Synthesis of PPAR-γ Activators Inspired by the Marine Natural Product, Paecilocin A

A series of N-substituted phthalimide derivatives were synthesized based on a pharmacophore study of paecilocin A (a natural PPAR-γ agonist) and synthetic leads. The introduction of hydrophilic and hydrophobic groups to the phthalimide skeleton yielded compounds 3–14. Compound 7 showed significant PPAR-γ activation in a luciferase assay using rat liver Ac2F cells. Docking simulations showed that a free hydroxyl group on the phthalimide head and a suitable hydrophilic tail, including a phenyl linker, were beneficial for PPAR-γ activation. Compound 7 and rosiglitazone concentration-dependently activated PPAR-γ with EC50 values of 0.67 μM and 0.028 μM, respectively. These phthalimide derivatives could be further investigated as a new class of PPAR-γ ligands.

Endogenous PPAR-γ ligands typically contain a free carboxylic acid head and an unsaturated alkyl chain terminus (the tail), whereas TZDs and L-tyrosine analogues have carbonyl, amino or carboxyl groups at their hydrophilic heads and a phenol moiety, which acts as a linker between the head and tail. Hydrophilic head groups form H-bonds with key amino acid residues (Tyr 473 , His 449 , His 323 and Ser 289 ) of the ligand-binding domain (LBD) of PPAR-γ and stabilize PPAR-γ in the conformation required for successful co-activator recruitment [12][13][14][15]. In our previous study, we isolated paecilocin A (Figure 1), a new type of PPAR-γ agonist, from the jellyfish-derived fungus, Paecilomyces variotii [16], and based on the results of a pharmacophore study on paecilocin A and rosiglitazone, we proposed that the 3-hydroxy phthalide moiety of paecilocin A functions as a hydrophilic head and forms H-bonds with the key amino acid residues of the LBD of PPAR-γ [13]. Paecilocin A contains a hydrophilic 3-hydroxy phthalide moiety and a hydrophobic octyl chain; both thiazolidinedione (TZD) and tyrosine derivatives employ a phenol moiety as a linker between their hydrophilic heads and hydrophobic tails. (B) The N-substituted phthalimide skeleton of PPAR-γ agonists; a 3-hydroxy phthalimide moiety acts as the head, a phenol moiety as the central linker and a hydrophobic or hydrophilic substituent as the tail.
The easily accessible phthalimide moiety has often been employed as a pharmacophore in drug development [17], and some phenethylphenylphthalimide derivatives have been reported to be PPAR-γ agonists. However, the phthalimide moiety of these derivatives does not contain a free hydroxyl group, and it was speculated to serve as a hydrophobic tail, which settles in the hydrophobic region of the PPAR-γ LBD [18]. On the contrary, the phthalide moiety of paecilocin A was speculated to behave as a polar head group that forms H-bonds with key amino acid residues in the hydrophilic pocket of PPAR-γ LBD ( Figure 1A) [13]. Considering the structural similarity between the phthalimide moiety and the phthalide moiety of paecilocin A, we postulated the introduction of hydroxyl groups would allow the phthalimide moiety to be utilized as a polar head group and that this modification could be further extended using linker and tail groups to generate potential PPAR-γ ligands ( Figure 1B). In the present study, phthalimide-derived molecules were designed, produced and evaluated with respect to PPAR-γ activation in rat liver Ac2F cells.

Chemistry
N-substituted phthalimides can be prepared by heating phthalic anhydride with various N-containing reagents [19]. In the present study, this was performed by treating phthalic anhydrides (1, 2) with various amines in the presence of acetic acid to generate corresponding N-substituted phthalimides (Scheme 1). Compounds 3 and 4 were synthesized to imitate the skeletons of lipidic PPAR-γ agonists and paecilocin A by treating phthalic anhydrides with oleylamine, whereas compounds 5-7 were synthesized to imitate the topology of synthetic PPAR-γ agonists (e.g., rosiglitazone and farglitazar) by connecting the phthalimide head with p-hydroxy-or p-bromo-phenethyl groups. Previous studies have shown that a large hydrophobic substituent, such as a hetero-aromatic group, provides a good tail for PPAR-γ agonists [20]. Accordingly, we speculated that connecting the phenoxyl moiety with a hydrophobic tail designed to occupy the hydrophobic binding pocket in the LBD of PPAR-γ might enhance binding affinity. Therefore, compound 7 was derivatized by treating it with iodomethane, iodoethane, iodopropane, iodobutane, iodooctane or benzyl chloride to produce the hydroxy-substituted analogues 8-14 (Scheme 2). The major product obtained was the result of the monoalkylation of the hydroxyl group on the phenethyl moiety (11), and the dialkylated product (10) was produced at low levels. The monobenzylated derivative (14) was also prepared. Scheme 2. Synthesis of phthalimide derivatives (8-10, yield: ~25%; 11 and 12, yield: ~75%; 13, yield: ~60%; 14, yield: ~60%). Reagents and conditions: (a) RI (MeI, EtI, n-PrI, n-BuI, n-OctI) Ag 2 O, stir, CH 3 CN, reflux for 12 h; (b) benzyl chloride, K 2 CO 3 , NaI, DMF, stir, RT for 4 h.

