1.0 mM clofibrate, 10 µM ETYA, 50 µM LA, and 50 µM DHA. Each
treatment was carried out in repeated wells (n =
8-12). The results for induction were expressed as -fold
activation relative to Me
SO (DMSO) vehicle
controls. Wy, Wy-14,643; CF, clofibrate. This
experiment was repeated at least three to four times with qualitatively
and quantitatively similar results.
Figure 3: Selective prostaglandins can activate PPARs. Transfected U2OS cells were treated with 10 µM each of the indicated prostaglandins in repeated wells (n = 4). 100 µM Wy-14,643 (Wy) and 50 µM DHA were used as positive controls. The effects of each prostaglandin were determined from the repeated wells. PGA, prostaglandin A; PGB, prostaglandin B; PGD, prostaglandin D; PGE, prostaglandin E; PGF, prostaglandin F; PGJ, prostaglandin J.
Figure 4:
Enantioselective activation of PPAR
by 8(S)-HETE. A, U2OS cells transfected with
pTetO-Luc and PPAR fusion receptors were screened for receptor
activation with HPLC mixtures. The (S)-HETE mixture contained
0.3 µM each of 5(S)-HETE, 8(S)-HETE,
11(S)-HETE, 12(S)-HETE and 15(S)-HETE. The
(±)-HETE mixture contained 0.3 µM each of
5(±)-HETE, 8(±)-HETE, 11(±)-HETE,
12(±)-HETE, and 15(±)-HETE. Each treatment was performed
in repeated wells (n = 4). WY-14,643 (WY) and
DHA were used as positive control activators. Luciferase assays were
performed 48 h later as described under ``Materials and
Methods.'' Note that the (S)-HETE mixture gave a 9-fold
activation, and the (±)-HETE mixture showed about half of the
activity (4-5-fold) B, stimulation of pTetO-Luc by
TetR-PPAR
was tested in the presence of 1.3 µM each
of the five pure HETEs: 5(S)-HETE, 8(S)-HETE,
11(S)-HETE, 12(S)-HETE, and 15(S)-HETE. C, activation of PPAR
activation by 8-HETE is
stereoselective. Cells were induced with 1.0 µM each of
the two stereoisomers, 8(S)- and 8(R)-HETE.
Luciferase activities determined 48 h after induction were shown
relative to the EtOH vehicle control. D, stimulation of
PPAR
-induced transcriptional activation of the bifunctional enzyme
PPAR response element-TK-luciferase reporter gene. The concentrations
of 8(S)-HETE, 8(R)HETE, Wy-14,643, and ETYA were 1,
5, 10, and 10 µM, respectively. Mean and range of
duplicate points are shown, normalized to the luciferase activity from
the reporter in the presence of PPAR
and absence of activator,
which was four to five times the activity of the reporter in the
absence of transfected PPAR
. The results shown are representative
of two separate experiments.
We next tested the effect of each of one of the five pure (S)-HETEs that were contained in the (S)-HETE
mixture. Fig. 4B shows that 8(S)-HETE was
responsible for almost all of the activity from the HPLC mixture.
9(S)-HETE, as well as its 9(R)-stereoisomer, were
also inactive (data not shown). 1.3 µM 8(S)-HETE
was as active as 100 µM of Wy-14,643 ( Fig. 4and
below). As suggested by the results using the (S)-HETE
mixture, none of the pure (S)-HETEs activated PPAR or
PPAR
(data not shown). To test the stereospecificity of PPAR
activation by 8(S)-HETE, we directly compared the activities
of 8(S)-HETE and 8(R)-HETE. The results in Fig. 4C show that while the 8(S)-enantiomer
was a strong activator, 8(R)-HETE showed very little activity.
These findings indicated that activation of PPAR
by
8(S)-HETE was stereoselective. The ability of only the
8(S)-enantiomer to activate PPAR
but not other PPARs was
confirmed using wild type PPARs along with a naturally occurring
PPAR-response element (data not shown).
