Synthesis and Characterization of a Bidirectional Photoswitchable Antagonist Toolbox for Real-Time GPCR Photopharmacology

Article abstract of DOI:10.1021/jacs.7b11422

Noninvasive methods to modulate G protein-coupled receptors (GPCRs) with temporal and spatial precision are in great demand. Photopharmacology uses photons to control in situ the biological properties of photoswitchable small-molecule ligands, which bodes

Full text of DOI:10.1021/jacs.7b11422

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Article
Synthesis and characterization of a bi-directional photoswitchable
antagonist toolbox for real-time GPCR photopharmacology
Niels J. Hauwert, Tamara A.M. Mocking, Daniel Da Costa Pereira, Albert J. Kooistra,
Lisa M. Wijnen, Gerda Vreeker, Noortje W.E. Verweij, Bert H. De Boer, Martine J. Smit,
Chris de Graaf, Henry F. Vischer, Iwan J. P. De Esch, Maikel Wijtmans, and Rob Leurs
J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.7b11422 • Publication Date (Web): 22 Feb 2018
Downloaded from http://pubs.acs.org on February 23, 2018
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Synthesis and characterization of a bi‐directional photoswitchable antagonist toolbox
for real‐time GPCR photopharmacology
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Niels J. Hauwert, Tamara A.M. Mocking, Daniel Da Costa Pereira, Albert J. Kooistra, Lisa M. Wijnen,
Gerda C.M. Vreeker, Noortje W.E. Verweij, Bert H. de Boer, Martine J. Smit, Chris De Graaf, Henry F.
Vischer, Iwan J.P. de Esch, Maikel Wijtmans*, Rob Leurs*
Email address: m.wijtmans@vu.nl, r.leurs@vu.nl
Division of Medicinal Chemistry, Amsterdam Institute for Molecules, Medicines and Systems, Faculty of
Sciences, Vrije Universiteit Amsterdam, De Boelelaan 1108, 1081 HZ Amsterdam, The Netherlands
Abstract
Non‐invasive methods to modulate G protein‐coupled receptors (GPCRs) with temporal and spatial
precision are in great demand. Photopharmacology uses photons to control in situ the biological
properties of photoswitchable small‐molecule ligands which bodes well for chemical biological precision
approaches. Integrating the light‐switchable configurational properties of an azobenzene into the ligand
core, we developed a bi‐directional antagonist toolbox for an archetypical family A GPCR, the histamine
H3 receptor (H3R). From 16 newly synthesized photoswitchable compounds, VUF14738 (28) and
VUF14862 (33) were selected as they swiftly and reversibly photoisomerize and show over 10‐fold
increased or decreased H3R binding affinities, respectively, upon illumination at 360 nm. Both ligands
combine long thermal half‐lives with fast and high photochemical trans‐/cis conversion, allowing their
use in real‐time electrophysiology experiments with oocytes to confirm dynamic photomodulation of
H3R activation in repeated second‐scale cycles. VUF14738 and VUF14862 are robust and fatigue‐
resistant photoswitchable GPCR antagonists suitable for spatio‐temporal studies of H3R signaling.
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Introduction
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Optical control of biological processes is one of the emerging fields in biomedicine due to its high spatio‐
temporal resolution1‐5. Initially, optogenetics revolutionized the field with major breakthroughs in e.g.
the field of neuroscience3. Lately, the field of photopharmacology1,2,5,6 has been providing a
complementary framework allowing the use of photoswitchable ligand molecules to dynamically
modulate native biological targets. In this framework, the photo‐induced configuration of the ligand has
either lower or higher affinity (directionality) than the 'dark' configuration, providing a means to
reversibly regulate the biological target with light. It has very recently been applied to e.g. native
membrane‐bound ion channels7‐9 and G protein‐coupled receptors (GPCR) 10‐14. Most GPCR examples
have so far focused on single ligands typically incorporating a photosensitive group in the periphery of
the GPCR ligands10‐14, successfully giving reversible mono‐directional photoswitches. In this work, we
present a bi‐directional photoswitchable GPCR antagonist toolbox, i.e. a set of ligands from the same
scaffold amenable to switching in two directions, showing either a light‐induced increase or decrease in
GPCR affinity, by positioning the photoswitchable moiety in the core of the new ligands.
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To this end, we used a prototypic family A GPCR, the histamine H3 receptor (H3R). The H3R is
expressed mainly in the central nervous system (CNS) and involved in a variety of CNS processes15‐17
.
Recently, the first H3R antagonist (pitolisant, Wakix™) has been approved for the treatment of
narcolepsy18. The medicinal chemistry of small‐molecule H3R ligands is well‐developed19,20 and provides
interesting chemical opportunities to incorporate photoswitchable groups in order to obtain
photoswitchable H3R antagonists. The general H3R antagonist pharmacophore includes a positively
charged amine group21, which is considered to interact with either the amino acid residuesD1143.32 or
E2065.46 in the transmembrane part of the H3R protein19,22,23. A linker, usually an alkylether, allows the
ligand to make a slight kink towards the bottom of the orthosteric H3R binding site. The ether oxygen
atom is connected to an aromatic core (e.g. phenyl, naphthyl, and many others19,20), which allows for ‐
 stacking interactions with aromatic H3R residues, like the Y1153.33, Y3746.51, and F3987.39 residues 24‐26
.
The aromatic core can be further substituted with a variety of groups including lipophilic groups,
hydrogen bond acceptors, basic and acidic moieties.27 Various photoswitchable moieties were evaluated
for replacement of the H3R antagonist aromatic core5 and the photoisomerizable azobenzene moiety
was deemed the best fit, because of the narrow, flat and elongated aromatic nature of its trans isomer.
Although a naphthalene moiety has not been coined as an “azostere”1, our design started from a
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naphthalene‐containing H3R antagonist (1) developed by Roche et al.28, which matches the previously
mentioned pharmacophore criteria. We postulated that, due to the relatively small nature of the
naphthalene core, additional room for ‐ stacking interactions is present in the hydrophobic part of the
H3R binding site, allowing incorporation of the azobenzene photoswitch in the core of the H3R scaffold
(Fig. 1) itself rather than in the periphery, as most often observed for GPCR photo‐ligands.
In this paper, we show how this core‐centered strategy led to the new H3R antagonists
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VUF14738 and VUF14862 that represent a bi‐directional set of robust and fatigue‐resistant
photochemical GPCR antagonists and are unique assets to further detail H3R pharmacology. We envision
that these ligands are useful molecular tools for e.g. probing H3R mediated responses in physiology in a
similar fashion as stereoisomers like R‐ and S‐α‐methylhistamine29,30, but having control of H3R affinity
using an external trigger. Moreover, in view of the importance of the H3R in various brain functions17,31
spatiotemporal photopharmacology approaches will bring new experimental options (next to
optogenetics) in order to delineate the importance of local neuronal circuits in H3R pharmacology.
,
Figure 1: General design and concept of bi‐directional photoswitchable H3R antagonists. In this SAR study, the azobenzene
moiety is incorporated in the core of compound 1 to give a series of differently substituted trans isomers (cyan). Illumination
leads to their corresponding cis isomers (magenta). Within this study, we aimed to discover photoswitchable H3R antagonists
that show at least a 10‐fold increase or decrease in GPCR affinity upon illumination.
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Results & Discussion
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Synthesis and photochemical characterization of azobenzene‐derived H3 ligands
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Three synthesis routes were developed which allowed for great versatility in the substitution patterns of
the azobenzene moiety (Scheme 1). A set of anilines was obtained by an amide coupling from 2‐4 to 5‐7.
The anilines underwent diazotization followed by a quench with phenol to afford 8‐11 and Williamson
ether synthesis to yield the para‐ether substituted H3R photoswitchable ligands 12‐17.
