A Photohormone for Light-Dependent Control of PPARα in Live Cells

Article abstract of DOI:10.1021/acs.jmedchem.1c00810

Photopharmacology enables the optical control of several biochemical processes using small-molecule photoswitches that exhibit different bioactivities in theircis- andtrans-conformations. Such tool compounds allow for high spatiotemporal control of biolog

Full text of DOI:10.1021/acs.jmedchem.1c00810

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Article
A Photohormone for Light-Dependent Control of PPARα in Live
Cells
Sabine Willems,§ Johannes Morstein,§ Konstantin Hinnah, Dirk Trauner,* and Daniel Merk*
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ABSTRACT: Photopharmacology enables the optical control of
several biochemical processes using small-molecule photoswitches
that exhibit different bioactivities in their cis- and trans-
conformations. Such tool compounds allow for high spatiotempo-
ral control of biological signaling, and the approach also holds
promise for the development of drug molecules that can be locally
activated to reduce target-mediated adverse effects. Herein, we
present the expansion of the photopharmacological arsenal to two
new members of the peroxisome proliferator-activated receptor
(PPAR) family, PPARα and PPARδ. We have developed a set of
highly potent PPARα and PPARδ targeting photohormones
derived from the weak pan-PPAR agonist GL479 that can be
deactivated by light. The photohormone 6 selectively activated
PPARα in its trans-conformation with high selectivity over the related PPAR subtypes and was used in live cells to switch PPARα
activity on and off in a light- and time-dependent fashion.
The PPARs, belonging to the subfamily I of NRs (NR1C),
comprise three isoforms PPARα, PPARδ, and PPARγ, which are
fatty acid- and lipid-activated transcription factors essentially
involved in metabolic balance and inflammatory processes.27
The PPARα subtype is mainly expressed in tissues with high β-
oxidation rates, such as the liver, brown adipose tissue, heart, and
kidney, and is considered the master regulator of fatty acid
catabolism, gluconeogenesis, and ketone body synthesis in a
nutrition-dependent manner.27,28 Comparably, PPARδ is
involved in fatty acid uptake and oxidation, blood glucose
homeostasis, and thermogenesis. It is ubiquitously expressed
with main functions in skeletal muscles and brown adipose
tissue.27,29,30 Both PPARα and PPARδ are closely linked to
metabolic disorders, such as diabetes, cardiovascular disorders,
nonalcoholic fatty liver disease (NAFLD), dyslipidemia, and
obesity.30−32
INTRODUCTION
■
Photopharmacology utilizes photoswitchable small molecules as
tools to obtain optical control of biological activity and modulate
cellular processes with unprecedented spatiotemporal resolu-
tion.1−4 While this approach has been successfully established to
control ion channels,5,6 membrane receptors,7−9 transport-
ers,10−13 and enzymes,14−16 nuclear receptors (NRs) are fairly
new to this proceeding. Upon binding of mainly amphiphilic
ligands, NRs interact with specific DNA response elements to
regulate the transcription of their target genes.17,18 The success
of photoswitchable lipids19 suggests that fatty acid mimetic20
NR ligands could also be suitable for photopharmacology. This
could indeed be demonstrated with a few classes of NRs,
including retinoic acid receptor α (RARα), which can be
optically controlled by a photoswitchable retinoic acid
derivative.21 Subsequently, we have developed dedicated
photohormones for the bile acid-activated transcription factor
farnesoid X receptor (FXR)22 and the fatty acid-sensing
peroxisome proliferator-activated receptor γ (PPARγ),23 and
Tsuchiya et al. have reported a photoswitchable estrogen
receptor (ER) agonist.24 Still, in light of their value as chemical
tools for pharmacology, the collection of available photo-
switchable ligands for transcription factors is scarce.
Here, we report the development of potent photoswitchable
ligands for PPARα and PPARδ, extending the reach of
photopharmacology to the remaining members of the important
PPAR subfamily of nuclear receptors. Using a novel cellular test
system embedding fluorescent reporter genes, we demonstrate a
Received: May 4, 2021
Published: July 2, 2021
The NRs’ function governs multiple physiological processes,
comprising embryonal development, cell proliferation, differ-
entiation, metabolism, and homeostasis.25 NR modulation by a
wide variety of endogenous ligands, including thyroid and
steroid hormones, bile acids, fatty acids, and vitamins causes
intermediate (minutes) to prolonged (hours−days) actions.17,26
https://doi.org/10.1021/acs.jmedchem.1c00810
© 2021 American Chemical Society
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photoswitchable, time-resolved control of PPAR activation with
these new photohormones in living cells.
on all PPARs but with a slight functional preference (higher
efficacy) for the PPARδ subtype (Table 1). Using light of
wavelength λ = 365 nm for photoswitching, we also probed
PPAR activation by the cis-isomer of 1, which exhibited similar
activity on all subtypes except a slight trans-preference for
PPARδ.
RESULTS AND DISCUSSION
■
Owing to their superior photophysical properties, stability over
multiple light-switching cycles, compatibility with various
scaffolds, and ease of synthesis, azobenzenes have emerged as
the most widely used compound class in photopharmacology.2
Additionally, the photoswitching of azobenzenes employed in
amphiphilic molecules is ideally suited to achieve reversible
photoswitchable properties.33 Based on these considerations, we
employed GL47934,35 (1, Chart 1) as a lead compound, since it
Co-crystal structures of the PPARα (PDB ID: 4CI4) and
PPARγ (PDB ID: 4CI5)36 ligand binding domains (LBDs) in
complex with 1 were available for structure-based optimization.
To obtain structural insights for PPARδ, we employed molecular
docking using a high-resolution PPARδ co-crystal structure with
a close analogue of the PPARδ selective agonist GW50151637
(PDB ID: 5Y7X)38,39 as a template. Initially, we studied the
chain length of the alkoxy linker between the aromatic systems
for potential optimization of the scaffold (Figure S1). Short-
ening by one carbon (2) appeared to be well-tolerated, while the
further shortened diphenylether failed to span the ample binding
pocket. As a result, the methyl propionate side chain was not
properly placed in the lipophilic subpocket of PPARα formed by
Phe273, Val444, Leu456, and Leu460, but its methyl decoration
clashed with His440. The extended chain of the phenoxypropyl
analogue of 1, in contrast, was too long causing a twisted
conformation and a clash with Ser280. The phenoxyethyl (1)
and phenoxymethyl (2) linkers were hence favored according to
the docking studies.