To confirm that these
results were not an artifact related to the use of fusion proteins or
the TetO element, wild type PPAR was transfected into JEG3 human
choriocarcinoma cells along with a luciferase reporter containing a
naturally occurring PPAR-response element from the
hydratase-dehydrogenase (bifunctional enzyme) gene (30, 31) . As mentioned earlier, PPAR
activated
this reporter gene approximately 5-fold in the absence of exogenous
activator. Fig. 4D shows that this level of activation
was doubled by 8(S)-HETE, but not by 8(R)-HETE.
Indeed, the magnitude of activation of PPAR
by 8(S)-HETE
was about the same as that induced by maximal concentrations of
Wy-14,643 and ETYA.
Figure 5:
8(S)-HETE induces adipocyte
differentiation. A, morphological differentiation. Confluent
3T3-L1 preadipocytes were treated with 50 µM 8(S)-HETE, 50 µM 8(R)-HETE, 0.51
mM Wy-14,643 (Wy), or EtOH alone for 7 days. Phase
contrast microscopy is shown. B, induction of aP2, a molecular
marker of adipocyte differentiation. Northern analysis of 3 µg of
total RNA prepared from cells treated as in A. -
indicates control (EtOH) treatment. Northern analysis of -actin
mRNA expression is shown for comparison.
Figure 6:
Comparative activities of naturally
occurring and synthetic PPAR activators. U2OS cells transfected with
each PPAR fusion were tested with various concentrations of activators.
The upper panels, activators of PPAR; middle
panels, activators of PPAR
; and the lower panels,
activators of PPAR
. Each treatment was performed in repeated wells (n = 8), and the means are shown as relative -fold
activation compared to the Me
SO or EtOH vehicle control
values. This experiment was performed twice with similar results. CF, clofibrate.
The relative abilities to activate PPAR were PGA1
> PGD2 > DHA > LA and Wy-14,643 (Fig. 6, middle
panels). Significantly, the highest concentrations of PGA1 (50
µM) and DHA (100 µM) caused a greater fold
activation of PPAR
than did 100 µM Wy-14,643 (Fig. 6, middle panels; note differences in scale).
Studies of PPAR
indicated relative activities of PGD2 > PGA1,
DHA > clofibrate, Wy-14,643, and LA (Fig. 6, lower
panels). From the analyses conducted here, DHA, PGD2, and PGD1 ( Fig. 3and Fig. 6) appear to be the best activators of
PPAR
. The effects of Wy-14,643 and clofibrate on PPAR
were
much less than on PPAR
. LA was also a very poor activator for
PPAR
relative to PPAR
and PPAR
.
We have established a screen for PPAR activators which has the advantage of greatly reduced background when compared to assays involving transient transfection of wild type PPARs. The use of the TetR/TetO system also allows direct comparison of the magnitude of transactivation between nuclear hormone receptors, such as the ER and PPAR, as shown here. The use of chimeric PPARs involving fusion to a heterologous DNA-binding domain could lead to differences resulting from altered DNA binding and/or heterodimerization with RXR. However, parallel studies with wild type receptors and natural PPAR-response elements confirmed the validity of the screen. It is also possible that we would have obtained somewhat different results had we studied different cell types. Nevertheless, application of this assay to known PPAR activators confirmed the suggestion by others that different PPAR subtypes are differentially regulated by peroxisome proliferators and fatty acids. Furthermore, use of this assay as a screen led to the finding of differential PPAR activation by naturally occurring eicosanoids. The differential activation of PPARs is consistent with the fact that the C-terminal ligand-binding domains are less highly conserved among PPAR subtypes than among thyroid hormone receptor and retinoic acid receptor subtypes(10, 13, 14) .