The second route was designed to overcome the undesired para‐selectivity in the diazotization and
comprises oxidation of aniline 18 and 19 to a nitroso‐intermediate, condensation with an aniline to
furnish 20‐24 and a Pd/RockPhos‐catalyzed ether synthesis32 to yield meta‐ether substituted ligands 25‐
30. For the synthesis of constrained ligands, either a Mitsunobu reaction (for 31 and 33) or a
Pd/RockPhos‐catalyzed ether synthesis32 (for 32) was employed instead of the Williamson ether
synthesis. The Mitsunobu approach was unable to yield 34 and instead a tosylate‐displacement strategy
was used for this compound.33
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Scheme 1: General synthetic scheme for photoswitchable H3R antagonists. Reagents and conditions: (a) pyrrolidine,
EDCI.HCl, DIPEA, HOBt.H2O, DMF, RT, 16 hrs, 57 – 95%; (b) (I) NaNO2, 1M aq. HCl, 0oC, 5 min; (II) phenol, aq. NaOH, RT, 30
min; (III) 1M aq. HCl, aq. sat. NH4Cl, RT, 10 min, 30 – 81%; (c) NaI, K2CO3, Pip‐(CH2)n‐Cl.HCl, DMF, 130oC, 16 hrs, 31 – 76%; (d)
Oxone™, H2O:DCM 4:1, RT, 3 hrs; (e) R‐Ar‐NH2, AcOH:DCM 1:1, RT, 16 hrs, 30 – 76% (over 2 steps); (f) R‐OH, RockPhos,
[PdCl(C3H5)]2, Cs2CO3, PhMe, 90oC, 23 hrs, 12 – 47%; (g) R‐OH, DEAD, PPh3, THF, 0oC RT, 18 hrs, 73 – 33%; (h) 1‐
methylimidazole, 4‐MePhSO2Cl, DCM, RT, 48 hrs, 43%; (i) (I) Oxone™, H2O:DCM 4:1, RT, 6 hrs; (II) 3‐OTBMDS‐aniline, AcOH,
RT, 16 hrs; (III) TBAF, THF, 0oC, 10 min, 13%; (j) (I) NaH, DMF, RT, 30 min; (II) 36, 90 oC, 16 hrs, 59%; Pip=1‐piperidino.
We first investigated the photoisomerization of all final compounds as well as their thermal relaxation at
20 C as determined by the method of Priimagi et al34 (Table 1, Fig. S1). Clear trends in the
photochemistry could be deduced. It was observed that para‐ether derivatives (scaffold A) have a
distinctly higher max for the ‐* transition of the trans isomers than meta‐ether derivatives (scaffold
B). This trend can be explained due to a better “push‐pull” profile in the electron density of the
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azobenzene unit of scaffold A, which is known to red shift the max of electronic transitions35,36. It was
found that upon continuous illumination at 10 mM in DMSO‐d6 with 360 ± 25 nm light a photostationary
state (PSS) containing over 85% cis isomer (LC area at 254 nm) could be obtained for all compounds,
with several ligands showing a PSS containing more than 95% cis isomer.
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Various trends can be observed regarding the thermal relaxation half‐lives of the compounds. In
line with literature, we observed a 3 to 5‐fold difference in thermal relaxation half‐life between the
para‐ethers (12, 13, 15, 17) and their respective meta‐ether (25, 26, 28, 30) analogs having a propyl‐
piperidine side chain.37 Such an effect can be attributed to increased conjugation in the para‐
derivatives38. In the series of ethyl‐piperidine analogs (14 versus 27, 16 versus 29) this effect of para‐
versus meta‐ether substitution is even more pronounced, increasing the differences in thermal
relaxation half‐life to 360 and 212‐fold, respectively. The propyl, cyclobutyl and 4‐piperidinyl spacers
appear to have a comparable influence on the thermal relaxation half‐lives (compare 15 versus 31‐33,
and 15/28 versus 33/34). If the amide group is placed at the ortho (13, 26) or para position (17, 30) the
thermal relaxation half‐life is comparable to its non‐substituted counterpart (12, 25). However, having
the amide in the meta position as in 15/28 yields a modest increase in thermal relaxation half‐life39.
Table 1: Structure‐photochemistry relationship of the photoswitchable antagonist series.
A
B
t @ 20 oC
(days)b
11.2
Compound
Amide
PSSa ± SEM
(area % cis)
96.6 ± 0.33
93.2 ± 0.29
88.2 ± 2.5
88.9 ± 1.3

max trans
(nm)
344
max cis
½
Scaffold
R
Number
12
position
(nm)
432
430
435
431
c
A
B
A
B
‐
c
25
‐
317
52.2
17
4
4
352
9.52
30
323
29.3
16
A
4
349
434
0.23
86.6 ± 2.5
29
15
B
A
4
3
324
347
429
432
48.8
26.5
91.3 ± 1.4
95.4 ± 0.50
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B
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318
345
427
431
122
86.8 ± 0.63
90.6 ± 0.77
0.69
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27
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3
318
425
248
89.3 ± 0.50
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425
161
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aPhotostationary state area percentages after illumination at 10 mM in DMSO‐d6 and as determined by LCMS analysis at 254
nm. bThermal relaxation half‐life times were determined according to the method of Priimagi et al.34 Arrhenius plots are
available in Fig. S1 cUnsubstituted phenyl moiety.
Compounds 28 and 33 were selected for further photochemical characterization based on their
favorable pharmacological characteristics (vide infra). The thermal relaxation half‐life times for these
compounds in 50 mM Tris‐HCl pH 7.4 buffer at room temperature were also directly determined,
resulting in half‐lives of 114 days for compound 28 (Fig. S2) and 26 days for compound 33 (Fig. S3).
These long thermal relaxation half‐lives are similar to the values obtained from the Arrhenius plots at 20
C and uniquely allowed for a thorough analysis of the photoswitching and relaxation process by 1H‐
NMR spectroscopy in conjunction with LC‐MS chromatography and UV‐Vis spectroscopy. We first
investigated isomerization of 28 and 33 at 10 mM in DMSO‐d6 by monitoring both 1H‐NMR and LC‐MS
profiles in time. While multiple 1H‐NMR signals change upon photoswitching, the signals for the CH or
CH2 group adjacent to the ether O‐atom provided a clearly resolved signal for quantification (Fig. 2A, C).
For both 28 and 33 the percentages of isomerization as analyzed by 1H‐NMR spectroscopy and LC‐MS
analysis at 254 nm (Fig. 2B, D) are comparable yet slightly different, reflecting most likely some
differences in ‐values of the trans and cis isomers (Table S1). Despite these minor differences, LC‐MS
was subsequently used as a routine analysis in view of the higher throughput.
Next, the photoisomerization rate was investigated at 25 M in 50 mM Tris‐HCl pH 7.4 buffer
supplemented with 1% DMSO‐d6 using UV‐Vis spectroscopy (Fig. 2E, F). Rapid photoisomerization was
observed for both compounds during illuminations at both 360 ± 25 nm and 430 ± 17 nm to obtain high
amounts of cis isomer or considerable amounts of trans isomer, respectively (Fig. S4‐S6, S10‐S12). After
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30 seconds of illumination, isomerization of both key compounds had reached isomerization
percentages within 5% of their PSS. Thus, illuminating for 300 seconds was always sufficient to reach
PSS. Arguably due to both solvent and concentration effects36,40,41 the measured isomerization rates are
considerably higher in the experiments performed at 25 M in buffer, compared to the measurements
at 10 mM in DMSO‐d6 of the two H3R ligands.
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We subsequently studied the real‐time isomerization behavior of both compounds at a
concentration of 25 M in 50 mM Tris‐HCl pH 7.4 buffer supplemented with 1% DMSO‐d6 using a setup
which allowed the illumination perpendicular to the path of the UV‐Vis spectrometer light source. Under
these conditions, we determined a photoisomerization half‐life for 33 of 4 ± 0.2 seconds under
illumination with 360 ± 25 nm and 8 ± 0.6 seconds under illumination with 430 ± 17 nm (Fig. 2G). For 28
the values were found to be 16 ± 0.9 seconds under illumination at 360 ± 25 nm and 11 ± 0.3 seconds
under illumination at 430 ± 17 nm (Fig. 2H).
Both photoswitchable compounds proved to be very photostable. Even after 48 hours of
illumination at 360 ± 25 nm with an intensity of 0.77 mW/mm2 no apparent photodegradation was
visible in both 1H‐NMR and LC‐MS analyses of 28 and 33. To exemplify the resistance to fatigue of 28 and
33 over 1000 isomerization cycles switching between illumination at 360 ± 25 and 430 ± 17 nm with 20‐
second intervals were executed. The absorbance at max of the ‐* transition of the trans isomer did
not alter considerably (Fig. 2I, 2J, S16 and S17) indicating that both key photoswitches are highly
resistant to photobleaching.