Chart 1. Lead Compound pan-PPAR Agonist GL479 (1)
already contains an azobenzene linker but has not been
previously investigated in a light-dependent fashion and was
exclusively studied in the dark-adapted trans-form. GL479 (1) is
a moderately potent pan-PPAR agonist with balanced potencies
a
Table 1. PPAR Modulatory Activity of GL479 (1) and Derivatives 2−11
EC50 [μM] (max. rel. act. [%])
PPARγ
#
R1
R2
−H
n
conf.
PPARα
PPARδ
1 (GL479)
−H
−H
2
trans
cis
trans
cis
trans
cis
trans
cis
trans
cis
trans
cis
trans
cis
trans
cis
1.09 0.02 (31 1)
0.8 0.2 (29 2)
3.7 0.3 (40 3)
3.5 0.7 (28 3)
0.66 0.07 (24 1)
1.1 0.2 (22 3)
2.67 0.16 (25 1)
6.0 0.6 (28 2)
5.40 0.07 (23 1)
6.6 0.3 (22 1)
EC50 > 10 μM (toxic ≥ 6 μM)
EC50 > 10 μM
2.2 0.1 (61 2)
6
1 (34 6)
2
−H
1
2
2
2
2
2
2
2
2
2
2.8 0.3 (40 3)
4.1 0.2 (26 1)
1.42 0.09 (33 1)
3.8 0.8 (31 5)
2 (39 6)
8.1 0.4 (28 1)
1.77 0.07 (40 1)
3.4 0.2 (22 1)
0.54 0.04 (45 1)
3.0 0.5 (41 4)
0.24 0.02 (46 1)
0.35 0.06 (44 2)
0.14 0.01 (45 2)
0.31 0.03 (37 2)
0.12 0.01 (47 1)
0.8 0.1 (32 1)
0.113 0.004 (43 1)
0.41 0.09 (40 6)
0.12 0.02 (65 4)
0.38 0.03 (35 2)
3
−CH3
−Cl
−H
4
−H
0.79 0.09 (26 1)
1.1 0.2 (24 2)
3.2 1.3 (14 1)
EC50 > 10 μM
8
5
−CH(CH3)2
−H
−H
0.47 0.01 (25 1)
0.28 0.05 (19 2)
0.0070 0.0006 (38 1)
0.24 0.02 (43 1)
0.029 0.004 (49 2)
0.040 0.003 (45 1)
0.009 0.002 (46 2)
0.071 0.007 (42 1)
0.036 0.006 (34 2)
0.64 0.08 (26 2)
0.07 0.01 (87 5)
0.19 0.01 (41 1)
0.04 0.01 (76 8)
0.29 0.05 (41 4)
2.1 0.2 (54 3)
2.6 0.2 (17 1)
1.2 0.1 (20 1)
1.7 0.8 (24 1)
0.82 0.07 (14 1)
0.92 0.08 (22 1)
0.6 0.1 (34 2)
0.77 0.07 (32 1)
2.3 0.1 (77 2)
1.2 0.3 (23 2)
1.2 0.2 (65 5)
1.3 0.2 (33 2)
4.5 0.2 (68 2)
4.8 0.4 (42 2)
6
−CH3
−Cl
7
−H
8
−H
−CF3
−CH3
−CF3
−Cl
9
−CH3
−CH3
−CH3
trans
cis
trans
cis
trans
cis
10
11
a
Activities were determined in uniform Gal4 hybrid reporter gene assays in HEK293T cells. Maximum relative activation (max. rel. act.) refers to
the activity of the respective reference agonist (PPARα: GW7647; PPARγ: pioglitazone; PPARδ: L165,041; each at 1 μM). Data are the mean
standard deviation (SD), n ≥ 3.
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Figure 1. Molecular docking of 1, 3, and 6 to reveal optimization potential. Trans-1 (teal) is shown in all models for comparison. (A−D) PPARα
binding site (gray) from the co-crystal structure with lead 1 (teal, PDB ID: 4CI436) and (E−H) PPARδ pocket (light blue) with the docked lead
structure 1 for comparison (PDB ID: 5Y7X38). The insets show overlays of the top 10 binding poses for each structure with their respective root-mean-
square deviation (RMSD) range to the pose shown in the binding site. Docking of cis-1 (green, A and E) suggested that binding of the cis-counterpart is
not favored. Terminal elongation with a p-methyl group enables improved binding modes of 3 in trans- (purple, B and F) and cis-states (magenta, G).
An o-methyl residue introduced in 6 (orange, D and H) extends into a lipophilic cavity. Also see Figures S1 and S2.
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In line with the activities observed in vitro, docking of 1 in
trans- and cis-states revealed slightly unfavorable binding modes
for the cis-counterpart with clashes of the azo linker (Figure 1A)
and the terminal phenyl ring (Figure 1E). No clashes were
observed for the shortened analogue cis-2, and several sound
binding poses were predicted (Figure S1). These computational
observations suggested 1 as a preferable lead for the develop-
ment of trans-selective analogues and that trans-binding might
be optimized by extension in the para-position of the terminal
ring to hinder cis-binding (Figure 1C,G). Introduction of a
methyl group in the para-position was well-tolerated by PPARα
as trans-3 and the co-crystallized ligand aligned well (Figure 1B).
However, molecular docking of the extended trans-3 to the
PPARδ ligand binding site revealed its cis-counterpart cis-3 as
the two top-ranked poses (Figures 1F,G and S2A). This
observation together with high-energy barriers for cis/trans-
isomerization of 43−52 kcal/mol in the forcefield used for the
calculations (Amber10:EHT, Figure S3A,B) suggested less
preference of PPARδ for trans-binding. Further structural
analysis indicated that introduction of methyl, chlorine, or
trifluoromethyl modifications in the ortho-position of the central
phenyl ring would address unoccupied space in the binding sites
of PPARα and PPARδ to enhance affinity (Figure 1D,H).
Especially in PPARα, introduction of a methyl group in the
ortho-position (6) seemed favorable as the moiety extended
toward a lipophilic subpocket (Figures 1D and S2B).
The biological evaluation of the photoswitchable ligands for
PPAR modulation was performed in uniform hybrid Gal4
reporter gene assays in HEK293T cells. These test systems
utilize chimeric constructs composed of the respective human
nuclear receptor LBD and the DNA-binding domain of the Gal4
protein derived from yeast. Gal4-responsive firefly luciferase was
used as a reporter gene, and constitutively expressed Renilla
luciferase served for normalization and to monitor test
compound toxicity. GW7647, Pioglitazone, and L165,041
(each at 1 μM) served as reference agonists for the three
PPAR isoforms α, γ, and δ, respectively, to obtain the relative
activation efficacy of the test compounds. To characterize both
isomers of 1−11 individually, all assays were conducted with the
trans-isomers in the dark and with the preilluminated cis-isomers
(λ = 365 nm) using a Cell DISCO40,41 system during incubation.
