S29434, a quinone reductase 2 inhibitor: Main biochemical and cellular characterization

Article abstract of DOI:10.1124/mol.118.114231

Quinone reductase 2 (QR2, E.C. 1.10.5.1) is an enzyme with a feature that has attracted attention for several decades: in standard conditions, instead of recognizing NAD(P)H as an electron donor, it recognizes putative metabolites of NADH, such as N-methyl- and N-ribosyl-dihydronicotinamide. QR2 has been particularly associated with reactive oxygen species and memory, strongly suggesting a link among QR2 (as a possible key element in pro-oxidation), autophagy, and neurodegeneration. In molecular and cellular pharmacology, understanding physiopathological associations can be difficult because of a lack of specific and powerful tools. Here, we present a thorough description of the potent, nanomolar inhibitor [2-(2-methoxy-5H-1,4b,9-triaza(indeno[2,1-a]inden-10-yl)ethyl]-2-furamide (S29434 or NMDPEF; IC₅₀5 5–16 nM) of QR2 at different organizational levels. We provide full detailed syntheses, describe its cocrystallization with and behavior at QR2 on a millisecond timeline, show that it penetrates cell membranes and inhibits QR2-mediated reactive oxygen species (ROS) production within the 100 nM range, and describe its actions in several in vivo models and lack of actions in various ROS-producing systems. The inhibitor is fairly stable in vivo, penetrates cells, specifically inhibits QR2, and shows activities that suggest a key role for this enzyme in different pathologic conditions, including neurodegenerative diseases.

Full text of DOI:10.1124/mol.118.114231

Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
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Title page
S29434, a quinone reductase 2 inhibitor: main biochemical and cellular characterization
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,
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Jean A. Boutin *, Frederic Bouillaud , Elzbieta Janda , István Gacsalyi **, Gérald
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Guillaumet , Etienne C. Hirsch , Daniel A. Kane , Françoise Nepveu , Karine Reybier ,
Philippe Dupuis , Marc Bertrand , Monivan Chhour , Thierry Le Diguarher , Mathias
Antoine , Karen Brebner , Hervé Da Costa , Pierre Ducrot , Adeline Giganti , Vishalgiri
Goswami , Hala Guedouari , Patrick P. Michel , Aakash Patel , Jérôme Paysant , Johann
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Stojko , Marie-Claude Viaud-Massuard , Gilles Ferry
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Pôle d’Expertise Biotechnologie, Chimie & Biologie, Institut de Recherches SERVIER,
7
8290 Croissy-sur-Seine, France
2
Institut Cochin, INSERM U1016, CNRS-UMR8104, Université Paris Descartes 75014
Paris, France
3
Department of Health Sciences, Magna Graecia University, 88100 Catanzaro, Italy
4
Egis Pharmaceuticals PLC, 1165 Budapest, Hungary.
5
Institut de Chimie Organique et Analytique (ICOA), Université d'Orléans, UMR CNRS
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311, 45067 Orléans Cedex 2, France
6
Institut du Cerveau et de la Moelle épinière, ICM, Inserm U 1127, CNRS UMR 7225,
Sorbonne Université, F-75013, Paris, France
7
Departments of Human Kinetics (DK) and Psychology (KB), St. Francis Xavier University,
Antigonish, NS, Canada
8
UMR 152 Pharma-Dev, Université de Toulouse, IRD, UPS, 31062 Toulouse, France
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EUROFINS-CEREP SA, 86600 Celle L'Evescault, France
1
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0
Technologie Servier, Orléans, France
1
CNRS-UMR 7292 « GICC Innovation Moléculaire et Thérapeutique », Université de
Tours, 37200 Tours, France
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Oxygen Healthcare Pvt Ltd, 382213 Ahmedabad, Gujarat, India
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Pôle d’Innovation Thérapeutique de Cardiologie, Institut de Recherches SERVIER, 92150,
Suresnes, France
*
*
: present address Institut de Recherches Internationales Servier, 92150 – Suresnes
*: present address : ATRC Aurigon Ltd., Budapest, Hungary
1

Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
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Running title page:
Running title: Quinone reductase 2 inhibitor: S29434
Corresponding author:
*
Jean A. Boutin, Institut de Recherches Internationales Servier, 50 rue Carnot, 92284
Suresnes France
Telephone : +33(0)155724400
Email : jean.boutin@servier.com
Abbreviations: BNAH: N-benzyldihydronicotinamide; CLQ: chloroquine;; MPP+: 1-methyl-
4
-phenylpyridinium; MPTP: 1-methyl-4-phényl-1,2,3,6-tetrahydropyridine;; NRH: N-
ribosyldihydronicotinamide; Psf: permeability surface filter ; Pst: permeability surface total;
QR1: NAD(P)H dehydrogenase (quinone 1), EC 1.6.5.2; QR2: quinone reductase 2 or
ribosyldihydronicotinamide dehydrogenase (quinone), formerly E.C. 1.10.99.2 now E.C.
1
.10.5.1; ROS: reactive oxygen species; S29434: [2-(2-methoxy-5H-1,4b,9-triaza(indeno[2,1-
a]inden-10-yl)ethyl]-2-furamide.
•
•
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•
Number of text pages: 31
Number of tables: 1
Number of figures: 13
Number of references: 81
Words in Abstract: 192
Words in Introduction: 677
Words in Discussion: 1491
2

Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
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MOL #114231
Abstract: Quinone reductase 2 (QR2, E.C. 1.10.5.1) is an enzyme with a feature that has
attracted attention for several decades: in standard conditions, instead of recognizing
NAD(P)H as an electron donor, it recognizes putative metabolites of NADH such as N-
methyl- and N-ribosyl-dihydronicotinamide. QR2 has been particularly associated with
reactive oxygen species and memory, strongly suggesting a link among QR2 (as a possible
key element in pro-oxidation), autophagy, and neurodegeneration. In molecular and cellular
pharmacology, understanding physiopathological associations can be difficult because of a
lack of specific and powerful tools. Here we present a thorough description of the potent,
nanomolar inhibitor S29434 (IC = 5 to 16 nM) of QR2 at different organizational levels. We
5
0
provide full detailed syntheses; describe its co-crystallization with and behavior at QR2 on a
millisecond timeline; show that it penetrates cell membranes and inhibits QR2-mediated ROS
production within the 100 nM range; and describe its actions in several in vivo models, and
lack of actions in various ROS-producing systems. The inhibitor is fairly stable in vivo,
penetrates cells, specifically inhibits QR2, and shows activities that suggest a key role for this
enzyme in different pathological conditions including neurodegenerative diseases.
3

Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
This article has not been copyedited and formatted. The final version may differ from this version.
MOL #114231
Introduction
Quinone reductases, described as early as 1954 in peas, are pyridine nucleotide–dependent
enzymes involved in the detoxification of natural quinones (Wosilait and Nason, 1954).
Conover and Ernster (1960) first described the reference quinone reductase enzyme, initially
designated as DT-diaphorase, in mammals. In the 1990s, another quinone reductase was
cloned: quinone reductase 2 (QR2 or NQO2), with an unusual history (Zhao et al., 1997). The
initial description indicated that QR2 was a strict analogue of quinone reductase 1 (a.k.a., DT-
diaphorase, or NAD(P)H dehydrogenase (quinone 1), QR1, NQO1, E.C. 1.6.5.2), with similar
properties. Later findings revealed, however, that it was quite different. Research in the 1960s
identified QR2 as the only reductase using non-canonical co-substrates such as N-ribosyl- and
N-methyl-dihydronicotinamide and para-quinones as substrates (Liao and Williams-Ashman,
1
961; Liao et al., 1962), but not NADH or NADPH (Zhao et al., 1997; Ferry et al., 2010).