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Figure 2: (A) A representative part of 1H‐NMR spectra of 10 mM of compound 33 in DMSO‐d6 illuminated at 360 ± 25 nm
displayed at various time points (seconds). The presented peak belongs to the hydrogen atom explicitly drawn in the
structure shown above the spectrum. The full 1H‐NMR spectra are available in supplementary figure S18. (B) A
representative part of LC‐MS chromatograms belonging to the illuminated NMR samples shown in Fig. 2A. The full
chromatograms are available in supplementary figure S19. (C) A representative part of 1H‐NMR spectra of 10 mM of
compound 28 in DMSO‐d6 illuminated at 360 ± 25 nm displayed at various time points (seconds). The presented peak belongs
to the hydrogen atom explicitly drawn in the structure shown above the spectrum. The full 1H‐NMR spectra are available in
supplementary figure S20. (D) A representative part of LC‐MS chromatograms belonging to the illuminated NMR samples
shown in Fig. 2C. The full chromatograms are available in supplementary figure S21. (E) UV‐Vis spectra of 25 M of
compound 33 (trans) in 50 mM Tris‐HCl pH 7.4 buffer containing 1% DMSO‐d6. PSS cis represents a sample which has been
illuminated for 300 seconds using 360 ± 25 nm light. PSS trans represents subsequent illumination for 300 seconds using 430
± 17 nm. (F) UV‐Vis spectra of 25 M of compound 28 (trans) in 50 mM Tris‐HCl pH 7.4 buffer containing 1% DMSO‐d6. PSS cis
represents a sample which has been illuminated for 300 seconds using 360 ± 25 nm. PSS trans represents subsequent
illumination for 300 seconds using 430 ± 17 nm. (G) Absorbance at 344 nm of 25 µM of compound 33 in 50 mM Tris‐HCl pH
7.4 buffer + 1% DMSO‐d6. UV‐Vis spectra were obtained with 1‐second intervals under alternating illumination with 360 ± 25
nm and 430 ± 17 nm perpendicular to the light source of the UV‐Vis spectrometer. (H) Absorbance at 319 nm of 25 M of
compound 28 in 50 mM Tris‐HCl pH 7.4 buffer + 1% DMSO‐d6. UV‐Vis spectra were obtained with 1‐second intervals under
alternating illumination with 360 ± 25 nm and 430 ± 17 nm perpendicular to the light source of the UV‐Vis spectrometer. (I)
Repeated isomerization cycles of 25 M of compound 33 in a pH 7.4 buffer containing 15 mM HEPES, 64 mM NaCl, 25 mM
KCl, 0.4 mM CaCl2 and 0.8 mM MgCl2 containing 1% DMSO‐d6 analyzed at 349 nm. PSS cis was obtained by using illuminations
for 20 seconds at 360 ± 25 nm. PSS trans was obtained by using illuminations for 20 seconds at 430 ± 17 nm. An extended
figure is available in supplementary figure S16. (J) Repeated isomerization cycles of 25 M of compound 28 in a pH 7.4 buffer
containing 15 mM HEPES, 64 mM NaCl, 25 mM KCl, 0.4 mM CaCl2 and 0.8 mM MgCl2 containing 1% DMSO‐d6 analyzed at 319
nm. PSS cis was obtained by using illuminations for 20 seconds at 360 ± 25 nm. PSS trans was obtained by using illuminations
for 20 seconds at 430 ± 17 nm. An extended figure is available in supplementary figure S17.
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Photopharmacological characterization
Because of their long thermal relaxation half‐lives, the newly synthesized photoswitchable GPCR
ligands are uniquely compatible with competition radioligand‐binding experiments. All new azobenzene‐
derived H3R ligands, either illuminated using 360 ± 25 nm to PSS (as determined by LC‐MS) or kept in the
dark, were evaluated for their affinity for the human H3R by a [3H]‐N‐methylhistamine competition
binding experiments performed in the dark. Both histamine and, as previously reported28, the parent
naphthalene‐compound 1 potently displace [3H]‐N‐methylhistamine binding to membranes
overexpressing the H3R (Table 2) and their affinity values were not affected by illumination of the
compound solutions at 360 ± 25 nm (data not shown). Replacement of the naphthalene core of 1 with
the azobenzene core resulted in a series of moderate to highly potent H3R ligands with pKi values
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between 6.01 and 8.76 as trans isomers (i.e. without illumination). The direct azobenzene analog of 1,
compound 17, bound the H3R with only a slightly lower affinity (pKi = 7.96 ± 0.06, Table 2) compared to 1
(pKi = 8.29 ± 0.01, Table 2). Furthermore, the affinity for H3R was modulated effectively by shifting the
position of the amine side chain, with scaffold A (para‐substitution) always resulting in better affinities
than scaffold B (meta‐substitution) (Table 2). In general, the propyl spacer proved advantageous over
the ethyl spacer for H3R affinity. The effects of amide positioning varied amongst the different
subclasses.
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Rather than absolute affinities of the trans isomers for H3R, the shift in affinity upon illumination
was considered the prime parameter for photopharmacology applications and selection of key
compounds. An increase in binding affinity upon photoisomerization (illumination at 360 ± 25 nm) was
observed for 12, 25, 27, 28 and 30. Interestingly, most of these compounds contain a meta‐ether‐
substituted azobenzene core (scaffold B) with either a 3‐carboxamide (27, 28), a 4‐carboxamide (30) or
no carboxamide substituent (25). The ethyl‐spaced 27 has an 8.3‐fold decrease in H3R affinity compared
to the propyl‐spaced analog 28. Within the set of compounds lacking a carboxamide, meta‐ and para‐
ether‐substituted azobenzene compounds 25 and 12 (scaffolds B and A) show similar shifts in H3R
affinity upon isomerization. The 13.5‐fold increase in H3R affinity upon photoswitching of 28 (Fig. 3A) led
to its identification as one of the key compounds of this study (VUF14738).
Compounds that show decreased H3R binding affinity upon isomerization were primarily found
to have a para‐ether‐substituted azobenzene (scaffold A). The trend that ethyl‐spaced compounds
perform less well than their respective propyl‐spaced analogs was also observed for the pairs 14/15 and
16/17. Derivatives with a 2‐carboxamide substituents (13, 26) gave smaller shifts in H3R affinity
compared to their 3‐carboxamide counterparts (15, 28). The observed decreases in H3R binding affinity
upon isomerization observed for these compounds (13‐17, 26 and 29) were deemed insufficient for
proper photopharmacological application. To address this issue, the core azobenzene‐substituted
scaffold was maintained but the propyl spacer between the ether and piperidine moiety was
constrained using isopropylpiperidinyl and 1,3‐cyclobutyl spacers. Such linker rigidification is a known
strategy to increase the binding affinity of H3R antagonists33,42. Importantly, we reasoned that the
limited degrees of freedom associated with the linker rigidification would hamper the cis isomer from
(partially) readjusting its binding mode in the GPCR protein. Upon linker rigidification, the H3R binding
affinities for 31 and 33 were considerably increased compared to their flexible counterpart 15 (Table 2).
Additionally, a pronounced increase in the shift in H3R affinity between the trans and cis analogs was
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observed. Of note is the lower H3R affinity of stereoisomer 32 compared to 33, which is in accordance
with previously reported data33. Compound 33 outperformed 31 based on having a larger shift in H3R
affinity (11.2‐fold lower, Fig. 3B) upon isomerization and higher absolute H3R affinity. Moving the trans‐
cyclobutyl side‐chain to the meta‐position (34), effectively arriving at a constrained version of key
compound 28, virtually abolished the pKi shift and did not increase the binding affinity. All this led to the
designation of 33 as second key compound (VUF14862).
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Compounds 28 and 33 either gain or lose at least 10‐fold H3R affinity upon illumination (Fig. 3A,
B). Thus, using the same photoswitchable core and by proper side chain substitution in the periphery of
the scaffold, we developed bi‐directional GPCR photoswitchable ligands. Moreover, 28 and 33 show a 10
to 100‐fold H3R selectivity over the H1R and no measurable affinity for H2R and H4R (Table S2). With the
cautious use of 28 in view of its residual H1R affinity, 28 and 33 offer a bi‐directional set of
photopharmacological tools for H3R.