The biological activity of 1−11 is shown in Table 1.
Profiling of the lead compound GL479 (1) revealed agonism
on all three PPAR isoforms with low micromolar EC50 values in
consistence with the previously reported data for PPARα and
γ.34,35 The trans-1 and cis-1 isomers exhibited comparable
activity, except for a weak preference for trans-1 on PPARδ.
Aiming to improve upon the potency, subtype-selectivity, and
trans-preference of this new photohormone chemotype, we
systematically varied the structural features of 1 suggested by
molecular docking to hold optimization potential. Chain
shortening by one carbon in cis-/trans-2 slightly reduced
potency, thereby not evolving as a preferred modification and
was not further pursued. Extension in the para-position of the
terminal phenyl motif of 1 with methyl- (3) and chloro- (4)
substituents hardly affected potency but slightly improved the
desired trans-preference on PPARα. The isopropyl analogue 5,
in contrast, was slightly more active on PPARα than 1 with a
preference for the cis-conformation. Additionally, the para-
modifications (3−5) moderately improved selectivity over
PPARγ. Overall, 3−5 failed to provide a marked structural
optimization. Hence, we focused our attention on the ortho-
position of the central phenyl motif (6−8). As suggested by
molecular docking, introduction of a methyl group (6) provided
a remarkable improvement in PPARα agonism, which was
accompanied by a pronounced preference for trans-6. This
modification was also favored by PPARδ in terms of potency and
trans-preference despite a smaller impact. Activity on PPARγ
was not affected, resulting in strongly improved selectivity over
this PPAR subtype. A chlorine atom in the ortho-position (7)
increased potency on PPARα and PPARδ as well, but trans-
preference was lost. Trifluoromethyl analogue 8 was also very
potent on PPARα and exhibited a 10-fold preference for the
trans-state but was not superior to 6. On PPARδ, trifluoromethyl
substitution (8) evolved as the most favored modification in
terms of potency. In an attempt to fuse the terminal (3−5) and
central modifications (6−8) of the GL479 scaffold, we first
combined methyl groups in both positions (9), resulting in a
potent dual PPARα/δ agonist with pronounced trans-preference
but moderate activation efficiency. The combination of the
terminal methyl motif with a trifluoromethyl group on the
central ring (10) markedly improved activation efficiency with a
slight loss in potency on PPARα and less trans-preference while
a chlorine atom on the central ring (11) resulted in balanced
high potency on PPARα/δ with strong activation efficiency and
higher trans-preference. Additionally, 11 was selective for
PPARα/δ over PPARγ. Overall, the computer-aided structural
refinement of GL479 (1) as a photohormone yielded the highly
potent and selective photoswitchable PPARα agonist 6 (Figure
Following these computational observations, we studied
analogues 2−11 for improved optical control of PPARα and
PPARδ (Scheme 1). All photohormone candidates 1−11 were
a
Scheme 1. Synthesis of 1−11
a
Reagents and conditions: (a) ethyl 2-bromo-2-methylpropanoate,
K2CO3, DMF, reflux, 4 h, 47−80%; (b) azobenzenes 16−25, PPh3,
DEAD, THF, room temperature (rt), 16 h; and (c) NaOH, EtOH, rt,
16 h, 20−93% over two steps.
obtained from the respective esters 14 and 15 following an SN2
reaction of the phenols 12 and 13 with ethyl 2-bromo-2-
methylpropanoate and K2CO3 in dimethylformamide (DMF) as
previously reported.34 The Mitsunobu reaction of 14 and 15
with the commercially available azobenzenes 16−25 using
triphenylphosphine and diethyl azodicarboxylate (DEAD) in
tetrahydrofuran (THF) and subsequent saponification of the
obtained esters under basic conditions gave the desired
carboxylic acids 1−11 (Scheme 1).
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2A,B) and the dual PPARα/δ photohormone 11, which was
equally equipped with favorable potency and selectivity (Figure
pulses every 15 s) served to maintain the cis-conformation while
using a minimal nontoxic light dose. mCherry fluorescence was
measured every hour starting 8 h after addition of 6 (10 nM).
When cells were treated with trans-6, a constant increase in
mCherry fluorescence was observed between 8 and 24 h after
incubation (Figure 3A, blue curve). When cells were initially
treated with cis-6 for 8 h before switching to the trans-
counterpart, the mCherry signal was markedly delayed (Figure
3A, red curve). By contrast, when cells were initially treated with
trans-6 before switching to the cis-form after 8 h, the mCherry
signal initially increased similar to pure trans-6 treatment for 2 h
and then reached a low plateau with no further increase (Figure
3A, green curve). Similar observations were made with
photohormone 11 on Gal4-PPARδ (Figure S6A). Hence,
photohormones 6 and 11 enable spatiotemporal control of the
transcription factors PPARα and PPARδ.
Next, we studied selective and dual optical control of PPARα
and PPARδ in a similar setting (Figure 3B). HEK293T cells
were transiently transfected with Gal4-responsive mCherry and
Gal4-PPARα or Gal4-responsive eGFP and Gal4-PPARδ. The
cells were then pooled 5 h after transfection and treated with
DMSO (0.1%), cis-/trans-6, or cis-/trans-11, and the fluo-
rescence of mCherry and eGFP was determined after 40 h using
a fluorescence microscope (Figures 3B, S6B, and S7). DMSO-
treated cells revealed almost no mCherry or eGFP fluorescence.
Trans-6 at 10 nM markedly induced mCherry expression while
cis-6 had almost no effect, demonstrating optical control of
PPARα activity. At a higher concentration (300 nM), the
mCherry signals were similar for cis- and trans-6, and slight
eGFP fluorescence indicated weak PPARδ activation. Hence,
the photohormone 6 allows for strong, selective, and light-
dependent PPARα activation at a low 10 nM concentration,
qualifying 6 as an attractive tool for photopharmacology.
Figure 2. Characterization of photohormone 6. Dose−response curves
in cellular Gal4-PPARα (A) or Gal4-PPARδ (B) hybrid reporter gene
assays in HEK293T cells. Cis-6 was preilluminated (λ = 365 nm, 3 min)
and maintained in cis-state with a Cell DISCO system.40,41 Relative
activation refers to 1 μM GW7647 (PPARα) or L165,041 (PPARδ).
