Natural ligands of RXR receptors

CHAPTER NINE

Patricia García, Paula Lorenzo, Angel R. de Lera∗

Departamento de Qu´ımica Orga´nica, Facultade de Qu´ımica, CINBIO and IBIV, Universidade de Vigo,

Campus As Lagoas-Marcosende, Vigo, Spain

^∗Corresponding author: e-mail address: qolera@uvigo.es

Contents

1. Introduction. Structure of RXR receptors

2. Structural determinants for ligand binding to RXR receptors

3. Natural RXR ligands

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3.1 Dietary compounds. Natural rexinoids

3.2 Rexinoids from plants

3.3 Endogenous RXR ligands

4. Diastereo- and enantioselective synthesis of (R)-9-cis-13,14-dihydroretinoic

acid 34

5. Materials

5.1 Reagents and solvents

6. Methods

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6.1 Method for formation of BT-sulfide and BT-sulfone

6.2 Method for formation of dienylstannane

6.3 Method for stannane-iodine exchange

6.4 Method for Suzuki cross-coupling

6.5 Method for saponification

6.6 Method for enantiomeric excess determination of (9Z,13R)-13,14-

dihydroretinoic acid (9Z,13R)-34

Acknowledgments

230

References

Abstract

Given the role of retinoid X receptors (RXRs) as promiscuous partners of heterodimeric

complexes with other members of the Nuclear Receptor (NR) superfamily, RXR ligands

(rexinoids) play fundamental roles in gene transcription, since upon ligand binding

either transcriptionally activate the “permissive” subclass of heterodimers or synergize

with partner ligands in the “non-permissive” subclass of heterodimers. The collection of

natural products thus far reported to bind RXR are described, including those discov-

ered by high-throughput screening (HTS), mere serendipity, and a combination of

those. Detailed protocols for the diastereo- and enantioselective synthesis of (R)-9-

cis-13,14-dihydroretinoic acid, a putative natural RXR ligand, are provided.

#

Methods in Enzymology, Volume 637

ISSN 0076-6879

https://doi.org/10.1016/bs.mie.2020.02.006

2020 Elsevier Inc.

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Abbreviations

AF

activation function

ATRA

CADD

CAR

all-trans-retinoic acid

computer-aided drug design

constitutive androstane receptor

ChEMBL European Molecular Biology Laboratory Chemical Database

CoA

co-activator

Co-R

DHA

EBI

co-repressor

docosahexaenoic acid

European Bioinformatics Institute

farnesoid X receptor

FXR

GPCR

GGA

HETE

LBD

LBP

G-protein coupled receptor

geranyl geranoic acid

hydroxyeicosatetraenoic acid

ligand binding domain

ligand binding pocket

LXR

NOE

NR

liver X receptors

nuclear Overhauser effect

nuclear receptor

PPAR

PXR

RAR

RXR

RetSat

SAR

peroxisome proliferator-activated receptor

pregnane X receptor

retinoic acid receptor

retinoid X receptor

retinol saturase

structure–activity relationships

silencing mediator for retinoid and thyroid hormone receptors

thyroid hormone receptor

vitamin D receptor

SMRT

TR

VDR

1. Introduction. Structure of RXR receptors

Retinoid X receptors RXRα, β, and γ (NR2B1, NR2B2 and NR2B3)

are members of the nuclear receptor (NR) superfamily of ligand-dependent

transcriptional regulators that bind DNA response elements associated with

target genes (Germain et al., 2006; Laudet & Gronemeyer, 2002). Similar

to other NRs, RXRs feature a modular structure, which can be dissected

in several domains with associated functions, most notably the N-terminal

activation function (AF-1), the central DNA-binding domain (DBD) and

the C-terminal ligand-binding domain (LBD).

The LBD is a single protein domain of 11–13 helices and a short β-turn

arranged in three layers to form the so-called anti-parallel alpha-helical

RXR natural modulators

211

sandwich (Huang, Chandra, & Rastinejad, 2014). The multifunctional

nature of this NR domain is shown not only by the control of ligand binding

and receptor dimerization, but also by the interaction with the so-called

co-regulators (co-repressors, CoRs, or co-activators, CoAs) through a

ligand-dependent activation function (AF-2). Mechanistically, ligand bind-

ing induces the repositioning of the C-terminal helix H12 of the LBD due to

alterations on the interaction surfaces with co-regulators (Huang et al., 2014;

le Maire et al., 2012).

In addition, agonist ligand binding induces an exchange of CoR by CoA

protein complexes and also of chromatin-remodeling enzymes. As a conse-

quence, the condensed transcriptionally inactive state is transformed into the

activated accessible state, leading ultimately to gene transcription. Other

structural alterations on these complexes due to the action of ligands that

act as antagonists, partial agonists and inverse agonists have also been rev-

ealed in the last few years (le Maire et al., 2012; Xu et al., 2001).

Being the common heterodimerization partner of multiple NRs, RXR

occupies a central position in NR signaling (Mangelsdorf & Evans, 1995),

since it can act as an active or as a silent transcription partner, and therefore

the corresponding RXR-NR heterodimers are called permissive or non-

permissive, respectively, if they are activated by the RXR ligand or by

the partner’s ligand (although synergistic effects are usually observed if both

partners of the heterodimer are bound to their ligands), or only by the part-

ner’s ligand whereas ligand-bound RXR is silent or subordinated (Germain,

Iyer, Zechel, & Gronemeyer, 2002).