Prostaglandins are lipid regulators of a number of important
cellular processes. Much of the prostaglandin literature has focused on
the role of cell surface receptors in mediating the pleiotropic effects
of these compounds(38, 39) . However, given their
circulating concentrations, low molecular weights, and lipophilicity,
it seems plausible that a subset of prostaglandins could activate
nuclear receptors directly or indirectly after diffusion into (or
production within) target cells. Indeed, certain prostaglandins such as
PGA1, PGD2, and PGJ2 have anti-tumor effects on human cancer cells,
including those derived from melanoma(40) ,
leukemia(41) , and ovarian carcinoma(42) . The ability
of these agents to regulate cell proliferation and apoptosis at least
partially involves nuclear
mechanisms(43, 44, 45) . It is therefore of
particular interest that these prostaglandins are the same subset which
activated PPARs. Peroxisome proliferators are hepatocarcinogens in
rodents, but the relationship between the anti-tumor effects of
prostaglandins described above and hepatic tumorigenicity is not clear.
The above-mentioned prostaglandins were generally equal or more
effective PPAR activators than the fatty acids and peroxisome
proliferators. Not all prostaglandins tested activated PPARs, and the
failure of PGE and PGF to activate was consistent with earlier studies
of Xenopus PPARs by Keller et al.(22) . However, that
group also reported that xPPAR was not activated by PGD2 while we
found that it did activate the mammalian PPAR
used in the present
study. PGD2 and the other prostaglandins that had activity in our
system activated all three PPAR subtypes. Since the structures of
inactive and active prostaglandins are not extremely different,
analysis of common features of the active prostaglandins may provide
clues to the structural requirements for activation of PPARs, whether
direct or indirect.
The ability of indomethacin to inhibit some of
the effects of peroxisome proliferators and fatty acids suggested that
these agents could act by a mechanism which is convergent with that of
the prostaglandins. Indeed, inhibitors of fatty acid oxidation can also
activate PPARs(46) . It should be cautioned, however, that
although metabolism of LA is consistent with a potential involvement of
cyclooxygenase pathways, DHA is an -3 fatty acid not
traditionally considered to be a precursor of arachidonic acid or
prostaglandins. To explain this, we speculate that either the effects
of indomethacin were not due entirely to inhibition of cyclooxygenase
or, alternatively, that DHA indirectly influenced prostaglandin
synthesis or metabolism. However, consistent with its function as a
cyclooxygenase inhibitor, indomethacin did not inhibit activation of
any of the PPAR subtypes by PGD2 (data not shown). The ability of both
ETYA, an inhibitor of arachidonate metabolism, as well as eicosanoids,
which are arachidonic acid metabolites, to activate PPARs also appears
paradoxical, but suggests that either ETYA has other cellular effects
or that certain arachidonic acid metabolites exert indirect effects
which mimic those of ETYA. Furthermore, the mechanism of the
interaction between Wy-14,643 and cyclooxygenase pathways is not
presently clear.
The ability of a HETE compound to activate PPAR is
of significance. HETEs are lipoxygenation products of arachidonate,
whose synthesis is cyclooxygenase-independent. In this regard, it is
noteworthy that indomethacin was least effective in inhibiting the
activation of PPAR, the PPAR that was activated by
8(S)-HETE. The best studied HETEs, 5-, and 12-, and
15-hydroxyeicosatetraenoic acids, are involved in a variety of
biological processes including inflammation, blood pressure regulation,
renal function, and respiratory airway smooth muscle
tone(33, 34, 35) . There is evidence that
these compounds function in part through cell surface receptors, but it
seems possible that they could exert a subset of their effects via
nuclear receptors. However, these compounds did not activate PPARs in
our experiments. 8(S)-HETE, in contrast, was a strong
activator of transcription by PPAR
. To our knowledge,
8(S)-HETE is the first example of a compound which
stereoselectively activates PPAR.
8(S)-HETE has not been as
thoroughly studied as other HETEs. It is a naturally occurring
compound, and a 8(S)-lipoxygenase activity involved in
8(S)-HETE biosynthesis has been shown to be present in mouse
epidermis(47, 48) . The tumor promoter
12-O-tetradecanyolphorbol-13-acetate induces this enzymatic
activity and causes a large increase in 8(S)-HETE (but not its
8(R)-enantiomer) in skin (47, 48) . The
normal function of 8(S)-HETE in skin is unknown. Since
PPAR is the primary mediator of peroxisome proliferator action in
liver(49) , and peroxisome proliferators cause liver tumors in
rodents(50) , it is of interest to consider whether
8(S)-HETE has similar effects in liver, where
PPAR
-inducible cytochrome p450 enzymes provide an alternative
pathway for eicosanoid biosynthesis(51) .