Data are the mean SD, n ≥ 3. ** p <0.01 and *** p <0.001 (t-test vs
cis-6). Photophysical characteristics of 6: UV−vis absorption scans (C,
50 μM in dimethyl sulfoxide (DMSO)) in dark-adapted (trans, black),
460 nm adapted (trans, blue), and 365 nm adapted (cis, gray)
photostationary states. (D) Reversible cycling (50 μM in DMSO) with
alternating illumination at 365 and 460 nm.
CONCLUSIONS
■
Photohormones have evolved as a new attractive type of tool
compounds that enable light-dependent modulation of tran-
scription factors to achieve spatiotemporal control of transcrip-
tional activity. As shown by our live-cell fluorescence reporter
gene assay, this allows for photoswitching between transcrip-
tionally active and inactive states, hence presenting photo-
hormones as valuable in vitro tools for functional studies. In the
field of nuclear receptors, temporal control appears particularly
relevant since these (hormone) receptors are endogenously
activated by short-lived molecules such as hormones and
reactive metabolites. Hence, nuclear receptors are shortly
stimulated under physiological conditions, which can be
unmatchably mimicked with photohormones. The importance
of time in nuclear receptor modulation is further highlighted by
their involvement in the circadian clock. Beyond this, optical
control of nuclear receptor activity may also gain therapeutic
relevance as it opens an avenue to local activation and
inactivation, for example, with wirelessly powered and
biodegradable optoelectronics to convert photohormones
between active and inactive conformations in certain tissues.42
This may enable reduction of target-mediated adverse effects
that cannot be avoided by conventional structural optimization
to improve selectivity. Photohormones, in contrast, can be on-
site activated or deactivated to spare tissues in which the target
of interest mediates side effects. In vivo studies have shown that
such optical control has therapeutic potential in metabolic
diseases43 and cancer.40,44
S4A,B). The PPARα selective photohormone 6 additionally
revealed high stability against microsomal degradation with an in
vitro half-life of 103 20 min (Figure S5), suggesting favorable
metabolic stability and potential for in vivo applications.
Photophysical characterization of the new photohormones 6
and 11 by UV−vis spectroscopy upon illumination with λ = 460
nm (trans, blue) and λ = 365 nm (cis, gray) confirmed
wavelength-dependent switching (Figures 2C and S4C), and
repeated photoswitching over multiple cycles demonstrated
high photostability (Figures 2D and S4D; 1 in Figure S3C,D).
Intrigued by the favorable activity profile and photophysical
properties of the photophormones 6 and 11, we studied their
applicability as tools in a cellular setting. To reflect
spatiotemporal control as a key feature of photopharmacology,
we established a new cellular assay to observe time-resolved
nuclear receptor activation and reversible activity of photo-
hormones in intact living cells over a long time period.
For this, we used Gal4-responsive fluorescent mCherry or
enhanced green fluorescent protein (eGFP) expression
constructs as reporter genes in HEK293T cells. After transient
transfection with the reporter and Gal4-PPARα, cells were
treated with photohormone 6 (Figure 3A). Light (λ = 460 and
365 nm) was used for switching between cis- and trans-
conformations, and a Cell DISCO system40,41 (75 ms light
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Figure 3. Spatiotemporal control of PPARα by 6. (A) Time-dependent PPARα activation by 6 using an mCherry fluorescence reporter in live
HEK293T cells. Trans-6 was maintained in the dark (blue). Switching between the cis- and trans-isomers after 8 h was performed by illumination with λ
= 460 nm (red) or λ = 365 nm (green) for 3 min, with subsequent maintenance of the cis-state using the Cell DISCO system.40,41 Fold fluorescence
intensity refers to 0.1% DMSO. Lines, mean; shadows, standard error of the mean (SEM); n = 3. (B) Live-cell imaging of PPARα-dependent mCherry
reporter induction by trans-6 (dark) and cis-6 (λ = 365 nm). The PPARδ-dependent eGFP fluorescence is negligible. The insets show mean SEM
relative reporter expression compared to GW7647 (PPARα) or L165,041 (PPARδ), n = 2.
residual protium in the NMR solvent (CDCl3: δ = 7.26; DMSO-d6: δ =
2.50; THF-d8: δ = 1.72). Carbon chemical shifts are expressed in ppm
(δ scale) and are referenced to the carbon resonance of the NMR
solvent (CDCl3: δ = 77.16; DMSO-d6: δ = 39.52; THF-d8: δ = 25.31).
High-resolution mass spectra (HRMS) were obtained with an Agilent
6224 accurate mass time-of-flight (TOF) liquid chromatography/mass
spectrometry (LC/MS) system using either an electrospray ionization
(ESI) or atmospheric pressure chemical ionization (APCI) ion source.
All reported data refer to the positive ionization mode. The compound
purity was analyzed by high-performance liquid chromatography-
ultraviolet (HPLC-UV) detection, and all compounds used for
biological characterization had a purity ≥95%.
Following these considerations, we have developed a highly
potent and selective PPARα photohormone from the weak pan-
PPAR agonist GL479 (1) in a structure-guided fashion. The
resulting analogue 6 not only exceeds the template compound 1
in terms of potency (factor > 150) and selectivity on PPARα but
is also equipped with the ability to be switched off by light-
induced isomerization to its cis-conformer, which is less active
on the intended target by a factor of 35. By enabling selective
optical control of PPARα, the photohormone 6 valuably
expands the photopharmacology toolbox for nuclear receptors,
enabling unprecedented in vitro and in cellulo experiments.
Synthesis and Analytical Characterization of 6 and 11 and Their
Precursor 14. (E)-2-Methyl-2-(4-(2-(2-methyl-4-(phenyldiazenyl)-
phenoxy)ethyl)phenoxy)propanoic acid (6). Ethyl 2-(4-(2-
hydroxyethyl)phenoxy)-2-methylpropanoate45 (14, 20.0 mg, 79.3
mmol, 1.00 equiv), (E)-2-methyl-4-(phenyldiazenyl)phenol (20, 21.9
mg, 103 mmol, 1.30 equiv), and triphenylphosphine (27.0 mg, 103
mmol, 1.30 equiv) were dissolved in anhydrous THF (0.5 mL). DEAD
(44.6 mg, 103 mmol, 1.30 equiv, 40% solution in toluene) was added
dropwise at 0 °C. The mixture was stirred at rt overnight. The solvent
was removed under reduced pressure. The crude was dissolved in
EtOH, and 1M NaOH was added. The reaction was stirred at rt for 16 h.
The solvent was removed under reduced pressure, and water was added
and acidified with 1 M HCl. The mixture was extracted with CH2Cl2,
dried over Na2SO4, filtered, and concentrated under reduced pressure.