2. Structural determinants for ligand binding to RXR

receptors

As in other NRs, the CoR- or the CoA-bound states of RXR are

exquisitively sensitive to ligand action (as agonists, antagonists or inverse

agonists) and therefore, to their structure (Bourguet & Moras, 2015;

Dominguez, Alvarez, & de Lera, 2017; Huang et al., 2014; Mendoza-

Parra, Bourguet, de Lera, & Gronemeyer, 2015).

The crystal structures of RXR-ligand complexes (Bourguet & Moras,

2015; Huang et al., 2014) clearly show that the most selective RXR ligands

feature a bent structure (see Fig. 1), which is enforced by a tetrahedral car-

bon, a 1,1-disubstituted olefin, or a twisted polyene side-chain for optimiz-

ing binding to the L-shaped RXR LBP. This sharp turn on the LBP volume

is induced by the (highly conserved) conformation of a lysine residue in H11

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Fig. 1 RXRα–9-cis-retinoic acid (31, see Fig. 4) crystal structure (PDB code: 1k74)

(Germain et al., 2009).

(residue L436 in human RXRα) (Huang et al., 2014). As a consequence, the

shape of the RXR LBP is very restrictive to ligand binding, and selects those

ligands that are either conformationally or configurationally twisted or suf-

ficiently flexible to twist around single bonds. Such bending is not structur-

ally attained by the elongated fully conjugated polyenes (although not for

partially unsaturated analogs), such as all-trans-retinoic acid.

More than 500 RXR agonists and antagonists (with IC/IC₅₀lower than

50μM) have been annotated in ChEMBL, the database dependent on the

European Bioinformatics Institute (EBI). From the collection of RXR

modulators, going from naturally occurring compounds, those discovered

by high throughput screening or mere serendipity, only a few belong to

the first group. The rationale for the structure-based design of the most

important classes of RXR modulators has been described elsewhere

(Bourguet & Moras, 2015; Dominguez et al., 2017; Hiebl, Ladurner,

Latkolik, & Dirsch, 2018; Mendoza-Parra et al., 2015).

Natural ligands are compounds present in Nature, which bind their

receptors with high potency (ideally, in the nanomolar range) usually leading

to conformational changes in the complex structure, inducing thereby rel-

evant physiological activities. We will discuss the biological relevance of the

natural ligands reported for RXR. Needless to say, the relevant question of

their presence at the right concentration on cells and tissues remains in most

cases uncertain. Some of these compounds have been detected under

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supraphysiological conditions or nutritional intervention and their relevance

for the control of biological processes dependent on RXR remains

uncertain.

3. Natural RXR ligands

3.1 Dietary compounds. Natural rexinoids (Fig. 2)

A collection of flexible unsaturated natural fatty acids acquired in the diet

have been reported to bind to and activate RXR (Dominguez et al.,

2017; Hiebl et al., 2018). The promiscuity of RXR on binding several

metabolites indicates a putative role for this receptor as an intracellular sensor

of the cell metabolic status, binding either the natural ligand or fatty acids,

depending upon their supply and local concentration.

Within the series of flexible unsaturated fatty acids, both the endogenous

and those acquired from the diet, that bind to and activate RXR, doco-

sahexaenoic acid (DHA) 1 (Fig. 2) isolated from mouse brain, was the first

reported activator of RXR in cell-based assays (de Urquiza et al., 2000).

DHA 1 is a major constituent of nutrients and is considered as a conditionally

essential fatty acid. DHA 1 is highly abundant (up to 30–50% of the total

fatty acid content) in mammalian brain cells, where it is found associated

to membrane phospholipids, and accumulates in the retina. It is required

for brain maturation in rodents and humans, and its deficiency results in

abnormalities similar to those in RXRα-deficient mice. Despite these find-

ings, it has not been demonstrated that DHA 1 is required for RXR-RAR

retinoid signaling or any other RXR-NR nonretinoid signaling pathway in

which RXR may be involved (Mic, Molotkov, Benbrook, & Duester,

2003; Wietrzych-Schindler et al., 2011).

DHA 1 easily adapts to the LBP of RXR, and the crystal structure of

RXRα bound to DHA 1 and a co-activator peptide in a homodimeric asso-

ciation (Egea, Mitschler, & Moras, 2002), indicates an occupancy of about

80% of the LBP volume (compared to 72% for 9-cis-retinoic acid 31, see

Fig. 1). All atoms of the ligand are in direct van der Waals contact with

the protein and in particular those filling the hydrophobic cavity occupied

by the β-ionone ring-binding subpocket of 9-cis-retinoic acid (31, see Fig. 4)

bound to RXRα. Since the endogenous levels of DHA (which range from

0.1 to 0.01μM in animal and human serum or tissue samples) are not suffi-

cient for transcriptional activation, which requires at best a concentration

range of 5–10μM (Lengqvist et al., 2004), its role as physiological ligand

Fig. 2 Natural and dietary (unsaturated) fatty acids. Arbitrary conformations have been selected.

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is questionable, despite the reported ability of vertebrates to synthesize DHA

from α-linolenic acid as precursor of plant origin.

Additional fatty acids isolated from rat tissue total lipid extracts were also

shown to activate RXR (Goldstein, Dobrzyn, Clagett-Dame, Pike, &

DeLuca, 2003). Out of the large number of family members only the highly

unsaturated fatty acids were able to bind the receptor with high affinity as

shown by competition experiments with tritiated 9-cis-retinoic acid for

binding to mRXRγ. Notably, linoleic acid 2, which is one of the essential

fatty acids, and also oleic acid 3, palmitoleic acid 4, arachidonic acid 5 and

docosapentaenoic acid 6, which have been isolated mainly from testis

and heart, were found to activate mRXRγ at high (EC₅₀>200μM) con-

centrations.(Lengqvist et al., 2004) Other eicosanoids, most notably

hydroxyeicosatetraenoic acids such as (S)-5-HETE 7 and (S)-12-HETE 8

were reported to mimic some actions of the retinoids in cell-based assays

(Eager, Brickell, Snell, & Wood, 1992).