Later, Talalay and colleagues cloned and crystallized QR2 (Zhao et al., 1997), but its
physiologic role and specificity were poorly documented and understood, except that it
recognized non-quinone compounds such as CB1954 as a substrate (Wu et al., 1997). This
finding was the basis of efforts to develop CB1954 as a prodrug with interesting cytotoxic
properties (Knox et al., 2003). QR2 has been implicated in effects distantly related to quinone
metabolism, such as antimalarial action (Kwiek et al., 2004; Cassagnes et al., 2017), the
AMPA-signaling pathway (Rappaport et al., 2015), paraquat-mediated toxicity (Janda et al.,
2
013; 2015), oxidative stress sensing (Leung and Shilton, 2013), and oocyte maturation (Chen
et al., 2017).
In the search for the melatonin MT3 binding site, described by Duncan et al. (1988) and
our team (Paul et al., 1999), we found that the binding site was, in fact, in QR2 (Nosjean et
al., 2000), see also discussion in Boutin & Ferry (2018). A screen of about 20,000 compounds
from our library, comprising natural products such as flavonoids (Boutin et al., 2005), added
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Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
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further candidates to the list of QR2 inhibitors. Already on this list were resveratrol
(
Buryanovskyy et al., 2004), tacrine (den Braver-Sewradj et al., 2017), chloroquine (CLQ)
Kwiek et al., 2004), dabigatran (a prescription thrombin inhibitor; Michaelis et al., 2012),
(
and tetracyclic compounds (Boussard et al., 2006). Some products of quinone reduction are
fairly unstable, especially in the absence of conjugating system(s), so the reaction products
(quinols) may spontaneously yield back the original quinones in the presence of oxygen.
Similar quinone cycling was described for QR1 (Bindoli et al., 1990; Nutter et al., 1992; Bian
et al., 2017). Without the cellular conjugating capacities that neutralize the process by
conjugating the quinol (e.g., menadiol) with glucuronic acid (Kappus and Sies, 1981; Bolton
et al., 2000; Nishiyama et al., 2010), the by-product of this futile cycle is the production of
reactive oxygen species (ROS). Because electron paramagnetic resonance spectroscopy (EPR)
can detect ROS generation (Reybier et al., 2011), we used this method to show that under
some conditions, the pro-oxidant property of QR2 can be confirmed and measured.
Many organs and cells express QR2 (Nosjean et al., 2001), where it might have a key
role in several pathological conditions. Given this potential, we assessed the specificity of the
QR2 inhibitor S29434 and its actions in different contexts, adding to the already available
data to consolidate its use as an established QR2 inhibitor tool. The present paper gives the
full description of the characteristics of S29434. Despite several publications on this inhibitor,
no report has described its synthesis, specificity for alternative enzymes or receptors, or
characteristics of its stability in living systems. Thus, we generated as much information as
possible about S29434 to make its use more appropriate for QR2 pharmacology. Here we
cover its most important characteristics, from its chemistry and molecular and cellular
pharmacology to its specificity for other ROS-generating systems and its in vivo activities.
These data illustrate that S29434 or unknown metabolites reach QR2 and inhibit this enzyme
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Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
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in situ. Furthermore, its specificity seems to be limited to QR2, making this compound a
valuable tool for understanding the role of QR2 in multiple scenarios, even in vivo.
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Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
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Materials and Methods
Chemical syntheses. S29434 synthesis has never been published. Two different paths were
explored for its synthesis. Both are summarized in Fig. 1 & 2. The synthetic routes are given
in full details in the supplementary materials, as well as the various steps permitting to obtain
the compounds are detailed in the Supplementary data 1. The final analyses of the product
obtained with the second route are available as Supplemental Data 1 (spectral analyses of
compounds 3, 4 and S29434 obtained by the first synthetic route). All data obtained
(analytical, spectral, biophysical) on the compound were consistent with its structure. S29434
was obtained with purity above 99% as adjudged by LC-MS and NMR (see Supplemntary
analytical Data),. Analytical details of the second route Supplementary Figure S2 is also
given as Supplementary Data 2, 3, 4, 5, 6 and 7.
QR2 inhibition measurements. The potency of S29434 was measured in standard kinetic
assays. Briefly, QR2 enzymatic activity was measured under FAD saturation using substrate
(menadione) and 100 μM co-substrate BNAH or NRH. An oxidoreduction reaction was
performed at 25 °C in a Tris–HCl 50 mM, pH 8.5, FAD 500 nM, octyl-glucopyranoside 1
mM, dimethylsulfoxide (DMSO) 5% buffer. Enzymatic kinetics measured the decrease in co-
substrate absorbance corresponding to its oxidation. This reaction was followed either at 350
nm (BNAH) or 340 nm (NRH) on a FLUOstar 384-well plate reader (BMG, Offenburg,
Germany). The slope of absorbance decrease was determined using FLUOstar Optima
software (BMG) and expressed in units of optic density (UDO).s-1 then in M.s-1 using the
Beer–Lambert law. This measurement was next corrected from its corresponding non-
enzymatic (spontaneous) oxidation rate measured in the absence of enzyme, which
corresponds to the specific maximum-activity oxidoreduction reaction. For the inhibitory
hQR2 enzymatic activity assay, S29434 was used in the range of 50 pM to 5 µM. The
inhibitory concentration 50 % (IC ) and the inhibition percentage of co-substrate oxidation
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Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
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were determined using the PRISM program (GraphPad Software Inc., San Diego, CA, USA).
These experiments were conducted over 50 times (inhibition, Fig. 3A) or in triplicate (Fig.
3
B).
Binding kinetics of S29434 and resveratrol to QR2 FADox (oxidized FAD). Transient
kinetics were measured at 25 °C (S29434) or 4 °C (resveratrol) with a SFM-300 stopped-flow
mixer fitted with a FC-15 cell and coupled to a MOS-200 M rapid spectrophotometer
equipped with a 150 W Xe−Hg lamp (Bio-Logic Science Instruments, Seyssinet-Pariset,
France). Typical mixing dead time was 2.6 ms. QR2 tryptophans were excited at 295 nm to
minimize photobleaching and excitation of tyrosines. Fluorescence emission was recorded
using a 320-nm cutoff filter combined with a UG11 band-pass filter to block S29434
fluorescence. Binding experiments were performed under pseudo-first-order conditions by
mixing an equal volume (75 μl) of QR2 and ligand in Tris–HCl 50 mM, β-octyl-D-
glucopyranoside 1 mM, DMSO 5%, pH 8.5. QR2 monomer concentration after mixing was 1
μM. Fluorescence traces were first analyzed individually with the BioKine software (Bio-
Logic, v4.80) as the sum of up to two exponential terms, as described by equation 1, with n
being the number of exponentials, ai the amplitude of the ith exponential, kobsꢀi the rate
constant of the ith exponential, c the trace end point, and bt accounting for the slow linear
drift caused by photobleaching. Fig. 5 was generated with Prism (GraphPad, v7.03).
n


(
-kobsi t)
y   aie
  bt  c

(1)

i1

The kobs values were plotted against ligand concentration. When a linear increase of kobs
was observed, data were fitted to equation 2 describing a one-step binding mechanism, using
Prism (GraphPad, v7.03).