EXPERIMENTAL SECTION
■
Chemistry. General. All reagents and solvents were purchased from
commercial sources (Sigma-Aldrich, TCI Europe N.V., Strem
Chemicals, etc.) and used without further purification unless otherwise
noted. Reactions were monitored by TLC on precoated, Merck silica
gel 60 F254 glass-backed plates. Flash silica gel chromatography was
performed using silica gel (SiO2, particle size 40−63 μm) purchased
from Merck. All NMR spectra were measured on a Bruker Avance III
HD 400 spectrometer (equipped with a CryoProbe). Multiplicities in
the following experimental procedures are abbreviated as follows: s =
singlet, d = doublet, t = triplet, q = quartet, quint = quintet, sext = sextet,
hept = heptet, br = broad, and m = multiplet. Proton chemical shifts are
expressed in parts per million (ppm, δ scale) and are referenced to the
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The crude product was purified using flash column chromatography
(CH2Cl2 to 20% MeOH in CH2Cl2) to yield 6 (28.2 mg, 67.4 μmol,
85%) as an orange solid. 1H NMR (400 MHz, THF-d8): δ = 7.84 (d, J =
7.3 Hz, 2H), 7.74 (s, 2H), 7.46 (t, J = 7.4 Hz, 2H), 7.40 (d, J = 7.1 Hz,
1H), 7.19 (d, J = 8.5 Hz, 2H), 7.05−6.99 (m, 1H), 6.84 (d, J = 8.5 Hz,
2H), 4.30−4.19 (m, 2H), 3.06 (t, J = 6.9 Hz, 2H), 2.25 (s, 3H), 1.52 (s,
6H). 13C NMR (100 MHz, THF-d8): δ = 175.6, 160.8, 155.7, 153.9,
147.4, 132.6, 131.0, 130.5, 129.8, 128.1, 124.8, 124.6, 123.3, 120.1,
default settings for each tool/function unless stated otherwise.
Amber10:EHT was used as the default forcefield for all calculations.
Molecular Docking. Docking was performed using the X-ray
structures of the PPARα LBD complexed with GL479 (PDB ID:
4CI436) and the PPARδ LBD complexed with GW501516 derivative
(PDB ID: 5Y7X38). Protonation states of the complexes were adjusted
using the MOE QuickPrep tool. The compounds were prepared using
the Energy minimize tool and MOE Wash tool: protonation state
dominant at pH 7.0. Docking was performed using the following
settings in the MOE Dock tool: receptor: receptor + solvent; site: ligand
atoms; placement: Triangle matcher; score: London dG; poses: 100;
refinement: induced fit; refinement score: GBVI/WSA dG; poses: 10.
As a pharmacophore query, the carboxylate oxygen of the respective
crystallized ligand was set as an anionic H-bond acceptor feature with a
1.0 radius and one H-bond acceptor projection feature with a radius of
1.4 from the same atom. All 10 superimposed binding poses are shown
as insets. The RMSD values between docked poses were calculated with
the mol_rmsd SVL script in MOE and are displayed as min−max
ranges. Redocking of the crystallized ligand GL479 (1) in PPARα (PDB
ID: 4CI436) resulted in an RMSD value of 0.1565 (range 0.1565−
1.1580, mean 0.6668).
+
111.5, 79.5, 70.1, 35.8, 25.9, 16.6. HRMS: m/z calcd for C25H27N2O4
([M + H]+): 419.1965; found, 419.1963.
(E)-2-(4-(2-(2-Chloro-4-(p-tolyldiazenyl)phenoxy)ethyl) phe-
noxy)-2-methylpropanoic acid (11). Ethyl 2-(4-(2-hydroxyethyl)-
phenoxy)-2-methylpropanoate45 (14, 21.9 mg, 87.0 mmol, 1.00
equiv), (E)-2-chloro-4-(p-tolyldiazenyl)phenol (25, 30.0 mg, 113
mmol, 1.30 equiv), and triphenylphosphine (29.6 mg, 113 mmol,
1.30 equiv) were dissolved in anhydrous THF (0.5 mL). DEAD (49.1
mg, 113 mmol, 1.30 equiv, 40% solution in toluene) was added
dropwise at 0 °C. The mixture was stirred at rt overnight. The solvent
was removed under reduced pressure. The crude was dissolved in
EtOH, and 1 M NaOH was added. The reaction was stirred at rt for 16
h. The solvent was removed under reduced pressure, and water was
added and acidified with 1 M HCl. The mixture was extracted with
CH2Cl2, dried over Na2SO4, filtered, and concentrated under reduced
pressure. The crude product was purified using flash column
chromatography (CH2Cl2 to 20% MeOH in CH2Cl2) to yield 11
Torsion Profile. The minimized energies (in kcal/mol) of the
associated lowest-energy structure were calculated for the conversion
from trans- to cis-state of 1 and vice versa using the MOE torsion profile
tool in bidirectional mode with both nitrogen atoms selected from the
azo (NN) double bond.
1
(17.6 mg, 38.9 μmol, 45%) as an orange solid. H NMR (400 MHz,
In Vitro Pharmacological Characterization. Hybrid Reporter
Gene Assays. Plasmids. The Gal4-fusion receptor plasmids pFA-
CMV-hPPARα-LBD, pFA-CMV-hPPARγ-LBD, and pFA-CMV-
hPPARδ-LBD coding for the hinge region and the ligand binding
domain of the canonical isoform of the respective nuclear receptor have
been reported previously.46 pFR-Luc (Stratagene, La Jolla, CA) was
used as the reporter plasmid and pRL-SV40 (Promega, Madison, WI)
for normalization of transfection efficiency and cell growth.
DMSO-d6): δ = 7.94−7.85 (m, 2H), 7.78 (d, J = 8.3 Hz, 2H), 7.39 (dd, J
= 8.6, 3.8 Hz, 3H), 7.27 (d, J = 8.7 Hz, 2H), 6.78 (d, J = 8.6 Hz, 2H),
4.34 (t, J = 6.9 Hz, 2H), 3.11−3.01 (m, 2H), 2.40 (s, 3H), 1.49 (s, 6H).
13C NMR (101 MHz, DMSO-d6): δ = 175.1, 156.0, 153.9, 149.9, 145.9,
141.6, 130.9, 130.0, 129.9, 124.9, 122.5, 122.5, 122.1, 118.4, 113.8, 78.3,
+
69.8, 33.9, 25.0, 21.0. HRMS: m/z calcd for C25H26ClN2O4 ([M +
H]+): 453.1576; found, 453.1573.