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Table 2: Affinity for the human H3R of photoswitchable azobenzene‐derived H3R ligands
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7
A
B
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9
10
11
12
13
14
15
16
17
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28
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Compound
number
Amide
PSSa ± SEM
pKi trans ±
SEMb
pKi at PSSa ±
Scaffold
R
pKi shiftc
n
position (area % cis)
SEMb
Histamine
‐
7.89 ± 0.05
‐
‐
‐
12
3
1
‐
8.29 ± 0.01
‐
d
12
25
17
30
A
B
A
B
‐
‐
96.6 ± 0.33
93.2 ± 0.29 6.42 ± 0.11e 7.04 ± 0.09
7.33 ± 0.11
8.06 ± 0.06
0.74
0.62
5
6
3
6
d
4
4
88.2 ± 2.5
88.9 ± 1.3
7.96 ± 0.06
6.01 ± 0.13
7.36 ± 0.02 −0.59
6.69 ± 0.15 0.68
16
29
A
B
4
4
86.6 ± 2.5
91.3 ± 1.4
6.92 ± 0.12
6.24 ± 0.02
6.56 ± 0.06 −0.36
5
3
5.96 ± 0.06 −0.28
7.26 ± 0.08 −0.63
15
28
A
B
3
3
95.4 ± 0.50
86.8 ± 0.63
7.90 ± 0.07
6.19 ± 0.11
5
3
7.32 ± 0.04
5.85 ± 0.14 −0.30
6.38 ± 0.01 0.19
1.13
14
27
A
B
3
3
90.6 ± 0.77
89.3 ± 0.50
6.16 ± 0.06
6.19 ± 0.02
4
4
13
26
A
B
2
2
88.9 ± 2.2
88.4 ± 1.8
7.97 ± 0.13
7.29 ± 0.08
7.55 ± 0.10 −0.41
7.09 ± 0.05 −0.19
4
4
31
A
3
90.5 ± 1.3
8.39 ± 0.09
7.40 ± 0.15 −0.98
5
32
33
A
A
3
3
96.8 ± 0.21
94.6 ± 1.8
7.10 ± 0.12
8.76 ± 0.09
6.48 ± 0.03 −0.62
7.71 ± 0.09 −1.05
3
4
34
B
3
89.3 ± 0.40
7.25 ± 0.04
7.15 ± 0.07 ‐0.10
4
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aPhotostationary state area percentages after illumination at 10 mM in DMSO‐d6 and as determined by LCMS analysis at 254
nm. bMeasured by displacement of [3H]‐N‐methylhistamine on HEK293T cell homogenates transiently expressing the H3R.
cDefined as pKi PSS cis – pKi trans. dUnsubstituted phenyl. eTrans sample contained 3.9 area % cis compound.
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Binding mode characterization
Molecular modeling studies were performed for tool compounds 28 and 33 in order to gain
insights into the molecular mechanism of the observed affinity changes upon isomerization. Molecular
docking of 28 and 33 into a series of H3R homology models yielded three potential binding modes in
which the ligands form either an ionic interaction with the D1143.32 or E2055.46 residue in transmembrane
helices (TM) 3 and 5, respectively24,43,44. The predicted binding modes share similarity with the binding
orientation of ergotamine as observed in crystal structures of serotonin receptors 5‐HT1B and 5‐HT2B
(PDB IDs: 4IAR, 4IB4)45,46, aminergic GPCRs that share 54‐62% sequence similarity for the binding site
with H3R (Fig. S22). Each of the 12 resulting H3R‐ligand complexes (2 ligands x 2 isomers x 3 binding
modes) was investigated using 50 ns molecular dynamics (MD) simulations. The MD trajectories
indicated that only for one binding mode (Fig. S22A) i) a consistent interaction fingerprint (IFP)47 was
observed throughout the simulation, and ii) the stable ligand binding mode was in line with the
observed SAR data (Table 2). In this binding mode (Fig. 3C, D) the ligands form a stable salt bridge with
D1143.32 and move upward along TMs 2, 3, and 7 into the extracellular vestibule (ECV). The trans isomer
of 33 adopts an elongated binding mode and makes multiple aromatic stacking interactions
with Y912.61, Y942.64, Y3947.35, and F3987.39 while the carboxamide substituent is solvent exposed between
the extracellular loops 1 and 2 (Fig. 3D). The trans isomer of 28, on the other hand, adopts an arched
binding mode within the extracellular vestibule (Fig. 3C) due to its meta‐substitution. The binding modes
of the trans isomers of both 28 and 33 are similar to the previously proposed binding modes for bitopic
ligands of the aminergic dopamine D2 receptor48. The cis isomers of both 28 and 33 adopt an almost
identical binding mode (Fig. 3C, D) in which the carboxamide is buried in the ECV in close proximity to
Y3947.35, folded against the azobenzene, and located at the same position as the pyrrolidine moiety of
ergotamine bound to 5‐HT1B and 5‐HT2B crystal structures (Fig. S23).
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The opposite effects of trans to cis isomerization on the H3R binding affinities of 28 and 33 can
be explained by differences in: i) the complementarity of polar or apolar surface areas of the ligand and
the protein binding site49, and ii) the loss of conformational degrees of freedom as the result of ligand
binding (conformer focusing50). Both isomers of 33 are expected to be less affected by conformer
focusing due to their rigid cyclobutyl‐spacer compared to the flexible propyl spacer of 28. The decreased
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affinity of the trans isomer versus the cis isomer of 28 (Table 2) can be explained by the fact that the
apolar surface area of 28 is >4 times more solvent exposed in the trans isomer binding mode than in the
cis isomer binding mode in H3R. Conversely, the trans isomer of 33 has a reduced buried polar surface
area compared to the folded cis isomer, while the apolar surface area of 33 is able to form a
complementary fit with the apolar surface area of the H3R binding site. In addition, the extended trans
isomer of 33 allows more degrees of conformational freedom than the folded cis isomer, altogether
resulting in an increased H3R affinity for the trans isomer of 33 (Table 2).
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Figure 3: (A‐B) Representative curves of [3H]‐N‐methylhistamine binding displacement by increasing concentrations (A) 28
or (B) 33. The curve indicated as PSS cis consists of predominantly (>90%) cis compound. (C‐D) The proposed binding modes
of (C) 28 and (D) 33 in an H3R homology model based on representative MD snapshots. The cyan and magenta carbon atoms
of the ligands correspond to the trans and cis isomer respectively and for clarity purposes parts of ECL1/TM3 and ECL2 are
not shown.
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Dynamic photochemical modulation of H3R
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As the new azobenzene‐containing H3R photoswitches can be dynamically and reversibly isomerized
with light, we applied the two key H3R photoswitches in a dynamic intact cell system with a real‐time
readout. Two‐electrode voltage clamp (TEVC) using Xenopus laevis oocytes co‐expressing H3R and a GIRK
channel composed of human GIRK1 (Kir3.1) and GIRK4 (Kir3.4) subunits (Fig. 4A)51 allows for a real‐time
H3R readout. In all experiments, H3R‐expressing oocytes were used that evoked a high current (>500 nA)
upon histamine superfusion in 64 mM NaCl, 25 mM KCl, 15 mM HEPES, 0.8 mM MgCl2 and 0.4 mM CaCl2
pH 7.4 at a membrane potential of ‐80 mV. Under these conditions, histamine induced GIRK channel
activation with a pEC50 value of 7.3 ± 0.06 (Fig. 4A). No currents were observed after histamine
application to non‐injected oocytes (Fig. S24). The GIRK channel activation by histamine (1 M), could be
fully inhibited upon simultaneous superfusion with the H3R antagonist clobenpropit (1 M) or by
pretreating the oocytes with the Gi‐protein inhibitor pertussis toxin (1.37 ng/oocyte) (Fig. S25 and
S26). This shows that the histamine‐induced current in Xenopus laevis oocytes co‐expressing H3R and a
GIRK channel composed of human GIRK1 (Kir3.1) and GIRK4 (Kir3.4) subunits is H3R mediated through
Gi‐protein coupling to GIRK.
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We first ensured that photoisomerization behavior of 28 and 33 in HEPES pH 7.4 buffer was
similar to behavior in 50 mM Tris‐HCl pH 7.4 buffer (Fig. S4‐S15). Next, we superfused Xenopus oocytes
expressing H3R and a GIRK channel with either trans‐28 or trans‐33 and histamine for dynamic receptor
inhibition studies. Superfusion of only 28 or 33 on both non‐injected and H3R‐GIRK expressing oocytes
did not result in changes in current, confirming that these H3R ligands act as antagonists (Fig S27 – S30).