It is not unusual to find unsaturated fatty acids bound to the LBP in the

corresponding crystal structures of holo NRs (Huang et al., 2014), which are

present due to the competing biosynthetic pathways of the bacterial expres-

sion host. This is the case of compound 3, which was present in the crystal

structure of the LBP of the ligand-bound RARα/F318A RXRα

heterodimer complex. Oleic acid 3 conformationally adapts to the

U-shape and resembles 9-cis-retinoic acid in the extended volume of the

RXRα binding pocket due to the F318A mutation (Bourguet et al., 2000).

Several other natural products termed protectins, resolvins and maresins

(9–11, Fig. 2), which are bis-, and tris-hydroxylated DHA metabolites

(Serhan & Petasis, 2011), have also been found endogenously in vertebrates

including humans (Mas, Croft, Zahra, Barden, & Mori, 2012), some of them

even at higher levels than the parent DHA. They have been reported to dis-

play a large variety of biological activities through blocking or activation of

various GPCRs, as well as anti-inflammatory and neuro-protective effects,

similar to those regulated by RXRs (Bazan, 2018; Krishnamoorthy et al.,

2010). Different mechanisms have been proposed to explain the numerous

biological activities of these compounds, termed collectively Specialized

Proresolving Mediators (SPMs) although binding and transactivation of

RXRs has been rarely investigated (Krishnamoorthy et al., 2010;

Serhan, 2014).

Phytanic acid 12 (Fig. 2), a metabolite of the phytol fragment of chlo-

rophyll that can be obtained from dietary sources, was reported to bind

RXR, but not RAR, and elicit transactivation, most likely through

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RXR homodimers (Kitareewan et al., 1996; LeMotte, Keidek, & Apfel,

1996). It promoted transactivation of RXRα activities at 10μM, and entered

into competition with [³H]-9-cis-retinoic acid for RXRα binding with a Ki

of 4μM, ca. 200-fold less efficient than 9-cis-retinoic acid (Zomer et al.,

2000). RXR transactivation by phytanic acid 12 (at 4–10μM) could take

place under physiological levels in plasma (1–6μM) but in mouse tissue

10–100 lower levels were measured. Therefore, physiological RXR-

activation could be cell-type specific and depend on subcellular availability

(including nucleus) of both this natural product and its putative metabolites.

Since humans do not efficiently absorb chlorophyll, the metabolic precursor

of phytanic acid, its presence in the human body is due to the digestion of

meat and milk products on the diet, and therefore it cannot be considered as

an endogenous ligand in humans. Thus, phytanic acid 12 should be consid-

ered as a RXR ligand candidate of nutritional origin, but its physiological

role in RXR activation remains uncertain.

Its unsaturated precursor phytenic acid 13 (Fig. 2), which has been

detected in bovine serum extracts, is likewise considered as RXR ligand

since it was found to also displace tritiated 9-cis-retinoic acid x from the

receptor in competition assays, whereas its Z isomer was inactive

(Kitareewan et al., 1996; LeMotte et al., 1996).

Additional C20-fatty acids, like geranyl geranoic acid derivatives (GGA,

14), which are present in diet relevant herbs have been reported as potent

RXR activators (Araki, Shidoji, Yamada, Moriwaki, & Muto, 1995),

although they could not be identified endogenously.

In summary, there is a strong link between DHA 1 and RXR-mediated

signaling, although a clear picture of how it operates in vivo is still missing.

Since the potency of these unsaturated compounds is not sufficient to make

them qualify as true endogenous RXR ligands, they should instead be con-

sidered as partial RXR agonists (Bourguet et al., 2000). It appears that the

promiscuity of RXR with several unsaturated metabolites reflects its funda-

mental role as intracellular sensor of the cell metabolic status, binding fatty

acids and analogous compounds depending on their supply and their local

concentrations. All together, these results underline the potential involve-

ment of RXR in lipid homeostasis through complex feedback mechanisms

in potent association with other NRs (such as PPAR or FXRs).

3.2 Rexinoids from plants (Fig. 3)

Natural product honokiol 15 (Fig. 3), isolated from Magnolia obovata, and

positional isomer magnolol 16, from Magnolia officinalis, used in traditional

Fig. 3 Naturally occurring RXR ligands from plants with phyto-pharmacological relevance.

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Chinese medicine to cure cough, diarrhea and allergic rhinitis (also

metabolic syndrome) have been reported as RXR modulators (Kotani,

Tanabe, Mizukami, Makishima, & Inoue, 2010; Zhang et al., 2011).

Honokiol 15 was found to induce the transactivation of the PPARγ/

RXRα heterodimer, but not that of the RXRα/RXRα homodimer

(Kotani et al., 2010). The crystal structure of the ligand-bound RXRα rev-

ealed that the allyl phenol moieties of magnolol 16 occupy the L-shaped

arms of the LBD and, since it does not contain a carboxylic acid, stabilization

involves formation of a hydrogen bond with Asn306 on N-terminal helix

H5 by one of the phenol groups. Ligand binding to RXRα and PPARγ

was estimated as EC₅₀ꢀ40μM and EC₅₀ꢀ2μM, respectively, for 16.