Ethyl 2-(4-(2-hydroxyethyl)phenoxy)-2-methylpropanoate45 (14).
4-(Hydroxyethyl)phenol (12, 200 mg, 1.45 mmol, 1.00 equiv) and
K2CO3 (2.00 g, 14.5 mmol, 10.0 equiv) were dissolved in DMF (4.5
mL). Ethyl 2-bromo-2-methylpropanoate (875 mg, 4.48 mmol, 3.10
equiv) was added, and the mixture was stirred for 4 h under reflux.
Water was added, phases were separated, and the aqueous layer was
extracted with EtOAc, washed with brine, dried over Na2SO4, filtered,
and concentrated under reduced pressure. The crude product was
purified using flash column chromatography (hexanes/EtOAC (8:2))
to yield 14 (292 mg, 1.16 mmol, 80%) as a colorless liquid. 1H NMR
(400 MHz, CDCl3): δ = 7.13−7.05 (m, 2H), 6.83−6.76 (m, 2H), 4.24
(q, J = 7.1 Hz, 2H), 3.82 (t, J = 6.5 Hz, 2H), 2.80 (t, J = 6.6 Hz, 2H),
1.58 (s, 6H), 1.25 (d, J = 7.1 Hz, 3H). 13C NMR (100 MHz, CDCl3): δ
= 174.5, 154.2, 132.1, 129.8, 119.6, 79.2, 63.9, 61.5, 38.5, 25.5, 14.2.
Procedure. HEK293T cells were cultured in Dulbecco’s modified
Eagle’s medium (DMEM), high glucose with 10% fetal calf serum
(FCS), sodium pyruvate (1 mM), penicillin (100 U/mL), and
streptomycin (100 μg/mL) at 37 °C and 5% CO2. Twenty-four
hours before transfection, the cells were seeded in transparent 96-well
plates (3 × 104 cells/well). Before transfection, the medium was
changed to Opti-MEM without supplements. Transient transfection
was carried out using Lipofectamine LTX reagent (Invitrogen,
Carlsbad, CA) according to the manufacturer’s protocol with pFR-
Luc (Stratagene), pRL-SV40 (Promega), and the corresponding Gal4-
fusion nuclear receptor plasmid. Five hours after transfection, the
medium was changed to Opti-MEM supplemented with penicillin (100
U/mL) and streptomycin (100 μg/mL) and additionally containing
0.1% dimethyl sulfoxide (DMSO) and the respective test compound or
0.1% DMSO alone as the untreated control. Each concentration was
tested in duplicate, and each experiment was repeated independently at
least three times. After incubation overnight (14−16 h), the cells were
assayed for luciferase activity using the Dual-Glo Luciferase Assay
System (Promega) according to the manufacturer’s protocol.
Luminescence was measured with a Tecan Spark 10M luminometer
(Tecan Deutschland GmbH, Crailsheim, Germany). Normalization of
transfection efficiency and cell growth was done by dividing the firefly
luciferase data by Renilla luciferase data and multiplying the value by
1000, resulting in relative light units (RLUs). Fold activation was
obtained by dividing the mean RLU of the test compound by the mean
RLU of the untreated control. Max. relative activation refers to the max
fold activation by a test compound divided by the fold activation of the
respective reference agonist (at a concentration of 1 μM). All hybrid
assays were validated with the respective reference agonists (PPARα:
GW7647; PPARγ: pioglitazone; PPARδ: L165,041), which yielded
EC50 values in agreement with the literature. Characterization of the
respective cis-counterparts was performed in the same way with
preirradiated compounds (irradiation for 3 min at λ = 365 nm before
incubation). To maintain the compound in the cis-adapted state, the
Cell DISCO system was used during incubation with 75 ms light pulses
(λ = 370 nm) every 15 s. For dose−response curve fitting and
+
HRMS: m/z calcd for C14H19O3 ([M − H2O]+): 235.1329; found,
235.1335.
Photophysical Characterization. UV−vis spectra were recorded
using a Varian Cary 50 Bio UV−visible spectrophotometer with
BRAND Ultra-Micro UV-Cuvettes (10 mm light path). Switching was
achieved using λ = 365 or 460 nm light-emitting diode (LED) light
sources. The LEDs were pointed directly into the top of the sample
cuvette. An initial spectrum of all photohormones (50 μM in DMSO)
was recorded (dark-adapted state, black) and then again following
illumination at λ = 365 nm for 1 min (cis-adapted state, gray). A third
spectrum was recorded after irradiation at λ = 460 nm for 1 min (trans-
adapted state, blue). To obtain the reversible trans ← → cis spectrum,
absorption at λabs = 340 nm was constantly measured, while alternating
illumination at λ = 365 or 460 nm for the indicated times allowed for
rapid isomerization of the photohormones (50 μM in DMSO). A
mercury lamp with a power of 75 watts connected to a monochromator
was directly pointed into the top of the sample cuvette, providing
irradiation via an optic fiber cable with the two distinct wavelengths λ =
365 and 460 nm for 1 min each (absorption was read at λabs = 340 nm).
Computational Methods. General. Calculations were performed
in Molecular Operating Environment (MOE, version 2020.09,
Chemical Computing Group ULC, Montreal, QC, Canada) using
10399
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J. Med. Chem. 2021, 64, 10393−10402

Journal of Medicinal Chemistry
pubs.acs.org/jmc
Article
calculation of EC50 values, the equation “[Agonist] vs response−
variable slope (four parameters)” was performed with mean relative
photoswitching of the test compounds was conducted after 8 h of
incubation with irradiation for 3 min at λ = 365 or 460 nm, respectively.