Little is also
known about the mechanism of 8(S)-HETE action. In our
experiments 8(S)-HETE selectively activated PPAR as well
or better than any natural or synthetic compound tested to date. Using
a protease protection assay that has been used to detect ligand binding
by other nuclear receptors(52) , we have been unable to
demonstrate direct binding of 8(S)-HETE to PPAR
. (
)However, a negative result in this assay may be due to
failure of binding to induce a conformational change which alters
protease sensitivity, rather than actual failure to bind to PPAR
.
Nevertheless, the stereoselectivity of 8(S)-HETE raises the
possibility that this compound, or a closely related metabolite, binds
directly to PPAR
. Interestingly, the hydroperoxyeicosatetranenoic
acid related to 8(S)-HETE (8(S)-HPETE) did not
activate xPPAR
in a previous study in which the 8-HETEs were not
evaluated(22) .
In the present studies we found that
8(S)-HETE, which appeared PPAR-specific in transient
transfection experiments, induced endogenous aP2 gene expression and
adipocyte differentiation of cultured 3T3-L1 cells. In this regard it
is interesting that epidermal expression of a dominant negative RAR
which also blocked PPAR action led to loss of multilamellar lipid
structures from the stratum corneum of mouse
skin(53, 54) . The requirement for higher
concentration of 8(S)-HETE for adipocyte differentiation than
for transactivation of PPAR
may be due to the very low expression
of PPAR
in 3T3-L1 preadipocytes(18) . Another possibility
is that 8(S)-HETE is metabolically inactivated during 3T3-L1
cell culture. Similar discrepancies between the ED
for
induction of adipocyte differentiation and PPAR activation have been
observed for WY-14,643 and ETYA(18) . It should also be noted
that although WY-14,643 caused a greater extent of adipocyte
differentiation than 8(S)-HETE in the present studies, the
concentration of Wy-14,643 was 10 times higher than the highest
concentration of 8(S)-HETE.
The predominant PPAR in
adipocytes is PPAR whose expression is itself highly specific for
adipocytes(15, 16) . The potential role of activated
PPAR
as a primary determinant of adipocyte differentiation is
underscored by the observation that ectopic expression of PPAR
is
sufficient for adipose conversion of fibroblast cell
lines(19) . Furthermore, while this paper was under review,
antidiabetic thiazolidinediones which induce adipocyte differentiation (55, 56) were found to specifically activate PPAR
(57) . (
)It is possible that 8(S)-HETE,
which was PPAR
-specific in other cell lines, or its
metabolite(s) induce adipocyte differentiation by activating
PPAR
in 3T3-L1 cells. However, it is conceivable that PPARs other
than PPAR
may also play a role in adipocyte differentiation.
Compounds such as WY-14,643 and ETYA also induced adipocyte
differentiation yet were poor activators of PPAR
, and there is a
significant time lag between commitment to adipocyte differentiation
after exposure to PPAR activators and the induction of
PPAR
(15, 16) . Both PPAR
and PPAR
are
also induced during adipocyte differentiation(18) . PPAR
is present in preadipocytes and activates transcription of the
adipocyte-specific aP2 gene, leading to the suggestion that PPAR
may be important for the earliest events in induction of adipose
differentiation by PPAR activators(20) . PPAR
has also
been shown to induce aP2 gene expression(19) . The present
results support the notion that induction and maintenance of adipocyte
differentiation by activation of PPARs is likely to be a complex
process involving multiple PPAR subtypes. The availability of
subtype-specific PPAR activators may allow further dissection of the
mechanism of this and other important biological processes regulated by
peroxisome proliferators, fatty acids, and eicosanoids.