(2)
k  k [ligand]  k
obs
1
-1
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Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
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Specificity tests. General specificity. Most of the tests were subcontracted to Eurofins. All
experimental details can be found online on their site (www.eurofinsdiscoveryservices.com).
Specificity tests. Kinases. The panel of kinases for screening is available from Eurofins
(ExpressDiversityKinase). It comprises 46 different kinases chosen to cover a maximal
structural diversity among this large family of over 500 different enzymes. S29434 was tested
at 100 nM on the enzymatic activity of all the kinases using a standard radiometric assay.
Specificity tests Sirtuin experiments. The use of this original technology has been reported in
a technical note by Agilent (see https://www.agilent.com/cs/library/applications/5990-
9
345en_lo.pdf). In brief, recombinant sirtuin 1 (SIRT1) was produced in E. coli and purified.
The non-acetylated and acetylated Foxo-3 substrates (hFoxo-3a-290-K(Ac): Ac-DSPSQLS-
K(Ac)-WPGSPTS-NH2) were synthesized by GENEPEP SA (Saint-Jean-de-Védas, France).
Deacetylase reactions were carried out in reaction buffer (Tris–HCl 50 mM, NaCl 137 mM,
KCl 2.7 mM, MgCl2 1 mM, bovine serum albumin 0.05%, TCEp 1 mM, NAD+ 0.8 mM) and
2
0 nM SIRT1 for 30 min at room temperature. The reaction was initiated by the addition of
peptide substrate and quenched by the addition of 2% formic acid. The transitions of peptides
detection: substrate: (excitation at 829.1 nm; emission at 544.6 nm) and product (excitation at
8
08.4 nm; emission at 544.5 nm) were detected.
Specificity tests. NAD(P)H oxidase experiments. This enzyme activity was measured using
the lucigenin assay as described by Sasaki et al. (2013) and by Kuribayashi et al. (2008). This
assay was adapted to the U937 cell line as biological source.
ROS production in acellular and cellular experiments using electron paramagnetic
resonance (EPR). 5,5’-dimethyl-1-pyrroline N-oxide (DMPO) were purchased from
Interchim (Montluçon, France). Menadione, adrenochrome, 3,3’-methylene-bis(4-
hydroxycoumarin) (dicoumarol), NADH, and PBS were purchased from Sigma-Aldrich-Fluka
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Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
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Co. (Saint Quentin Fallavier, France). BNAH was purchased from TCI Europe (Zwijndrecht,
Belgium). NRH was custom synthesized by O2h (Ahmedabad, India). DMSO was purchased
from Fisher (UK, Loughborough). The DMPO stock solution (1 M) was prepared in water
and stored at -80 °C until required. The inhibitors (S29434 and dicoumarol) were prepared as
1
mM solutions in DMSO and diluted in PBS to yield 0.2 mM. The substrates and co-
substrates stock solutions were prepared in DMSO. Experiments on pure enzyme (QR1 or
QR2) were performed with demineralized (18 MΩ) and deaerated water containing
tris(hydroxymethyl)amino-methane 50 mM and ß-octyl-glucoside 1 mM, final pH 8.5. The
EPR spectra were recorded immediately after mixing the pure enzyme with the inhibitor when
needed (dicoumarol or S29434), DMPO (final concentration 125 mM), the co-substrate
(NADH or NADPH for QR1 and NRH or BNAH for QR2, final concentration 3 mM) and
finally the quinone (menadione, 125 µM). Naïve Chinese Hamster Ovary cells (CHO-k1-NT)
with basal expression of QR1 and QR2 and cell lines overexpressing QR1 (CHO-k1-QR1) or
overexpressing QR2 (CHO-k1-QR2) were custom-made and purchased fromVectalys
6
(
Ramonville,France) (Cassagnes et al., 2015). The experiments were performed on 5.10 cells
suspended in a final volume of 400 µl. EPR spectra were obtained at X-band and at room
temperature on a Bruker EMX-8/2.7 (9.86 GHz) equipped with a high-sensitivity cavity
(4119/HS 0205) and a gaussmeter (Bruker, Wissembourg, France). The analyses were
performed with a flat quartz cell FZKI160-5 X 0.3 mm (Magnettech, Berlin, Germany).
WINEPR and SIMFONIA software programs (Bruker) were used for EPR data processing
and spectrum computer simulation. Typical scanning parameters were as follows: scan
number, 5; modulation amplitude, 1 G; modulation frequency, 100 kHz; microwave power, 1
mW; sweep width, 100 G; sweep time, 41.94 s; time constant, 20.48 ms; and magnetic field
3
465–3560 G. The intensities of the EPR signals were evaluated by measuring the amplitude
peak to peak of the second line.
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Liver and brain mitochondrial respiration. The animal experimentation component of the
studies was conducted with the approval of the local Animal Care and Use Committee and
according
to
the
applicable
guidelines
of
the
CNRS
(http://www.cnrs.fr/infoslabos/reglementation/euthanasie2.htm). Rats (male, 150 g) were
sacrificed by cervical dislocation followed by decapitation (fast bleeding). The liver and brain
were quickly removed and placed in ice-cold mitochondria isolation buffer (sucrose 300 mM,
Tris 10 mM, EGTA 1 mM, pH 7.4). Mitochondria were prepared by differential
centrifugation, and the final mitochondrial pellet was resuspended in isolation buffer (20–80
mg/mL). Mitochondria were resuspended in 5 mL of mitochondrial respiration buffer (KCl
1
00 mM, sucrose 40 mM, TES 10 mM, MgCl 5 mM, EGTA 1 mM, 5 mM phosphate, fatty
2
acid–free bovine serum albumin 0.4 %, pH 7.2), and this suspension was distributed in each
of the two chambers (2 mL final volume) of the Oxygraph 2K (Oroboros Instruments,
Innsbrück, Austria). Temperature was set to 25 °C. Respiration was initiated with addition of
glutamate and malate (5 mM each), and a fully stimulated phosphorylating state 3 was
obtained by addition of 1.25 mM ADP. Increasing amounts of the S29434 were added to one
chamber using 1,000 × concentrated working solutions in DMSO, and simultaneously the
same volume (2 µl) of DMSO was added in the other chamber. For the liver experiments, 6
experiments, in 3 different days, with different mitochondria preparations were used. For the
brain experiments, il was 3 different preparation that were used.
Muscle mitochondrial respiration. With the exception of Amplex Red Ultra (Life
Technologies) and BNAH (see above), all chemicals and reagents were purchased from
Sigma-Aldrich. Male Wistar rats were purchased from Charles River Laboratories, Inc.
(Montréal, QC) and acclimated for approximately 2 weeks in the Animal Care Facility at St.
Francis Xavier University (Canada). During the acclimation period, the rats were housed in
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Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
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pairs in standard cages with a plastic tunnel to provide environmental enrichment. All rats had
access to standard rat chow and water, ad libitum. The room in which the rats were housed
was maintained at 20 °C/22 °C and 40%/60% relative humidity and on a reversed 12-h
light/dark cycle. Approval was granted by the institutional Animal Care Committee at St.