Biological Activity
Synthesized compounds were subsequently evaluated for PPAR-γ activation using a luciferase assay in Ac2F cells (a rat liver cell line). The potencies of compounds 6 and 7 were comparable to rosiglitazone at 10 μM ( Figure 2). In docking simulations [21], the hydrophilic head of compound 7 was found to form H-bonds with the key amino acid residues (Tyr 473 , His 323 and Ser 289 ) of the LBD of PPAR-γ ( Figure 3), in the same manner as rosiglitazone [13]. Compounds 3, 4 and 6-14 also appeared to bind to the LBD of PPAR-γ and to interact with key amino acid residues (see Appendix Table A1). Alkylated members (3 and 4) showed low binding affinities, whereas 6 and 7 showed high binding affinities, which suggested that the aromatic linker provided better binding than a long alkyl chain, possibly because the alkyl chain is too bulky to fit into the binding pocket. Compounds 3 and 4 produced even lower levels of PPAR-γ activation than the control, possibly because of their cytotoxicities.
Secondary N-substituted phthalimide derivatives (11)(12)(13)(14) activated PPAR-γ more than the control (rosiglitazone) at 10 μM, except compound 13 ( Figure 4). As was expected, secondary derivatives with a free hydroxyl group on the phthalimide moiety (11)(12)(13)(14) were more effective than O-alkylated derivatives (8-10) (Figure 2). These findings indicated that the free 3-OH on the phthalimide moiety was essential for inducing PPAR-γ activity, and this notion was consistent with the strong docking affinities of 11-14 to the LBD of PPAR-γ (see Appendix Table A1). Interestingly, the PPAR-γ agonistic activity of a long octyl tail derivative (13) sharply decreased at a concentration of 10 μM, which was similar to that observed for compounds 3 and 4, which also have a long hydrophobic tail (Figures 2 and 4). These observations could be due to the cytotoxic effects of derivatives with long hydrophobic tails. Indeed, compound 13 exhibited strong cytotoxicity toward Ac2F cells at 10 μM ( Figure 5). Although 7 and 14 formed similar H-bond networks in docking simulations ( Figure 3B and Appendix Figure A1), compound 14, which possess a benzyl group, seemed reluctant to locate in the binding site, based on its smaller NP (the number of modes located in the binding pocket) and AI (affinity of the top-ranked mode) values than compound 7 (see Appendix Table A1).

Figure 2.
In vitro assay of PPAR-γ activation by phthalimides 3-10 and by rosiglitazone at 10 μM in rat liver Ac2F cells. Con., the negative control, transfected with a plasmid containing PPAR response element (PPRE) and pcDNA3; Rosi (rosiglitazone) was used as the positive reference control. Treated cells were transiently transfected with PPRE plus full-length human PPAR-γ1 expression vector (pFlag)-PPAR-γ1. Luciferase expressions (folds of the control) are the means ± SDs (n = 3). * p < 0.05.  Figure 4. In vitro assay of PPAR-γ activation by phthalimides 7, 11-14 and by rosiglitazone at 1 μM or 10 μM in rat liver Ac2F cells. Con., the negative control, transfected with plasmid containing PPRE and pcDNA3. Rosi, rosiglitazone. Rosiglitazone was used as the positive reference control to monitor the activation of the luciferase reporter. Compound-treated cells were transiently transfected with PPRE plus pFlag-PPAR-γ1. Luciferase expressions (folds of the control) are the means ± SDs (n = 3). * p < 0.05. Based on the above-mentioned results, 7 was selected as a potential lead compound and further evaluated at different concentrations (0.0098 μM~10 μM) versus rosiglitazone ( Figure 6). Both compound 7 and rosiglitazone exhibited concentration-dependent PPAR-γ activation. The activity of compound 7 was comparable to that of rosiglitazone at the concentration of 10 μM. Cell proliferation enhancement by compound 7 was compared with that of rosiglitazone at different concentrations ( Figure 7). No significant change in cell viability was induced by either compound after 6 h of treatment at concentrations of one and 10 μM, which exclude the possibility that enhanced proliferation was responsible for the observed increase in luciferase expression. Furthermore, the low cytotoxicity of compound 7 toward Ac2F cells demonstrated its potential safety, a key factor for a lead development compound. Figure 6. Dose-dependent PPAR-γ agonistic activities of rosiglitazone and compound 7. Ac2F cells were stimulated with rosiglitazone or compound 7 at various concentrations (0.0098 μM~10 μM). Con., the negative control, transfected with plasmid containing PPRE and pcDNA3. Rosi (rosiglitazone) was used as the positive reference control to monitor the activation of luciferase reporter. Treated cells were transiently transfected with PPRE plus pFlag-PPAR-γ1. Luciferase expressions (fold versus the control) are presented as the means ± SDs (n = 3). * p < 0.05.