Fluorescence Cell Imaging. Plasmids. The Gal4-responsive
fluorescence reporters mCherry and eGFP were expressed from the
plasmids pUAS-mCherry-NLS (Addgene, entry 87695)49 and pGRE-
GFP (Addgene, entry 12516),50 respectively. The Gal4-fusion receptor
plasmid pFA-CMV-hPPARα-LBD and pFA-CMV-hPPARδ-LBD
coding for the hinge region and the ligand binding domain of the
canonical isoform of the respective nuclear receptor have been reported
previously.46
Procedure. HEK293T cells were cultured in DMEM, high glucose
with 10% FCS, sodium pyruvate (1 mM), penicillin (100 U/mL), and
streptomycin (100 μg/mL) at 37 °C and 5% CO2. Twenty-four hours
before transfection, cells were seeded in 24-well plates (1.5 × 105 cells/
well). Before transfection, the medium was changed to Opti-MEM
without supplements. Transient transfection was carried out using
Lipofectamine LTX reagent (Invitrogen) according to the manufac-
turer’s protocol with the combination of pUAS-mCherry-NLS
(Addgene, entry 87695) and pFA-CMV-hPPARα-LBD or pGRE-
GFP (Addgene, entry 12516) and pFA-CMV-hPPARδ-LBD. Five
hours after transfection, the cells transfected with pUAS-mCherry-NLS
and pFA-CMV-hPPARα-LBD or pGRE-GFP and pFA-CMV-hPPARδ-
LBD were trypsinized and pooled in a 50 mL tube, centrifuged, and
resuspended in Opti-MEM supplemented with penicillin (100 U/mL)
and streptomycin (100 μg/mL). The transfected cells were then
reseeded in 96-well plates already containing Opti-MEM with
supplements and 0.1% DMSO and the respective test compound or
0.1% DMSO alone as the untreated control. Each concentration was
performed in duplicate, and each experiment was repeated
independently two times. Characterization of the respective cis-
counterparts was performed in the same way with preirradiated
compounds (irradiation for 3 min at λ = 365 nm before incubation). To
maintain the compound in the cis-adapted state, the Cell DISCO
system was used during incubation with 75 ms light pulses (λ = 370 nm)
every 15 s.
Imaging. For the analysis of fluorescence reporter gene expression,
images were acquired 40 h after incubation with the test compounds
using a Zeiss Axio Observer Z1 (Carl Zeiss AG, Oberkochen,
Germany) microscope equipped with a 10× objective lens. The filter
sets had the following wavelengths/bandwidths: mCherry, excitation
545/40 nm ET bandpass; beam splitter T570 LPXR; and emission
620/60 nm ET bandpass; and eGFP, excitation 470/40 nm ET
bandpass; beam splitter T495 LPXR; and emission 525/50 nm ET
bandpass. Image analysis was performed with ImageJ software (version
1.53e). Integrated densities were measured from representative images
in separated stacks (mCherry and eGFP channel) as RGB color images
for the whole area, and relative integrated densities refer to mean
integrated densities divided by the mean integrated density of the
respective reference agonist (at a concentration of 1 μM; PPARα:
GW7647; PPARδ: L165,041).
activations
SD using GraphPad Prism (version 7.00, GraphPad
Software, La Jolla, CA).
Microsomal Stability Assay. The solubilized test compound 6 or
7-ethoxycoumarin as a reference (5 μL, final concentration 10 μM) was
preincubated at 37 °C in 432 μL of phosphate buffer (0.1 M, pH 7.4)
together with 50 μL of an NADPH regenerating system (30 mM
glucose-6-phosphate, 4 U/mL glucose-6-phosphate dehydrogenase, 10
mM NADP, 30 mM MgCl2). After 5 min, the reaction was started by
the addition of 13 μL of the microsome mix from the liver of Sprague−
Dawley rats (Invitrogen; 20 mg protein/mL in 0.1 M phosphate buffer)
in a shaking water bath at 37 °C. The reaction was stopped by adding
500 μL of ice-cold methanol at 0, 15, 30, and 60 min. The samples were
centrifuged at 5000g for 5 min at 4 °C, and the test compound was
quantified from the supernatants using HPLC-UV detection. The
composition of the mobile phase was adapted to the test compound in a
range of MeOH 40−90% and water (0.1% formic acid) 10−60%; flow-
rate: 1 mL/min; stationary phase: Purospher STAR, RP18, 5 μm, 125 ×
4; precolumn: Purospher STAR, RP18, 5 μm, 4 × 4; detection
wavelength: 254 and 280 nm; and injection volume: 50 μL. Control
samples were performed to check the test compound’s stability in the
reaction mixture: the first control was without NADPH, which is
needed for the enzymatic activity of the microsomes, the second control
was with inactivated microsomes (incubated for 20 min at 90 °C), and
the third control was without the test compound (to determine the
baseline). The amounts of the test compound were quantified by an
external calibration curve. Data are expressed as the mean SEM of the
remaining compound from three independent experiments. In vitro
half-life was calculated by a logarithmic linear transformation of the
remaining amounts of nonmetabolized test compound versus time in
GraphPad Prism 7 as described previously.47,48
Fluorescence Reporter Gene Assay. Plasmids. The Gal4-
responsive fluorescence reporter mCherry was expressed from plasmid
pUAS-mCherry-NLS (Addgene, entry 87695, Watertown, MA,
U.S.A.).49 The Gal4-fusion receptor plasmid pFA-CMV-hPPARα-
LBD and pFA-CMV-hPPARδ-LBD coding for the hinge region and the
ligand binding domain of the canonical isoform of the respective
nuclear receptor have been reported previously.46
Procedure. HEK293T cells were cultured in DMEM, high glucose
with 10% FCS, sodium pyruvate (1 mM), penicillin (100 U/mL), and
streptomycin (100 μg/mL) at 37 °C and 5% CO2. Twenty-four hours
before transfection, the cells were seeded in black cell culture 96-well
microplates with μClear flat bottom (3 × 104 cells/well; Greiner Bio-
One GmbH, Frickenhausen, Germany). Before transfection, the
medium was changed to Opti-MEM without supplements. Transient
transfection was carried out using Lipofectamine LTX reagent
(Invitrogen) according to the manufacturer’s protocol with pUAS-
mCherry-NLS (Addgene, entry 87695) and the corresponding Gal4-
fusion nuclear receptor plasmid. Five hours after transfection, the
medium was changed to Opti-MEM supplemented with penicillin (100
U/mL) and streptomycin (100 μg/mL) and additionally containing
0.1% DMSO and the respective test compound or 0.1% DMSO alone as
the untreated control. Each concentration was tested in duplicate, and
each experiment was repeated independently three times. After
incubation for 8 h, the living cells were assayed for fluorescence
reporter intensity every hour until 24 h and at 36 h. The fluorescence
intensity (FI) was measured after excitation at 585/10 nm with the
emission wavelength of 610/10 nm in bottom reading mode with a
Tecan Spark luminometer (Tecan Deutschland GmbH). Fold FI was
obtained by dividing the mean FI of the test compound by the mean FI
of the untreated control. Hybrid fluorescence assay performance was
monitored with the respective reference agonists at a concentration of 1
μM (PPARα: GW7647; PPARδ: L165,041). Characterization of the
respective cis-counterparts was performed in the same way with
preirradiated compounds (irradiation for 3 min at λ = 365 nm before
incubation). To maintain the compound in the cis-adapted state, the
Cell DISCO system was used during incubation with 75 ms light pulses
(λ = 370 nm) every 15 s. For two of the three study arms,
ASSOCIATED CONTENT
■
sı
* Supporting Information
The Supporting Information is available free of charge at
https://pubs.acs.org/doi/10.1021/acs.jmedchem.1c00810.