Francis Xavier University prior to commencing the study, in accordance with guidelines of
the Canadian Council on Animal Care guidelines. Variations of the permeabilized myofiber
preparation are published elsewhere (Kuznetsov et al., 2008; Perry et al., 2011; Pesta and
Gnaiger, 2012; Perry et al., 2013). In brief, immediately following euthanasia with sodium
pentobarbital, two cuts were made on the right gastrocnemius muscle of each animal. Red
(i.e., oxidative) portions of the gastrocnemius were extracted and placed in ice-cold buffer X,
consisting of (in mM): 50 MES, 7.23 K EGTA, 2.77 CaK EGTA, 20 imidazole, 0.5 DTT, 20
2
2
taurine, 5.7 ATP, 14.3 phosphocreatine, and 6.56 MgCl .6H O (pH 7.1). Four fiber bundles
2
2
from each gastrocnemius section were separated along the longitudinal axis, using needle
tipped forceps (Fine Science Tools, Inc., Vancouver, CB, Canada) in ice-cold buffer X under
magnification (Discovery V8, Carl Zeiss, Oberkochen, Germany). Following dissection, fiber
bundles were placed in vials containing saponin (50 μg/mL) dissolved in 2.0 mL of buffer X.
These vials were then placed on a nutating mixer (VWR International, Radnor, PA, USA) and
kept at 4 °C for 30 min. After permeabilization, fiber bundles were transferred to a wash
buffer solution, MiR05, consisting of (in mM): 20 taurine, 0.5 EGTA, 3 MgCl , 60 K-
2
lactobionate, 10 KH PO , 20 HEPES, 110 sucrose, 20 creatine, 1 g/L fatty acid–free bovine
2
4
serum albumin, pH 7.1. Fiber bundles remained in MiR05 for approximately 15 min with
regular inversion on the nutating mixer until experimentation, after which the samples were
placed into the Oxygraph-2k (Oroboros). Blebbistatin was included in the permeabilization
buffer, wash buffer, and assay to inhibit spontaneous contraction of muscle fibers (Perry et al.,
2
011; Perry et al., 2013). Substrate-dependent respiratory oxygen consumption was measured
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with the Oxygraph 2K (Oroboros). H O production was monitored in permeabilized rat red
2
2
gastrocnemius muscle fibers simultaneously with respirometry using an amperometric add-on
LED module to the Oroboros Oxygraph-2k. Prior to the first substrate addition of the
respirometric protocol, 10 µM Amplex Red Ultra, 5 U/mL superoxide dismutase, and 1 U/mL
horseradish peroxidase were added to the oxygraph chambers containing the fibers. H O
2
2
calibration experiments were performed under the same parameters used for experimental
data collection. The relationship between H O and fluorescence intensity was linear to at
2
2
least 0.7 µM. Next the following were added, in order: 4 mM malate, 0.2 mM
octanoylcarnitine, 5 mM ADP, 20 mM L-lactate, 5 mM NAD+, 5 mM pyruvate, and 10 μM
UK-5099, and 10 mM glutamate was then added to test glutamate oxidation. Then, 10 mM
succinate, 1 μM auranofin (thioredoxin reductase inhibitor), and 100 μM carmustine (BCNU;
inhibitor of thioredoxin and glutathione reductases) were added to the chambers consecutively
to inhibit the H O scavenging systems mitochondria. Finally, H O was titrated to internally
2
2
2
2
calibrate the resorufin signal for H O . Following conclusion of the respirometry experiments,
2
2
fibers were placed directly into water to clear out any remaining substrates in the cytosol.
After 5 min, fiber bundles were then placed in a clean dry Eppendorf tube and freeze-dried
using Labconco® FreeZone1. Following freeze-drying for 5 h, fiber bundles were weighed on
a Mettler Toledo® Excellence Plus XP6 Ultra-Microbalance.
Results represent means ± SD; n = 4. A two-way, repeated measures ANOVA was completed
for each oxygen flux and hydrogen peroxide production to identify any interaction among the
three groups of exercise and histamine receptor antagonists, exercise only, and control. This
step was followed by a one-way ANOVA with repeated measures (Bonferroni post hoc
analysis) to examine differences between different substrate additions. Data analysis was
completed using Prism 7 (GraphPad). The α-level for statistical significance was set at 0.05.
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Measurement of mitochondrial ROS levels by flow cytometry in cells. MitoSox (Thermo
Fisher ScientificLife Tech., Molecular probes, Waltham, MA, USA) was used to measure
mitochondrial superoxide levels in cells by flow cytometry. HepG2 were a kind gift from Dr.
Eugenio Arcidiacono (University Magna Graecia, Catanzaro, Italy), were cultured in high-
glucose (4.5 g/L) Dulbecco’s modified essential medium (DMEM), as described (Lascala et
al., 2018). U373 cells were cultured in low-glucose DMEM. Both types of DMEM were
supplemented with 10 % fetal bovine serum (FBS), 1 % penicillin and streptomycin solution
(Pen-Strep) and 1 % glutamine, all from Carlo Erba srl (Milan, Italy). HepG2 and U373 cells
3
3
seeded on 24-well plates at the density 80 x 10 and 60 x 10 per well, respectively, 2 days
before starting treatments. Cells were exposed to S29434 or vehicle (DMSO) for 6, 18 and 24
h before the end of the experiment. Next, the cells were washed with pre-warmed serum-free
DMEM (high and low-glucose, depending on the cell type) and then incubated with MitoSox
(2.5 µM, 15 min) diluted in DMEM as above. The experiment was finished by washing the
cells twice with pre-warmed phosphate-buffer saline followed by trypsin-mediated
detachment of cells and flow cytometry analysis according to a previously published
protocols (Janda et al., 2013)
Transient silencing of QR2 expression and autophagy assay in hepatic cells. HepG2 cells,
were cultured as described above. Cells were seeded on 12-well plates one day before
-2
transfection at a density of 30,000 cm . Non-silencing or control (Ctrl) siRNA (All-stars) and
a set of human QR2-targeting siRNA were purchased from Qiagen (Hilden, Germany).
Transfection of HepG2 cells was performed using Lipofectamine 2000 (L2000, Life
Technologies, Invitrogen, Monza MB, Italy) and Optimem (Life Technologies, Invitrogen)
according to the manufacturer’s instructions. Briefly, 360 pmol siRNA QR2 or control siRNA
and 18 µl of L2000 were mixed and incubated in 600 µl of Optimem for 15–20 min at room
temperature. Each well containing HepG2 cells in 500 µl of serum-free medium was overlaid
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with 100 µl of siRNA-Lipofectamine complexes and incubated in a cell-culture incubator.
After 5 h, the medium was changed for DMEM supplemented with FBS without Pen-Strep.
At 24 h later, the cells were treated with S29434 or DMSO for 24 h in regular medium with
Pen-Strep. CLQ 25 µM was added 3 h before the end of the experiment to half of the samples.
The cells were lysed, and the lysates were run on 12% SDS-PAGE gels and assayed for LC3
and QR2 expression by western blotting, as previously described (Janda et al., 2015). The
antibodies were polyclonal rabbit anti-LC3A/B (MBL,Enzo Life Sciences, Farmingdale, NY,
USA) used at 1:2000, anti–glyceraldehyde-3-phosphate dehydrogenase (Santa Cruz Biotech,
Dallas, TX, USA) used at 1:500and anti-QR2 (Sigma-Aldrich) used at 1:1000).