Illumination directly on the oocytes (Fig. 4A) in the absence of 28 or 33 did not change the magnitude of
histamine evoked currents (Fig. S31). Yet, in the presence of either 33 or 28, clear photoswitchable
antagonism of the histamine‐induced current was observed under alternating illumination. We
confirmed that both compounds were able to show this behaviour in a concentration dependent fashion
(Fig. S32, S33), yielding Ki values for 33 that nicely match the observed Ki values, obtained in the
radioligand binding experiments (TEVC pKi >99% trans: 8.72 ± 0.11, PSS cis: 7.98 ± 0.15, radioligand
binding: pKi >99% trans: 8.76 ± 0.09, PSS cis: 7.71 ± 0.09). An increase in histamine‐induced GIRK
activation was observed when trans‐33 was illuminated with 360 ± 25 nm, confirming that the H3R
antagonistic effect of 33 is reduced upon isomerization to its cis isomer. When subsequently the
superfused oocytes were illuminated at 430 ± 17 nm, a decrease in H3R‐induced GIRK activation was
observed due to a stronger H3R antagonistic effect by the trans isomer of 33 as a consequence of
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increased H3R binding affinity. This effect could be repeated several times in subsequent illumination
cycles in the same experiment (Fig. 4B). For 28, the reciprocal light‐induced H3R affinity shift compared
to 33 translated well to a reciprocal effect upon illumination in the TEVC setup (Fig. 4C). That is, 28
reached the highest H3R inhibition, measured as reduced histamine‐induced GIRK channel activation,
upon illumination with 360 ± 25 nm. Also, 28 could be effectively switched back in the TEVC setup upon
illumination with 430 ± 17 nm, and again the GPCR blocking properties could be modulated by
alternating illumination (Fig. 4C). These experiments demonstrate the rapid bi‐directional modulation of
the H3R using the two complementary photoswitchable ligands 28 and 33.
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Figure 4: (A) Schematic drawing of the TEVC setup used for dynamic H3R and GIRK current (in)activation and concentration‐
response curve of histamine evoked currents in Xenopus oocytes expressing H3R and GIRK. The insert shows the current
increase upon continuous histamine (1 M) perfusion in time. (B) Representative part of a GIRK‐mediated current trace
during continuous perfusion with 5 M histamine in competition with 1 M 33 under illumination of the Xenopus oocyte
with alternating 360 ± 25 and 430 ± 17 nm wavelength as measured by TEVC. An extended time trace is available in Figure
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S34. (C) Representative part of a GIRK mediated current trace during continuous perfusion with 5 M histamine in
competition with 1 M 28 and illumination of the oocyte with alternating 360 ± 25 and 430 ± 17 nm wavelength as measured
by TEVC. An extended time trace is available in Figure S35.
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A robust bi‐directional photochemical toolbox for dynamic antagonism of the H3R has been successfully
developed, using a core‐centered approach replacing a naphthalene moiety in a known scaffold by a
properly substituted azobenzene unit. SAR exploration delivered 16 compounds that show a shift in
binding affinity for the H3R upon photoisomerization. Two key compounds, 28 and 33, were selected
based on their 13.5‐fold increase or 11.2‐fold decrease in H3R binding affinity upon photoisomerization,
respectively. The long thermal relaxation half‐lives and resistance to fatigue of these compounds
allowed in‐depth spectroscopic analysis (NMR, LC‐MS, UV‐Vis) of the photochemical properties. The
rapid photochemical isomerization directly translated to dynamic, light‐modulated H3R blockade in real‐
time electrophysiology experiments using H3R and GIRK co‐expressing Xenopus oocytes. In this dynamic
experimental setup, histamine‐induced GPCR activation could be modulated on a time scale of seconds
and we were able to show real‐time, light‐sensitive blockade of the H3R activation by both new tool
compounds. In our view 28 (VUF14738) and 33 (VUF14862) are highly useful photopharmacological
tools for temporal studies to dissect the complex signaling cascade of H3R. Moreover, the presented bi‐
directional modulation of H3R protein by closely related photosensitive analogs of the same core
scaffold further emphasizes the exceptional opportunities of conformational modulation of the GPCR
protein family.
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Experimental section
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Synthesis and characterization of compounds
All starting materials were obtained from commercial suppliers (primarily being Sigma‐Aldrich, Acros Organics, Fluorochem and
Strem Chemicals) and used without purification. 4‐hydroxyazobenzene (8) was obtained from Sigma‐Aldrich. Cis‐3‐piperidin‐1‐
ylcyclobutanol (35) was synthesized according to the literature procedure of Wijtmans et al.52. Anhydrous THF and DCM were
obtained by passing it through an activated alumina column prior to use. Anhydrous toluene was freshly distilled over CaH2, or
dried over activated 4Å molecular sieves, which were added at least 24 hrs before use. Anhydrous DMF was purchased from
Acros Organics (Geel, Belgium) and used without prior purification. All reactions were carried out under nitrogen atmosphere
unless mentioned otherwise. TLC analyses were performed using Merck F254 aluminum‐backed silica plates and visualized 254
nm UV‐light or a potassium permanganate stain. Flash column chromatography was executed using Silicycle Siliaflash F60 silica
gel, or by means of a Biotage® Isolera equipment using Biotage® SNAP columns. All HRMS spectra were recorded on a Bruker
micrOTOF mass spectrometer using ESI in positive ion mode. All NMR spectra were recorded on either a Bruker Avance 250,
Bruker Avance 400, Bruker Avance 500 or Bruker Avance 600 spectrometer. The peak multiplicities are defined as follows: s,
singlet; d, doublet; t, triplet; q, quartet; p, pentet; dd, doublet of doublets; dt, doublet of triplets; td, triplet of doublets; bs,
broad singlet; m, multiplet. The spectra were referenced to the internal solvent peak as follows: CDCl3 (1H = 7.26 ppm, 13C =
77.16 ppm), DMSO‐d6 (1H = 2.50 ppm, 13C = 39.52 ppm), CD3OD (1H = 3.31 ppm, 13C = 49.00 ppm)53. IUPAC names were adapted
from ChemBioDraw Ultra 14.0 (Perkin‐Elmer). Purities were measured with the aid of analytical LC‐MS using a Shimadzu LC‐
20AD liquid chromatography pump system with a Shimadzu SPDM20A diode array detector with the MS detection performed
with a Shimadzu LCMS‐2010EV mass spectrometer operating in both positive and negative ionization mode. The column used
was an Xbridge (C18) 5 μm column (50 mm × 4.6 mm or 100 mm × 4.6 mm). The following solutions are used for the eluents.
Solvent A: water: formic acid 999:1 and solvent B: acetonitrile: formic acid 999:1. The eluent program used is as follows: flow
rate: 1.0 mL/min, start 95% A in a linear gradient to 10% A over 4.5 min, hold 1.5 min at 10% A, in 0.5 min in a linear gradient to
95% A, hold 1.5 min at 95% A, total runtime: 8.0 min. Compound purities were calculated as the percentage peak area of the
analyzed compound by UV detection at 254 nm. All chemistry and analyses of photosensitive compounds were carried out
under dimmed or red light.
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Detailed chemical experimental procedures and chemical analyses are supplied in the Supporting Information.
Photochemistry
UV‐Vis spectra were obtained using a Thermo‐scientific Evolution 201 PC spectrophotometer or a Shimadzu UV‐2401 PC
spectrophotometer. Fits of UV‐Vis spectroscopy data were generated using Mathworks Matlab R2014A (8.3.0.532). Illumination
was executed using a Sutter instruments Lambda LS with a 300 Watt full‐spectrum lamp connected to a Sutter instruments
Lambda 10‐3 optical filter changer equipped with 430 ± 17 nm and 360 ± 25 nm filters. For photochemical analyses
illuminations were performed in Hellma Suprasil™ quartz 114‐QS cuvettes. Thermal relaxation experiments and Arrhenius
extrapolations were performed according to Priimagi et al.34 using a compound concentration of 25 µM in 50 mM Tris‐HCl pH
7.4 buffer + 1% DMSO‐d6 and temperatures of 60 oC, 70 oC and 80 oC. Illuminations for pharmacological experiments were
performed in cylindrical clear glass vials with a volume of 4.5 mL. The typical distance between light source and vial or cuvette
was 2 cm. In illuminations during TVEC experiments the light‐source was positioned at 5 cm from the chamber containing the
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oocyte. The focused beam of the light source has a diameter of 1.8 cm and the beam was pointed such that it illuminated the
full oocyte. The light intensity on the oocyte is 0.77 mW/mm2 using the 360 ± 25 nm filter and 0.57 mW/mm2 for the 430 ± 17
nm filter as measured using a Thorlabs PM16‐401 power meter.