The data might explain its preference for activation of RXR/PPARγ tran-

scription at EC₅₀ꢀ10μM (Zhang, Xu, et al., 2011). Likewise, the reported

anti-inflammatory, neuroprotective or antitumor activities of honokiol 15

in animal models of Alzheimer’s or Parkinson’s disease (Kotani et al.,

2010) may be related to the proposed stimulation of RXRα (Wang,

Dong, & Wang, 2018).

More recent Molecular Modeling and NMR spectroscopy concluded

that honokiol 15 targets RXR at both sides of the interface, acting on

the coactivator side of the dynamic activation function (AF2) or alternatively

it switches from one to the other side of the interface (Scheepstra et al.,

2014). Rational design allowed to split the dual-binding properties of the

natural product honokiol 15 at the dynamic nuclear receptor interface

AF2, and led to derivatives that either stabilize AF2 through binding at

the LBP or destabilize AF2 through binding at the coactivator solvent-

exposed side of the interface (Scheepstra et al., 2014).

Drupanin 17 (Fig. 3) isolated from green propolis (bee glue), a resinous

mixture produced by honey bees, has shown selective, dual RXR/PPARγ

agonistic profile in transactivation and binding assays (no binding to RARs,

LXRs, VDR was observed), and also increased Ap2/FABP4 expression and

accelerated lipid accumulation in differentiated 3T3-L1 cells. The dual

RXR/PPARγ agonistic profile was similar to that described for magnolol

16, although drupanin 17 can bind with higher efficacy to all RXR isotypes

(EC₅₀¼2–7μM) than to PPARγ (EC₅₀ꢀ15μM) (Nakashima, Murakami,

Tanabe, & Inoue, 2014).

Naturally occurring anthraquinones danthron and rhein (18, 19), iso-

lated from Chinese rhubarb Rheum palmatum have been reported to function

as antagonists of 9-cis-retinoic acid 31 (Fig. 4) in RXRα-LBD, Gal4-

luciferase transactivation assays with IC₅₀values of 10 and 70μM,

Fig. 4 Structures of apocarotenoids (including retinoids).

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respectively (Zhang et al., 2011; Zhang, Chen, Chen, Jiang, & Shen, 2011).

A crystal structure of tetrameric RXR was found to be stabilized by the

anthraquinones as an inactive form with the recruitment of SMRT (silenc-

ing mediator for retinoid and thyroid hormone receptors) corepressor, and

showed a strong displacement of H12 helix, making it inaccessible for for-

mation of transcriptionally functional homo- or heterodimers. Danthron 18

and rhein 19 inhibited RXR-mediated transactivation through not only

RXR-homodimers, but also through heterodimers of RXR with FXR,

LXR or PPARγ.

Sesquiterpene lactone bigelovin 20 (Fig. 3), isolated from the flowers of

Inula helianthus-aquatica or Inula hupehensis, which are used in folk medicine

for their anti-cancer potential, binds to RXRα-LBD (no binding to LXRα,

FXR or PPARγ was observed), but transactivates through the RXRα-

PPARγ heterodimers (not through RXRα-FXR heterodimers) and

suppresses RXR-homodimers or RXRα-LXR heterodimers on their

respective response elements in the 1–10μM concentration range (Zhang

et al., 2011). The ligand-mediated remodeling of LBD observed in the crys-

tal structure of RXR-LBD was proposed to rationalize the discriminatory

activities between RXR-homodimer and heterodimer interactions, and

the opposing activities of specific RXR heterodimers.

Computer-aided drug design (CADD) has also been applied to RXR

receptors, leading to the discovery of rexinoids that modulate the activities

of RXR in a canonical and in a non-canonical way, through binding to

the receptor surface or attachment to the co-regulatory binding surface

(co-activator binding site and dynamic protein interfaces).

Screening the Dictionary of Natural Products based on isomimetics of

natural RXR ligands led to identification of valerenic acid 21 as new natural

RXR ligand (Merk, Grisoni, Friedrich, Gelzinyte, & Schneider, 2018). This

sesquiterpenoid, isolated from Valeriana officinalis (a flowering plant mainly

present in Europe and Asia), showed agonistic activity in the micromolar

range, which was reflected in the efficient induction of the RXR transcrip-

tion target genes ABCA1 and ApoE. The Gal4-based reporter assays per-

formed in HEK293T cells revealed intriguing partial selectivity for

RXRβ, since valerenic acid showed better EC₅₀(5μM) and about

10-times higher transactivation of RXRβ as compared to RXRα

(EC₅₀¼27μM), and RXRγ (EC₅₀¼43μM). In addition, no activation

of RARs, PPARs, LXRs, FXR, CAR, VDR and PXR was observed.

Molecular modeling studies suggested similar binding modes of 21 to

RXRα and RXRβ as classical rexinoids (Merk et al., 2018).

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221

Among other natural products analogs isolated from Cratoxylum formosum

ssp. pruniflorone R 22 (Fig. 3) was found to be active on RXRα, when

CV-1 cells were transiently transfected with the TREpal-tk-CAT reporter

(Duan et al., 2010). Xanthone 23 was the most potent of the family of ana-

logues isolated from Cratoxylum formosum ssp. pruniflorum and assayed using

the transcriptional activities of RXRα in CV-1 cells co-transfected with

TREpal-tk-CAT (Duan et al., 2011). Neriifolone B 24, isolated from the

stems of Cratoxylum cochinchinense, was also reported as an inhibitor of the

RXRα transcriptional activity (Shen et al., 2014). Similar to other deriva-

tives and analogues, it showed concentration-dependent RXRα inhibitory

activities.