Testing S29434 against neuronal degeneration in vitro. Animals were housed, handled,
and taken care of in accordance with recommendations of the Guide for the Care and Use of
Laboratory Animals of the National Institutes of Health (NIH Publication no. 85-23, revised
1
996) and the European Union Council Directives (2010/63/EU). Experimental procedures
were authorized by the ethical committee on animal experiments (Comité Charles Darwin #5).
Cultures were prepared from the ventral midbrain of Wistar rat embryos at gestational age
1
5.5 days (Janvier LABS, Le Genest-St.-Isle, France). Dissociated cells in suspension
obtained by mechanical trituration of midbrain tissue pieces were seeded at a density of 1.2–
5
2
1
.5 × 10 cells/cm onto Nunc 48-well multi-dish plates precoated with 1 mg/mL
polyethylenimine diluted in borate buffer, pH 8.3, as previously described (Toulorge et al.,
011). In some experiments, the cultures were maintained in N5 medium supplemented with 5
2
mM glucose, 5 % horse serum, and 0.5 % fetal calf serum, except for the first 3 days in vitro,
when the concentration of fetal calf serum was set at 2.5 % to favor initial culture maturation
(Guerreiro et al., 2008). In the other experiments, we used a chemically defined serum-free
medium consisting of equal volumes of Dulbecco’s minimal essential medium and Ham’s
F12 nutrient mixture (DMEM/F12, Life Technologies, Invitrogen), supplemented with 10
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µg/mL insulin, 30 mM glucose, and 100 IU/mL Pen-Strep (Troadec et al., 2001). Dopamine
neurons were detected by tyrosine hydroxylase immunofluorescence staining using
procedures previously described (Toulorge et al., 2011). These neurons represented
approximately 2-3% of the total number of neuronal cells present in these cultures after
plating. The cultures, fixed for 12 min using 4% formaldehyde in Dulbecco’s phosphate-
buffered saline (PBS), were washed twice with PBS before an incubation step at 4 °C for 24 -
7
1
2 h with primary antibodies. A monoclonal anti- tyrosine hydroxylase antibody diluted
/5000 (ImmunoStar, Inc., Hudson, WI, USA) or a polyclonal anti- tyrosine hydroxylase
antibody diluted 1/1000 (US Biologicals, Salem, MA, USA) was used to assess survival of
dopamine neurons. Cell counting was performed with a Nikon TE 2000 inverted microscope
(Nikon, Champigny-sur-Marne, France) at 200 time magnification, using a 20 × objective
matched with a 10 × ocular. The number of tyrosine hydroxylase+ neurons in each culture
well was estimated after counting visual fields distributed along the x- and y-axes. The
functional integrity and synaptic function of dopamine neurons were evaluated by their ability
3
to accumulate [ H]-dopamine (50 nM; 40 Ci/mmol; PerkinElmer, Courtaboeuf, France), as
previously described (Guerreiro et al., 2008). When using N5 medium supplemented with
serum, dopamine neurons in culture degenerate spontaneously and progressively during the
maturation process of these cultures (Toulorge et al., 2011). Thus, in this setting,
pharmacological treatments were initiated immediately after plating and were then renewed
daily after replacing a fraction (two thirds) of culture medium. Dopamine cell survival was
assessed in 9 days in vitro cultures, i.e., at a stage where about 70 % of dopamine neurons
have already died (Toulorge et al., 2011). Some sets of cultures were treated with veratridine
(0.8 µM), a depolarizing agent used as a positive control for neuroprotection in this setting
+
(
Salthun-Lassalle et al., 2004). Treatments with the mitochondrial toxin MPP (1-methyl-4-
phenylpyridinium) were performed in cultures in which the spontaneous death process was
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+
prevented by supplementing the N5 medium with a depolarizing concentration of K (30 mM)
in the presence of 1 µM of the N-methyl-D-aspartate receptor antagonist dizocilpine
([5R,10S]-[1]-5-methyl-10,11-dihydro-5H-dibenzo[a,d]cyclohepten-5,10-imine, a.k.a. MK-
8
01), as previously described (Salthun-Lassalle et al., 2004). Note that this treatment did not
+
+
interfere with the neurotoxic effects of MPP . MPP was applied to midbrain cultures between
4
and 6 days in culture, whereas protective treatments were added after 2 days in vitro culture
and renewed thereafter until fixation of the cultures at 6 days in culture. Finally, oxidative
stress–mediated dopamine cell death was achieved by placing the cultures in a defined serum-
free DMEM/F12 medium containing 1.5 µM of ferrous iron (Troadec et al., 2001). The
antimitotic compound ARA-c (1.5 µM) was added to the cultures after plating to prevent glial
cell proliferation. Dopamine cell survival was assessed at 5 days in culture in these
conditions. Experimental values expressed as mean ± SEM were derived from triplicates of
three independent experiments. Data were analyzed using the SigmaPlot 12.5 software (Systat
Software Inc, San Jose, CA, USA) with one-way analysis of variance (ANOVA) followed the
Student–Newman–Keuls post hoc test for all pairwise comparisons.
Object recognition in mice. Experiments were performed in a quiet, dimly lit room in a dark
Plexiglas chamber (L × W × H: 25 × 35 × 25 cm). On the day before the test day (day 0,
familiarization phase), the animals were allowed to explore the test apparatus for 2.5 min.
Twenty-four hours later (day 1, acquisition phase), after a 30-min pretreatment time after drug
treatment (1 and 15 mg/kg, i.p.), each test animal was placed in the middle of the test box
from the previous day, and the 5-min acquisition trial was begun. In the first trial, mice were
allowed to explore two identical objects (10 sec per object within a total 5-min period).
During test time, the exploration time (ET) was measured with a stopwatch. ET is defined as
direct, active olfactory exploration of objects 1 and 2. In general, it consists of nosing and
sniffing of the edge and the top, as well as approaching and crossing the line approximately 1
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cm around the objects. Posturing and mounting are not included in measures of investigation.
On day 2 (24 h later, retention phase), a new object and a familiar object were placed in the
test box, and the animals were replaced in the box for 4 min and ET measured again.
Discrimination index = [Exploration time of new object in seconds (N) – Exploration time of
familiar object in seconds (F)]/[Exploration time of new object in seconds (N) + Familiar
object exploration time in seconds (F)] = [N-F]/[N+F]. A thorough description of these
methods can be found in previous publications (Ennaceur and Delacour, 1988; Bartolini et al.,
1
996; Bevins and Besheer, 2006). For in vivo data, one-way ANOVA was performed
(followed by Dunnett’s multiple comparisons test) for the DI data.
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Results
Chemistry. S29434 was synthesized by Guillaumet and colleagues in the late 1990s as a
compound to explore the melatoninergic system(s). We characterized it as an interesting tool
for QR2 studies after we discovered that MT3, the unconventional melatonin binding site we
and others described (Duncan et al., 1988; Paul et al., 1999), was indeed in QR2 (Nosjean et
al., 2000). The first synthesis, as is typical, aimed at producing several examples of tetracyclic
chemicals that could be assimilated into two indolic structures fused together. The goal at that
stage was to document the chemical series, of which S29434 was the most potent. The route
led to tens of milligrams of material and was not meant to yield greater quantity. The
synthetic route is summarized in Fig. 1. The second synthesis scheme was intended to
rationalize the yields of each step, even if alternative synthetic steps had to be substituted in
the original protocols, as summarized in Fig. 2. This alternative route was meant to deliver
gram amounts of the compound. Full details of the synthetic routes are given as
supplementary data. Crucially, the final product, used in most of the experiments presented
here, is 99% pure, based on LC-MS and NMR spectra data (Supplementary data). The mass
spectrometry data as well as the NMR data confirmed the structure of the final product.