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Computational chemistry
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Homology model construction. The sequence of H3R was obtained from UniProt (accession code Q9Y5N1) and aligned to the
unique sequences of crystallized aminergic GPCRs (PDB accessed at 3 Nov. 2015). The sequence similarity and identity were
assessed for the full sequence, each TM helix, extracellular loop 2, and the binding pocket as previously defined23. Based on this
analysis the sequence alignment of H3R to H1R, M3R, D3R, 5‐HT1BR and 5‐HT2BR and their crystal structures (PDB‐codes 3RZE,
4U15, 3PBL, 4IAR, 4IB4) was used as input for MODELLER (version 9.15)54. 500 different homology models were constructed
from which 36 were selected based on visual inspection. Compounds trans‐28, cis‐28, trans‐33, and cis‐33 were subsequently
docked into each model using PLANTS55 (settings: speed 1, scoring function ChemPLP, 10‐fold docking, 50 proposed poses per
fold) into each of the models while treating the side chains of E2065.46 and F3987.39 as flexible. The resulting 26,000 docking
poses were filtered using IFP47 on either and ionic and H‐bond interaction with D1143.32 or E2065.46 from which the top 5
ChemPLP‐scored poses were kept for further processing yielding 396 H3R‐ligand complexes. Visual inspection highlighted 3
clusters of potential binding modes. From each ligand in each binding mode cluster one model was selected for further
investigation with molecular dynamics simulations.
Molecular dynamics simulations. Each ligand‐H3R complex was aligned to the membrane‐aligned H1R structure from the OPM
database. A sodium ion was added in the sodium binding site based on high‐resolution sodium‐bound 1‐adrenoceptor crystal
structure (PDB‐code 4BVN). Subsequently the rotamers from the sodium binding site residues were manually
optimized56. The H3R‐ligand complexes were embedded in a pre‐equilibrated box containing a lipid bilayer (134 POPC lipids),
explicit solvent (~11,000 water molecules) and a 0.15 M concentration of NaCl (~70 ions). Using MOE (v2015.1001; Chemical
Computing Group Inc., Montreal, Quebec, Canada) the ligand‐H3R complex was solvated, and waters between the receptor and
the membrane and outside the receptor were removed. The ligands were parameterized for GAFF using Amber's antechamber
with AM1‐BCC partial charges. The resulting parameters of the ligand were checked, and the azobenzene parameters were
optimized in line with the parameters reported by Schäfer et al.57. The AMBER99SB*‐ILDN force field was used for the protein
and the lipids and waters were described according to the Berger lipid parameters and the TIP3P model, respectively58. After
neutralizing the system, a 1000‐step steepest descent minimization of the system was performed and after which the complex
was equilibrated for 75 and 300 ps using a 1 and 2 fs time step, respectively. A subsequent production run of 50 ns as an NVT
ensemble was performed for each combination. The trajectories were analyzed using IFP47 resulting in a time‐based interaction
profile for each H3R‐ligand complex. These MD simulations were performed using GROMACS 5.159.
Pharmacology
General. Within this paper, the 445‐isoform of the human H3R is referred to as wild‐type receptor. [3H]‐Mepyramine (specific
activity 20 Ci/mmol‐1), [125I]‐Iodoaminopotentidine (specific activity 2200 Ci/mmol‐1), [3H]‐N‐methylhistamine (specific activity
78.3 Ci/mmol‐1) and [3H]‐histamine (specific activity 17.5 Ci/mmol‐1) were purchased from PerkinElmer (Groningen, the
Netherlands). All other chemicals were of analytical grade and obtained from commercial sources.
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Cell culture & transfection. Human embryonic kidney 293T cells (HEK293T cells) were cultured in Dulbecco’s modified eagle
medium (DMEM) supplemented with 10% fetal bovine serum (FBS), penicillin (50 IU/mL) and streptomycin (50 g/mL).
HEK293T cells were transiently transfected with H3R cDNA using the polyethylenimine (PEI) method. In brief, 24 hrs prior to
transfection 2∙106 cells were seeded on a 10 cm dish. Cells were transfected with 5 g H3R cDNA and 20 g PEI in 500 L 150
mM NaCl‐solution. DNA‐PEI mixture was incubated for 15‐30 min at 22 °C before dropwise addition to the cells. Cells were
maintained for 48 hrs at 37 °C with 5% CO2 before harvesting.
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Preparation of cell homogenates. 48 hrs after transfection cells were detached and collected from the plates by vigorous
pipetting using phosphate buffered saline (PBS). Cells were centrifuged at 3000 rpm for 10 min and cell‐pellet was stored at ‐20
°C until the day of experiment.
Radioligand binding assay. For radioligand binding experiments samples were pretreated to obtain trans and cis samples. In
brief, 1 mM ligand solution was divided in two samples which were either illuminated at 360 ± 25 nm until their
photostationary state was reached as analyzed by LC‐MS, or kept in the dark for the same duration as the illuminated samples.
All subsequent handling was performed in the dark or under near‐infrared light. [3H]‐N‐methylhistamine binding assays were
performed in a total volume of 100 l consisting of cold ligand (increasing concentration ranging from 10‐5 M to 10‐12 M)
prepared in assay buffer [50 mM Tris‐HCl pH 7.4], 1.6 nM [3H]‐N‐methylhistamine and HEK293T cell homogenates transiently
expressing the human H3R (>25 g/50 l). The mixture was incubated for 2 hrs at 22 °C before termination by rapid filtration
over a 96 well GF/C filter plate pre‐coated with 0.5% PEI using Perkin Elmer 96‐well filtermate‐harvester (Perkin Elmer). The
filter was washed 5 times with ice‐cold wash buffer [50mM Tris‐HCl pH 7.4, 4 °C]. 5 hrs after addition of Microscint O
scintillation liquid filter‐bound radioactivity was measured using a microbeta wallac trilux scintillation counter (Perkin Elmer).
Binding selectivity screening. H1R radioligand binding assays were performed according to Kuhne et al.60 using an incubation
time of 2 hours instead of 4 hours. H2R radioligand binding assays were performed according to Leurs et al.61 using HEK293T
cells instead of CHO cells and a total volume of 400 L instead of 200 L. H4R radioligand binding assays were performed
according to Nijmeijer et al.62
Data analysis. All experiments were analyzed using Graphpad prism 6.05 (Graphpad software inc., San Diego, USA) IC50 values
obtained from competition binding experiments were converted to Ki values using the Cheng‐Prusoff equation63.
Two electrode voltage clamp (TEVC) assay
RNA synthesis. The Kir3.1 and Kir3.4 were both supplied in the pcDNA3.1 vector and were a kind gift of K. Sahlholm (Karolinska
Institute, Stockholm, Sweden). The H3R was cloned into the pCIneo vector. Restriction enzymes BAMHI and NdeI were used to
linearize the constructs pCIneo‐H3R and pcDNA3.1‐Kir3.1 and 3.4, respectively. After the digestion, 0.5% SDS and Proteinase K
(200 µg/mL) were added and the mixture was incubated at 50 oC for 30 min. DNA was extracted by phenol/chloroform and the
precipitation method. The quality and digestion was checked on 1% agarose gel. The cRNA synthesis was performed with the T7
mMessage mMachine kit (Ambion, Austin, TX, USA). After the synthesis, the cRNA quality and amount was checked on a
denaturating gel for RNA as described by Almeida et al.64.