RXR agonistic activity has been reported for two prenylated flavonones

isolated from Sophora tonkinensis, which is used in traditional Chinese medi-

cine (Inoue, Tanabe, Nakashima, Ishida, & Kotani, 2014). Both natural prod-

ucts (25, 26) were found to bind at the nanomolar range (EC₅₀0.77 and

0.78μM, respectively) to the three RXR isotypes (no binding to RARα,

LXRα or PPARα, β/δ or γ), and to transactivate through PPARγ/RXR,

PPARδ/RXR, RXR/LXR, and RXR/RAR heterodimers, with the

ability to potentiate the activity of agonists for the partner NRs. However,

the concentration required for inducing a similar transcriptional activity than

bexarotene (LDG1069) was 10 times higher. In addition, the prenylated

flavonones were shown to induce the preferential expression of lipoprotein

lipase, angiopoietin-like protein 4, and heme oxygenase-1, which are

involved in prevention of heart diseases, diabetes, insulin resistance, and ath-

erosclerosis. Therefore, the transcriptional activity of these ligands on RXR

heterodimers with LXR, PPARβ/δ, PPARγ are different to those induced by

bexarotene, and further studies on these prenylated flavonones are required in

order to clarify their potential utility.

Xanthones 27 and 28 were reported as RXRα agonists, with K_Dto

RXRα-LBD of 68.3 and 14.0μM, respectively, and showed selective cyto-

toxicity against HeLa cells, most likely by extrinsic pathways. Apoptosis was

shown to be promoted through PARP cleavage and caspase-8 activation.

A fluorescence quenching assay suggested that these two xanthones can

interact with the Arg316 of the RXRα-LBD and directly bind to

RXRα. Molecular docking studies taking into account the topology of

these two compounds indicated their structural adaptation to occupy the

L-shaped binding pocket of RXRα (Qiu et al., 2019). The proposal is in

agreement with SAR studies showing that the hydroxyl group at C5 and

an aliphatic chain at C2 in chromone-type compounds as well as a free

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3-OH and an aliphatic chain substituted at the 7-OH in these xanthones are

essential for their potent RXRα transcriptional inhibitory activities.

Likewise, it is consistent with the finding that prenylated xanthones

exhibited more potent effects on the RXRα transcriptional inhibitory activ-

ities than simple xanthones (Duan et al., 2010, 2011; Shen et al., 2014).

3.3 Endogenous RXR ligands (Fig. 4)

β-Apo-13-carotenone 29 (Fig. 4), has been recently demonstrated to

stabilize inactive tetrameric RXR, instead of the transcriptionally active

RXRα dimers (Eroglu, Hruszkewycz, Curley, & Harrison, 2010; Sun,

Narayanasamy, Curley, & Harrison, 2014). Ki values for RXRα of 8nM

in a 9-cis-retinoic acid 31 competitive displacement assay were measured,

although different modes of binding are suggested by in silico ligand docking

studies with RXRα-LBD (Sun et al., 2014). In addition, this apocarotenoid

appears to also act as a competitive inhibitor of RARs with a Ki of 4nM

(Eroglu et al., 2012), and to reduce ATRA-RAR-mediated activity by

about 70%. Lower levels of β-apo-13-carotenone 29 (1nM) following

chronic consumption of β-carotene-enriched diets has been reported

(Cooperstone et al., 2017). Since β-apo-13-carotenone 29 is generated

by thermal degradation and oxidation of β-carotene during food processing

(it is also found only in some fresh fruits), it cannot be produced in mammals

(Harrison & Quadro, 2018). Similarly, β-apo-14⁰-carotenal 30 (Fig. 3) has

been reported to act as a potential inhibitor of RXRα as well as of selected

heterodimeric partners (PPARα, PPARβ/δ, LXRα and LXRβ, but not

RARs). This apocarotenoid can be found in several melons, but its detection

in human plasma was not consistent (Harrison & Quadro, 2018). It is there-

fore unlikely that those apocarotenoids are physiologically meaningful,

despite their in vitro inhibitory activities.

Although it is traditionally considered as a natural RXR ligand, it is

important to indicate that 9-cis-retinoic acid 31 (Fig. 4) has not been

conclusively shown to be endogenously present, except in pancreatic

β-cells, and recently in human serum in very low concentrations (Arnold,

Amory, Walsh, & Isoherranen, 2012). Either its concentration is under

the detection limit, it is present transiently or it accumulates in local cell

populations (Kane, 2012).

Additional biologically potent endogenous vitamin A derivatives, among

them 9-cis-3,4-didehydroretinoic acid 32 and 9-cis-4-oxo-retinoic acid 33,

have been demonstrated to bind to and activate RXR in vitro

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(Mangelsdorf & Evans, 1995; Mangelsdorf et al., 1995). 9-cis-3,4-

didehydroretinoic acid 32 could be biosynthetically derived from the pre-

cursor vitamin A₂(all-trans-3,4-didehydroretinol) following similar routes

to those of 31 from vitamin A (all-trans-retinol). 9-cis-4-oxo-retinoic acid

33 has been isolated from Xenopus embryos, where it was found to modulate

axial pattern formation (Pijnappel et al., 1998), after its likely formation by

isomerization of the all-trans isomer, which is endogenously present in the

same organism (Pijnappel et al., 1993). Metabolite 33 was shown to bind

RARα,β,γ and RXRα,β,γ, with IC₅₀values about 5–10 times higher than

those of 9-cis-retinoic acid 31, and to induce a synergistic activation of the

RAR/RXR heterodimers, but not the RXR homodimers (in contrast to

9-cis-retinoic acid 31) (Pijnappel et al., 1998).