Inhibition of QR2. Various reports show that depending on the substrate/co-substrate couples
used to measure the catalytic activity of QR2, the recorded IC values range from 1 to 40 nM
5
0
(Table 1). We stabilized our system in a standard configuration (see materials and methods
section) for all the internal needs for screening of this enzyme (S29434 was the reference
compound) and identified an IC of 16 ± 3.2 nM (mean ± S.D., n = 51). The considerable
5
0
amount of data accumulated on this material render this information quite robust, at least
when using the human cloned enzyme, BNAH as co-substrate, and menadione as substrate. A
typical sigmoid curve of the concentration/activity relationship is presented in Fig. 3A. We
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also present a comparison of S29434 inhibition of the enzyme catalytic activity while using
BNAH, a synthetic co-substrate, or NRH, a natural candidate co-substrate of QR2 (Fig. 3B).
With the natural co-substrate, the inhibition curve is shifted slightly leftward, from an IC of
5
0
1
7 ± 3 nM in the presence of BNAH to 5 ± 1 nM with NRH (mean ± S.D.). Nevertheless, the
use of BNAH is recommended because the compound is commercially available and slightly
more robust than its ribosylated counterpart (Boutin et al., 2005). S29434 is a competitive
inhibitor, as reported previously (Mailliet et al, 2005; Ferry et al., 2010; Pegan et al, 2011).
Inhibitor specificity: Standard panel of molecular targets (Eurofins-CEREP profiles). We first
checked the specificity of the inhibitor S29434 for possible off-target activities (from the QR2
perspective). First, we assessed its activity in a series of standard assays designed to evaluate
its behavior against a selection of receptors and enzymes (see, for example, Millan et al.,
2
012). All tests were done in duplicate, independently, at two concentrations: 100 nM and 10
µM. With one exception, the results indicated no capacity of S29434 to inhibit binding at or
activity of the selected targets, even at high concentrations (>10 µM) (data not shown). The
notable exception was that S29434 could displace binding of 2-iodomelatonin at the
melatonin receptor MT2, showing an affinity of 0.14 µM. This result is not surprising given
that S29434 was initially synthesized as part of a melatonin receptor agonist discovery
process. Finally, S29434 showed no cytotoxic activities towards several standard cancer cell
lines (Caco-2, HCT116, HT-29, and L1210) up to 10 mM, based on MTT assays.
Inhibitor specificity: Kinases. Duncan et al. (2008) reported the toxicity of two casein kinase
2
inhibitors, TBBz (4,5,6,7-1H-tetrabromobenzimidazole) and DMAT (2-dimethylamino-
,5,6,7-tetrabromo-1H-benzimidazole), presumably because of their inhibition of QR2
4
catalytic activity. Similarly, Rix et al. (2007) reported that QR2 is the only non-kinase target
of the BCR-ABL inhibitors imatinib and nilotinib. Thus, we assessed whether the QR2
inhibitor could inhibit one or several catalytic activities of a panel of kinases. Of the 46
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included kinases, no inhibition greater than 20% was recorded for S29434 at 100 nM (data not
shown). Because of the high potency of S29434, no higher concentrations were tested.
Inhibitor specificity: QR1. The best candidate for possible specificity of S29434 would be
QR1, the closest relative of QR2. We also wanted to confirm that dicoumarol, the standard
QR1 inhibitor (Ernster et al., 1962) in wide use, could not inhibit QR2. Based on previous
catalytic activity measurement comparisons between the two enzymes (Ferry et al., 2010),
S29434 shows good selectivity for QR2 over QR1, with the reverse result for dicoumarol. In a
new set of experiments (Fig. 4), we used EPR analysis and showed that S29434 at 10 µM did
not inhibit QR1, compared to full inhibition by dicoumarol. Indeed, S29434 clearly had no
effect on the intensity of the EPR spectrum, as represented by the six-lined spectrum
characteristic of the radical adduct [DMPO-CH3]•. This radical adduct originates from the
spin trapping of •CH3 formed by reaction of •OH (produced during the re-oxidation of
menadiol) with DMSO.
Inhibitor specificity: sirtuin. Because of the link between sirtuin-mediated NAD degradation
(Jiang et al., 2017) and QR2’s use of a possible NADH catabolite, we sought to ensure that
the inhibitor’s activity did not lead to upstream inhibition of sirtuin, diminishing the
availability of its co-substrate. Even at 10 µM, S29434 did not inhibit sirtuin, whereas
superoxide dismutase and diphenyleneiodonium chloride were potent inhibitors (not shown).
Inhibitor specificity: NADPH oxidase. In addition to mitochondria, a main cellular source of
ROS production is NADPH oxidase (Sorce et al., 2017; Teixeira et al., 2017). Thus, we
checked the capacity of S29434 to inhibit this family of enzymes, using the NADPH oxidase
inhibitor diphenyleneiodonium. The reference compound led to a complete inhibition, but
S29434 remained poorly active up to 10 µM, at which point it began to inhibit about 20% of
ROS production from those cells (Supplemental Figure 3).
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S29434/quinone reductase 2 characteristics: Binding kinetics. Resveratrol has been described
as a potent inhibitor of QR2 (Buryanovskyy et al., 2004) and thus might serve as a reference
compound in some conditions. The binding of either S29434 or resveratrol leads to quenching
of the intrinsic fluorescence of QR2 (FADox). We used pre-equilibrium stopped-flow
spectroscopy to measure the real-time binding kinetics of the two compounds and compare
their binding mechanisms (Fig. 5). Resveratrol binding kinetics were too fast at 25 °C, so we
decreased the temperature to 4 °C to slow the rate. The results were consistent with a single-
6
-1
-1
step binding mechanism, governed by an association rate constant k of 19 10 s M and a
1
-1
dissociation rate constant k of 27 s . A K estimate of 1.4 µM was calculated from the ratio
-1
D
k /k , which is in good agreement with the isothermal titration calorimetric data obtained on
-1
1
the same enzyme batch, yielding a K of 0.6 µM at 25 °C (data not shown). In contrast,
D
S29434 displayed slower but biphasic binding transients, consistent with a two-step binding
mechanism involving a conformational change. Re-plotting the kobs rate constants for the fast
phase yielded a linear plot, diagnostic of a bimolecular association step, governed by a k of
1
6
-1
-1
-1
-1
1
4 10 s M and a dissociation rate constant k of 61 s .M . A K estimate for this
-1 D
encounter complex of 4.4 µM can be calculated from the ratio k /k . The observed rate of the
-1
1
slow phase was independent of S29434 concentration in the investigated range, staying close
-1
to 30 s . This step therefore likely reflects a conformational rearrangement, either following
binding (induced fit) or preceding it (conformational selection). These preliminary kinetic
data strongly suggest the evolution of the binding mechanism from the low-affinity single-
step binder resveratrol to the high-affinity two-step binder S29434. They also are in
agreement with results from our mass spectrometry studies of the inhibitor/enzyme binding
(Antoine et al., 2012).