Supporting Information containing Figures S1−S7, Table
S1, synthesis and analytical characterization of 1−11 and
their precursors, HPLC traces, and NMR spectra (PDF)
Molecular formula strings containing molecular struc-
tures of 1−11 in trans and cis conformations and
associated activity data (csv)
AUTHOR INFORMATION
■
Corresponding Authors
Dirk Trauner − Department of Chemistry, New York
University, New York, New York 10003, United States;
10400
https://doi.org/10.1021/acs.jmedchem.1c00810
J. Med. Chem. 2021, 64, 10393−10402

Journal of Medicinal Chemistry
pubs.acs.org/jmc
Article
of a G Protein-Coupled Receptor with a SNAP-Tethered Photo-
chromic Ligand. ACS Cent. Sci. 2015, 1, 383−393.
orcid.org/0000-0002-6782-6056; Email: dirktrauner@
nyu.edu
Daniel Merk − Institute of Pharmaceutical Chemistry, Goethe
University Frankfurt, 60438 Frankfurt, Germany;
orcid.org/0000-0002-5359-8128; Email: merk@
pharmchem.uni-frankfurt.de
(8) Morstein, J.; Dacheux, M. A.; Norman, D. D.; Shemet, A.;
Donthamsetti, P. C.; Citir, M.; Frank, J. A.; Schultz, C.; Isacoff, E. Y.;
Parrill, A. L.; Tigyi, G. J.; Trauner, D. Optical Control of
Lysophosphatidic Acid Signaling. J. Am. Chem. Soc. 2020, 142,
10612−10616.
(9) Prischich, D.; Gomila, A. M. J.; Milla-Navarro, S.; Sanguesa, G.;
̈
Authors
Diez-Alarcia, R.; Preda, B.; Matera, C.; Batlle, M.; Ramírez, L.; Giralt,
E.; Hernando, J.; Guasch, E.; Meana, J. J.; de la Villa, P.; Gorostiza, P.
Adrenergic Modulation With Photochromic Ligands. Angew. Chem.,
Int. Ed. 2021, 60, 3625−3631.
Sabine Willems − Institute of Pharmaceutical Chemistry,
Goethe University Frankfurt, 60438 Frankfurt, Germany
Johannes Morstein − Department of Chemistry, New York
University, New York, New York 10003, United States
Konstantin Hinnah − Department of Chemistry, New York
University, New York, New York 10003, United States
(10) Quandt, G.; Höfner, G.; Pabel, J.; Dine, J.; Eder, M.; Wanner, K.
T. First Photoswitchable Neurotransmitter Transporter Inhibitor:
Light-Induced Control of γ-Aminobutyric Acid Transporter 1 (GAT1)
Activity in Mouse Brain. J. Med. Chem. 2014, 57, 6809−6821.
(11) Cheng, B.; Morstein, J.; Ladefoged, L. K.; Maesen, J. B.; Schiøtt,
B.; Sinning, S.; Trauner, D. A Photoswitchable Inhibitor of the Human
Serotonin Transporter. ACS Chem. Neurosci. 2020, 11, 1231−1237.
(12) Cheng, B.; Shchepakin, D.; Kavanaugh, M. P.; Trauner, D.
Photoswitchable Inhibitor of a Glutamate Transporter. ACS Chem.
Neurosci. 2017, 8, 1668−1672.
Complete contact information is available at:
https://pubs.acs.org/10.1021/acs.jmedchem.1c00810
Author Contributions
§S.W. and J.M. contributed equally to this study.
Notes
The authors declare no competing financial interest.
(13) Bonardi, F.; London, G.; Nouwen, N.; Feringa, B. L.; Driessen, A.
J. M. Light-Induced Control of Protein Translocation by the SecYEG
Complex. Angew. Chem., Int. Ed. 2010, 49, 7234−7238.
ACKNOWLEDGMENTS
■
(14) Velema, W. A.; Van Der Berg, J. P.; Hansen, M. J.; Szymanski, W.;
Driessen, A. J. M.; Feringa, B. L. Optical Control of Antibacterial
Activity. Nat. Chem. 2013, 5, 924−928.
pUAS-mCherry-NLS was a gift from Robert Campbell
(Addgene plasmid no. 87695; http://n2t.net/addgene:87695;
RRID:Addgene_87695). pGRE-GFP was a gift from Bert
Vogelstein (Addgene plasmid no. 12516; http://n2t.net/
addgene: 12516; RRID:Addgene_12516). J.M. and K.H.
thank the German Academic Scholarship Foundation for a
fellowship. J.M. thanks the New York University for a Margaret
and Herman Sokol fellowship and the NCI for an F99/K00
award (1F99CA253758-01). D.M. thanks the Aventis Founda-
tion for financial support.
(15) Tsai, Y.-H.; Essig, S.; James, J. R.; Lang, K.; Chin, J. W. Selective,
Rapid and Optically Switchable Regulation of Protein Function in Live
Mammalian Cells. Nat. Chem. 2015, 7, 554−561.
(16) Kol, M.; Williams, B.; Toombs-Ruane, H.; Franquelim, H. G.;
Korneev, S.; Schroeer, C.; Schwille, P.; Trauner, D.; Holthuis, J. C.;
Frank, J. A. Optical Manipulation of Sphingolipid Biosynthesis Using
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ABBREVIATIONS USED
■
(18) Aranda, A.; Pascual, A. Nuclear Hormone Receptors and Gene
Expression. Physiol. Rev. 2001, 81, 1269−1304.
DEAD, diethyl azodicarboxylate; eGFP, enhanced green
fluorescent protein; ER, estrogen receptor; FI, fluorescence
intensity; FXR, farnesoid X receptor; LBD, ligand binding
domain; NAFLD, nonalcoholic fatty liver disease; NR, nuclear
receptor; PPAR, peroxisome proliferator-activated receptor;
RAR, retinoic acid receptor; RLU, relative light units; rt, room
temperature
(19) Morstein, J.; Impastato, A. C.; Trauner, D. Photoswitchable
Lipids. ChemBioChem 2021, 22, 73−83.
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and Challenges for Fatty Acid Mimetics in Drug Discovery. J. Med.
Chem. 2017, 60, 5235−5266.
(21) Morstein, J.; Awale, M.; Reymond, J. L.; Trauner, D. Mapping the
Azolog Space Enables the Optical Control of New Biological Targets.
ACS Cent. Sci. 2019, 5, 607−618.
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