The endogenous presence of 9-cis-13,14-dihydroretinoic acid 34 and its

all-trans isomer, has recently been proven in several mice organs (liver,

serum, brain) by HPLC-MS-MS and UV and quantified (quantitative bind-

ing affinity of K_d90ꢁ20nM; cf. 20ꢁ10nM for 9-cis-retinoic acid 31 by

fluorescence quenching assays) to be sufficient to maintain RXR activities

(Ru€hl et al., 2015). Titration experiments monitored by ESI-MS showed

that the affinity of RXR for the S enantiomer was about 65% lower than

that for the R enantiomer. Crystal structure studies on the R enantiomer

bound to hRXRα LBD and a 13-residue peptide comprising the nuclear

receptor binding surface NR2 of NCoA2 confirmed binding to the canon-

ical binding pocket, adopting a similar orientation than 9-cis-retinoic acid

31, which is consistent with their similar transcriptional activities on several

nuclear receptor-signaling pathways, possibly through the corresponding

permissive heterodimers. Therefore, 9-cis-13,14-dihydroretinoic acid 34

was considered as the first endogenous and physiologically relevant retinoid

RXR ligand in mammals (Ru€hl et al., 2015). This ligand and other

dihydroretinoids are presumably produced by retinol saturase (RetSat)

(Moise et al., 2009). The structure of 9-cis-13,14-dihydroretinoic acid 34

had previously been assigned to a metabolite detected in rats, together with

its taurine derivative, after being fed with high doses of 9-cis-retinoic acid 31

(Shirley et al., 1996), but in-depth characterization of its structure and func-

tion was not pursued. 9-cis-retinol might be the metabolic precursor of

RetSat action to generate 34 following similar biosynthetic pathways as

(R)-all-trans-13,14-dihydroretinol to all-trans-13,14-dihydroretinal (Moise

et al., 2008; Moise, Kuksa, Blaner, Baehr, & Palczewski, 2005; Moise,

Kuksa, Imanishi, & Palczewski, 2004), and further metabolism to the car-

boxylic acid. The specific all-trans to 9-cis isomerization process is likely

224

Patricia García et al.

to occur in a fully conjugated polyene precursor, namely a carotenoid or a

C9-C10-dihydrocarotenoid. Nevertheless, the precise mechanistic details

and the possibility that the isomerization step could be enzymatically driven

are yet unknown.

Aputativemetaboliteof34,namely(S)-9-cis-4-oxo-13,14-dihydroretinoic

acid 35 has been shown to be present in the liver and other tissues of both

wild-type mice and rats fed a standard laboratory diet, as well as in humans

(Schmidt, Volland, Hamscher, & Nau, 2002). This compound activated

RAR-dependent gene transcription via RAR–RXR heterodimers, although

it was less potent than all-trans-retinoic acid, despite its higher concentration in

mice liver, but showed no transcriptional activity with either RXRα orRXRβ

(Schuchardt et al., 2009).

4. Diastereo- and enantioselective synthesis

of (R)-9-cis-13,14-dihydroretinoic acid 34

The preparation of (R)-9-cis-13,14-dihydroretinoic acid (R)-34 was

based on the Suzuki cross-coupling of enantiopure trienyliodide 40 and

boronic acid 41 (Scheme 1). The synthesis of (R)-3 started with (2Z,4E)-

stannyldienol 35 (Dom´ınguez, Pazos, & de Lera, 2000; Pazos & de Lera,

1999) which was transformed into the benzothiazolyl allyl sulfide 36 by

Mitsunobu reaction with the corresponding thiol and subsequently oxidized

to sulfone 37 with H₂O₂and a peroxymolybdate (VI) reagent (Schultz,

Freyermuth, & Buc, 1963) at ꢂ10°C. The Julia–Kocienski olefination

(Aıssa, 2009; Blakemore, 2002) was performed using a slight excess of base

¨

Scheme 1 Reagents and conditions: (a) PPh₃, BTSH, DIAD, CH₂Cl₂, 2h (98%).

(b) (NH₄)₆Mo₇O₂₄ꢃ4H₂O, 30% H₂O₂, EtOH, ꢂ10ºC, 17h (66%). (c) NaHMDS, THF,

ꢂ78ºC, 30min (93%). (d) I₂, CH₂Cl₂, 25ºC, 30min. (e) Pd(PPh₃)₄, 10% aq. TlOH, THF,

25ºC, 3h (42%). (f ) 2M KOH, MeOH, 80ºC, 45min (84%).

RXR natural modulators

225

(NaHMDS, 1.15 equivalents) and 1.7 equivalents of enantiopure aldehyde

(R)-38 (Leonard, Mohialdin, Reed, Ryan, & Swain, 1995; Moise et al.,

2008). As anticipated from previous findings on the stereoselectivity of

the Julia–Kocienski reactions of allylsulfones and aldehydes (Sorg &

Bru€ckner, 2005; Vaz, Alvarez, Souto, & de Lera, 2005), the geometry of

the newly formed olefin of trienyl ester (R)-39 is Z (which was confirmed

by NOE experiments). Treatment of the precursor stannane with a solution

of iodine in CH₂Cl₂produced the iodide (R)-40 via Sn-I exchange and

iodine-promoted isomerization of the Z,Z,E triene to the desired E,Z,E

geometric isomer (as confirmed by NOE experiments). The Suzuki reaction

(Miyaura & Suzuki, 1995; Suzuki, 2011) of freshly prepared boronic acid 41

and trienyl iodide (R)-40 using Pd(PPh₃)₄as catalyst and 10% aqueous

TlOH as base in THF at ambient temperature, followed by immediate

work-up afforded ethyl (R)-9-cis-13,14-dihydroretinoate (R)-42 in 78%

yield. Saponification of (R)-42 provided in 84% yield the desired carboxylic

acid (R)-34 (Ru€hl et al., 2015).