S29434/QR2 characteristics: Co-crystallization. Through various collaborations, we
generated co-crystals of S29434 in QR2 that we deposited in the Protein Data Bank
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(www.rcsb.org) (Pegan et al., 2011; Gerard et al., 2018) including several unpublished
crystals at 1.4 Å resolution (4QOD;); however, the interactions and orientation of the
compound in the crystal were not discussed in great detail. In brief, as shown in Fig. 6, the
ligand S29434 fits deeply into a QR2 hydrophobic cavity (PDB: 3OX3), interacting with
subunits A and B with its amido-furan side chains pointing toward the solvent. It makes π-
edge interactions with Trp105 and Phe126, π-π interactions with Phe178, and a water-
mediated H-bond with Asn161. Additional π-π interactions exist with co-factor FAD.
Although the furan moiety makes an additional hydrophobic interaction with Ile194 from
QR2, this part of the molecule also seems to hang out of the catalytic site. This feature
potentially offers the possibility of modification of this part of the molecule with, for instance,
a fluorophore moiety, producing associated properties, as it has been done for chromen-2-one
derivatives as inhibitors of QR1 (Bian et al, 2017).
Transit of S29434 across the cellular membrane and blood–brain barrier. We next
conducted a series of experiments to characterize the properties of S29434 in terms of
pharmacokinetics and cellular penetration. Firstly, we measured the behavior of S29434 in an
in cellulo blood–brain barrier model (Cechelli et al., 1999), based on co-cultures of bovine
brain micro-vessel endothelial cells and rat astrocytes (Booher and Sensenbrenner, 1972)
showing expression of the pGP efflux transporter. For this purpose, we measured the
permeability surface total (PSt) (filter + collagen + cells) and the permeability surface filter
(PSf) (filter + collagen without cells) expressed in milliliters per minute. These values were
4
.58 cm/min and 3.78 cm/min, respectively. Therefore, the ratio PSt/PSf (permeability class)
for S29434 was determined to be 121%, showing no barrier effect of the BBB cells compared
with the filter+collagen alone, classifying S29434 as a high permeable compound (Cechelli et
al., 1999). Secondly, S29434 was tested on the well-known Caco2 model, expressing the main
efflux transporters present at the intestinal and the BB barriers (pGP, BCRP and MRP2). The
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-6
apical to basolateral (A2B) apparent permeability (Papp) was 33 10 cm/s (incubation
concentration at 20µM) corresponding also to a high permeable compound, in the same range
of that of propranolol, the high passive permeability reference. The basolateral to apical
(B2A) permeability was not quantified in these experiments but the high A2B permeability
suggested no efflux transporter involvement.
QR2 and ROS production. QR2 activity can be followed by EPR (Reybier et al., 2011;
Cassagnes et al., 2015). As noted, this functional assay is indirectly linked to the enzyme.
Indeed, the reaction product, a quinol, has poor stability in standard conditions. In the
presence of oxygen, the quinol is oxidized back into the original quinone, which in turn serves
as substrate for the enzyme. The result is a futile cycle between the two chemical species with
the concomitant production of ROS [see Reybier et al. (2011) for a complete discussion]. This
process seems to occur in acellular as well as cellular contexts (Cassagnes et al., 2017). Here,
we completed and supported these findings, showing by EPR spectroscopy that pure QR2 in
the presence of its co-substrate (BNAH) produces ROS during catalytic activity (Fig. 7);
S29434 inhibits this effect. Furthermore, under those experimental conditions, neither NADH
nor NAD(P)H could substitute for BNAH. Under basal conditions (no substrate or co-
substrate of QR2), very low levels of ROS were recorded.
QR2 and ROS production: Mitochondrial respiration in liver and brain. Because S29434
diminishes the ROS produced during the futile cycle between quinol and quinone in aerobic
conditions (Reybier et al., 2011), we wanted to ensure that this effect did not result from
direct interference of S29434 with ROS even though the compound is inactive in other ROS-
generating systems (such as mitochondria). Our results confirmed that S29434 cannot trap
●
●-
hydroxyl radicals ( OH) produced via the Fenton reaction or superoxide radicals (O )
2
produced from crown ether and KO (not shown). We also tested another source of cellular
2
ROS production: the mitochondria. To evaluate the mitochondrial effect of the drugs,
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mitochondrial respiration was settled with complex I substrates (glutamate/malate) and in
phosphorylating conditions (1.25 mM ADP). The relative effect of increasing amounts of
S29434 is indicated by the ratio of respiratory rate compound/solvent, with a value of 1
indicating a lack of effect. With this approach, S29434 produced no effect up to 10 µM
concentrations and revealed no statistically significant changes in respiratory rate with rat
liver or rat brain mitochondria (Fig. 8). The system was validated with experiments with
rotenone or oligomycin, two known inhibitors of the mitochondrial respiratory chain, with
results similar to those reported in the literature (data not shown).
QR2 and ROS production: Mitochondrial respiration in muscle. The next step involved
independent assessment of mitochondria in saponin-permeabilized rat gastrocnemius muscle
fibers. As illustrated in Fig. 9A, the conditions resulted in no differences in respiration,
confirming that neither BNAH nor S29434 had an effect in these samples. Likewise, BNAH
and S29434 did not affect H O production (Fig. 9B). Expression of H O as a percentage of
2
2
2
2
O flux yielded a statistically significant difference between BNAH and BNAH + S29434
2
(Fig. 9C) in the absence of ADP only. This finding suggested that BNAH may increase
mitochondrial H O in non-phosphorylating (i.e., State 4 or LEAK) conditions. However,
2
2
because neither respiration nor H O alone was affected, a more plausible explanation is that
2
2
the difference in proportional H O production is the result of non-mitochondrial sources of
2
2
H O in the samples, and not of bulk O flux.
2
2
2
Effects of S29434 on cells. Nervous system cells. S29434 was described as having a beneficial
effect against paraquat-induced toxicity in cellulo and in vivo (Janda et al., 2013, 2015),
suggesting a potential role of QR2 in Parkinson’s disease. This finding prompted us to
analyze its effect in models of neuronal degeneration in cellulo. Primary midbrain dopamine
neurons in culture were established from fetal rat ventral midbrains. In this system, S29434
(up 100 µM) did not reduce spontaneous loss of dopamine neurons in culture (Fig. 10 A),
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while the reference neuroprotective molecule, veratridine, efficiently rescued these cells.
S29434 also failed to protect dopaminergic cells from iron-induced oxidative stress (with iron
in the culture medium) caused by a Fenton-type reaction occurring spontaneously (Fig. 10 B).
At 100 µM, S29434 even tended to become slightly toxic. As expected, however, the
chelation of iron with deferoxamine (10 µM) led to robust protection under these conditions.
In contrast to these observations, S29434 showed concentration-dependent protective effects
+
against the mitochondrial complex I inhibitor MPP (Fig. 10C). This effect was already
present at 0.1 µM, peaked at 1 µM, and declined progressively between 10 and 100 µM.
+
Nicotine, used at 10 µM as a positive control, also rescued dopamine cells. Because MPP is
selectively taken up by the dopamine transporter to exert its selective toxicity towards
dopamine neurons, we tested whether S29434 operated as an inhibitor of dopamine uptake.
We thus performed this routine experiment according to details already published in our
laboratory (Guerreiro et al., 2008). S29434 does not bind to the ad hoc transporter (not
+
shown), a finding that strongly suggests that it blocks the intrinsic mechanism of MPP
toxicity, not its transport.