Experimental Section
3. A mixture of amine (1.2 equivalent) and phthalic anhydride in aqueous glacial acetic acid (1 M) was stirred and heated under reflux overnight. Products were precipitated by adding water, filtered and washed thoroughly with water. Residues were diluted with MeOH, dried with MgSO 4 and evaporated to provide the crude products, 3-7 (yield: ~90%).  (14) To a suspension of 3-hydroxy-N-(p-benzyloxy-phenethyl) phthalimide 7 (13.4 mg, 0.047 mmoL), K 2 CO 3 (6.5 mg, 0.047 mmoL) and NaI (2.6 mg, 0.02 mmoL) in DMF (2 mL) was added benzyl chloride (6.0 μL, ca. 0.06 mmoL). The mixture was then stirred for 30 min at 0 °C and for 4 h at room temperature, acidified with aqueous 6 M HCl and extracted with EtOAc. The organic layer was successively washed with H 2 O and brine, dried with MgSO 4 and evaporated to give the crude product, which was purified by RP-HPLC using 85% aqueous MeOH as the eluant to give 14 (yield 95%): white powder; 1

Luciferase Assay
Rat liver Ac2F cells were obtained from the American Type Culture Collection (ATCC, Rockville, MD, USA). Cells were grown in Dulbecco's Modified Eagle Medium (DMEM, Nissui, Tokyo) containing 2 mM L-glutamine, 100 mg/mL streptomycin, 2.5 mg/L amphotericin B and 10% heat-inactivated fetal bovine serum (FBS) and maintained in a humidified atmosphere containing 5% CO 2 at 37 °C. The TK-PPRE × 3-luciferase reporter plasmid containing three copies of the PPAR response element (PPRE) in acyl CoA oxidase promoter was generously donated by Dr. Christopher K. Glass (University of California, San Diego, CA, USA). The pcDNA3 expression vector and full-length human PPAR-γ1 expression vector (pFlag-PPAR-γ1) were generously donated by Dr. Chatterjee (University of Cambridge, Addenbrooke's Hospital, Cambridge, UK). For luciferase assays, plasmids were transfected into Ac2F cells in a 48-well plate (5 × 10 4 cells/well) with effector plasmids and the TK-PPRE × 3-luciferase reporter plasmid (1 μg/well) plus pcDNA3(0.1 μg/well) or pFlag-PPAR-γ1 (0.1 μg/well) using Lipofectamine™ 2000 (Invitrogen Co., Carlsbad, CA, USA), according to the manufacturer's instructions. After transfection for 4 h, conditioned media was replaced with complete medium, and cells were incubated for an additional 20 h. The medium was then removed, and cells were exposed in serum-free media to rosiglitazone or test compounds for 6 h, washed with PBS and assayed using the ONE-Glo™ Luciferase Assay System (Promega, Madison, WI, USA). Luciferase activities were measured using a GloMax ® -Multi Microplate Multimode Reader (Promega Co., Sunny Vale, CA, USA).

Cell Proliferation Assay
Cell viabilities were evaluated using a WST (EZ-CyTox, Daeil Lab Service Co., Ltd, Seoul, South Korea). Ac2F cells (a rat liver cell line) were harvested and plated into 96-well microtiter plates at optimal seeding density (1 × 10 4 cells per well) and preincubated in complete medium in a humidified atmosphere containing 5% CO 2 at 37 °C for 24 h. The complete medium was removed and test substances dissolved in serum-free medium (100 μL/well) added, and cells were incubated for 6 h, 12 h, or 24 h. WST reagent (10 μL/well) was then added and incubated at 37 °C for 1 h. Absorbances were read using a iMark Microplate Absorbance Reader (Bio-Rad Laboratories, Hercules, CA, USA) at a test wavelength of 450 nm and a reference wavelength of 655 nm.

Molecular Docking Study
Docking calculations were performed using AutoDock Vina 1.

Conclusions
In conclusion, based on the results of our pharmacophore study of the marine natural product, paecilocin A, and of synthetic PPAR-γ agonists, we designed a series of N-substituted phthalimides. A free hydroxyl group on the phthalimide moiety and a hydrophilic tail were found to promote PPAR-γ activation. Furthermore, docking simulation results produced some interesting correlations between 3D structures and biological activities. We believe that further intensive optimization and evaluation of this new PPAR-γ agonist scaffold are likely to be rewarding.