5. Materials

Silica gel column chromatography: Merck Kieselgel 60 (230–400 mesh

particle size and C18-silicagel.

Aluminum plates for tlc: Merck Kieselgel 60F254.

Visualization for analytical tlc: UV irradiation (254nm) or staining with an

ethanolic solution of phosphomolybdic acid.

5.1 Reagents and solvents

2-Mercaptobenzothiazole, 97%.

Triphenylphosphine (PPh₃), ꢄ95%.

Tetrahydrofuran (THF), dried and distilled before use.

Acetonitrile, ACS reagent, max 0.0075% H₂O.

Diisopropyl azodicarboxylate (DIAD), 98%.

Ethanol (EtOH), reagent grade.

Ammonium heptamolybdate tetrahydrate [(NH₄)₆Mo₇O₂₄ꢃ4H₂O],

>99%.

Hydrogen peroxide (35% in H₂O).

Diethyl ether (Et₂O), ACS reagent.

Diethyl ether (Et₂O), for synthesis, stabilized with 7ppm BHT.

Methanol (MeOH), for HPLC ꢄ99.9%.

Sodium bis(trimethylsilyl)amide solution (NaHMDS, 1M in THF).

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Patricia García et al.

Iodine (I₂), ꢄ99.8%.

Dichloromethane (CH₂Cl₂) for synthesis, stabilized with amylene.

Sodium dithionite (Na₂S₂O₃), ꢄ82%.

Sodium sulfate (Na₂SO₄), ꢄ99%, anhydrous powder.

Hexane, synthesis grade.

Triethylamine (Et₃N), ꢄ99%.

Tetrakis(triphenylphosphine)palladium [Pd(PPh₃)₄], 99%.

Thallium hydroxide (TlOH); a 10% aqueous solution, prepared from

barium hydroxide, Ba(OH)₂(ꢄ98%) and thallium sulfate, Tl₂SO₄(ꢄ99.5%).

Celite^®, ꢄ95%.

Potassium hydroxide (KOH), a 2M aqueous solution.

Hydrochloric acid (HCl), a 10% aqueous solution.

tert-Butyl methyl ether (t-BuOMe), ꢄ99.8%.

Trifluoroacetic acid (TFA), 99%.

Phosphomolybdicacidhydrate, Na₃[P(Mo₃O₁₀)₄]ꢃxH₂O, technicalgrade.

6. Methods

6.1 Method for formation of BT-sulfide and BT-sulfone

6.1.1 (2Z,4E)-1-(Benzothiazol-2-yl)sulfonyl-5-(tri-n-butylstannyl)-3-

methylpenta-2,4-diene (37)

1. A solution of (2Z,4E)-3-methyl-5-(tributylstannyl)penta-2,4-dien-1-ol

35 (1.0g, 2.58mmol), 2-mercaptobenzothiazole (0.65g, 3.87mmol) and

PPh₃(1.10g, 4.21mmol) in THF (14mL) in a 50mL round bottom flask

equipped with a magnetic stirring bar and stirred for 5min at 0°C.

2. A solution of DIAD (0.77mL, 3.87mmol) in THF (5mL) was added

dropwise to the above flask via syringe and the mixture was stirred for

30min at 25°C.

3. The solvent was removed and the residue was purified by column

chromatography (C18-silica gel, CH₃CN) to afford 1.11g (78%) of

(2Z,4E)-1-(benzothiazol-2-yl)sulfanyl-5-(tri-n-butylstannyl)-3-meth-

ylpenta-2,4-diene 36 as a colorless oil.

4. To a solution of 36 (0.48g, 0.89mmol) in EtOH (9mL), at ꢂ10°C, was

added a solution of (NH₄)₆Mo₇O₂₄ꢃ4H₂O (0.44g, 0.36mmol) in aque-

ous hydrogen peroxide (35%, 7.7mL, 89.1mmol).

5. Stirring was maintained for 17h at ꢂ10°C.

6. Work-up: the mixture was quenched with H₂O and extracted with

Et₂O (3ꢅ). The combined organic layers were washed with brine

(3ꢅ) and dried (Na₂SO₄), and the solvent was removed.

RXR natural modulators

227

7. Purification: the residue was purified by chromatography (C18-silica gel,

MeOH) to afford 0.33g (66%) of a colorless oil, which was identified

as 37.

6.2 Method for formation of dienylstannane

6.2.1 Ethyl (3S,4Z,6Z,8E)-3,7-dimethyl-9-(tri-n-butylstannyl)nona-4,6,8-

trienoate ((S)-39)

1. A solution of (2Z,4E)-1-(benzothiazol-2-yl)sulfonyl-5-(tri-n-butylstannyl)

3-methylpenta-2,4-diene 37 (0.115g, 0.20mmol) in THF (9mL) in a

50mL round bottom flask equipped with a magnetic stirring bar was

cooled down to ꢂ78°C and then was treated with NaHMDS (0.23mL,

1M in THF, 0.23mmol).

2. After stirring for 30min at this temperature, a solution of ethyl (S)-3-

methyl-4-oxobutanoate 38 (0.044g, 0.30mmol) in THF (4.5mL) was

added and the resulting mixture was stirred for 1h at ꢂ78°C.

3. It was then carefully allowed to reach room temperature for 3h.

4. After lowering the temperature to 0°C, Et₂O and water were added and

the mixture was allowed to warm up to room temperature.