H3R and GIRK expression in oocytes. Xenopus laevis oocytes were supplied as “topgrade” oocytes (Ecocyte, Castrop‐Rauxel,
Germany). Each oocyte was injected with a volume of 46 nL, containing 3 ng cRNA of each GIRK subunit and 50 ng cRNA of the
H3R or with 46 nL of RNAse free water as control with the Nanoject II (Drummond Scientific Company, Broomall, PA, USA). The
oocytes were incubated for 4‐6 days at 12 °C in modified Barth solution (88 mM NaCl, 1 mM KCl, 2.4 mM NaHCO3, 15 mM
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HEPES, 0.33 mM Ca(NO3)2, 0.41 mM CaCl2, 0.82 mM MgSO4, 2.5 mg pyruvic acid, 100 g/mL Streptomycin, 50 g/mL
Gentamycin, adjusted with Tris to pH 7.4), as described by Sahlholm et al.51.
Two‐electrode voltage clamp. Two‐electrode voltage clamp (TEVC) was performed with the Axoclamp 900A amplifier and the
Digidata 1550 Digitizer. The currents were recorded and analyzed with pClamp 10.6 (Molecular Devices, Sunnyvale, CA, USA).
Glass micropipettes were pulled from borosilicate capillaries (GC150‐10, Harvard Apparatus, Edenbridge, UK) with the P‐1000
micropipette puller (Sutter Instrument, Novato, CA, USA,), to have a resistance of 1‐3 M when filled with 3 M KCl solution. The
oocyte was positioned in the recording chamber (RC‐1Z, Warner Instruments, Hamden, CT, USA) and by gravity flow, high
potassium solution (64 mM NaCl, 25 mM KCl, 0.8 mM MgCl2, 0.4 mM CaCl2, 15 mM HEPES and adjusted with Bis‐Tris Propane to
pH 7.4) was perfused through the recording chamber. For the study of H3R ligands, the membrane potential was clamped at ‐80
mV and the evoked currents were measured at room temperature (~20 °C). The ligands (histamine, clobenpropit, 28 and 33)
were diluted, in the high potassium solution to obtain the appropriate concentrations. For the photoswitchable ligands (28 and
33), the TEVC was performed in the dark and the different wavelengths (360 ± 25 nm and 430 ± 17 nm) were applied directly on
the oocyte in the recording chamber for 1 min (with 28) or 2 min (with 33) with a Sutter instruments Lambda LS with a 300 watt
full spectrum lamp connected to a Sutter instruments Lambda 10‐3 optical filter changer equipped with 430 ± 17 nm and 360 ±
25 nm filters to observe the (functional) switch of the ligands. For the Pertussis toxin experiment, 46 nL (1.38 ng) of PTX or
water as control was injected in H3R/GIRK expressing oocytes (4‐6 days after cRNA injection) with the Nanoject II The oocytes
were incubated for another 7 hrs before measuring the currents evoked by 1 M histamine to the oocytes.
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Associated content
Detailed synthetic procedures, chemical analyses and spectra, and additional data as noted in text.
Acknowledgements
All authors acknowledge the Netherlands Organisation for Scientific Research (NWO) for financial
support (TOPPUNT, “7 ways to 7TMR modulation (7‐to‐7)”, 718.014.002). EWEV is supported by NWO
ECHO project 711.013.014. All authors participate in the European Cooperation in Science and
Technology Action CM1207 [GPCR‐Ligand Interactions, Structures, and Transmembrane Signaling: A
European Research Network (GLISTEN)]. We thank Hans Custers for HRMS analyses, Yara Huppelschoten
for determining the selectivity of the key compounds amongst the histamine receptor family, and
Andrea Van de Stolpe and Ken Lion for setting up the photochemical equipment. Kristoffer Sahlholm
(Karolinska institute) is kindly acknowledged for providing the pcDNA3.1‐Kir3.1 and ‐Kir3.4 plasmids.
Competing financial interests
The authors declare no competing financial interests.
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References
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6
7
8
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(1)
(2)
Broichhagen, J.; Frank, J. A.; Trauner, D. Acc. Chem. Res. 2015, 48, 1947.
Lerch, M. M.; Hansen, M. J.; van Dam, G. M.; Szymanski, W.; Feringa, B. L. Angew. Chem. Int. Ed. Engl. 2016,
55, 10978
(3)
(4)
(5)
(6)
(7)
Szobota, S.; Isacoff, E. Y. Annu. Rev. Biophys. 2010, 39, 329.
Beharry, A. A.; Woolley, G. A. Chem. Soc. Rev. 2011, 40, 4422.
Szymanski, W.; Beierle, J. M.; Kistemaker, H. A.; Velema, W. A.; Feringa, B. L. Chem. Rev. 2013, 113, 6114.
Velema, W. A.; Szymanski, W.; Feringa, B. L. J Am Chem Soc 2014, 136, 2178.
Stein, M.; Middendorp, S. J.; Carta, V.; Pejo, E.; Raines, D. E.; Forman, S. A.; Sigel, E.; Trauner, D. Angew.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Chem. Int. Ed. Engl. 2012, 51, 10500.
(8)
(9)
Frank, J. A.; Moroni, M.; Moshourab, R.; Sumser, M.; Lewin, G. R.; Trauner, D. Nat. Commun. 2015, 6, 7118.
Broichhagen, J.; Schonberger, M.; Cork, S. C.; Frank, J. A.; Marchetti, P.; Bugliani, M.; Shapiro, A. M.; Trapp,
S.; Rutter, G. A.; Hodson, D. J.; Trauner, D. Nat. Commun. 2014, 5, 5116.
(10)
Pittolo, S.; Gomez‐Santacana, X.; Eckelt, K.; Rovira, X.; Dalton, J.; Goudet, C.; Pin, J. P.; Llobet, A.; Giraldo, J.;
Llebaria, A.; Gorostiza, P. Nat. Chem. Biol. 2014, 10, 813.
(11)
(12)
Schönberger, M.; Trauner, D. Angew. Chem. Int. Ed. Engl. 2014, 53, 3264.
Bahamonde, M. I.; Taura, J.; Paoletta, S.; Gakh, A. A.; Chakraborty, S.; Hernando, J.; Fernandez‐Duenas, V.;
Jacobson, K. A.; Gorostiza, P.; Ciruela, F. Bioconjug. Chem. 2014, 25, 1847.
(13)
Broichhagen, J.; Johnston, N. R.; von Ohlen, Y.; Meyer‐Berg, H.; Jones, B. J.; Bloom, S. R.; Rutter, G. A.;
Trauner, D.; Hodson, D. J. Angew. Chem. Int. Ed. Engl. 2016, 55, 5865.
(14)
Westphal, M.; Schafroth, M. A.; Sarott, R.; Imhof, M.; Bold, C.; Leippe, P.; Dhopeshwarkar, A.; Grandner, J.;
Katritch, V.; Mackie, K.; Trauner, D.; Carreira, E. M.; Frank, J. A. J Am Chem Soc 2017.
(15)
(16)
(17)
Haas, H.; Panula, P. Nat. Rev. Neurosci. 2003, 4, 121.
Leurs, R.; Bakker, R. A.; Timmerman, H.; de Esch, I. J. P. Nat. Rev. Drug. Discov. 2005, 4, 107.
Panula, P.; Chazot, P. L.; Cowart, M.; Gutzmer, R.; Leurs, R.; Liu, W. L.; Stark, H.; Thurmond, R. L.; Haas, H. L.
Pharmacol. Rev. 2015, 67, 601.
(18)
(19)
(20)
(21)
Schwartz, J. C. Brit. J. Pharmacol. 2011, 163, 713.
Lebois, E. P.; Jones, C. K.; Lindsley, C. W. Curr. Top. Med. Chem. 2011, 11, 648.
Leurs, R.; Vischer, H. F.; Wijtmans, M.; de Esch, I. J. P. Trends Pharmacol. Sci. 2011, 32, 250.
Celanire, S.; Wijtmans, M.; Talaga, P.; Leurs, R.; de Esch, I. J. Drug Discov Today 2005, 10, 1613.
Uveges, A. J.; Kowal, D.; Zhang, Y.; Spangler, T. B.; Dunlop, J.; Semus, S.; Jones, P. G. J. Pharmacol. Exp. Ther.
(22)
2002, 301, 451.
(23)
Kooistra, A. J.; Kuhne, S.; de Esch, I. J.; Leurs, R.; de Graaf, C. Br. J. Pharmacol. 2013, 170, 101.
Axe, F. U.; Bembenek, S. D.; Szalma, S. J. Mol. Graph. Model. 2006, 24, 456.