Effects of S29434 on cells. Autophagy. Autophagy is a vesicle-mediated pathway that ensures
recognition and transport of defective organelles and protein aggregates to lysosomes, where
they are digested. We tested if S29434 could induce autophagy in human HepG2 cells, a cell
line derived from liver, the organ with the highest level of QR2 expression (Nosjean et al.,
2
000). S29434 dose-dependently induced LC3-II, a marker of autophagy vesicles (Fig. 11A).
Silencing of QR2 by more than 50% increased basal LC3-II levels and suppressed LC3-II
induction by S29434. This result indicated that the autophagy response to this compound does
not depend on its off-target effects but is directly mediated by QR2 inhibition (second panel in
Fig. 11A, B). Of importance, LC3-II was further accumulated in S29434-treated cells when
autophagy flux was blocked by the lysosomal inhibitor ClQ, indicating true induction of
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autophagy by the QR2 inhibitor (Fig. 11B). Further details are given in Supplementary Data
, including discussion of the whole blots. These data confirm that QR2 plays a role in
8
regulation of autophagy. Oxidative stress and redox reactions have a strong impact on
autophagy machinery (Janda et al., 2012; 2015). Because QR2 inhibition does not reduce
basal ROS levels in HepG2 cells (Fig. 12A) or U373 astrocytes (Fig. 12B), however, the
mechanism by which S29434 triggers autophagy should be mitochondrial ROS-independent
and QR2-dependent, as suggested by above results (Fig. 11A, B; Supplementary Data 8).
Effects of S29434 in vivo. Object recognition tests in CH3 wild-type mice. An interplay
between QR2 and memory has been reported (Brouillette and Quirion, 2008; Benoit et al.,
2
010; Rappaport et al., 2015). To further assess this relationship, we used the inhibitor
S29434 in the object recognition test. The test trial with the new object was carried out 24 h
after the acquisition trial with two identical objects. S29434 was given intraperitoneally, 30
min before the acquisition trial (1 and 15 mg/kg, respectively). Fig. 13A shows the
discrimination index, where N is the time spent inspecting the new object and F is the time
spent exploring the familiar object during the test trial. Both concentrations of S29434 led to
an enhanced time of exploration of the unfamiliar object, although the effects of the two doses
(1 and 15 mg/kg) did not differ. Under these circumstances and experimental conditions, the
object recognition memory of wild-type mice is enhanced by a factor of ~16 by i.p. injection
of S29434 over control mice performance in the same test. As reported for other models
(
Rappaport et al., 2015), this finding suggests that by inhibiting QR2, S29434 has a positive
effect at least on this particular aspect of memory.
In vivo considerations on S29434: Object recognition memory in the KO CH3QR2 mice.
-/-
-/-
The same experiments were carried out on KO CH3QR2 mice, and the results are presented
in Fig. 13B. For QR2 KO animals, the results strongly reinforce the previous findings. Indeed,
treated (1 or 15 mg/kg) and control animals did not differ, indicating that the product has no
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Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
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further effect on this parameter. Because the QR2 KO animals were genetically deficient in
this enzyme only (Mailliet et al., 2004), the lack of S29434 activity strongly suggests that the
compound is specific to QR2. Furthermore, we know that these animals did not lose their
learning capacity compared to the wild-type mice, as demonstrated in several tests including
the Morris water maze, object recognition, and rotarod performance test. They performed
better than their wild-type counterparts on all of these tests (Benoit et al., 2010).
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Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
This article has not been copyedited and formatted. The final version may differ from this version.
MOL #114231
Discussion
General. Molecular pharmacology is based on either the manipulation (including the
mutagenesis) of targets or their inhibition/activation by synthetic or natural chemicals. A good
example is certainly dicoumarol, the QR1 reference inhibitor. This compound has been
widely used in the literature (over 2700 references), yet many of its off-target effects bias the
interpretation of the data obtained [see discussion in Scott et al. (2011)]. To avoid a similar
scenario with S29434, here we sought to fill in information gaps related to this compound,
including assessments of its in cellulo and in vivo activity, stability, permeability, and overall
specificity and to publish a clear synthesis method.
Chemistry. S29434 belongs to a family of flat tetracyclic compounds, some that we (Boussard
et al., 2006) and others have described, e.g., cryptolepines (Onyeibor et al., 2005; Lavrado et
al., 2008; Whittell et al., 2011) or other types of indolizino-indolones (Bhattacharya et al.,
2
001). Compounds that inhibit QR2 activity also have been reported, including
pyrroloquinoline ammosamides (Reddy et al., 2011), indolequinones (Dufour et al, 2011; Yan
et al, 2011) and furan-amidines (Alnabulsi et al., 2018). Because few, if any, have been
characterized beyond their enzymatic test activities, we believe that the present report is
noteworthy because it describes S29434 activity in different in cellulo and in vivo situations.
Acellular behavior, specificity. In an acellular system, S29434 is a potent inhibitor of QR2
activity (IC ~10 nM). It marginally inhibits QR1, if at all. Furthermore, S29434 does not
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potently inhibit more than 100 other targets, with the sole exception of the melatonin receptor
, for which it has a binding affinity in the micromolar range. Of course, the notion of
2
specificity is limited to targets for which measures are available. It is not possible to know if
S29434 will uniquely recognize QR2 among the protein products of the 26,000 genes in the
human genome. The mechanism by which S29434 binds to and interacts with QR2 has been
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Molecular Pharmacology Fast Forward. Published on December 19, 2018 as DOI: 10.1124/mol.118.114231
This article has not been copyedited and formatted. The final version may differ from this version.
MOL #114231
described (Pegan et al., 2011; Antoine et al., 2012). Here, we add information about the way
the compound occupies the catalytic site of the enzyme. We also identify ways to measure
S29434’s affinity for QR2, in complement with its capacity to inhibit it. These two notions
(affinity for the enzyme and inhibition potency) are too often conflated, and it is important to
distinguish them. Furthermore, the biophysical measure allowed us to evaluate how long the
compound sits in the catalytic site, which can vary widely across a series of analogue
compounds for a given enzyme, ranging from seconds to hours.
Stability in vivo, membrane crossing. S29434 is demonstrated here to be highly permeable on
both an in vitro BBB model that express the pGP and on the Caco2 model expressing the 3
main efflux transporters present at both intestinal and BBB barriers (pGP, BCRP, MRP2); this
suggests no efflux involvement even if not directly demonstrated by bi-directional
experiments. It was already clear that S29434 could penetrate different types of primary or
cultivated cells (Benoit et al., 2010; Janda et al., 2013; Chen et al., 2017). Even if an in vivo
pharmacokinetic study would have help to determine the PK parameters such as T1/2 or
bioavailability, the fact that the compound was active in rodent models when administered by
intraperitoneal route (Janda et al., 2015; Rappaport et al., 2015) demonstrate that at least
sufficient in vivo brain exposures (level and duration) were reached in mammals.
Nevertheless, it is clear for us that a complete assessment of the plasma stability, as well as a
quantification of the part of the injected molecule that reaches the brain of an animal after
intraperitoneal treatment would greatly help the future studies.
Cellular aspects, ROS. As noted, QR2 is closely linked with ROS production. Thus, our main
goal was to ensure that S29434 was not a scavenger per se and that the reduction in ROS
bursts in cellular experiments was the result of S29434 interference with QR2. Indeed, the
results showed that S29434 can inhibit QR2-dependent production of ROS in various cell
types, including hepatoma HepG2 cells, the astroglial cell line U373 (Janda et al., 2013), and
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