5. Work up: the reaction mixture was diluted with Et₂O and the layers

were separated. The aqueous layer was extracted with Et₂O (3ꢅ), the

combined organic layers were dried (Na₂SO₄) and the solvent was

removed.

6. Purification: the residue was purified by column chromatography (C-18

silica gel, MeOH) to afford 0.94g (93%) of (S)-39 as pale yellow oil.

6.3 Method for stannane-iodine exchange

6.3.1 Ethyl (3R,4E,6Z,8E)-9-iodo-3,7-dimethylnona-4,6,8-trienoate

((R)-40)

1. To a solution of ethyl (3R,4Z,6Z,8E)-3,7-dimethyl-9-(tri-n-butylstannyl)

nona-4,6,8-trienoate (R)-39 (0.060g, 0.121mmol) in CH₂Cl₂(5.3mL) in

a 25mL round-bottomed flask equipped with a magnetic stirring bar was

added dropwise a solution of iodine (0.046g, 0.182mmol) in CH₂Cl₂

(2.8mL).

2. The resulting mixture was stirred for 30min at 25°C.

3. Work-up: a saturated Na₂S₂O₄solution was then added and the reaction

mixture was extracted with Et₂O (3ꢅ), the combined organic layers

were dried (Na₂SO₄) and the solvent was removed.

228

Patricia García et al.

4. Purification: the residue was purified by column chromatography (silica

gel, 97:3 hexane/Et₃N) to afford 0.037g (92%) of (R)-40 as pale

yellow oil.

6.4 Method for Suzuki cross-coupling

6.4.1 Ethyl (9Z,13R)-13,14-dihydroretinoate ((R)-42)

1. To a solution of ethyl (3R,4E,6Z,8E)-9-iodo-3,7-dimethylnona-4,6,8-

trienoate (R)-40 (0.036g, 0.107mmol) in THF (2.3mL) in a 5mL

round-bottomed flask equipped with a magnetic stirring bar was added

Pd(PPh₃)₄(0.013g, 0.011mmol).

2. After 5min at room temperature, 2,6,6-trimethylcyclohex-1-enylboronic

acid 41 (0.027g, 0.161mmol) was added in one portion followed by a 10%

aqueous solution of TlOH (0.75mL, 0.407mmol).

3. Stirring was maintained for 3h at 25°C.

4. Work-up: Et₂O was added and the reaction mixture was filtered through

a short pad of Celite^®. The filtrate was washed with NaHCO₃(aqueous

saturated solution), the organic layer was dried (Na₂SO₄) and the solvent

was removed.

5. Purification: the residue was purified by column chromatography (silica

gel, 97:3 hexane/Et₃N) to afford 0.028g (78%) of a pale yellow oil,

which was identified as ethyl (9Z,13R)-13,14-dihydroretinoate (R)-42.

6.5 Method for saponification

6.5.1 (9Z,13R)-13,14-dihydroretinoic Acid ((R)-34)

1. To a solution of ethyl (9Z,13R)-13,14-dihydroretinoate (R)-42

(0.023g, 0.069mmol) in MeOH (4.7mL) in a 5mL round-bottomed

flask equipped with a magnetic stirring bar was added KOH (2M aque-

ous solution, 1.1mL, 2.27mmol).

2. The reaction mixture was allowed to stir at 80°C for 45min.

3. The reaction was allowed to cool down to room temperature.

4. Work-up: CH₂Cl₂and brine were added and the layers were separated.

The aqueous layer was washed with H₂O (3ꢅ). The combined aqueous

layers were acidified with 10% HCl and extracted with CH₂Cl₂(3ꢅ).

The combined organic layers were dried (Na₂SO₄) and the solvent

was removed.

5. Purification: the residue was purified by column chromatography (silica

gel, gradient from 95:5 to 90:10 CH₂Cl₂/MeOH) to afford 0.017g

(84%) of (R)-34 as pale yellow oil (Fig. 5).

RXR natural modulators

229

0.020

Sample: 13S

310 nm

0.015

0.010

0.005

0.000

0.020

310 nm

Sample: 13S +13R

0.015

0.010

0.005

0.000

0.00 2.00

4.00 6.00

8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00 24.00 26.00 28.00 30.00 32.00

Minutes

0.0025

0.0020

0.0015

0.0010

0.0005

0.0000

Sample: 13R

310 nm

0.006

0.005

0.004

0.003

0.002

0.001

0.000

310 nm

Sample: 13S +13R

0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00 24.00 26.00 28.00 30.00 32.00 34.00

Minutes

Fig. 5 Separation of 9-cis-13,14-dihydroretinoic acid enantiomers.

230

Patricia García et al.

6.6 Method for enantiomeric excess determination

of (9Z,13R)-13,14-dihydroretinoic acid (9Z,13R)-34

1. Stationary phase: Chiralpak IA column.

2. Eluent: 92.5:7.5 v/v solvent mixture of hexane (+0.1% TFA)/

t-BuOMe.

3. Isocratic mode at a flow of 2mL/min.

4. Detection by absorption at 310nm and 254nm.

Acknowledgments

We thank Dr. Belꢀen Vaz and Dr. Susana Alvarez for the optimization of the enantioselective

synthesis of 9-cis-13,14-dihydroretinoic acid, and the Spanish MINECO (SAF2016-77620-

R-FEDER and BIO2016-78057-R; Xunta de Galicia (Consolidacio´n GRC ED431C

2017/61 from DXPCTSUG; ED-431G/02-FEDER “Unha maneira de facer Europa” to

CINBIO, a Galician research center 2016–19) for financial support.

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Article Doi

Natural ligands of RXR receptors

DOI: 10.1016/bs.mie.2020.02.006

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