Lorenzi, S.; Mor, M.; Bordi, F.; Rivara, S.; Rivara, M.; Morini, G.; Bertoni, S.; Ballabeni, V.; Barocelli, E.; Plazzi,
(24)
(25)
P. V. Bioorg. Med. Chem. 2005, 13, 5647.
(26)
(27)
Kim, S.‐K.; Fristrup, P.; Abrol, R.; Goddard, W. A. J. Chem. Inf. Model. 2011, 51, 3262.
Sander, K.; von Coburg, Y.; Camelin, J. C.; Ligneau, X.; Rau, O.; Schubert‐Zsilavecz, M.; Schwartz, J. C.; Stark,
H. Bioorg Med Chem Lett 2010, 20, 1581.
(28)
(29)
Roche, O.; Nettekoven, M.; Vifian, W.; Sarmiento, R. M. Bioorg. Med. Chem. Lett. 2008, 18, 4377.
Arrang, J. M.; Garbarg, M.; Lancelot, J. C.; Lecomte, J. M.; Pollard, H.; Robba, M.; Schunack, W.; Schwartz, J.
C. Nature 1987, 327, 117.
(30)
Alves‐Rodrigues, A.; Leurs, R.; Wu, T. S.; Prell, G. D.; Foged, C.; Timmerman, H. Br J Pharmacol 1996, 118,
2045.
(31)
Nieto‐Alamilla, G.; Marquez‐Gomez, R.; Garcia‐Galvez, A. M.; Morales‐Figueroa, G. E.; Arias‐Montano, J. A.
Mol Pharmacol 2016, 90, 649.
(32)
(33)
Wu, X.; Fors, B. P.; Buchwald, S. L. Angew. Chem. Int. Ed. Engl. 2011, 50, 9943.
Wijtmans, M.; Denonne, F.; Celanire, S.; Gillard, M.; Hulscher, S.; Delaunoy, C.; Van Houtvin, N.; Bakker, R.
A.; Defays, S.; Gerard, J.; Grooters, L.; Hubert, D.; Timmerman, H.; Leurs, R.; Talaga, P.; de Esch, I. J. P.; Provins, L.
Medchemcomm 2010, 1, 39.
(34)
(35)
(36)
Ahmed, Z.; Siiskonen, A.; Virkki, M.; Priimagi, A. Chem Commun 2017, 53, 12520.
Bleger, D.; Hecht, S. Angew. Chem. Int. Ed. Engl. 2015, 54, 11338.
Dong, M.; Babalhavaeji, A.; Hansen, M. J.; Kalman, L.; Woolley, G. A. Chem. Commun. (Camb) 2015, 51,
12981.
(37)
Talaty, E. R.; Fargo, J. C. Chem Commun 1967, 65.
23
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Page 24 of 25
1
2
3
4
(38)
(39)
Garcia‐Amoros, J.; Velasco, D. Beilstein J Org Chem 2012, 8, 1003.
Asano, T.; Okada, T.; Shinkai, S.; Shigematsu, K.; Kusano, Y.; Manabe, O. Journal of the American Chemical
Society 1981, 103, 5161.
5
(40)
(41)
(42)
Serra, F.; Terentjev, E. M. Macromolecules 2008, 41, 981.
Whitten, D. G.; Wildes, P. D.; Pacifici, J. G.; Irick, G. J. Am. Chem. Soc. 1971, 93, 2004.
Dvorak, C. A.; Apodaca, R.; Barbier, A. J.; Berridge, C. W.; Wilson, S. J.; Boggs, J. D.; Xiao, W.; Lovenberg, T.
6
7
8
9
W.; Carruthers, N. I. J. Med. Chem. 2005, 48, 2229.
(43)
(44)
Rai, B. K.; Tawa, G. J.; Katz, A. H.; Humblet, C. Proteins 2010, 78, 457.
Levoin, N.; Labeeuw, O.; Krief, S.; Calmels, T.; Poupardin‐Olivier, O.; Berrebi‐Bertrand, I.; Lecomte, J. M.;
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Schwartz, J. C.; Capet, M. Bioorg. Med. Chem. 2013, 21, 4526.
(45)
Wang, C.; Jiang, Y.; Ma, J.; Wu, H.; Wacker, D.; Katritch, V.; Han, G. W.; Liu, W.; Huang, X. P.; Vardy, E.;
McCorvy, J. D.; Gao, X.; Zhou, X. E.; Melcher, K.; Zhang, C.; Bai, F.; Yang, H.; Yang, L.; Jiang, H.; Roth, B. L.; Cherezov, V.; Stevens,
R. C.; Xu, H. E. Science 2013, 340, 610.
(46)
Wacker, D.; Wang, C.; Katritch, V.; Han, G. W.; Huang, X. P.; Vardy, E.; McCorvy, J. D.; Jiang, Y.; Chu, M.; Siu,
F. Y.; Liu, W.; Xu, H. E.; Cherezov, V.; Roth, B. L.; Stevens, R. C. Science 2013, 340, 615.
(47)
(48)
Marcou, G.; Rognan, D. J. Chem. Inf. Model. 2007, 47, 195.
Lane, J. R.; Donthamsetti, P.; Shonberg, J.; Draper‐Joyce, C. J.; Dentry, S.; Michino, M.; Shi, L.; Lopez, L.;
Scammells, P. J.; Capuano, B.; Sexton, P. M.; Javitch, J. A.; Christopoulos, A. Nat. Chem. Biol. 2014, 10, 745.
(49)
(50)
(51)
(52)
Schmidtke, P.; Luque, F. J.; Murray, J. B.; Barril, X. J. Am. Chem. Soc. 2011, 133, 18903.
Tirado‐Rives, J.; Jorgensen, W. L. J. Med. Chem. 2006, 49, 5880.
Sahlholm, K.; Nilsson, J.; Marcellino, D.; Fuxe, K.; Arhem, P. Eur. J. Pharmacol. 2007, 567, 206.
Wijtmans, M.; Denonne, F.; Célanire, S.; Gillard, M.; Hulscher, S.; Delaunoy, C.; Van houtvin, N.; Bakker, R.
A.; Defays, S.; Gérard, J.; Grooters, L.; Hubert, D.; Timmerman, H.; Leurs, R.; Talaga, P.; de Esch, I.; Provins, L. Med. Chem.
Commun. 2010, 1, 39.
(53)
Fulmer, G. R.; Miller, A. J. M.; Sherden, N. H.; Gottlieb, H. E.; Nudelman, A.; Stoltz, B. M.; Bercaw, J. E.;
Goldberg, K. I. Organometallics 2010, 29, 2176.
(54)
(55)
(56)
Sali, A.; Blundell, T. L. J. Mol. Biol. 1993, 234, 779.
Korb, O.; Stutzle, T.; Exner, T. E. J. Chem. Inf. Model. 2009, 49, 84.
Miller‐Gallacher, J. L.; Nehme, R.; Warne, T.; Edwards, P. C.; Schertler, G. F.; Leslie, A. G.; Tate, C. G. PLoS
One 2014, 9, e92727.
(57)
(58)
(59)
(60)
Schäfer, L. V.; Muller, E. M.; Gaub, H. E.; Grubmuller, H. Angew. Chem. Int. Ed. Engl. 2007, 46, 2232.
Cordomi, A.; Caltabiano, G.; Pardo, L. J. Chem. Theory Comput. 2012, 8, 948.
Berendsen, H. J. C.; van der Spoel, D.; van Drunen, R. Comput. Phys. Commun. 1995, 91, 43.
Kuhne, S.; Kooistra, A. J.; Bosma, R.; Bortolato, A.; Wijtmans, M.; Vischer, H. F.; Mason, J. S.; de Graaf, C.; de
Esch, I. J.; Leurs, R. J Med Chem 2016, 59, 9047.
(61)
(62)
(63)
(64)
Leurs, R.; Smit, M. J.; Menge, W. M.; Timmerman, H. Br J Pharmacol 1994, 112, 847.
Nijmeijer, S.; Vischer, H. F.; Rosethorne, E. M.; Charlton, S. J.; Leurs, R. Mol Pharmacol 2012, 82, 1174.
Yung‐Chi, C.; Prusoff, W. H. Biochem. Pharmacol. 1973, 22, 3099.
Almeida, P.; de Boer, G.‐J.; de Boer, A. H. J. Plant Physiol. 2014, 171, 438.
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